Comprehensively Characterizing the Thioredoxin Interactome In Vivo Highlights the Central Role Played by This Ubiquitous Oxidoreductase in Redox Control

Abstract

Thioredoxin (Trx) is a ubiquitous oxidoreductase maintaining protein-bound cysteine residues in the reduced thiol state. Here, we combined a well-established method to trap Trx substrates with the power of bacterial genetics to comprehensively characterize the in vivo Trx redox interactome in the model bacterium Escherichia coli. Using strains engineered to optimize trapping, we report the identification of a total 268 Trx substrates, including 201 that had never been reported to depend on Trx for reduction. The newly identified Trx substrates are involved in a variety of cellular processes, ranging from energy metabolism to amino acid synthesis and transcription. The interaction between Trx and two of its newly identified substrates, a protein required for the import of most carbohydrates, PtsI, and the bacterial actin homolog MreB was studied in detail. We provide direct evidence that PtsI and MreB contain cysteine residues that are susceptible to oxidation and that participate in the formation of an intermolecular disulfide with Trx. By considerably expanding the number of Trx targets, our work highlights the role played by this major oxidoreductase in a variety of cellular processes. Moreover, as the dependence on Trx for reduction is often conserved across species, it also provides insightful information on the interactome of Trx in organisms other than E. coli.

Thioredoxins (Trxs)¹ are small antioxidant enzymes that catalyze the reduction of disulfide bonds that form in substrate proteins either as part of a catalytic cycle (see below) or following exposure to reactive oxygen species (ROS). As such, Trxs are involved in many different cellular processes, ranging from the defense against oxidative stress to the regulation of numerous signal transduction pathways and the modulation of the inflammatory response (1, 2).

Trxs have been identified in most living organisms, including archaea, bacteria, plants and mammals (1). They all share a canonical WCGPC catalytic motif located on a highly conserved fold, which consists of five β-strands surrounded by four α-helices (Fig. 1A) (3). The cysteine residues of the WCGPC motif are the key players used by Trxs to break disulfide bonds in substrate proteins. In this reaction, the first cysteine performs a nucleophilic attack on an oxidized substrate, which results in the formation of a mixed-disulfide intermediate between Trx and its substrate. This intermediate is then resolved by a nucleophilic attack of the second cysteine of the WCGPC motif, leading to the oxidation of thioredoxin and the release of the reduced substrate (1) (Fig. 1B). The catalytic cysteine residues are then converted back to the reduced state by the flavoenzyme thioredoxin reductase at the expense of the reduced form of nicotinamide adenine dinucleotide phosphate (NADPH).

Fig. 1.

Trapping of Trx1 substrates in a ΔtrxAΔtrxC E. coli strain in vivo. A, Structure of reduced E. coli thioredoxin (PDB code: 1XOB). All thioredoxins share a canonical WCGPC catalytic motif located on a highly conserved fold, which consists of five β-strands (light orange) surrounded by four α-helices (cyan). The cysteines of the catalytic motif are represented in orange. The figure was generated using MacPyMol (Delano Scientific LLC 2006). B, Reaction mechanism of disulfide reduction by thioredoxin. The reaction takes off with a nucleophilic attack of the N-terminal cysteine of the conserved WCGPC motif targeting the disulfide. As a result, an intermediate mixed disulfide complex is formed between Trx and the substrate protein, which in turn is reduced by a nucleophilic attack of the C-terminal cysteine of the WCGPC motif. C, To trap proteins linked to Trx1, Trx1_WCGPA was expressed in a ΔtrxAΔtrxC mutant. The proteins were TCA-precipitated and analyzed by Western blotting using an anti-His antibody. Induction of Trx1_WCGPA led to the accumulation of several high-molecular weight complexes, which disappeared following addition of DTT. The molecular mass markers (in kDa) are indicated on the left. D, The mixed-disulfide complexes were purified by affinity chromatography using Ni-NTA agarose. After concentration, the proteins trapped by Trx1_WCGPA were separated by SDS-PAGE (nonreducing dimension). The complexes were then separated in a second, reducing dimension. Proteins were identified by mass spectrometry. The spots analyzed by mass spectrometry were numbered as indicated (see Table I). The molecular mass markers (in kDa) are indicated on the left. The inset shows the reducing pathway that was genetically modified. Here, by deleting the trxA and trxC genes, it is the Trx pathway that was impaired. Trx, thioredoxin; TR, thioredoxin reductase; Grx, glutaredoxin; gshA encodes the gamma-glutamylcysteine synthetase; gshB encodes the glutathione synthetase; GR, glutaredoxin reductase.

Escherichia coli Trx1, encoded by the gene trxA, is the first Trx that was discovered (4, 5). Being constitutively expressed in the cytoplasm, it reduces disulfide bonds that form during the catalytic cycle of important enzymes such as ribonucleotide reductase (RNR) (4), 3′-phosphoadenosine 5′-phosphosulfate (PAPS) reductase (6), the methionine sulfoxide reductases MsrA, MsrB and fRMsr (7–9), the inner membrane protein DsbD (10, 11), and the peroxidases Tpx (12) and BCP (13). In addition to its role in enzyme recycling, Trx1 also controls the redox state of proteins with cysteine residues prone to oxidation. When exposed to ROS, sensitive cysteines are first modified to sulfenic acids (-SOH), which then often react with another thiol present in the vicinity to form a disulfide (14). Being able to reduce both sulfenic acids and disulfides (1, 15), Trx1 plays an active role in the protection of proteins from oxidative damages. Differential thiol trapping experiments led to the identification of a handful of proteins that depend on this protective activity of Trx1 (16).

