Redox regulation of glutenin subunit assembly in the plant endoplasmic reticulum

Authors

  • Alessio Lombardi,

    1. Istituto di Biologia e Biotecnologia Agraria, Consiglio Nazionale delle Ricerche, Milano, Italy
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    • These authors contributed equally to this work.

  • Richard S. Marshall,

    1. Istituto di Biologia e Biotecnologia Agraria, Consiglio Nazionale delle Ricerche, Milano, Italy
    Current affiliation:
    1. Department of Genetics, University of Wisconsin, Madison, WI, USA
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    • These authors contributed equally to this work.

  • Chiara L. Castellazzi,

    1. Istituto di Biologia e Biotecnologia Agraria, Consiglio Nazionale delle Ricerche, Milano, Italy
    Current affiliation:
    1. Departamento de Biologia Estructural y Computacional, Instituto de Investigación Biomédica, Barcelona, Spain
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  • Aldo Ceriotti

    Corresponding author
    • Istituto di Biologia e Biotecnologia Agraria, Consiglio Nazionale delle Ricerche, Milano, Italy
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(e-mail ceriotti@ibba.cnr.it).

Summary

The glutenin fraction of wheat storage proteins consists of large polymers in which high- and low-molecular-weight subunits are connected by inter-chain disulfide bonds. We found that assembly of a low-molecular-weight glutenin subunit in the endoplasmic reticulum is a rapid process that leads to accumulation of various oligomeric forms, and that this assembly is sensitive to perturbation of the cellular redox environment. In endoplasmic reticulum-derived microsomes, low-molecular-weight glutenin subunits are subjected to hyper-polymerization, indicating that cytosolic factors play an important role in limiting polymer size. Addition of physiological concentrations of reduced glutathione is sufficient to maintain the original polymerization pattern of the glutenin subunits upon cytosol dilution. Furthermore, we show that a low-molecular-weight glutenin subunit can be glutathionylated in endoplasmic reticulum-derived microsomes, and that it can be directly reduced by glutathione in vitro. These results indicate that glutenin polymerization is sensitive to changes in the redox state of the cell, and suggest that the presence of a reducing cytosolic environment plays an important role in regulating disulfide bond formation in the endoplasmic reticulum of plant cells.

Introduction

Hexaploid and tetraploid wheats (Triticum spp.) are amongst the most widely grown and important cereal crops in the world. The unique ability of wheat flour to form dough that possesses the necessary rheological properties required for production of various foodstuffs is largely due to the characteristics of proteins that accumulate in the endosperm cells during seed development, which are the major component of gluten (Gianibelli et al., 2001; She et al., 2011). Gluten proteins play key roles in determining the unique baking quality of wheat by conferring water absorption capacity, cohesivity, viscosity and elasticity to dough. These proteins contain a high proportion of glutamate and proline residues, and may be divided into two functionally distinct groups on the basis of their polymerization status and solubility in aqueous alcohol: soluble monomeric gliadins and insoluble polymeric glutenins.

The precise number of individual gluten protein components is yet to be fully determined, but 2D gel analysis suggests that 100 is a reasonable estimate (Gianibelli et al., 2001). Gliadins are classified into four groups according to their primary sequences and electrophoretic mobility at low pH (α, β, γ and ω), with disulfide bonds being either absent (ω-type) or present only as intra-molecular disulfides (α-, β- and γ-types). By contrast, glutenins form heterogeneous mixtures of polymers, in which individual subunits are connected by inter-molecular disulfide bonds. The glutenins are also divided into four classes, based on their electrophoretic mobility and molecular weight (Payne and Corfield, 1979), namely the A-type high-molecular-weight glutenin subunits (molecular weight range 80–100 kDa) and the B-, C- and D-type low-molecular-weight glutenin subunits (LMW-GS; molecular weight range 20–45 kDa). These subunits cross-link to form glutenin polymers, which are amongst the largest protein molecules known in nature (Wrigley, 1996).

In common with other seed storage proteins, gliadin and glutenin subunits are secretory proteins that are synthesized on the rough endoplasmic reticulum (ER) and co-translationally imported into the ER lumen. Once within the lumen, wheat gluten proteins may follow two principal trafficking routes: a Golgi-dependent route that leads to deposition within protein bodies of vacuolar origin, or a Golgi-independent route in which protein deposits form in the ER lumen, and may ultimately fuse with the vacuole (Levanony et al., 1992; Rubin et al., 1992; Vitale and Ceriotti, 2004; Tosi et al., 2009, 2011; Francin-Allami et al., 2011).

A number of models of glutenin subunit assembly have been proposed (Gianibelli et al., 2001), but determination of the precise structure of glutenin polymers has been hampered by the complexity of the situation in wheat endosperm, in which many different subunits are simultaneously expressed. The study of glutenin polymer formation has thus taken advantage of heterologous expression systems, in which the behaviour of individual subunits may be more easily monitored. Previously, assembly of some high-molecular-weight glutenin subunits was studied in transgenic tobacco (Robert et al., 1989; Shani et al., 1994), and polymerization of a LMW-GS from Triticum aestivum cv. Cheyenne, known as B11-33, has also been investigated by utilizing transient expression in tobacco protoplasts (Lombardi et al., 2009). Inter-chain disulfide bonds were shown to be crucial for B11-33 polymer formation. However, rather than being governed by specific pairing between subunits, assembly appears to rely on a relaxed specificity with regard to disulfide bond formation, whereby each cysteine residue may form a disulfide bond with a variety of companion cysteines present on identical or different subunits (Lombardi et al., 2009). This would drive the formation of a large number of complex glutenin polymers. Together with data obtained from direct analysis of disulfide bonds present in gluten proteins (reviewed by Tatham and Shewry, 1997), these results have shed some light on the interactions that may control the structure of glutenin polymers. However, several aspects of the assembly process, such its kinetics, the source of oxidizing equivalents and the enzymes involved, remain to be clarified.

