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. 2021 Sep 27;188(1):241–254. doi: 10.1093/plphys/kiab457

Two protein disulfide isomerase subgroups work synergistically in catalyzing oxidative protein folding

Fenggui Fan 1,2, Qiao Zhang 3, Yini Zhang 4,5, Guozhong Huang 1, Xuelian Liang 3, Chih-chen Wang 4,5,2, Lei Wang 4,5,2,#, Dongping Lu 1,3,2,✉,#
PMCID: PMC8774737  PMID: 34609517

Abstract

Disulfide bonds play essential roles in the folding of secretory and plasma membrane proteins in the endoplasmic reticulum (ER). In eukaryotes, protein disulfide isomerase (PDI) is an enzyme catalyzing the disulfide bond formation and isomerization in substrates. The Arabidopsis (Arabidopsis thaliana) genome encodes diverse PDIs including structurally distinct subgroups PDI-L and PDI-M/S. It remains unclear how these AtPDIs function to catalyze the correct disulfide formation. We found that one Arabidopsis ER oxidoreductin-1 (Ero1), AtERO1, can interact with multiple PDIs. PDI-L members AtPDI2/5/6 mainly serve as an isomerase, while PDI-M/S members AtPDI9/10/11 are more efficient in accepting oxidizing equivalents from AtERO1 and catalyzing disulfide bond formation. Accordingly, the pdi9/10/11 triple mutant exhibited much stronger inhibition than pdi1/2/5/6 quadruple mutant under dithiothreitol treatment, which caused disruption of disulfide bonds in plant proteins. Furthermore, AtPDI2/5 work synergistically with PDI-M/S members in relaying disulfide bonds from AtERO1 to substrates. Our findings reveal the distinct but overlapping roles played by two structurally different AtPDI subgroups in oxidative protein folding in the ER.

Two subgroups of Arabidopsis protein disulfide isomerases exhibit differences but work synergistically in catalyzing the formation of native disulfide bonds in proteins in the endoplasmic reticulum.

Introduction

As the entrance to the protein secretory pathway, the endoplasmic reticulum (ER) is the organelle where secreted and plasma membrane (PM) proteins are synthesized, modified, folded, and subjected to quality control, which ensures that only correctly folded proteins exit the ER. Accumulation of unfolded proteins in the ER will cause ER stress, which results in activation of unfolded protein response (UPR) to promote folding capacity of the ER and restore stressed cells to their normal states (Vitale and Boston, 2008; Howell, 2013; Sun et al., 2021).

It is known that disulfide bond formation plays important role in protein folding in the ER (Bulleid and Ellgaard, 2011; Smith et al., 2011). In eukaryotic cells, protein disulfide isomerase (PDI) (EC 5.3.4.1) is an enzyme that catalyzes the disulfide bond formation and isomerization in substrate proteins. It contains four thioredoxin (Trx) domains in the order of a–b–b′–a′, of which, both the a and a′ domains have -CGHC- active sites (Hatahet and Ruddock, 2009). The Arabidopsis (Arabidopsis thaliana) genome encodes at least 14 PDI family members that are grouped into 6 PDI subfamilies: A, B, C, L, M, and S (Figure 1). The PDI-L subfamily consists of AtPDI1, AtPDI2, AtPDI3, AtPDI4, AtPDI5, and AtPDI6, sharing the standard structure a–b–b′–a′, with two catalytic Trx domains a and a′ and two noncatalytic domains b and b′. Both PDI-M and PDI-S members contain three domains, with the a domain and the a′ domain adjacent to each other. The PDI-M members, AtPDI9 and AtPDI10, share the domain arrangement a–a′–b (Houston et al., 2005; Lu and Christopher, 2008). Arabidopsis PDI-S consists of only one member, AtPDI11, with the domain structure a–a′–D. The D domain of AtPDI11 is similar to the C-terminal domain of human ERp29/ERp28, which is responsible for binding to the P domain of the lectin chaperones calreticulin and calnexin in human (Sakono et al., 2014; Kozlov et al., 2017; Nakao et al., 2017). AtPDI11 has no orthologs in yeast and animals. Recently, we found that AtPDI11 has protein oxidative activity in vitro (Fan et al., 2018). In addition, there are five AtPDIs (AtPDI7, AtPDI12, AtPDI13, AtPDI8, and AtPDI14) that have only one putative catalytic Trx domain. AtPDI8, the only member of the PDI-B subfamily, contains one putative transmembrane domain and was found to be a type I membrane protein (Selles et al., 2011; Yuen et al., 2016a, 2016b).

Figure 1.

Figure 1

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Schematic representation of AtPDIs. Boxes marked with a and a′ represent the catalytic Trx domains of AtPDIs that contain the indicated active sites; b and b′ represent the noncatalytic Trx domains of AtPDIs. TMD: transmembrane domain. D: the D domain of AtPDI11.

ER oxidoreductin-1 (Ero1) proteins (Ero1p for yeast; Ero1α and Ero1β for mammals) serve as the major disulfide donors for PDIs in the ER (Meyer et al., 2019). The a′ domain of human PDI could be oxidized by human Ero1α (Baker et al., 2008; Wang et al., 2009; Chambers et al., 2010) or Ero1β (Wang et al., 2011). Recently, we found that Arabidopsis Ero1s, AtERO1, and AtERO2 are required to mediate oxidative protein folding in the ER; and AtERO1 likely acts as the primary sulfhydryl oxidase. Furthermore, AtPDI9 is oxidized by the outer active site of AtERO1, in turn, the outer active site is re-oxidized by AtERO1’s inner active site (Fan et al., 2019). In yeast, a hierarchy of interaction exists between Ero1p and the multiple Pdi1p homologs (Vitu et al., 2010). Mammalian Ero1s have a high specificity for PDI (Wang et al., 2011; Araki et al., 2013; Zhang et al., 2014). However, it remains unclear how oxidizing equivalents are transferred from AtERO1 to structurally diverse PDIs in Arabidopsis; and in turn, how AtPDIs function to guarantee efficient and faithful disulfide formation.