Although a number of proteins that interact with Trx1 have been identified using copurification experiments (17), a comprehensive survey of proteins that depend on this oxidoreductase for reduction is still missing. The goal of the present study was to fully grasp the importance of Trx1 in controlling the redox state of intracellular proteins by characterizing the redox interactome of this major oxidoreductase. A powerful approach to identify Trx substrates consists in trapping the covalent intermediates that form between this protein and its substrates when the second catalytic cysteine of the WCGPC motif is mutated to an alanine. In this case, dissociation of the mixed-disulfide intermediate is prevented, stabilizing the complexes between thioredoxin and its substrates (1). This approach, which has never been applied to E. coli Trx1, already led to the identification of putative Trx substrates in a variety of organisms, including Chlorobaculum tepidum and Synechocystis sp (18–21), algae and plants (22–34), parasites (35–37), and mammals (38). However, to our knowledge, in all studies carried out so far, the Trx mutants were used to capture potential target proteins in cellular extracts, being sometimes immobilized on a resin (18–20, 22–37). Hence, substrate trapping occurred in vitro. As proteins that normally do not get oxidized in vivo may become oxidatively damaged following cell lysis, in vitro trapping is likely to lead to the identification of nonphysiological targets. Moreover, if certain physiological substrates unfold following overoxidation during extract preparation, they may not be recognized by the Trx trapping mutant and will be missed.

In this study, we decided to take advantage of E. coli being easily amenable to genetic manipulation and protein expression to comprehensively identify the substrates of Trx1 in vivo. To that end, we expressed a Trx1_WCGPA mutant in cells that were genetically engineered to optimize trapping by alteration of the cytoplasmic reducing pathways and that were exposed or not to an exogenous oxidative stress. This led us to the identification of a total of 268 putative substrates of Trx1, including 201 that had never been reported to depend on Trx for reduction and 78 that were found to interact with Trx1 only under severe oxidative stress conditions. Two of the new proteins involved in a disulfide complex with Trx1_WCGPA, the bacterial actin homolog MreB and the protein involved in sugar import PtsI, were studied in more detail. Our study significantly expands the number of Trx targets, further highlighting the role played by this major oxidoreductase in a variety of cellular processes. The dependence of a given protein on the Trx system for reduction is usually conserved. For instance, RNR, MsrA, and MsrB are Trx substrates in organisms ranging from bacteria and fungi to plants and mammals (39). Therefore, our work also provides insightful information on the interactome of Trx in organisms other than E. coli.

EXPERIMENTAL PROCEDURES

Bacterial Strains, Growth Conditions, and Plasmids

Bacterial strains and plasmids used in this study are listed in supplemental Table S1. Bacterial strains are E. coli MC4100 derivatives (40). All alleles, unless indicated, were moved by P1 transduction using standard procedures (41).

Strains IA195 and IA198, where ptsI was replaced by ptsI_C502S or ptsI_{C272AC324MC575S} on the chromosome, were constructed as follows. First, a cat-sacB cassette, encoding chloramphenicol acetyl transferase (cat) and SacB, a protein conferring sensitivity to sucrose, was amplified from strain CH1990 using primers ptsI-catsacB_Fw and ptsI-catsacB_Rv. The resulting PCR product shared a 40-bp homology to the 5′ UTR and 3′ UTR of ptsI at its 5′ and 3′ ends, respectively, and was used for lambda-red recombineering in strain IA186 as previously described (42). We verified that the cat-sacB cassette replaced the ptsI gene in the resulting strain (IA190) by sequencing across the junctions. The cat-sacB cassette was subsequently replaced by one of the mutated versions of ptsI (ptsI_C502S or ptsI_{C272AC324MC575S}) using the same technique. Briefly, ptsI_C502S and ptsI_{C272AC324MC575S} were amplified from plasmids pIA4 and pIA5 using primers catsacB-ptsI_Fw and catsacB-ptsI_{C502S_}Rv, or catsacB-ptsI_Fw and catsacB-ptsI_{C272AC324MC575S_}Rv, respectively, and the resulting PCR products were used for a second round of lambda-red recombineering in strain IA192 (IA190 transformed with pSIM5-Tet vector). Loss of the cassette in the resulting IA195 and IA198 strains was verified by positive (sucrose resistance) and negative (chloramphenicol sensitivity) selection and by PCR.

Unless indicated, bacteria were grown aerobically at 37 °C in LB medium. When necessary, growth media were supplemented with ampicillin (200 μg/ml), kanamycin (50 μg/ml) or chloramphenicol (10 μg/ml). For testing the ability to ferment glucose, MacConkey medium, supplemented with 0.5% glucose, was used.

Plasmid Constructions

The plasmids and primers used in the present study are listed in supplemental Tables S1 and S2. The pFB1 expression vector was constructed as follows. The region encoding trxA was amplified from the chromosome using primers Trx1_Fw and Trx1_Rv and inserted into the pHE43 vector restricted with NcoI and BglII (yciM_His was excised from the plasmid and replaced by trxA, allowing Trx1 to be His-tagged at the C terminus). The second cysteine of the WCGPC motif of Trx1 was mutated into an alanine by site-directed mutagenesis of pFB1 using primers Trx1_{WCGPA_}Fw and Trx1_{WCGPA_}Rv, generating plasmid pFB2. The pIA1 expression vector was constructed as follows. The region containing trxC was amplified from the chromosome using primers Trx2_Fw and Trx2_Rv and inserted into the pBAD-HisB vector restricted with NcoI and BglII. The second cysteine of the WCGPC motif of Trx2 was mutated into an alanine by site-directed mutagenesis of pIA1 using primers Trx2_{WCGPA_}Fw and Trx2_{WCGPA_}Rv, generating plasmid pIA2. The pIA3 expression vector was constructed as follows. ptsI was amplified from the chromosome using primers PtsI_Fw and PtsI_Rv and inserted into the pET28a vector. The cysteines of PtsI were mutated into serine, alanine, or methionine by site-directed mutagenesis of pIA3 using primers PtsI_{C502S_}Fw and PtsI_{C502S_}Rv, and primers PtsI_{C272A_}Fw, PtsI_{C272A_}Rv, PtsI_{C324M_}Fw, PtsI_{C324M_}Rv, PtsI_{C575S_}Fw, and PtsI_{C575S_}Rv, generating plasmids pIA4 and pIA5.