In mammalian cells, a number of oxidative pathways for disulfide bond formation are now known (Margittai and Bánhegyi, 2010; Bulleid and Ellgaard, 2011). The most thoroughly characterized to date involves oxidation of substrate proteins by protein disulfide isomerase (PDI), which is in turn oxidized by the endoplasmic reticulum oxidase Ero1, which was first identified in yeast (Frand and Kaiser, 1998; Pollard et al., 1998). It has recently been shown that plant homologues of Ero1 and PDI are also involved in storage protein deposition in rice (Onda et al., 2009, 2011; Satoh-Cruz et al., 2010), and that PDI plays an important role in glutenin polymerization during the breadmaking process (Koh et al., 2010). However, the pathways that mediate disulfide bond formation within the plant ER have not been characterized in detail.

Although glutathione was initially thought to act as an oxidant in the process of disulfide bond formation (Hwang et al., 1992), its actual role appears to be more complex (Chakravarthi et al., 2006; Appenzeller-Herzog et al., 2010). In both yeast and mammalian cells, glutathione has been proposed to play an important role as a reductant in the ER (Cuozzo and Kaiser, 1999; Jessop and Bulleid, 2004; Molteni et al., 2004), either directly reducing proteins as they fold, or maintaining ER oxidoreductases in the reduced state, allowing them to act in recovery of proteins containing non-native disulfides.

In wheat endosperm cells, polymer formation may continue after glutenins have been transported out of the ER. Indeed, although glutenin subunits have a large number of free thiol groups during the cell division and cell enlargement phases, these become oxidized during grain dehydration (Rhazi et al., 2003). This suggests that substantial oxidation of thiol groups, and hence polymer formation, involves subunits and small oligomers that have already been transported to the storage vacuole. Similar structural maturation of a multimeric protein by disulfide formation in storage vacuoles was also recently reported in the castor bean plant, Ricinus communis (Marshall et al., 2010). However, the mechanisms controlling these oxidation steps remain to be elucidated.

Here we have investigated the factors that control the polymerization of a LMW-GS in the plant cell ER. We found that oligomer formation is a rapid process, and that the ratio between the various polymeric forms is sensitive to treatment with reducing or oxidizing agents. We also present evidence indicating that the reducing cytosolic environment limits formation of high-molecular-weight polymers in the ER. These results indicate that mechanisms controlling the redox state of the ER play an important role in determining the polymerization state of LMW-GS subunits.

Results

Kinetics and redox regulation of LMW-GS assembly

The B11-33 clone from Triticum aestivum cv. Cheyenne encodes a B-type LMW-GS (Okita et al., 1985). The protein is synthesized as a precursor containing an N-terminal signal peptide for import into the ER, followed first by a short N-terminal region that contains the first cysteine residue (C25) involved in polymer formation, and then by a repetitive domain that is rich in glutamine and proline residues. The C-terminal part of the protein is stabilized by the presence of three intra-chain disulfide bonds, and contains one further cysteine residue (C230) that remains available for polymer formation (Figure 1) (Orsi et al., 2001). A triple-HA tag was added to the C-terminus of B11-33 to facilitate immunoprecipitation of the protein.

Figure 1.

Schematic structure of the B11-33-HA precursor. The positions of C25, C230 and the three intra-chain disulfide bridges are indicated. Non-repetitive regions are shaded. SP, signal peptide; HA, triple-HA tag.

We have previously shown that B11-33 assembly is mediated by formation of inter-chain disulfide bonds involving combinations of C25 and C230 (Lombardi et al., 2009). This results in accumulation of three types of dimer, together with a set of larger oligomeric forms. To monitor the kinetics of LMW-GS assembly, tobacco protoplasts expressing the B11-33-HA protein (Lombardi et al., 2009) were pulse labelled for 20 min with radioactive amino acids, and chased for various periods of time (Figure 2). Oligomeric forms were already present at the end of the pulse (Figure 2, lane 5), and the assembly pattern did not obviously change during the 5 h chase period (Figure 2, lanes 6–8). This indicates that assembly is a rapid process that reaches completion soon after synthesis. In addition to the various oligomeric forms of the B11-33-HA protein, other bands that are not present in the control lanes were evident at the 0 h chase point. The intensity of these bands decreased during the chase, suggesting that they represent adducts of the B11-33-HA monomer and oligomers with endogenous tobacco proteins. The identity of the proteins involved in these transient interactions was not studied further.

Figure 2.