In this work, we find that PDI-L and PDI-M/S subgroups have distinct but overlapping activities: it is likely that AtPDI2/5/6 mainly serve as an isomerase, while AtPDI9/10/11 are more efficient in transferring oxidizing equivalents from AtERO1 to catalyze disulfide formation in the substrates. Furthermore, AtPDI2/5 and AtPDI9/10/11 work synergistically in oxidative protein folding in the ER. Our work will advance our understanding of the redox networks in the ER of Arabidopsis.

Results

AtERO1 interacts with multiple PDIs

In yeast and mammals, a hierarchy of interaction or specificity of interaction exists between Ero1s and the multiple PDI homologs (Vitu et al., 2010; Benham et al., 2013). However, it is still unknown how many PDIs require AtERO1/2 to provide disulfide bonds in Arabidopsis. AtPDI3 and AtPDI4 contain the nonclassical type of active sites -CARS- in a domain, and -CXNC- in the a′ domain (X represents V or I) (Figure 1). The soybean AtPDI3 ortholog (GmPDIL-3) and its wheat ortholog have no oxidative folding activities in vitro (Iwasaki et al., 2009; Kimura et al., 2015). In the following studies, we focused our investigation on PDI-L members AtPDI1/2/5/6, and PDI-M/S members AtPDI/9/10/11.

We first performed co-immunoprecipitation (co-IP) assays to analyze the association of AtERO1 with multiple AtPDIs. The results showed that AtERO1 was associated with all tested AtPDIs (Figure 2, A and B). Of note, the size of some AtPDIs, like AtPDI1/2, is greater than their expected molecular weight, which could be caused by unknown post-translational modification (Supplemental Table S1; Figure 2B). However, AtERO1 was not associated with SP-RFP-FLAG-KDEL, an ER-localized red fluorescence protein with the signal peptide (SP) of PDI11 at the N-terminus and an ER retention signal KDEL (Lys-Asp-Glu-Leu) at the C-terminus (Huang et al., 2019; Figure 2, A and C).

Figure 2.

Figure 2

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Association of AtERO1 with multiple AtPDIs. A, Schematic representation of the constructs harboring AtPDI CDSs. B, Analysis of the association of AtERO1 with multiple AtPDIs by Co-IP assays. AtERO1 was fused to an HA tag, and PDIs were fused to a FLAG tag. AtERO1 and AtPDIs were co-expressed in Arabidopsis protoplasts. AtERO1-HA was co-expressed with an untagged GFP in Arabidopsis protoplasts, serving as the control. AtPDIs were immunoprecipitated with an anti-FLAG antibody; and the associated proteins were analyzed by immunoblotting with an anti-HA antibody (top two parts). The expression of PDI-FLAG and AtERO1-HA in Arabidopsis protoplasts (input of Co-IP assays) was analyzed by immunoblotting with an anti-FLAG and anti-HA antibody, respectively, and was shown in the bottom two parts. C, AtERO1 was not associated with SP-RFP-FLAG-KDEL. SP-RFP-FLAG-KDEL is an ER-localized red fluorescence protein with the SP derived from PDI11 at the N-terminus and an ER retention signal KDEL at the C-terminus. AtERO1-HA was co-expressed with AtPDI11-FLAG or SP-RFP-FLAG-KDEL in Arabidopsis protoplasts. Co-IP assays were performed as described in (B).

Moreover, we examined whether recombinant His–FLAG–AtPDIs could be pulled down (PD) by AtERO1 protein fused to a GST tag. The results of glutathione S-transferase (GST) pull-down (PD) assays demonstrated that AtERO1 can directly bind to all AtPDIs tested (Figure 3, A–G). These results suggest that at least AtPDI1, AtPDI2, AtPDI5, AtPDI6, AtPDI9, AtPDI10, and AtPDI11 could directly interact with AtERO1.

Figure 3.

Figure 3

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Direct interactions between AtERO1 and multiple AtPDIs. Recombinant full-length AtPDIs fused with His and FLAG tags were purified using Ni-NTA agarose. GST-ERO1 or GST proteins immobilized on GSH beads were incubated with purified His–FLAG–PDI proteins for 2 h; and GST PD assays were performed. Then the washed beads were immunoblotted with anti-FLAG antibodies to detect His–FLAG–AtPDI proteins binding to GST-ERO1 (upper part). GST-ERO1/GST proteins immobilized on GSH beads and His–FLAG–PDI proteins used in these PD assays were stained with Coomassie Brilliant Blue (lower part). A, GST-ERO1 + His–FLAG–PDI1. B, GST-ERO1 + His–FLAG–PDI2. C, GST-ERO1 + His–FLAG–PDI5. D, GST-ERO1 + His–FLAG–PDI6. E, GST-ERO1 + His–FLAG–PDI9. F, GST-ERO1 + His–FLAG–PDI10. G, GST-ERO1 + His–FLAG–PDI11. H, AtERO1 interacts with the a–a′ region of AtPDI11, but not the D domain of AtPDI11. GST PD assays were performed between GST-AtERO1 and His–FLAG–PDI11, His–FLAG–PDI11ΔD (a PDI11 truncation lacking the D domain), or His–FLAG–PDI11-D (the AtPDI11 D domain).

It was found that the b′xa′ region of human PDI provided the essential binding site for Ero1α (Wang et al., 2009). Although AtPDI11 has no b and b′ domain, we showed that it also interacted with AtERO1 (Figures 2, B and 3, G). To determine which regions of AtPDI11 are responsible for its binding to AtERO1, we examined the interaction between AtERO1 and the AtPDI11 D domain or a PDI11 truncation lacking the D domain (PDI11ΔD; Figure 1). GST PD assays showed that the a–a′ region of AtPDI11, but not the D domain of AtPDI11 provided the binding sites for AtERO1 (Figure 3H).