Trapping and Purification of Trx1_WCGPA-substrate Complexes

Strains FB32, IA170 and IA370 were grown in one liter LB medium at 37 °C to reach an A₆₀₀ of 0.5. Expression of Trx1_WCGPAwas induced by addition of 0.2% l-arabinose. After 2 h, the culture was treated, or not, with 2 mm hypochlorous acid (HOCl) for 10 min and then precipitated with 10% trichloroacetic acid (TCA) and placed at 4 °C overnight. Cells were then centrifuged at 11,325 × g for 45 min. 50 ml of cold 5% TCA were added on the pellet and another centrifugation was performed at 17,696 × g for 20 min. The proteins were then resuspended in 25 ml of 100 mm NaPi pH 8.0, 300 mm NaCl, 0.3% SDS, 8 m urea, and 10 mm iodoacetamide (IAM) to prevent any further disulfide bond rearrangement. The lysate was homogenized on a roller during 20 min and centrifuged at 23,708 × g for 45 min. The cleared lysate was finally diluted three times and loaded onto a 1 ml HisTrap FF column (GE Healthcare) equilibrated with 50 mm NaPi pH 8.0, 300 mm NaCl, and 0.3% SDS (buffer A). After washing with buffer A, proteins were eluted with a linear gradient from 0 to 300 mm imidazole in buffer A. Only one peak eluted from the column. This fraction was concentrated to 1.5 ml (using a Vivaspin Turbo 15 device (Sartorius, Goettingen, Germany)) and proteins were resolved on SDS-PAGE under nonreducing conditions (first dimension). The gel lane was cut, incubated in 20 ml of a buffer containing 10% SDS, 0.3 m Tris pH 7.5, 100 mm dithiothreitol (DTT), and 50% glycerol for 1 h, and placed on top of a second SDS-PAGE gel. After electrophoresis, proteins were visualized with PageBlue Protein Staining Solution (Thermo Scientific).

Mass Spectrometry Analysis of Two-dimensional Gels

Spots of interest were excised from the SDS-PAGE gel, digested with trypsin, and analyzed by liquid chromatographic tandem mass spectrometry (LC-MS/MS) using an LTQ XL as described (43). Briefly, peptides were separated by an acetonitrile gradient on a C18 column and the MS scan routine was set to analyze by MS/MS the five most intense ions of each full MS scan, dynamic exclusion was enabled to assure detection of coeluting peptides.

Protein identification was performed with SequestHT. In details, peak lists were generated using extract-msn (ThermoScientific) within Proteome Discoverer 1.4.1. From raw files, MS/MS spectra were exported with the following settings: peptide mass range 350–5000 Da, minimal total ion intensity 500. The resulting peak lists were searched using SequestHT against a target-decoy E. coli protein database obtained from Uniprot (July 30, 2015, 4339 entries). The following parameters were used: trypsin was selected with proteolytic cleavage only after arginine and lysine, number of internal cleavage sites was set to 1, mass tolerance for precursors and fragment ions was 1.0 Da, considered dynamic modifications were + 15.99 Da for oxidized methionine and +57.00 for carbamidomethyl cysteine. Peptide matches were filtered using the q-value and Posterior Error Probability calculated by the Percolator algorithm ensuring an estimated false positive rate below 5%. The filtered Sequest HT output files for each peptide were grouped according to the protein from which they were derived and their individual number of peptide spectral matches was taken as an indicator of protein abundance. The obtained data can be found in supplemental Tables S3, S4, S5, S6, S7, and S8. To build Tables I, supplemental Tables S9, and S10, we selected the cysteine-containing proteins for which at least two peptides were found in two independent experiments (data from supplemental Tables S3 and S4 were used to build Table I, data from supplemental Tables S5 and S6 were used to build supplemental Table S9, and data from supplemental Tables S7 and S8 were used to build supplemental Table S10). The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium (44) via the PRIDE partner repository with the data set identifier PXD003098 (username: reviewer30780@ebi.ac.uk, with password:0gnlCrwC).

Table I. Thioredoxin targets identified in a ΔtrxAΔtrxC E. coli strain. The proteins in this table were identified in two independent experiments with at least two peptides found in the indicated spot. They all contain cysteine residues. The full list of identified proteins for Experience 1 can be found in Table S3 and those for Experience 2 in Table S4. #PSM, number of peptide-spectrum matches; #pept, number of peptides; #cys, number of cysteines; MW, molecular weight. See supplemental Fig. S4A for the gel of Experience 2.