Polymerization pattern and kinetics of assembly of LMW-GS. Tobacco protoplasts were transfected with empty vector (pD3) or with a plasmid encoding the B11-33-HA protein. After overnight incubation, protoplasts were radiolabelled with [35S]-cysteine and [35S]-methionine for 20 min, and chased for the indicated periods of time, before immunoselection of polypeptides from cell homogenates using anti-HA antibodies and analysis by non-reducing SDS-PAGE and fluorography. The positions of B11-33-HA monomers, dimers and oligomers are indicated on the right, and the positions of molecular mass markers (kDa) are shown on the left.

We next investigated whether the observed oligomerization pattern was sensitive to the action of redox-active compounds. In the C230S-HA mutant of the B11-33-HA protein, one of the two cysteine residues that are normally involved in inter-chain disulfide bond formation is mutated to serine; consequently, the protein can only form dimers. This simplifies the analysis, making it possible to monitor the effect of various redox-active agents simply by measuring the ratio between dimers and monomers. Protoplasts expressing the C230S-HA mutant were pulse labelled for 90 min and then incubated for 30 min in the presence of the reducing agents β-mercaptoethanol (β-ME) or dithiothreitol (DTT), or the oxidizing agent diamide (Figure 3). Each of these agents had a clear effect on the dimer/monomer ratio. Increasing concentrations of β-ME led to a decrease in the dimer/monomer ratio (Figure 3, lanes 2–4, and Table S1). DTT treatment not only affected accumulation of the dimers, but also partially reduced intra-chain disulfide bonds, as evidenced by a set of bands that migrate more slowly than the fully oxidized monomer (Figure 3, lanes 5–7) (Orsi et al., 2001). This reflects the fact that DTT is a stronger reducing agent than β-ME (Lees and Whitesides, 1993; Valetti and Sitia, 1994). By contrast, treatment with diamide had the opposite effect, resulting in a clear increase in the dimer/monomer ratio (Figure 3, lanes 8–10, and Table S1). Together, these results indicate that LMW-GS assembly status is sensitive to the redox conditions of the cell.

Figure 3.

Redox control of LMW-GS assembly. Tobacco protoplasts were transfected with plasmid encoding the C230S-HA protein. After overnight incubation, protoplasts were radiolabelled with [35S]-cysteine and [35S]-methionine for 90 min and chased for 30 min in the presence of increasing concentrations of reducing agent [β-mercaptoethanol (β-ME) or dithiothreitol (DTT)] or oxidizing agent (diamide). Polypeptides were immunoselected from cell homogenates using anti-HA antibodies and analysed by non-reducing SDS-PAGE. The positions of fully oxidized monomers, partially or fully reduced monomers, and dimers are indicated on the right, and the positions of molecular mass markers (kDa) are shown on the left.

We then investigated whether the polymerization of B11-33-HA was a reversible process by assessing whether B11-33-HA subunits were able to re-polymerize following reduction with β-ME. Although the previously described polymerization pattern for B11-33-HA or the C230S-HA mutant was observed prior to β-ME treatment (Figure 4, lanes 5 and 9, respectively), only the monomeric form accumulated following treatment (Figure 4, lanes 6 and 10). However, upon β-ME washout, the polymerization pattern for both B11-33-HA and C230S-HA was completely restored after 30 min (Figure 4, lanes 7 and 11). Control samples (Figure 4, lanes 8 and 12) revealed that subunits that are newly synthesized following β-ME washout polymerize or dimerize to the same extent as those synthesized prior to β-ME treatment. Reduction of B11-33-HA oligomers thus appears to be completely reversible, with inter-molecular disulfide bonds reforming once the reducing agent has been removed. This observation is consistent with recent findings in mammalian cells suggesting that the ER steady-state disulfide content is rapidly re-established over a short time scale after reductive challenge (Appenzeller-Herzog et al., 2010).

Figure 4.

Covalent assembly of LMW-GS is a reversible process. Tobacco protoplasts were transfected with empty vector (pD3) or with a plasmid encoding the B11-33-HA or C230S-HA proteins. After overnight incubation, protoplasts were radiolabelled with [35S]-cysteine and [35S]-methionine for 60 min and, after removal of an aliquot (control), were chased for 30 min in the presence of 10 mm β-mercaptoethanol (β-ME). After removal of a second aliquot (+ βME), remaining cells were washed three times with W5 medium (Ceriotti et al., 2003) to remove β-ME, and incubated for another 30 min (+ wash-out). A set of control protoplasts (+ wash-out*) were treated identically, with the exception that they were labelled for 60 min after β-ME treatment and washout rather than before. Polypeptides were immunoselected from cell homogenates using anti-HA antibodies and analysed by non-reducing SDS-PAGE. The positions of monomers, dimers and oligomers are indicated on the right, and the positions of molecular mass markers (kDa) are shown on the left.