AtERO1 prefers to oxidize PDI-M/S rather than PDI-L members

To examine how oxidizing equivalents are transferred from AtERO1 to diverse AtPDIs, we examined the oxidative folding activities of PDI-L members AtPDI1/2/5/6, and PDI-M/S members AtPDI9/10/11 in the presence of AtERO1. The oxidative folding activities of AtPDIs were analyzed in vitro by monitoring oxygen consumption in the AtERO1-AtPDIs-GSH system, where reduced glutathione (GSH) was used as the substrate of these AtPDIs. AtERO1 oxidized AtPDI9 most efficiently and oxidized AtPDI10/11 to a lesser extent. AtERO1 also weakly oxidized PDI5/6 and only slightly oxidized AtPDI2 and AtPDI1. Overall, PDI-M/S members are oxidized by AtERO1 much more efficiently than PDI-L members (Figure 4, A and B; Supplemental Figure S1).

Figure 4.

Figure 4

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AtERO1 prefers to oxidize PDI-M/S rather than PDI-L members. A, Oxidation of AtPDIs by AtERO1 as determined by oxygen consumption assays. GSH was supplied as reducing equivalents. The reaction contains AtERO1, AtPDIs, GSH, and FAD; Ctrl, reaction without AtPDIs. B, Relative activities were calculated by measuring the slope of the linear phase of the oxygen consumption curve in (A) after subtracting the Ctrl. Values are the means of three biological repeats ± se (standard error) (n = 3). Statistical significance among PDIs was determined by one-way analysis of variance (ANOVA), followed by Tukey’s post hoc test (P < 0.05); and different letters indicate significant difference. C, Analysis of AtPDIs activities with gel-based RNase A refolding assay. RNase A was separated by 15% nonreducing sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) followed by Coomassie Blue staining. Dred: denatured and reduced RNase A; Pox: partially oxidized RNase A; Fox: fully oxidation RNase A. Ctrl: reaction without AtPDIs.

Native ribonuclease A (RNase A) contains four disulfide bridges. The denatured and reduced (Dred) RNase A has frequently been used as an artificial substrate of PDI (Ruoppolo et al., 1996). We performed gel-based Dred RNase A oxidation assays using the AtERO1-AtPDIs-RNase A in vitro system. The results showed that in the presence of AtERO1, PDI-M/S members AtPDI9/10/11 oxidized the Dred RNase A much faster than PDI-L members AtPDI1/2/5/6 (Figure 4C). These results suggest that AtERO1 oxidizes AtPDI9/10/11 more efficiently than AtPDI1/2/5/6, and AtPDI9/10/11 are more efficient in catalyzing disulfide formation in substrates, by using oxidizing equivalents from AtERO1. It is noteworthy that AtPDI1/2/5/6, especially AtPDI5, also exhibited weak activities to oxidize the Dred RNase A, and the partially oxidized RNase A (Pox) mainly appeared at 30 min of the reaction, as assayed by the gel-based Dred RNase A oxidation (Figure 4C).

PDI-L subfamily members AtPDI2/5/6 mainly serve as an isomerase

Because human PDI can catalyze both disulfide formation and isomerization, we then determined to analyze the isomerase activities of AtPDIs. We examined the reactivation of scrambled RNase A by different AtPDIs through monitoring the hydrolysis of cyclic cytidine 3',5'-monophosphate (cCMP), the substrate of RNase A. Scrambled RNase A contains a set of nonnative disulfide bonds, which was obtained by oxidizing the Dred RNase A in a nonenzymatic manner (Supplemental Figure S2), and has been employed as a model to investigate the refolding of disulfide-containing proteins (Lyles and Gilbert, 1991; Ruoppolo et al., 1996). In this system, reactivation of scrambled RNase A is determined by the conversion of oxidized intermediates into the native state by the isomerase activity of PDIs. Interestingly, PDI-L subfamily members AtPDI2/5/6 were more active than PDI-M/S members in terms of isomerase activity, with AtPDI1 as an exception (Figure 5, A and B). Notably, PDI-M/S members also had weak isomerase activity, with AtPDI10 being the lowest one (Figure 5, A and B).

Figure 5.

Figure 5

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AtPDI2/5/6 are much more active than AtPDI9/10/11 in terms of isomerase activity. A, Analysis of AtPDIs activities with scrambled RNase A reactivation assay using GSH/GSSG as an oxidizing equivalent. Reactivation of scrambled RNase A was determined by monitoring the hydrolysis of cCMP. The reaction containing only GSH/GSSG was used as the Ctrl. B, Relative isomerase activities were calculated by measuring the slope of the linear phase of the reactivation curve in (A) after subtracting the Ctrl. Data present the means of three biological repeats ± se (n = 3). statistical significance among AtPDIs was determined by one-way ANOVA, followed by Tukey’s post hoc test (P < 0.05), and was indicated by different letters. C, ANS fluorescence spectra assay of AtPDIs. A total of 50-μM ANS was incubated with 5-μM AtPDIs for 20 min at 25°C in the dark. ANS emission spectra were determined at excitation at 370 nm. Ctrl: without PDIs. D, Enhancement factor was quantified according to the ANS fluorescence spectra in (C). Values are the means of three biological repeats ± se (n = 3). Statistical significance among AtPDIs was determined by one-way ANOVA, followed by Tukey’s post hoc test (P < 0.05), and indicated by different letters.

In human and yeast PDI, the b′ domain contains a hydrophobic pocket which is the principal binding site for substrates, and is necessary for the isomerase activity of PDI (Pirneskoski et al., 2004). As PDI-M/S members lack the b′ domain, we want to know whether their lower isomerase activity is related to their substrate-binding abilities. We performed a 1-anilinonaphthalene-8-sulfonic acid (ANS) fluorescence assay, in which, the binding of ANS to the hydrophobic regions of proteins results in a marked increase in fluorescence. We found that the ANS fluorescence intensity of AtPDI1/2/6 was higher than that of AtPDI9/10/11. The ANS fluorescence of AtPDI5 was significantly higher than that of AtPDI9/11, and was close to that of AtPDI10 (Figure 5, C and D). These results indicate that overall AtPDI1/2/5/6 has more hydrophobic surfaces exposed than PDI-M/S members that lack the b′ domain.