Gene	Uniprot ID	Description	Experience 1			Experience 2			#cys	MW (kDa)
			spot	#PSM	#pept	spot	#PSM	#pept
Energy metabolism
ATP-proton motive force interconversion
atpAb,c	P0ABB0	ATP synthase subunit alpha	15	14	11	14	3	3	4	55.2
atpDb,c	P0ABB4	ATP synthase subunit beta	14	44	22	12	17	13	1	50.3
atpGb	P0ABA6	ATP synthase gamma chain	8	8	8	7	2	2	2	31.6
nuoE	P0AFD1	NADH-quinone oxidoreductase subunit E	4	10	5	6	3	3	5	18.6
Glycolysis
fbaAb,c	P0AB71	Fructose-bisphosphate aldolase class 2	12	18	11	10	12	7	4	39.1
gapAb,c	P0A9B2	Glyceraldehyde-3-phosphate dehydrogenase A	11	13	7	9	33	10	3	35.5
lpdA	P0A9P0	Dihydrolipoyl dehydrogenase	15	24	13	14	15	9	5	50.7
pfkAb	P0A796	ATP-dependent 6-phosphofructokinase isozyme 1	11	4	3	9	4	4	6	34.8
pgkb	P0A799	Phosphoglycerate kinase	13	8	8	11	23	13	3	41.1
pykFb,c	P0AD61	Pyruvate kinase I	15	3	3	14	4	4	6	50.7
Gluconeogenesis
sdaA	P16095	l-serine dehydratase 1	15	2	2	14	4	4	9	48.9
Pentose phosphate pathway
gndb	P00350	6-phosphogluconate dehydrogenase, decarboxylating	13	10	9	11	16	10	2	51.4
talBb,c	P0A870	Transaldolase B	11	23	12	9	13	10	3	35.2
TCA cycle
gltAb	P0ABH7	Citrate synthase	14	9	9	12	4	4	7	48
icdb,c	P08200	Isocitrate dehydrogenase [NADP]	14	6	5	12	7	7	6	45.7
mdhb	P61889	Malate dehydrogenase	10	25	16	8	16	13	3	32.3
sucCb,c	P0A836	Succinyl-CoA ligase [ADP-forming] subunit beta	13	18	13	11	2	2	5	41.4
sucDb,c	P0AGE9	Succinyl-CoA ligase [ADP-forming] subunit alpha	9	13	10	7	4	4	5	29.8
Carbohydrate metabolism (other)
ackA	P0A6A3	Acetate kinase	13	8	7	11	4	4	6	43.3
aldA	P25553	Lactaldehyde dehydrogenase	15	4	4	14	3	3	3	52.2
deoB	P0A6K6	Phosphopentomutase	13	9	8	11	8	8	6	44.3
deoC	P0A6L0	Deoxyribose-phosphate aldolase	8	62	14	7	5	5	4	27.7
galF	P0AAB6	UTP–glucose-1-phosphate uridylyltransferase	10	2	2	8	2	2	3	32.8
garR	P0ABQ2	2-hydroxy-3-oxopropionate reductase	8	9	7	7	2	2	2	30.4
gatY	B1X7I7	Tagatose-1,6-bisphosphate aldolase GatY	9	12	4	7	22	6	4	30.8
gatZ	P0C8J8	Putative tagatose 6-phosphate kinase GatZ	14	14	10	14	14	6	8	47.1
glpK	P0A6F3	Glycerol kinase	15	9	9	14	13	11	5	56.2
nanA	P0A6L4	N-acetylneuraminate lyase	10	6	6	8	4	4	4	32.6
Processes
Cell division/cellular structure
mreBc	P0A9X4	Rod shape-determining protein MreB	12	5	5	10	2	2	3	36.9
Chaperones
groLb	P0A6F5	60 kDa chaperonin	16	6	6	14	2	2	3	57.3
Detoxification/oxidative stress response
ahpCb	P0AE08	Alkyl hydroperoxide reductase subunit C	4	136	16	6	175	13	2	20.7
bcpa,b	P0AE52	Putative peroxiredoxin bcp	1	3	3	3	9	6	3	17.6
katGb,c	P13029	Catalase-peroxidase	16	10	9	14	4	4	1	80
msrBa,b	P0A746	Peptide methionine sulfoxide reductase MsrB	1	6	3	3	5	5	6	15.4
msrCa,b	P76270	Free methionine-R-sulfoxide reductase	3	9	5	4	7	4	3	18.1
sodB	P0AGD3	Superoxide dismutase [Fe]	4	20	7	6	3	3	1	21.3
tpxa,b,c	P0A862	Thiol peroxidase	3	41	10	4	139	8	3	17.8
trxBa,b	P0A9P4	Thioredoxin reductase	10	4	4	9	6	5	4	34.6
ucpA	P37440	Oxidoreductase UcpA	9	6	5	8	2	2	4	27.8
Regulatory functions
arcA	P0A9Q1	Aerobic respiration control protein ArcA	8	12	9	7	2	2	2	27.3
furc	P0A9A9	Ferric uptake regulation protein	2	2	2	3	4	3	4	16.8
Transport/binding proteins
araG	P0AAF3	Arabinose import ATP-binding protein AraG	15	5	4	14	2	2	3	55
ppab	P0A7A9	Inorganic pyrophosphatase	4	8	5	6	2	2	2	19.7
Small molecules biosynthesis and degradation
Amino acid biosynthesis
aspA	P0AC38	Aspartate ammonia-lyase	15	5	5	14	13	11	11	52.3
cysKb	P0ABK5	Cysteine synthase A	11	6	5	9	6	6	1	34.5
glnAb	P0A9C5	Glutamine synthetase	15	3	2	14	12	8	4	51.9
glyAb	P0A825	Serine hydroxymethyltransferase	13	5	5	11	2	2	3	45.3
iscS	P0A6B7	Cysteine desulfurase IscS	13	8	8	11	5	5	3	45.1
metKb	P0A817	S-adenosylmethionine synthase	13	7	6	11	2	2	4	41.9
Amino acid degradation
tnaAb,c	P0A853	Tryptophanase	14	53	22	12	67	18	7	52.7
Fatty acid biosynthesis
accC	P24182	Biotin carboxylase	14	4	4	12	2	2	8	49.3
fabAc	P0A6Q3	3-hydroxydecanoyl-[acyl-carrier-protein] dehydratase	2	11	5	4	2	2	2	19
fabB	P0A953	3-oxoacyl-[acyl-carrier-protein] synthase 1	13	10	7	12	3	3	6	42.6
fabDb	P0AAI9	Malonyl CoA-acyl carrier protein transacylase	10	4	3	9	5	3	5	32.4
fabH	P0A6R0	3-oxoacyl-[acyl-carrier-protein] synthase 3	10	3	3	8	2	2	5	33.5
fabZc	P0A6Q6	3-hydroxyacyl-[acyl-carrier-protein] dehydratase FabZ	1	4	4	3	3	3	2	17
Deoxyribonucleotide/nucleoside metabolism
cddb	P0ABF6	Cytidine deaminase	9	3	3	8	2	2	5	31.5
Purine/pyrimidine ribonucleotide biosynthesis
preT	P76440	NAD-dependent dihydropyrimidine dehydrogenase subunit PreT	13	4	4	11	4	4	13	44.3
prs	P0A717	Ribose-phosphate pyrophosphokinase	10	2	2	9	2	2	4	34.2
purAb	P0A7D4	Adenylosuccinate synthetase	13	3	2	12	2	2	4	47.3
Salvage of nucleotides and nucleosides
deoA	P07650	Thymidine phosphorylase	14	3	3	12	3	3	2	47.2
Molybdenum cofactor biosynthesis
moaB	P0AEZ9	Molybdenum cofactor biosynthesis protein B	2	9	5	4	2	2	2	18.7
Iron-sulfur cluster assembly
iscUb	P0ACD4	Iron-sulfur cluster assembly scaffold protein IscU	1	4	3	3	3	3	3	13.8
Macromolecules
DNA transcription
hns	P0ACF8	DNA-binding protein H-NS	1	11	6	3	2	2	1	15.5
rho	P0AG30	Transcription termination factor Rho	14	11	11	12	3	3	1	47
Protein translation and modification
asnS	P0A8M0	Asparagine–tRNA ligase	14	2	2	14	2	2	4	52.5
gltX	P04805	Glutamate–tRNA ligase	15	12	9	14	14	9	7	53.8
hisS	P60906	Histidine—tRNA ligase	14	2	2	12	2	2	5	47
infC	P0A707	Translation initiation factor IF-3	4	6	4	6	4	3	1	20.6
prfB	P07012	Peptide chain release factor 2	14	2	2	14	2	2	2	41.2
rplB	P60422	50S ribosomal protein L2	8	5	5	8	3	3	2	29.8
rplE	P62399	50S ribosomal protein L5	3	64	15	4	13	5	1	20.3
rplFb	P0AG55	50S ribosomal protein L6	3	30	10	6	3	2	1	18.9
rplJ	P0A7J3	50S ribosomal protein L10	1	23	7	3	3	3	1	17.7
rplK	P0A7J7	50S ribosomal protein L11	1	19	5	3	7	4	1	14.9
rplN	P0ADY3	50S ribosomal protein L14	1	22	5	3	3	3	2	13.5
rplQ	P0AG44	50S ribosomal protein L17	1	19	5	3	11	4	1	14.4
rpsAb	P0AG67	30S ribosomal protein S1	16	60	28	11	2	2	2	61.1
rpsBb	P0A7V0	30S ribosomal protein S2	8	42	11	7	3	2	1	26.7
rpsKb	P0A7R9	30S ribosomal protein S11	1	17	7	3	3	3	2	13.8
rpsLb	P0A7S3	30S ribosomal protein S12	1	15	4	3	3	2	4	13.7
rpsM	P0A7S9	30S ribosomal protein S13	1	29	6	2	9	5	1	13.1
serS	P0A8L1	Serine–tRNA ligase	15	8	8	14	6	5	5	48.4
tsfc	P0A6P1	Elongation factor Ts	10	35	13	8	9	8	2	30.4
tufBb	P0CE48	Elongation factor Tu	13	99	20	11	87	17	3	43.3
t-RNA and r-RNA processing
miaB	P0AEI1	tRNA-2-methylthio-N(6)-dimethylallyladenosine synthase	15	3	3	14	4	4	6	53.6
mnmA	P25745	tRNA-specific 2-thiouridylase MnmA	13	11	8	11	6	5	7	40.9
ATP binding
ychF	P0ABU2	Ribosome-binding ATPase YchF	13	26	15	11	37	14	6	39.6
Protein degradation/proteases
clpXb	P0A6H1	ATP-dependent Clp protease ATP-binding subunit ClpX	14	3	3	12	6	6	7	46.3
pepD	P15288	Cytosol non-specific dipeptidase	15	8	7	14	4	4	4	52.9
Unknown proteins
yfeX	P76536	Probable deferrochelatase/peroxidase YfeX	10	6	6	8	3	3	4	33