The cytosol protects LMW-GS from hyper-oxidation

Among thiol-containing compounds, reduced glutathione (GSH) is a highly abundant low-molecular-mass metabolite that can accumulate at millimolar concentrations in the cytoplasm of plant cells (Meyer et al., 2001). As GSH has been proposed to be important in maintaining an ER oxidoreductase in a reduced state (Jessop and Bulleid, 2004), and as the polymerization pattern of B11-33-HA may reflect establishment of an equilibrium with one or more of these oxidoreductases and/or other redox couples, we determined whether the reducing cytosolic environment was involved in determining the B11-33-HA polymerization pattern. To assess the role of the cytosol in determining the polymeric status of LMW-GS, we took advantage of the fact that the cytosol content may be effectively diluted by homogenizing protoplasts in sucrose buffer, a procedure that preserves the integrity of the ER membrane and leads to formation of sealed ER-derived vesicles known as microsomes. When microsomes were isolated in the presence of the alkylating agent iodoacetamide to block exposed thiol groups, the typical polymerization pattern was observed (Figure 5a, top panel, lane 8). Conversely, when iodoacetamide was not included in the extraction buffer, no monomeric protein was evident, and the amount of dimeric protein was also greatly reduced (Figure 5a, top panel, lane 7). Analysis under reducing conditions showed that considerably less B11-33-HA protein is immunoselected from microsomes isolated in the absence of alkylating agents (Figure 5a, lower panel, compare lanes 7 and 8). These results indicate that, if thiol/disulfide reactions are not blocked by inclusion of an alkylating agent, cytosol dilution leads to hyper-oxidation of LMW-GS within microsomes, and hence formation of large polymers that cannot be efficiently extracted or immunoprecipitated. This interpretation is supported by analysis of the behavior of the C230S-HA mutant. In this case, homogenization in the absence of an alkylating agent leads to a net increase in the dimer/monomer ratio, again indicating that cytosol washout favors disulfide bond formation between LMW-GS (Figure 5a, top panel, compare lanes 13 and 14). In this case, however, the total amount of protein recovered was not drastically affected (Figure 5a, lower panel, compare lanes 13 and 14), confirming that the reduction in recovery of the B11-33-HA protein in the absence of alkylating agents is indeed linked to the ability of this protein to form large protein polymers.

Figure 5.

Reducing agents are required to maintain a native redox state in isolated microsomes. Tobacco protoplasts were transfected with empty vector (pD3) or with a plasmid encoding the B11-33-HA or C230S-HA proteins. After overnight incubation, protoplasts were radiolabelled with [35S]-cysteine and [35S]-methionine for 90 min.(a) Protoplasts were homogenized in 12% sucrose buffer to maintain the integrity of ER microsomes but dilute the cytosol, in the presence or absence of iodoacetamide or indicated concentrations of GSH. Polypeptides were immunoselected from cell homogenates using anti-HA antibodies and analysed by non-reducing (top panel) or reducing (lower panel) SDS-PAGE and fluorography. The positions of monomers, dimers and oligomers are indicated on the right.(b) Protoplasts expressing the B11-33-HA protein were homogenized in 12% sucrose buffer (sucrose) or in protoplast homogenization buffer (detergent) in the presence (+) or absence (−) of iodoacetamide. B11-33-HA polypeptides were immunoselected from cell homogenates using anti-HA antibodies and analysed by reducing SDS-PAGE and fluorography. The position of B11-33-HA monomers is indicated on the right. In both panels, the positions of molecular mass markers (kDa) are shown on the left.

To assess whether GSH was able to counterbalance the effect of cytosol removal, homogenization was performed in the absence of alkylating agents and in the presence of various concentrations of GSH (Figure 5a, lanes 9–12 and 15–18). The results clearly show that, in the case of both the B11-33-HA and C230S-HA proteins, inclusion of increasing concentrations of GSH effectively counteracts the hyper-oxidation observed upon cytosol dilution, and leads to a decrease in the dimer/monomer ratio (Table S2). Indeed, for both the polymerization-competent protein and the C230S-HA mutant, inclusion of 10 mm GSH produced a polymerization (or dimerization) pattern that was similar to that obtained in the presence of iodoacetamide (Figure 5a, top panel, compare lanes 8 and 10 and lanes 14 and 16). A higher GSH concentration led to greater accumulation of the monomeric protein.

We next determined whether confinement of the LMW-GS within microsomes was required for their hyper-oxidation. Protoplasts expressing the B11-33-HA protein or the C230S-HA mutant were homogenized in either a 12% sucrose buffer, in order to preserve microsome integrity, or a detergent-containing buffer, to solubilize the ER membranes. As hyper-oxidation of B11-33-HA subunits reduces their recovery by immunoprecipitation (Figure 5a), this feature was used to monitor the effect of the various homogenization conditions. As expected, when protoplasts were homogenized in sucrose buffer without detergent, recovery of the B11-33-HA protein was poor unless an alkylating agent was included in the homogenization buffer (Figure 5b, compare lanes 3 and 4). Conversely, omitting the alkylating agent did not affect the recovery of the B11-33-HA protein when a detergent was present in the homogenization buffer (Figure 5b, lanes 7 and 8). We conclude that confinement of the LMW-GS within ER-derived microsomes is required for their hyper-polymerization upon cytosol dilution.

GSH directly reduces LMW-GS oligomers in vitro

Together, these results indicate that maintenance of a reducing environment is important for preventing hyper-polymerization of LMW-GS within microsomes. Cytosolic thiol-containing compounds may protect LMW-GS from hyper-oxidation by scavenging oxidizing equivalents and/or by directly reducing ER-resident oxidoreductases and LMW-GS oligomers. It was therefore of interest to determine whether GSH directly reduces B11-33-HA oligomers in vitro. Immunoprecipitated B11-33-HA oligomers were incubated in vitro with various concentrations of GSH (Figure 6a). At concentrations of 5 or 2 mm (Figure 6a, lanes 14 and 15), no reduction of B11-33-HA oligomers was observed. However, some reduction was observed in the presence of 20 or 10 mm GSH (Figure 6a, lanes 12 and 13), indicating that GSH does indeed act directly on B11-33-HA disulfides. To assess whether another low-molecular-weight thiol-containing compound similarly reduced B11-33-HA oligomers, the activity of cysteine was also monitored. While relatively high GSH concentrations were necessary to observe substantial polymer reduction, cysteine was active even at concentrations as low as 2 mm (Figure 6a, lanes 17–20). Despite this, a small proportion of dimers appeared to be resistant to reduction even at high concentrations of cysteine or GSH.