Growth phenotype analysis of AtPDI1/2/5/6 and AtPDI9/10/11 mutants under reducing conditions or ER stress

Dithiothreitol (DTT) is a dithiol reagent that can disrupt the disulfide bonds of plant proteins (Martinez and Chrispeels, 2003; Lu and Christopher, 2008). The plant growth was inhibited by DTT in a dose-dependent manner (Supplemental Figure S3). To study the function of AtPDIs genetically, we analyzed the phenotypes of different mutant lines of AtPDIs genes (Supplemental Figures S4 and S5) under DTT treatment. We first analyzed the mutants of genes encoding PDI-M/S subgroup members that mainly catalyze the disulfide formation. The overall growth phenotypes of the pdi9, pdi10, and pdi11 single mutants were similar to that of Col-0 under normal conditions (Supplemental Figure S6). When they were grown on Murashige Skoog (MS) medium containing 1.4-mM DTT, only the growth of pdi11 mutant plant was dramatically inhibited as previously reported (Fan et al., 2018; Supplemental Figure S6). Moreover, pdi9/10 double mutant grew as wild-type (WT) plants under 1.2 or even 2.4-mM DTT treatment (Figure 6). However, knockout of AtPDI9/10 in the pdi11 background exacerbated the growth inhibition of mutant plants by DTT, as the growth of pdi9/10/11 seedlings were inhibited by DTT more severely than that of pdi11 (Figure 6, C and D).

Figure 6.

Figure 6

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The growth phenotypes of PDI9/10/11 gene mutants under DTT treatment. A, The growth phenotypes of Col-0, pdi9, pdi10, and pdi9/10 double mutant grown on 1/2 MS medium with or without 2.4-mM DTT for 7 d. Scale bar, 1 cm. B, Quantitative analysis of the fresh weight of Col-0 and mutants shown in (A). Values are the means ± se (n = 30 seedlings). Statistical significance compared to Col-0 was determined using one-way ANOVA followed by Student–Newman–Keuls tests (P > 0.05), ns, not significant. C, Knockout of AtPDI9/10 exacerbated the growth inhibition of pdi11 mutant plants under DTT treatment. Col-0, pdi11, pdi9/10, and pdi9/10/11 triple mutant seedlings were grown on 1/2 MS medium with or without 1.2-mM DTT for 7 d. Scale bar, 1 cm. D, Quantitative analysis of the fresh weight of Col-0 and mutants shown in (C). Values are the means ± se (n = 30 seedlings). Statistical significance was determined using one-way ANOVA followed by Student–Newman–Keuls tests, ***P < 0.001.

For the mutants of genes encoding PDI-L subfamily members, the overall growth phenotypes of the pdi1, pdi2, pdi5, and pdi6 were similar to that of Col-0 under normal conditions or when grown on MS medium containing 1.4 or 2.4-mM DTT (Figure 7, A–D; Supplemental Figure S6). However, both pdi1/2 and pdi5/6 double mutants exhibited more severe growth inhibition by DTT compared with Col-0 plants (Figure 7, A–D). Furthermore, the growth of pdi1/2/5/6 quadruple mutant was inhibited by DTT more significantly than that of pdi1/2 or pdi5/6 (Figure 7, E and F). These results suggest that PDI1/2/5/6 are functionally redundant. Notably, under normal conditions, pdi1/2/5/6 seedlings were slightly smaller than Col-0 (Figure 7, E and F).

Figure 7.

Figure 7

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The growth phenotypes of PDI1/2/5/6 gene mutants under DTT treatment. A, The growth phenotypes of Col-0, pdi1, pdi2, and pdi1/2 double mutant grown on 1/2 MS medium with or without 2.4-mM DTT for 7 d. Scale bar, 1 cm. B, Quantitative analysis of the fresh weight of Col-0 and mutants shown in (A). Values are the means ± se (n = 30 seedlings). Statistical significance compared to Col-0 was determined using one-way ANOVA followed by Student–Newman–Keuls tests, ***P < 0.001. C, The growth phenotypes of Col-0, pdi5, pdi6, and pdi5/6 double mutant grown on 1/2 MS medium with or without 2.4-mM DTT for 7 d. Scale bar, 1 cm. D, Quantitative analysis of the fresh weight of Col-0 and mutants shown in (C). Values are the means ± se (n = 30 seedlings). Statistical significance compared to Col-0 was determined using one-way ANOVA followed by Student–Newman–Keuls tests, ***P < 0.001. E, The growth of pdi1/2/5/6 quadruple mutant was inhibited by DTT more significantly than that of pdi1/2 and pdi5/6 double mutants. Plants were grown on 1/2 MS medium with or without 2.4 mM DTT for 7 d. Scale bar, 1 cm. F, Quantitative analysis of the fresh weight of Col-0 and mutants shown in (E). Values are the means ± se (n = 30 seedlings). Statistical significance between the samples was determined by one-way ANOVA followed by Student–Newman–Keuls tests, **P < 0.01, ***P < 0.001.

We have shown that AtPDI2/5/6 and AtPDI9/10/11 exhibited differences in catalyzing oxidative protein folding (Figures 4 and 5). To distinguish the functions of these two PDI subgroups genetically, we compared the growth inhibition of pdi1/2/5/6 and pdi9/10/11 by DTT treatments. We found that the growth of pdi9/10/11 exhibited much stronger inhibition than pdi1/2/5/6 when grown on 1/2 MS medium containing 1.4-mM DTT (Figure 8, A and B).

Figure 8.