^a Genes encoding known substrates of Trx1 in E. coli (39).

^b Genes encoding known substrates of Trx1 in other organisms a (18–31, 33–37, 39, 105–112).

^c Genes encoding proteins that copurify with Trx1 in E. coli (17).

Impact of Thioredoxin Deletion on MreB Distribution in Living Cells Upon A22 Treatment

Strains IA90, IA94, IA396, and IA398 were grown in LB medium at 30 °C to reach an A₆₀₀ of 0.5. S-(3,4-dichlorobenzyl) isothiourea (A22) was then added at a concentration of 5 μg/ml and growth was continued for 2 h. For complementation studies, expression of Trx1 in trans in IA396 was induced by addition of 0.2% l-arabinose when cells reached an A₆₀₀ of 0.1.

Microscopy Image Acquisition

Cells were spotted on PBS + 1% agarose pads between a glass slide and a coverslip. Imaging was performed using an Axio Observer.Z1 inverted epifluorescence microscope (Carl Zeiss, Oberkochen Germany) equipped with an AxioCam 506 mono camera (Carl Zeiss), 1x/C-mount camera adaptor (Carl Zeiss), phase contrast objective Plan Apochromat 100×/1.40 Oil Ph3 (Carl Zeiss), and filter set 31 (Carl Zeiss) to image RFP-associated fluorescence. Images were acquired using the ZEN2012 (blue edition) software (Carl Zeiss) and processed with MetaMorph (Molecular Devices, Sunnyvale, CA). Exposure times were identical for compared conditions.