Figure 6.

GSH directly reduces LMW-GS polymers.(a) Tobacco protoplasts were transfected with empty vector (pD3) or with a plasmid encoding the B11-33-HA protein. After overnight incubation, protoplasts were radiolabelled with [35S]-cysteine and [35S]-methionine for 60 min, before immunoselection of B11-33-HA polypeptides from cell homogenates using anti-HA antibodies. Immunoprecipitated proteins were then treated for 45 min with the indicated concentrations of GSH or cysteine, and analysed by non-reducing SDS-PAGE.(b) Cells were treated as in (a), with the exception that immunoprecipitated proteins were treated for 45 min in the presence (+) or absence (−) of 5 mm cysteine or 5 mm GSH that, where indicated, had been pre-incubated for 15 min with 0.2 mm NADPH and/or 2 U ml−1 glutathione reductase (GR). Asterisks indicate samples in which glutathione reductase was heat-inactivated at 95°C for 5 min prior to pre-incubation with GSH. In both panels, the positions of B11-33-HA monomers, dimers and oligomers are indicated on the right, and the positions of molecular mass markers (kDa) are shown on the left.

Because GSH and cysteine have similar reduction potentials (Keire et al., 1992), we hypothesized that GSH-mediated reduction may be inefficient, either because of kinetic factors or because of the known contamination of commercial GSH stocks with glutathione disulfide (Rabenstein, 2009). To assess whether inclusion of a known catalyst of thiol/disulfide reactions led to reduction of glutenin oligomers in the presence of low concentrations of GSH, bovine PDI was included in the reaction. However, no reduction was observed in the presence of 3.5 μm PDI and 1 mm GSH (Figure S1a). Under identical treatment conditions, PDI is able to reduce the disulfide bond between the A and B subunits of the mature cytotoxin ricin (Figure S1b, lane 16) (Spooner et al., 2004). Reduction of ricin holotoxin to its individual subunits is observed only after a 5 h chase (Figure S1b, compare lanes 12 and 16), as it is necessary for the intervening linker peptide between the A and B chains to be removed, which can only take place following deposition in the protein storage vacuole (Butterworth and Lord, 1983; Hiraiwa et al., 1997; Frigerio et al., 1998).

We then investigated whether contamination by glutathione disulfide may explain the differential effect of commercial GSH and cysteine preparations on B11-33-HA oligomers. Glutenin oligomers were efficiently reduced by 5 mm GSH in the presence of glutathione reductase and NADPH (Figure 6b, lane 18), that together efficiently reduce contaminating glutathione disulfide. Such a reduction was not observed when glutathione reductase was inactivated by heating at 95°C for 5 min (Figure 6b, lane 19), indicating that the activity of glutathione reductase in reducing glutathione disulfide is crucial for allowing polymer reduction by GSH. Together, these results indicate that GSH and cysteine can be directly involved in reduction of inter-chain disulfide bonds in glutenin polymers. As determined from the lack of any major effect on monomer mobility, the same compounds do not appear to significantly affect the intra-chain disulfides that stabilize the C-terminal domain of the B11-33-HA protein (Orsi et al., 2001).

Biotinylated GSH forms mixed disulfides with LMW-GS in ER-derived microsomes

If GSH does indeed directly reduce LMW-GS oligomers in microsomes, then a mixed disulfide intermediate between LMW-GS and GSH should be formed as part of the reduction process. To determine whether this intermediate is formed, we isolated microsomes from protoplasts expressing B11-33-HA or C230S-HA and resuspended them in a buffer containing BioGEE, a soluble ethyl ester of biotinylated glutathione. Formation of a mixed disulfide would thus result in biotinylation of the glutenin subunits. To determine whether such a mixed disulfide is formed, microsomes were incubated in the presence of BioGEE for 60 min and then treated with iodoacetamide to trap any mixed disulfides and quench excess BioGEE. Biotinylated proteins were then isolated on streptavidin-agarose beads and resolved by reducing SDS-PAGE. In the presence of GSH rather than BioGEE, no B11-33-HA or C230S-HA subunits were isolated from cell homogenates (Figure 7, lanes 3 and 5), as expected due to the absence of a biotin moiety. However, in the presence of BioGEE, a band corresponding in size to B11-33-HA and C230S-HA subunits was isolated from protoplasts transfected with plasmids encoding these proteins (Figure 7, lanes 4 and 6) but not from control protoplasts (lane 2). This result indicates that the biotinylated reagent acted upon oxidized B11-33-HA and C230S-HA, forming a mixed disulfide. This provides further evidence that GSH directly reduces disulfide bonds in LMW-GS within ER-derived microsomes.

Figure 7.