Figure 8

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Comparison of growth phenotypes of pdi1/2/5/6 and pdi9/10/11 under reducing conditions and Tm treatment. A, Col-0, pdi1/2/5/6, and pdi9/10/11 were grown on 1/2 MS medium with or without 1.4 mM DTT for 7 d. Scale bar, 1 cm. B, Quantitative analysis of the fresh weight of Col-0 and mutants shown in (A). Values are the means ± se (n = 30 seedlings). Statistical significance compared to Col-0 was determined by one-way ANOVA followed by Student–Newman–Keuls tests, *P < 0.05, ***P < 0.001. C, Col-0, pdi1/2/5/6, pdi9/10/11, and bzip28/60 were grown on 1/2 MS medium with or without 50-ng/mL Tm for 7 d. Scale bar, 1 cm. D, The percentage of small yellow, small-green, and large-green seedlings in (A) were calculated. Values are the means ± se (n = 3). Statistical significance compared to Col-0 was determined by one-way ANOVA followed by Student–Newman–Keuls tests, ***P < 0.001.

The disruption of disulfide bonds caused by DTT treatment can lead to the accumulation of unfolded proteins in the ER and cause ER stress, which triggers UPR (Martinez and Chrispeels, 2003). To examine whether the phenotypes of pdi1/2/5/6 and pdi9/10/11 under DTT treatment were due to their impaired ability to respond to ER stress, we analyzed these mutants under Tunicamycin (Tm) treatment, which triggers UPR by inhibiting N-linked glycosylation, a prerequisite for the folding of glycoproteins (Takatsuki et al., 1971). We found that the UPR mutant bzip28 bzip60 (Deng et al., 2013) was more sensitive to Tm treatment compared with Col-0 (Figure 8, C and D). Interestingly, pdi1/2/5/6 plants were also more sensitive to Tm treatment than Col-0 plants, while the extent of pdi9/10/11 growth inhibition by Tm was comparable to that of Col-0 (Figure 8, C and D). Both AtERO1 and AtERO2 are involved in oxidative protein folding in the ER; and ero1-3 and ero2-2 were more sensitive to DTT treatment compared with Col-0 plants (Fan et al., 2018). However, the mutants of AtERO1/2 were inhibited by Tm to an extent comparable to Col-0 (Supplemental Figure S7).

Taken together, these results suggest that the phenotypes of pdi9/10/11 under DTT treatment are likely caused by the impaired capacity of oxidative protein folding, and AtPDI9/10/11 mainly function in cooperation with AtERO1 to introduce disulfides. On the other hand, Tm treatment causes the accumulation of misfolded proteins in the ER, and these proteins are prone to form non-native disulfide bonds (Asker et al., 1998). Therefore, the phenotypes of pdi1/2/5/6 under Tm treatment are likely the consequence of the impaired ability to catalyze the isomerization of these incorrect disulfide bonds, suggesting that AtPDI1/2/5/6 mainly serve as an isomerase that may help refold the misfolded proteins in the ER. Importantly, these observations support the results obtained by biochemical assays, except for AtPDI1 (Figures 4 and 5).

Synergistic relationship of different AtPDIs

Previous studies have revealed that there is a hierarchy of cooperative redox interactions among ER oxidoreductases in mammalian cells (Araki et al., 2013; Kojima et al., 2014; Zhang et al., 2014). Here, we investigated the cooperation of different AtPDIs of by examining the reactivation of the Dred RNase A. It is known that during the reactivation of Dred RNase A by the yeast PDI, Dred RNase A is first converted into the oxidized form by the oxidized PDI, and then into its native state by the PDI’s isomerase activity (Wilkinson et al., 2005). We first investigated the reactivation of Dred RNase A by individual AtPDIs. Among the PDI-M/S members, AtPDI9 and AtPDI11 showed much higher activities than AtPDI10 (Supplemental Figure S8). This could be due to the extremely weak isomerase activity of AtPDI10, although it has high oxidative activity (Figures 4 and 5). Among the PDI-L members, AtPDI5 was much more efficient than other PDIs to reactivate Dred RNase A (Supplemental Figure S8). AtPDI5 shares 58% amino acid sequence similarity and 34% identity with mammalian PDI, being the highest among all AtPDIs (Edgar, 2004); and human PDI catalyzes both the disulfide bond formation and isomerization in substrate proteins (Wallis and Freedman, 2013). We showed that AtPDI5 exhibited the second strongest isomerase activity among all AtPDIs tested (Figure 5, A and B). Relative to other PDI-L members, AtPDI5 had the highest oxidative activity (Figure 4). Therefore, it is very likely that the efficient reactivation of Dred RNase A catalyzed by AtPDI5 is due to the coordination of its oxidase and isomerase activities. As expected, AtPDI1 only exhibited very faint activity to reactivate Dred RNase A (Supplemental Figure S8). Together, these results suggest that several AtPDIs are able to coordinate the oxidase and the isomerase activity in disulfide bond formation and isomerization.

Next, we sought to determine whether the combination of two PDIs, especially a PDI-L subfamily member and a PDI-M/S subfamily member will have synergistic effect on their activities. In the presence of AtERO1, when AtPDI2 and one of PDI-M/S members were added together, the reactivation of Dred RNase A was much greater than with each PDI alone, and was also greater than the sum of the activities of two single PDIs (Figure 9, A–C; Supplemental Figure S9, A–C). Similar results were obtained when AtPDI5 was combined with one of PDI-M/S members (Figure 9, D–F; Supplemental Figure S9, D–F). However, when two PDI-M/S members, AtPDI9 and AtPDI10 were added together, no such synergistic effect on their activities was observed (Figure 9, G; Supplemental Figure S9, G). These results suggest that AtPDI2–AtPDI9/10/11 or AtPDI5–AtPDI9/10/11 may work synergistically in relaying disulfide bonds from AtERO1 to substrates. Surprisingly, when two PDI-L members, AtPDI5 and AtPDI2 were added together, they also function synergistically in reactivating Dred RNase A (Figure 9H;Supplemental Figure S9H). This could be due to the relatively high oxidative activity of AtPDI5 (although much lower than that of PDI-M/S members; Figure 4B), which synergistically works with AtPDI2 in the reactivation of Dred RNase A.

Figure 9.