Determination of MreB In Vivo Redox State by Reverse Trapping

Strains JAS27 and FB30 were grown in LB medium at 37 °C to reach an A₆₀₀ of 0.5. H₂O₂ was then added at a concentration of 1 mm and growth was continued for 30 min. At several time-points, 1.8 ml of culture were precipitated with 10% TCA. Samples were then incubated on ice for 20 min and centrifuged at 16,100 × g for 5 min. The resulting pellets were washed with 400 μl ice-cold acetone and resuspended in 100 μl denaturing buffer (6 m urea, 200 mm Tris-HCl pH 8.5, 10 mm EDTA) supplemented with 100 mm N-ethylmaleimide (NEM). After a 20 min incubation at 25 °C and with 600 rpm shaking, the reaction was stopped by adding 10 μl of 10% TCA and left on ice for 20 min. The NEM-modified proteins were centrifuged and the pellet washed as described above. The proteins were then dissolved in 100 μl of 10 mm DTT in denaturing buffer. After a 1 h incubation at 25 °C and with 1400 rpm shaking, the reaction was stopped by adding 10 μl of 10% TCA and left on ice for 20 min. The reduced proteins were centrifuged and the pellet washed as above. The proteins were then dissolved in 30 μl of 1× sample buffer (2% SDS, 12% glycerol, 60 mm Tris-HCl pH 7.4, 0.01% bromphenol blue) containing 2 mm of methoxyl polyethylene glycol maleimide (MalPEG) 2 kDa. After a 1 h incubation at 37 °C and with 1400 rpm shaking in the dark, samples were diluted three times in 1× sample buffer and incubated for 1 h at room temperature. Samples were then loaded onto 10% SDS-polyacrylamide gels under denaturing conditions and MreB was detected by Western blotting using an anti-MreB antibody.

Identification of the MreB Cysteine Involved in the MreB- Trx1_WCGPA Interaction by LC-MS/MS

Recombinant MreB (5 μg), expressed and purified as in (45), was incubated with Trx1_WCGPA (1.6 μg) for 1 h on ice, in 200 μl of a buffer containing 50 mm Tris-HCl pH 8, 0.1 mm EDTA, 100 mm KCl, and 20% glycerol. Proteins were then precipitated with 10% TCA after 10 μg of bovine serum albumin were added as a carrier. The pellet was resuspended in 50 μl of 100 mm NH₄HCO₃ pH 8.0 and the proteins digested overnight at 30 °C with 0.5 μg of sequencing grade trypsin (Promega, Madison, WI). The peptides were analyzed by capillary LC-MS/MS in a LTQ XL ion trap mass spectrometer (Thermo Scientific, San Jose, CA), fitted with a microelectrospray probe. The mass spectrometer was operated in the data-dependent mode and switched automatically between MS, Zoom Scan for charge state determination and MS/MS for the five most abundant ions. Peptides were identified by Sequest and the mixed disulfide between MreB and Trx1_WCGPA was identified using DBond (46). The modified peptides were manually validated.

PtsI_AMCS Expression and Purification

BL21 (DE3) competent cells were transformed with plasmid pIA5, yielding strain IA160. During overnight cultures in LB medium containing 200 μg/ml of ampicillin at 37 °C without shaking, cells reached an A_{600 nm} of ∼0.5. Cells were then cultured aerobically with shaking (200 rpm) at 37 °C. After 30 min, protein expression was induced by adding 1 mm isopropyl β-d-1-thiogalactopyranoside (IPTG). Cells were harvested by centrifugation (2800 × g, 4 °C, 20 min) after 3 h of induction. Cells were resuspended in 10 ml buffer A containing 50 mm NaPi pH 8.0 and 300 mm NaCl and disrupted by two passages through a French press (1500 psi). The cell lysate was then centrifuged during 40 min (23,600 × g, 4 °C). Supernatant was recovered and diluted about 3 times in buffer A and filtered with a Minisart High-Flow filter (Sartorius, pores of 0.2 μm diameter). The diluted supernatant was loaded on a HisTrap FF (1 ml, GE Healthcare) at 1 ml/min. After washing the column, proteins were eluted using an imidazole gradient (0 to 180 mm in buffer A). The fractions containing PtsI_AMCS were concentrated using a Vivaspin Turbo 15 device (Sartorius) and desalted on a PD-10 column (GE healthcare) equilibrated with buffer B (50 mm NaPi pH 8.0 and 150 mm NaCl). PtsI_AMCS was then reloaded on an affinity chromatography column to reach a higher degree of purity (using the same protocol as described above).

Identification of the Dimedone-modified Peptide of PtsI by LC-MS/MS

Recombinant PtsI_AMCS (10 μg) was incubated for 5 min at room temperature in the presence of 5 mm dimedone (DMD). The sample was then incubated with 2 mm H₂O₂ for 15 min, and then treated with 50 mm IAM for 10 min at room temperature to block reduced thiols. The protein was then precipitated with 10% TCA and the pellet resuspended in 50 μl of 100 mm NH₄HCO₃ pH 8.0, for overnight digestion at 30 °C with 0.5 μg of sequencing grade trypsin (Promega, Madison, WI). The peptides were analyzed by capillary LC-MS/MS in a LTQ XL ion trap mass spectrometer (Thermo Scientific), fitted with a microelectrospray probe. The mass spectrometer was operated in the data-dependent mode and switched automatically between MS, Zoom Scan for charge state determination and MS/MS for the five most abundant ions. Peptides were identified by Proteome Discoverer software (Thermo Scientific, version 1.4.0.288) considering dynamic modifications on cysteine residues of 138.0 Da for sulfenic dimedone. Modified peptides were manually validated.

Experimental Design and Statistical Rationale

In order to reliably identify proteins forming mixed-disulfides with Trx, all trapping experiments (in the ΔtrxAΔtrxC mutant, with or without HOCl, and in the ΔtrxAΔgshA mutant) were performed in biological duplicates. To prevent oxidation of cysteine residues as well as the disulfide rearrangements that could potentially occur following cells lysis, cellular samples were prepared by directly adding 10% TCA to the cells growing in culture media. This acidic treatment not only denatures cellular proteins but also ensures that the reduced cysteine residues become protonated and therefore poorly reactive. In addition, IAM was added to the buffer used to resuspend the precipitated proteins prior to the diagonal gel analysis. IAM covalently modifies reduced cysteine residues, therefore blocking the thiol-disulfide exchange reactions that could occur when the sample is brought to the more alkaline pH of the resuspension buffer (pH 8).