GSH forms a mixed disulfide with LMW-GS subunits in isolated microsomes. Tobacco protoplasts were transfected with empty vector (pD3) or with plasmid encoding either the B11-33-HA or C230S-HA proteins. After overnight incubation, protoplasts were radiolabelled with [35S]-cysteine and [35S]-methionine for 60 min, and microsomes were isolated by homogenization in 12% sucrose buffer in the absence of iodoacetamide, followed by fractionation through a 17% sucrose pad. Microsomes were resuspended in either 10 mm GSH (lanes 1, 3 and 5) or 10 mm BioGEE (lanes 2, 4 and 6), and incubated for 45 min at 25°C. Biotinylated proteins were isolated using streptavidin-agarose beads, and analysed by reducing SDS-PAGE. The arrowhead indicates the position of reduced B11-33-HA and C230S-HA monomers. The positions of molecular mass markers (kDa) are shown on the left.

Discussion

Despite the important role of wheat storage proteins in determining the functional characteristics of wheat flour, the mechanisms that control the structure and size of glutenin polymers are still poorly understood. As glutenin subunits are held together by inter-chain disulfide bonds, the mechanisms regulating disulfide bond formation in the ER must play a crucial role in the overall process. ER-resident oxidoreductases (Houston et al., 2005) and plant homologues of the mammalian and yeast Ero1 proteins (Dixon et al., 2003; Onda et al., 2009) are possible players in the pathway leading to covalent assembly of glutenin subunits, as is GSH, which has been proposed to play a role in thiol/disulfide exchange reactions, and in depolymerization of glutenins in wheat dough during the breadmaking process (reviewed by Joye et al., 2009).

Our results indicate that glutenin subunit assembly is a rapid process, leading to formation of a set of oligomeric forms in which the subunits are connected by inter-chain disulfide bonds. The ratio between these forms is very sensitive to perturbations of the cellular redox environment, and strong hyper-oxidation of glutenin subunits is observed when microsomes are prepared in the absence of an alkylating agent, suggesting that the equilibrium between oxidizing and reducing pathways is somehow altered under these conditions. Inclusion of iodoacetamide in the homogenization buffer is essential to prevent hyper-polymerization during microsome preparation, but not when protoplasts are homogenized in the presence of detergent. As an uncharged molecule, iodoacetamide is membrane-permeable, and, once within microsomes, will react, albeit slowly, with free thiol groups present in LMW-GS monomers and oligomers (Hansen and Winther, 2009). This will hamper any further disulfide-mediated assembly reaction. Presumably, when protoplasts are homogenized in the presence of detergent, the dilution of LMW-GS and all other ER components efficiently precludes further polymerization, even in the absence of an alkylating agent.

Studies performed in yeast (Cuozzo and Kaiser, 1999) and mammalian cells (Jessop and Bulleid, 2004; Molteni et al., 2004) support the conclusion that GSH delivers reducing equivalents to the ER, thus balancing the activity of oxidative systems that are active in this compartment. Although microsome preparation probably preserves the activity of ER-located sulfhydryl oxidases, such as the plant Ero1 homologues (Dixon et al., 2003), the very large dilution of cellular content would drastically affect the surrounding environment. Conceivably, this may result in both import of oxidizing equivalents into ER-derived microsomes and a large reduction in the import of cytosolic low-molecular-weight thiol-containing compounds such as GSH, which, in mammalian cells, is thought to permeate the ER membrane via facilitated diffusion (Bánhegyi et al., 1999), possibly taking advantage of the inherent permeability of this membrane to small molecules (Le Gall et al., 2004). The observation that inclusion of physiological concentrations of GSH is sufficient to maintain the native polymerization pattern within isolated microsomes is consistent with the view that GSH is one of the crucial cellular factors that is diluted during homogenization. Thus, the actual polymerization pattern observed in living cells is controlled by the activity of oxidizing systems present within the ER, but is also dependent on the presence of a reducing cytosol that may either constitute a protective environment or play a more active role as a source of reduced thiols for import into the ER. Imported thiol-containing low-molecular-weight metabolites may act by reducing ER oxidoreductases and/or by directly reducing glutenin polymers. GSH rapidly reduces mammalian PDI in vitro (Lappi and Ruddock, 2011), and in vivo direct reduction of an ER oxidoreductase has been demonstrated in mammalian cells (Jessop and Bulleid, 2004). We found that biotinylated GSH binds to LMW-GS in isolated microsomes, and that inter-chain, but not intra-chain, disulfide bonds in polymerized LMW-GS are readily reduced by GSH. Consistent with these results, cysteine residues involved in LMW-GS polymerization were found to be the main site for GSH binding in wheat glutenins (Hüttner and Wieser, 2001). While these findings indicate that GSH potentially affects the polymerization state of glutenin subunits, they do not exclude the possibility that the main role of GSH is exerted in the cytosol, where it may protect the ER from hyper-oxidation without directly acting on ER proteins.