Figure 9

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Synergistic relationships between different AtPDIs. Analysis of cooperative protein-refolding activities of two different AtPDIs. Two AtPDIs were added together with the denatured and reduced RNase A. Reactivation of the denatured and reduced RNase A was determined by monitoring hydrolysis of cCMP. AtPDI combinations are AtPDI2 + AtPDI9 (A), AtPDI2 + AtPDI10 (B), AtPDI2 + AtPDI11 (C), AtPDI5 + AtPDI9 (D), AtPDI5 + AtPDI10 (E), AtPDI5 + AtPDI11 (F), AtPDI9 + AtPDI11 (G), and AtPDI2 + AtPDI5 (H). “Mix” represents the data obtained under conditions in which two AtPDI members were mixed in the reaction system. “Calc” represents the sum of the values of two individual PDIs. The experiments were repeated for three times.

Discussion

In eukaryotic cells, to ensure that a secretory or a PM protein is properly folded and equipped with the correct disulfide bonds, the actions of a set of ER catalysts or residents are coordinated, including various oxidoreductases, chaperones, and other folding enzymes (Sevier and Kaiser, 2008). PDI family proteins are important for catalyzing correct disulfide bond formation in the ER. Although the activities and functions of mammalian PDIs have been extensively studied, the roles of AtPDIs are largely unknown. Based on the domain structures, AtPDIs are grouped into several subfamilies, of which PDI-L subfamily has the structure of a–b–b′–a′, while PDI-M/S has adjacent a and a′ domains but lacking the b′ domain (Figure 1). In yeast and human, the b′ domain supplies the principal substrate binding site and is necessary for the isomerase activity of PDI (Pirneskoski et al., 2004). The ANS fluorescence assay showed that PDI-L subfamily members AtPDI1/2/5/6 overall have more hydrophobic surfaces exposed than PDI-M/S members AtPDI9/10/11 (Figure 5, C and D). Accordingly, AtPDI2/5/6 are much more active than PDI-M/S members in terms of isomerase activity (Figure 5, A and B), while AtPDI9/10/11 are more efficient in transferring oxidizing equivalents from AtERO1 and in catalyzing disulfide formation (Figure 4). Notably, AtPDI9/10/11 also have relatively weak isomerase activity; and AtPDI2/5/6, especially AtPDI5, also display oxidative activity, albeit much weaker than PDI-M/S members (Figures 4 and 5). Furthermore, several AtPDIs, like AtPDI5, are able to coordinate the oxidase and the isomerase activity in disulfide bond formation and isomerization (Supplemental Figure S8). Although AtPDI1 also has the same domain structure as AtPDI2/5/6, it has very low activity, likely because the active site in the a domain of AtPDI1 is -CGAC-, instead of the canonical -CGHC- motif (Figures 1, 4, and 9). Taken together, these results suggest that the two structurally distinct subfamilies of AtPDIs exhibit different but overlapping functions in catalyzing oxidative protein folding.

Almost all AtPDIs tested in this work have oxidative activities and interact with AtERO1 (Figures 2–4). Notably, different AtPDIs display varying oxidative activities (Figure 4). However, all of them, including AtPDI1, associate with AtERO1 with very similar affinities (Figure 2). This suggests that the rate-limiting factor for oxidization of AtPDIs by AtERO1 is not the association between AtERO1 and AtPDIs, but the distinct structures of different AtPDIs, or the coordination of AtERO1 and AtPDIs for oxidative reactions.

Interestingly, we have found that the combination of AtPDI2/5 and AtPDI9/10/11 shows the maximum ability to reactivate Dred RNase A in the presence of AtERO1 (Figure 9), indicating that PDI-L and PDI-M/S family members work synergistically to guarantee efficient and faithful disulfide formation. Our observation is in line with previous report showing that soybean PDIs can also function synergistically in folding unfolded proteins. Although a PDI-L member GmPDIL-2 has no oxidative refolding activity, it associates with the PDI-M member GmPDIM and they cooperatively refold the substrates (Matsusaki et al., 2016). However, whether AtPDI-L members could associate with PDI-M/S members and how these numerous AtPDIs coordinate in catalyzing oxidative protein folding in vivo needs to be determined in the future.

Application of DTT leads to the disruption of disulfide bonds of plant proteins (Martinez and Chrispeels, 2003; Lu and Christopher, 2008). Therefore, when the plants are treated with DTT, the enzymes that mediate the oxidative protein folding are required to restore the disrupted disulfide bonds in proteins; and the plants with impaired capacity to catalyze oxidative protein folding will suffer from DTT treatment more severely and exhibit stronger growth inhibition compared with WT plants. Based on such a presumption, we verified genetically that AtERO1 and AtERO2 function in oxidative protein folding, as their mutants showed much more substantial inhibition by DTT than Col-0 plants (Fan et al., 2019). Therefore, the phenotypic analysis of plants upon DTT treatment could be applied to examine gene’s function in modulating oxidative protein folding. In this work, we found that the growth of both pdi9/10/11 triple mutant and pdi1/2/5/6 quadruple mutant was inhibited by DTT more severely than Col-0 plants. Furthermore, pdi9/10/11 exhibited much stronger inhibition by DTT than pdi1/2/5/6 (Figure 8, A and B).

It is known that DTT treatment not only disrupts the disulfide bonds in proteins, the consequent accumulation of unfolded proteins in the ER also triggers UPR (Lu and Christopher, 2008). Basic leucine zipper transcription factor 28 (bZIP28) and bZIP60 are two mediators of plant UPR signaling (Liu et al., 2007; Deng et al., 2011). It was found that the growth of bzip28 bzip60 double mutant was strongly inhibited by DTT-induced ER stress (Deng et al., 2013). Besides DTT, another chemical, Tm induces UPR in plants through inhibiting N-linked glycosylation, which is a prerequisite for the folding of glycoproteins (Takatsuki et al., 1971). bzip28 bzip60 was more sensitive to Tm treatment compared with WT (Figure 8, C and D). Similarly, pdi1/2/5/6 plants were also more severely inhibited by Tm than Col-0. In contrast, pdi9/10/11 plants were inhibited by Tm to an extent comparable to Col-0, and so were the mutants of AtERO1/2 (Figure 8, C and D;Supplemental Figure S7).