The complexes involving Trx1_WCGPA bound to its substrates were purified by affinity chromatography using Ni-NTA agarose. Untagged proteins nonspecifically binding to the column were also eluted when the imidazole gradient was applied. However, as these contaminants are not involved in a mixed disulfide complex, they are separated according to molecular weight in both the nonreducing and reducing dimensions of the diagonal gel. They migrate therefore on a diagonal in the second dimension, in contrast to the Trx substrates, allowing the discrimination between the Trx interacting partners and the nonspecific contaminants.

To build Table I, supplemental Tables S9, and S10, the data were filtered to include only proteins containing cysteine residues and for which at least two peptides were found in the biological duplicates.

The procedures carried out to probe the in vivo redox state of MreB in the wild type and in the ΔtrxAΔtrxC mutant were done in biological triplicates as well as the microscopy experiments performed to investigate the physiological importance of the Trx system in MreB function. All the tests aiming at determining the importance of the cysteine residues of PtsI were also done in biological triplicates.

RESULTS AND DISCUSSION

Identification of Trx1 Targets In Vivo

To trap E. coli Trx1 bound to its substrates, we used a mutant version of the protein lacking the second cysteine of the catalytic site and fused to a C-terminal His-tag (Trx1_WCGPA). This mutant was expressed in a strain in which the Trx system had been disrupted by deletion of both trxA (encoding Trx1) and trxC (encoding Trx2, an atypical, less abundant, thioredoxin (47–49); see below) in order to optimize trapping. As shown in Fig. 1C, induction of Trx1_WCGPA expression led to the formation of several high-molecular weight complexes recognized by the anti-His antibody. These complexes were DTT-sensitive, indicating that they corresponded to Trx1_WCGPA bound to unknown proteins via a disulfide. Trx1_WCGPA and its covalently-bound partners were then purified by affinity chromatography. Only one peak eluted from the column when an imidazole gradient was applied (not shown). The purified sample was then analyzed by two-dimensional gel electrophoresis, in which the first dimension is nonreducing whereas the second is reducing (50–52). In this experiment, proteins that were not linked to Trx1_WCGPA by a disulfide bond were separated according to molecular weight in both dimensions and were therefore found on a diagonal in the second dimension. In contrast, proteins that were in complex with Trx1_WCGPA were separated from it under the reducing conditions of the second dimension and were found off the diagonal (Fig. 1D). These proteins were subjected to MS/MS analysis and identified. Ninety-one cysteine-containing proteins for which at least two peptides were reproducibly found off the diagonal in two independent experiments were considered as putative Trx1 substrates (Table I, constructed from supplemental Tables S3 and S4). These proteins included several well-known Trx1 targets such as the enzymes Tpx, BCP, MsrB, and fRMsr, nicely validating our data. As expected, given the conservation of the Trx interactome across species, the list of Trx1 substrates also included 40 proteins that had been shown to interact with Trx in organisms other than E. coli (Table I). Remarkably, 51 proteins that were released from Trx1_WCGPA in the second dimension had never been reported to depend on the Trx system for reduction in any organism. The new thioredoxin targets are involved in a variety of cellular and metabolic processes, including ATP production, glycolysis, carbohydrate metabolism, protein translation, iron-sulfur cluster assembly, and protein homeostasis (see Table I). Unlike enzymes such as Tpx and MsrB that form a disulfide bond during catalysis and depend on Trx for recycling, most of the newly identified Trx substrates are well-characterized proteins that have not been reported to form catalytic disulfides. However, the fact that they were trapped in a mixed-disulfide complex with Trx1_WCGPA implies that these proteins become Trx1 targets following the oxidation of at least one sensitive cysteine residue, most likely to a sulfenic acid possibly rearranging into a disulfide. In agreement with this, several of the identified Trx substrates have been reported to contain redox-sensitive cysteine residues, such as pyruvate kinase I (pykF) (53, 54), glyceraldehyde 3-phosphate dehydrogenase (gapA) (55–58), isocitrate dehydrogenase (icd) (59–61), malate dehydrogenase (mdh) (62–65), and tryptophanase (tnaA) (66–68). In addition, other newly identified Trx substrates, such as aspartate ammonia-lyase (aspA) (69, 70), biotin carboxylase (accC) (71–73), ribose-phosphate pyrophosphokinase (prs) (74, 75), ferric uptake regulation protein (fur) (76), and 3-oxoacyl-[acyl-carrier-protein] synthase 1 (fabB) (77), contain cysteine residues that are important for activity, suggesting that Trx1 is involved in maintaining them active.