Various lines of evidence indicate that formation of glutenin polymers occurs during at least two independent stages of wheat grain maturation (Rhazi et al., 2003; Tosi et al., 2011). First, during the cell division and enlargement phase, SDS-extractable polymeric proteins are formed (known as level 1 polymers). These correspond to smaller oligomers, similar to those observed in this study. A second step, occurring during grain desiccation (approximately 33 days after anthesis), leads to formation of larger aggregates by entanglement. These are stabilized by hydrogen bonding and additional disulfide bridges. Correspondingly, free thiol groups in storage proteins increase during the cell division and expansion phases and decline during the desiccation/maturation phase. Previous results have implicated glutathione in the accumulation of insoluble protein polymers during the desiccation phase of grain development (Rhazi et al., 2003). Although GSH is mainly present in its reduced form during the cell enlargement phase, the desiccation phase is accompanied by a decrease in glutathione reductase activity and a rapid increase in glutathione disulfide content. In addition, the level of polymeric protein-glutathione mixed disulfide was found to be low during the cell enlargement phase, to increase rapidly during the desiccation phase, and to be negatively correlated with polymer size, fraction of SDS-unextractable polymeric protein, and breadmaking performance (Rhazi et al., 2003; Li et al., 2004). Our results suggest that GSH may play a role in the formation of level 1 polymers by limiting their overall size and thus controlling their solubility characteristics. This, in turn, may affect the deposition and intracellular transport of glutenin polymers. Direct analysis of the glutathionylation level of glutenin subunits in the ER of wheat endosperm cells would help to clarify the role of this low-molecular-weight thiol in the early stages of wheat storage protein deposition.

In general, our results indicate that changes in the redox status of the cell have an impact on polymer assembly. For example, although intra-chain disulfide bonds are resistant to treatment with the reducing agent β-ME, inter-chain disulfide bonds linking B11-33-HA subunits are easily reduced. Conversely, diamide treatment favours covalent subunit assembly. If the observed polymerization pattern reflects a redox-regulated equilibrium, it is possible that physiological changes in the pathways that control ER redox homeostasis affect the molecular weight distribution of glutenin oligomers in the ER of wheat endosperm cells. In yeast, different challenges to protein folding, including prevention of glycosylation, inositol deprivation and increased protein secretion, cause ER oxidative folding stress (Merksamer et al., 2008). Endosperm cells are subjected to an enormous protein load during seed maturation, and it is possible that changing environmental conditions affect the efficiency of protein folding or other biosynthetic pathways, thus having an effect on ER redox. While these changes may not affect the formation of stable disulfide bonds, they may affect less stable ones, such as those linking glutenin subunits within the ER, and hence influence the molecular weight distribution of level 1 oligomers. In yeast, a redox-sensitive probe was found to be in a more reduced state during ER stress (Merksamer et al., 2008). In Arabidopsis, heat activates an ER stress response (Gao et al., 2008; Deng et al., 2011). This raises the possibility that exposure of wheat endosperm cells to ER stress conditions may affect glutenin polymer formation. Heat stress has been shown to affect flour, dough and baking quality, and this has been related to both an increased gliadin/glutenin ratio (Blumenthal et al., 1991, 1993) and a decrease in the proportion of higher-molecular-weight glutenin polymers (Wardlaw et al., 2002). While heat may affect wheat grain quality through many different mechanisms, it would be interesting to investigate whether stress-induced changes in ER redox contribute to the reduction in the size of glutenin polymers that is observed when wheat plants are subjected to high temperatures during the grain filling period.

Experimental Procedures

Recombinant DNA

Clone B11-33 (EMBL/GenBank accession number M11077), which encodes a low-molecular-weight glutenin subunit from the bread wheat Triticum aestivum cv. Cheyenne, has been described previously (Okita et al., 1985). Plasmids for the transient expression of B11-33-HA and the C230S-HA mutant in the pD3 expression vector have also been described previously (Lombardi et al., 2009). A plasmid encoding preproricin (EMBL/GenBank accession number X02388) in the expression vector pDHA (Tabe et al., 1995) was also reported previously (Lamb et al., 1985; Frigerio et al., 1998).

Protoplast isolation, transfection and labelling

Isolation of tobacco protoplasts from 4 to 7 cm long leaves of axenic Nicotiana tabacum cv. Petit Havana SR1 (Maliga et al., 1973), polyethylene glycol-mediated transfection and labelling with [35S]-Promix in vitro cell labelling mix (GE Healthcare, www.gehealthcare.com) or EasyTag EXPRESS 35S protein labelling mix (Perkin Elmer, www.perkinelmer.com) were performed essentially as described previously (Pedrazzini et al., 1994; Ceriotti et al., 2003). For each transfection, 40 μg of plasmid per 106 protoplasts was used. Where indicated, unlabelled cysteine and methionine were added to cells following radioactive labelling, and protoplasts were incubated for 30 min at 25°C in K3 medium (Ceriotti et al., 2003) containing the indicated concentrations of β-ME, DTT or diamide (from 100× stocks in H2O).

Immunoprecipitation of radiolabelled proteins

Frozen samples were homogenized by addition of protoplast homogenization buffer (Ceriotti et al., 2003) supplemented with Complete protease inhibitor cocktail (Roche Applied Sciences, www.roche.com), 1.5 mm phenylmethylsulfonyl fluoride and 70 mm iodoacetamide. Samples were then clarified by centrifugation at 13 000 g for 5 min at 4°C, and diluted to a final volume of 1 ml in NET-Gel buffer (50 mm Tris/HCl pH 7.5, 150 mm NaCl, 1 mm EDTA, 0.1% Igepal CA-630 (Sigma-Aldrich, www.sigmaaldrich.com), 0.25% gelatin, 0.02% NaN3). Proteins were immunoselected from protoplast homogenates using 12CA5 anti-HA monoclonal antibodies (Roche Applied Sciences) or anti-ricin antiserum (Vector Laboratories, www.vectorlabs.com), and either Protein A-Sepharose CL4B (GE Healthcare) or Protein G-Agarose (Invitrogen, www.invitrogen.com). Immunoselected polypeptides were analysed by SDS-PAGE under reducing or non-reducing conditions (Orsi et al., 2001), and visualized by fluorography. Alternatively, bands were detected using an FLA-9000 image scanner (Fujifilm, www.fujifilm.com). Band intensity was determined using TotalLab Quant software (www.totallab.com).