Together, these results suggest that the phenotypes of pdi9/10/11 under DTT treatment are likely to be caused mainly by the impaired capacity of oxidative protein folding in this triple mutant plant. On the other hand, the phenotypes of pdi1/2/5/6 under Tm treatment are likely the consequence of the impaired ability of this quadruple mutant to catalyze isomerization of the nondisulfide bonds formed on the misfolded proteins. In contrast, the phenotypes of pdi1/2/5/6 under DTT treatment may be caused by the impaired isomerase activity of this quadruple mutant for dealing with overaccumulation of misfolded proteins in the ER, or by the impaired capacity of oxidative protein folding, because PDI/2/5/6 also showed weak oxidase activity (Figure 4). However, we also do not exclude the possibility that PDI1/2/5/6 can also act as molecular chaperones assisting the protein folding in the ER.

In plants, multiple seed storage proteins have been found to be the substrates of plant PDIs. In rice, the major seed storage protein glutelin may serve as the substrate of PDIL1-1, the putative rice ortholog of AtPDI5, which is involved in the maturation of glutelin (Takemoto et al., 2002; Houston et al., 2005). Native disulfide bonds are barely formed in glutelins in the seeds of the esp2 rice mutant that is deficient in PDIL1-1 (Onda et al., 2009). Rice PDIL2 and PDIL3 play important roles in the accumulation of a seed storage protein, Cys-rich 10-kDa prolamin (Onda et al., 2011). Here, we found that two Arabidopsis PDI subclasses PDI-L and PDI-M/S exhibit distinct activities in catalyzing oxidative protein folding in the ER, where secreted and PM proteins are folded. However, which secreted and PM proteins could serve as the substrates of the AtPDIs needs to be identified.

Materials and methods

Plant materials and growth conditions

WT Arabidopsis (A. thaliana; Col-0) and AtPDIs mutant [pdi1, SALK_150463; pdi2, SALK_115574; pdi5, SALK_136642C; pdi6, SAIL_430_A04; pdi9, SAIL_776_E01; pdi10, SALK_206219C; pdi11, SALK_148421 (Fan et al., 2018); pdi1/2, pdi5/6, pdi9/10, pdi1/2/5/6 and pdi9/10/11], ero1-3, ero2-2 (Fan et al., 2019), and bzip28 bzip60 UPR mutant (Deng et al., 2013) were grown on soil at 22°C in a growth chamber with a 12-/12-h light/dark photoperiod and 60% relative humidity. For seedlings grown on medium, seeds were germinated on 1/2 MS medium (1% [w/v] sucrose, 0.8% [w/v] agar, and 2.5-mM MES at pH 5.7). The plates were first kept at 4°C for 2 d to break dormancy and then moved to the growth chamber under the same conditions as the soil-grown plants. For DTT or Tm treatment, plants were grown on 1/2 MS medium with different concentrations of DTT or Tm for 7 d.

RNA extraction, cDNA synthesis, and reverse transcription-polymerase chain reaction (RT-PCR)

The total RNA was isolated using TRIzol (Invitrogen, Waltham, MA, USA) following the manufacturer’s instructions. The cDNA was synthesized by reverse transcriptase (TaKaRa, Dalian, China) from 1-μg DNase I-treated total RNA. The RT-PCR primer sequences are listed in Supplemental Table S2.

Plasmid construction, recombinant protein expression, and purification

For protoplast transfection, the full-length coding sequence (CDS) of AtPDI1, AtPDI2, AtPDI5, or AtPDI6 was amplified by RT-PCR from Col-0 RNA, and cloned into the pHBT plant expression vector fused to a FLAG tag at the BamHI and StuI sites. The KDEL ER retention signal was fused to the FLAG tag and inserted at the C-terminus, thereby generating the vectors Pro35S::PDI1/2/5/6-FLAG-KDEL. For AtPDI 10, Pro35S::PDI10-FLAG-KDDL was generated with its ER retention signal at the C-terminus at the BamHI and StuI sites. Pro35S::PDI9-FLAG-KDEL or Pro35S::PDI11-FLAG was generated as described previously (Fan et al., 2018, 2019). Pro35S::RFP-FLAG-KDEL was generated as described previously (Huang et al., 2019). The cloning of the full-length CDS of AtERO1 into a plant expression vector has been described previously (Fan et al., 2019).

For the expression of recombinant proteins in Escherichia coli, AtPDIs were subcloned into a pET-28a vector fused to the 6× His and FLAG tags at their N-terminus at the BamHI and StuI sites (Han et al., 2017). The CDSs of AtPDI11 D domain and a PDI11 truncation lacking the D domain (PDI11ΔD) were amplified from the full-length AtPDI11 cDNA. Then they were cloned into the aforementioned pET-28a vector. The AtERO1 CDS (without the SP) was inserted into the pGEX-6P-1 vector as described previously (Fan et al., 2019). All the primers are listed in Supplemental Table S2.

The recombinant GST-ERO1 proteins were expressed and purified as previously described (Wang et al., 2009). Then GST-ERO1 proteins were digested by PreScission Protease (Amersham Pharmacia Biosciences) to remove the GST moiety (Wang et al., 2009). The expression of recombinant His–FLAG–AtPDIs in E. coli cells (BL21) was induced by 0.5-mM isopropyl thiogalactoside at room temperature for 10 h. Then His–FLAG–AtPDI proteins were purified using a Ni-NTA agarose purification kit (QIAGEN, Hilden, Germany) following the manufacturer’s instructions.

Arabidopsis protoplast isolation, transfection, and co-IP assays

Arabidopsis protoplast isolation and transfection, co-IP assays were performed as described previously (Huang et al., 2019).