Preparation of ER microsomes

To monitor the effect of GSH on the polymerization state of glutenin subunits within ER microsomes, protoplast pellets (containing 500 000 cells) were resuspended in 550 μl 12% w/w sucrose in step gradient buffer (100 mm Tris/HCl pH 7.6, 10 mm KCl, 1 mm EDTA, supplemented with Complete protease inhibitor cocktail) containing 0 or 70 mm iodoacetamide or 0, 2, 10, 20 or 40 mm GSH, and homogenized by pipetting 40 times with a Gilson-type micropipette through a 200 μl tip, followed by passing the sample four times through a 25G needle. The samples were centrifuged at 100 g for 5 min at 4°C, and the supernatant (containing intact microsomes and cytosol) was diluted in protoplast homogenization buffer supplemented with Complete protease inhibitor cocktail and 1.5 mm phenylmethylsulfonyl fluoride. Samples were clarified by centrifugation at 13 000 g for 5 min, gelatin was added to a final concentration of 0.25% w/v, and proteins were immunoselected using 12CA5 anti-HA monoclonal antibody and analysed by SDS-PAGE as described above.

Detection of biotinylated glutathione (BioGEE) conjugates

To fully isolate microsomes, protoplast pellets were resuspended in 12% w/w sucrose in step gradient buffer, and disrupted by pipetting as described above. Intact cells and debris were removed by centrifugation at 500 g for 5 min at 4°C. The supernatant (400 μl) was loaded onto a 17% w/w sucrose pad in step gradient buffer, and centrifuged at 100 000 g for 30 min in an SW55 Ti rotor (Beckman Coulter, www.beckmancoulter.com) at 4°C. Pellets (microsomes) were resuspended in 30 μl of 12% w/w sucrose in step gradient buffer containing 10 mm GSH or 10 mm BioGEE (Molecular Probes, www.molecularprobes.com), and incubated at 25°C for 45 min. Iodoacetamide was added to a final concentration of 70 mm, and samples were again incubated as above. Samples were frozen in liquid N2 and stored at -80°C. After thawing, excess BioGEE was removed by precipitation of proteins using an equal volume of 20% trichloroacetic acid. Protein pellets were washed twice with ice-cold acetone, and resuspended in 100 μl denaturing alkylating lysis buffer (50 mm Tris/HCl pH 7.4, 5 mm EDTA, 1% SDS, supplemented with Complete protease inhibitor cocktail, 1 mm phenylmethylsulfonyl fluoride and 10 mm iodoacetamide). Samples were then heated at 95°C for 5 min, diluted with nine volumes of non-denaturing lysis buffer (50 mm Tris/HCl pH 7.4, 300 mm NaCl, 5 mm EDTA, 1% Triton X-100, 10 mm iodoacetamide), and insoluble material was removed by centrifugation at 13 000 g for 5 min at 4°C. Bovine serum albumin (BSA) was added to 0.1% final concentration, and samples were incubated with 2.5% w/v streptavidin-agarose (Sigma-Aldrich) for 2 h at 4°C. Beads were washed four times with protoplast homogenization buffer, and precipitated proteins were analysed by SDS-PAGE as described above.

In vitro treatment of immunoprecipitated proteins

Immune complexes bound to Protein A-Sepharose or Protein G-agarose beads were washed once each with NET-Gel buffer containing 0.5 m NaCl, NET-Gel buffer containing 0.1% SDS, and sterile PBS. Beads were then incubated at 25°C for 45 min with the indicated concentrations of GSH or cysteine. Iodoacetamide was added to a final concentration of 70 mm, and samples were again incubated as above. Where indicated, GSH was pre-incubated with 0.2 mm NADPH and/or 2 U ml−1 glutathione reductase (from baker's yeast; Sigma-Aldrich). Alternatively, after washing, beads were incubated at 30°C for 30 min in the presence or absence of 1 mm GSH and either 3.5 μm BSA or 3.5 μm bovine PDI, as previously described (Spooner et al., 2004), followed by a further incubation at 30°C for 30 min in the presence of 70 mm iodoacetamide. Radiolabelled proteins were then analysed by SDS-PAGE as described above.

Acknowledgements

We wish to thank Anna Paola Casazza, Stefania Masci (Department of Agriculture, Forests, Nature and Energy, Università degli Studi della Tuscia, Viterbo, Italy) and Jakob Winther (Department of Biology, University of Copenhagen, Denmark) for helpful discussion and critical reading of the manuscript. This work was supported by grants from the Ministero dell'Istruzione, dell'Università e della Ricerca, projects RBNE01TYZF (Fondo per gli Investimenti della Ricerca di Base) and AGROGEN (Fondo per le Agevolazioni alla Ricerca).

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