Glutathione-S-transferse (GST) pull-down (PD) assays

Recombinant full-length AtPDIs fused with His and FLAG tags were purified using Ni-NTA agarose. GST-ERO1 was used as a bait to PD His–FLAG–AtPDIs. About 5-μg His–FLAG–AtPDIs and 10-μg GST-ERO1 proteins immobilized on GSH beads were incubated in a buffer containing 20-mM Tris–HCl (pH 7.5), 3-mM EDTA, 150-mM NaCl, and 0.5% (v/v) Triton X-100 for 2 h at 4°C. Then, the GSH beads were washed 5 times with a buffer containing 20-mM Tris–HCl (pH 7.5), 3-mM EDTA, 150-mM NaCl. Purified GST protein was used as a negative control. Bound His–FLAG–AtPDI proteins were detected by immunoblotting using an anti-FLAG antibody (Sigma, St Louis, MO, USA).

Oxygen consumption assay

The oxygen consumption assays were performed in buffer B (100-mM Tris–HAc, 50-mM NaCl, and 1-mM EDTA, pH 8.0) as previously described (Wang et al., 2009; Fan et al., 2019). GSH was used as the substrate (Baker et al., 2008).

Preparation of denatured and reduced RNase A

The denatured and reduced RNase A (Dred RNase A) was obtained as described (Lyles and Gilbert, 1991).

Gel-based RNase A reoxidation assay

Gel-based RNase A reoxidation analyses were carried out as previously described (Wang et al., 2009; Fan et al., 2019).

Preparation of scrambled RNase A

Scrambled RNase A was prepared as described previously (Akiyama et al., 1992). The Dred RNase A was incubated in buffer C (6–M Gdn–HCl, 0.1-M Tris–HCl, pH 8.8) in dark at room temperature >60 h, then was desalted in Buffer D (50-mM Tris–HCl, pH 7.6, 150-mM NaCl, 2-mM EDTA). The concentration of scrambled RNase A was determined by its absorbance at 278 nm using a NanoDrop 2000 Spectrophotometer.

Dred RNase A reactivation assay

The Dred RNase A reactivation assay was performed by incubating 3-μM AtPDIs, 3-μM AtERO1, 4.5-mM cCMP, 100-μM FAD, and 8-μM Dred RNase A in buffer B at 25°C. RNase A reactivation was determined by monitoring the absorbance increase at 296 nm and 25°C due to the hydrolysis of cCMP, using a Shimadzu UV-2700 UV–Vis Spectrophotometer.

Scrambled RNase A reactivation assay

The scrambled RNase A reactivation assay was performed by incubating 3-μM AtPDIs, 4.5-mM cCMP, 100-μM FAD, 1-mM GSH, 0.2-mM GSSG, and 8-μM scrambled in buffer B at 25°C. RNase A reactivation was determined as described in “Dred RNase A reactivation assay.”

ANS fluorescence assay

The ANS fluorescence assay was performed by incubating 50-μM ANS in 50-mM Tris–HCl (pH 7.6) and 150-mM NaCl with or without 5-μM AtPDIs for 20 min at 25°C in the dark. The fluorescence emission spectra of ANS were detected in the range of 400–600 nm with excitation at 370 nm. The concentration of ANS was determined based on the extinction coefficient at 350 nm of 5,000 m−1cm−1. The enhancement factor was calculated as previously described (Yu et al., 2020).

Accession numbers

Sequence data from this article can be found in the GenBank data libraries under the following accession numbers: AtERO1, At1G72280; AtERO2, At2G38960; AtPDI1, At3G54960; AtPDI2, At5G60640; AtPDI5, At1G21750; AtPDI6, At1G77510; AtPDI9, At2G32920; AtPDI10, At1G04980; AtPDI11, At2G47470; Mutants used in this article can be obtained from ABRC under the following accession numbers: pdi1, SALK_150463; pdi2, SALK_115574; pdi5, SALK_136642C; pdi6, SALK_430_A04; pdi9, SAIL_776_E01; pdi10, SALK_206219C; pdi11, SALK_148421; bzip60, SAIL_283_B03; bzip28, SALK_132285C; ero1-3, SALK_096805; ero2-2, SALK_074861.

Supplemental data

The following materials are available in the online version of this article.

Supplemental Figure S1. Expression and purification of recombinant His–FLAG–AtPDIs fusion proteins.

Supplemental Figure S2. Scrambled RNase A, Dred RNase A, and oxidized RNase A after separation by nonreducing SDS–PAGE.

Supplemental Figure S3. Plant growth was inhibited by DTT in a dose-dependent manner.

Supplemental Figure S4. AtPDI gene structure and characterization of the AtPDI mutants.

Supplemental Figure S5. Detection of AtPDI transcripts in the mutant plants.

Supplemental Figure S6. Growth of AtPDI single mutants on medium with or without DTT.

Supplemental Figure S7. Growth of AtERO1/2 mutant plants on medium with or without Tm.

Supplemental Figure S8. Reactivation of Dred RNase A by AtPDIs in the presence of AtERO1.

Supplemental Figure S9. Synergistic relationships between different AtPDIs.

Supplemental Table S1. The predicted number of amino acids and expected size of each AtPDI.

Supplemental Table S2. Primers used in this study.

Funding

This work was supported by Chinese Ministry of Science and Technology (2017YFA0504000), the National Natural Science Foundation of China (31571260, 32022033), the Strategic Priority Research Program of Chinese Academy of Sciences (XDB37020303), China Postdoctoral Science Foundation (2019M663795), and the State Key Laboratory of Plant Genomics of China.

Conflict of interest statement. The authors declare that there is no conflict of interest.

Supplementary Material

kiab457_Supplementary_Data
Click here for additional data file. (2.7MB, pdf)

 

D.L., L.W., and C.W. conceived the project. F.F., L.W., and D.L. designed research. F.F., Q.Z., Y.Z., G.H., and X.L. performed research. F.F., L.W., and D.L. analyzed data. D.L., L.W., C.W., and F.F. wrote the manuscript.

The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (https://academic.oup.com/plphys/pages/general-instructions) is: Dongping Lu (dplu@sjziam.ac.cn).

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