The Actin Homolog MreB is a Trx1 Substrate

We were intrigued by the identification of the actin structural homolog MreB as a putative Trx1 substrate. MreB is a major rod-shape determinant in E. coli and many other nonspherical bacteria (78). It forms discrete patches underneath the cell membrane that move around the cell circumference, in a manner that requires cell wall synthesis (79–81). In the presence of ATP or GTP, MreB polymerizes and forms small antiparallel double filaments (78). As shown in supplemental Fig. S1A, E. coli MreB has three cysteine residues, two of which (Cys113 and Cys324) are conserved in homologous proteins ranging from the α, β, γ-proteobacteria, to Bacilli and Clostridia. Although the conserved cysteines do not seem to be required for MreB function (78, 82–84), they are present in the nucleotide-binding pocket (supplemental Fig. S1B) where their oxidation would likely impair nucleotide binding and inhibit MreB (85). The important role played by MreB in rod-shaped bacteria prompted us to further investigate its redox sensitivity and interaction with Trx1. First, we confirmed that Trx1 (12 kDa) and MreB (36 kDa) form a DTT-sensitive 48 kDa mixed-disulfide complex in vivo, using a specific anti-MreB antibody (Fig. 2A). We then carried out reverse thiol trapping experiments (16) in order to determine if MreB can become oxidized in vivo, as implied by the identification of the Trx1-MreB complex. In this approach, all reduced cysteine residues are first covalently modified with N-ethylmaleimide (NEM). Oxidized cysteines are then reduced with DTT and subsequently alkylated with MalPEG. MalPEG is a 2 kDa molecule that modifies free thiols leading to a major shift on SDS-PAGE. As seen in Fig. 2B, we found that a slow-migrating band of ∼45 kDa appears when samples from both the wild type and the ΔtrxAΔtrxC mutant are treated with MalPEG, indicating that MreB can be oxidized in vivo. In addition, exposure of cells to H₂O₂, an oxidizing agent, led to the appearance of two additional higher-migrating bands in the ΔtrxAΔtrxC mutant but not in the wild type, which indicates that MreB is further oxidized when the Trx system is impaired. It is difficult to accurately predict the number of modified cysteine residue(s) from the migration of MalPEG-modified proteins in SDS-PAGE and attempts to identify the MreB cysteines getting oxidized in vivo failed. However, MS analysis of purified, air-oxidized MreB led to the identification of a disulfide bond between Cys113 and Cys324, the two cysteine residues found in the nucleotide binding pocket (not shown). Furthermore, LC-MS/MS analysis of the Trx1-MreB complex reconstituted in vitro using purified proteins identified Cys113 of MreB as the cysteine involved in the mixed-disulfide with Trx1_WCGPA (Fig. 2C). Altogether, these results reveal that the cytoskeletal protein MreB contains cysteine residues that are susceptible to oxidation. They also involve Trx1 in the recycling of MreB and suggest that the activity of MreB is redox-regulated.

Fig. 2.

The actin homolog MreB is a Trx1 substrate. A, After expression of Trx1_WCGPA in a ΔtrxAΔtrxC mutant and purification and concentration of the complexes, the elution fraction was analyzed by Western blotting using an anti-MreB antibody. The bands corresponding to MreB alone or in complex with Trx1_WCGPA are indicated. Upon DTT addition, the band corresponding to the complex disappeared whereas the MreB band increased, indicating that Trx1_WCGPA is released from MreB under reducing conditions. The molecular mass markers (in kDa) are indicated on the left. The fact that MreB was observed in the nonreduced sample indicates that a portion of this protein co-purified with Trx1_WCGPA on the Ni-NTA agarose column. B, To determine if MreB can become oxidized in vivo, a reverse thiol trapping experiment was performed in a wild-type (WT) strain and a ΔtrxAΔtrxC mutant strain. All reduced cysteine residues were first covalently modified with NEM. Oxidized cysteines are then reduced with DTT and subsequently alkylated with MalPEG. MalPEG modifies free thiols leading to a major shift on SDS-PAGE. Samples were analyzed by Western blotting using an anti-MreB antibody. The band corresponding to reduced MreB (MreB_red) is indicated. Upon MalPEG addition, a slow-migrating band (*) appears when samples from both the wild type and the ΔtrxAΔtrxC mutant are treated, indicating that MreB can be oxidized in vivo. Exposure (exp.) of cells to 1 mm H₂O₂ led to the appearance of two additional higher migrating bands (**) in the ΔtrxAΔtrxC mutant but not in the wild type, which indicates that MreB is further oxidized when the Trx system is impaired. The molecular mass markers (in kDa) are indicated on the left. Both parts of the figure come from a single membrane. C, Trx1_WCGPA forms a mixed disulfide complex with MreB. Mass spectrometry analysis of a triply charged parent ion of [M+3H]³⁺ = 1139.3 Da shows fragmentation characteristics of a disulfide linkage between Cys33 of Trx and Cys113 of MreB, as determined by the DBond software (46). P*, one strand of a dipeptide; p*, the other strand of a dipeptide; capital letters, fragment ions from peptide P*; lowercase letters, fragment ions from peptide p*. D, Wild-type (WT) and ΔtrxAΔtrxC (ΔAΔC) cells expressing an MreB-RFP-MreB sandwich fluorescent fusion were observed using phase contrast (left) and fluorescence microscopy (right). MreB localized similarly in the ΔtrxAΔtrxC mutant and in the wild type when cells were grown under normal conditions. Upon A22 treatment, an abnormal MreB localization pattern was observed in the ΔtrxAΔtrxC double mutant but not in the wild type. Expression of Trx1 (but not of the empty plasmid) in trans restored an MreB normal localization pattern. Bar, 5 μm.

To further investigate this unsuspected link between redox homeostasis and cell shape, we transduced a functional mreB-RFP-mreB sandwich fluorescent fusion (86) at the native mreB chromosomal locus in wild-type and ΔtrxAΔtrxC cells. As shown in Fig. 2D, we found no difference of MreB localization between the ΔtrxAΔtrxC mutant and the wild type when cells were grown under normal conditions. However, upon sub-lethal addition of S-(3,4-dichlorobenzyl) isothiourea (A22), a widely used MreB inhibitor targeting the active site (83), an abnormal MreB localization pattern was observed in the ΔtrxAΔtrxC double mutant but not in the wild type (Fig. 2D). Expression of Trx1 in trans restored MreB localization in the ΔtrxAΔtrxC double mutant (Fig. 2D). Thus, these data support the idea that Trx1 protects MreB from oxidative damages, possibly the formation of a disulfide bond in the nucleotide-binding pocket.

Probing the Impact of Exogenous Oxidative Stress on the Trx Interactome

The fact that the proteins listed in Table I were found in complex with Trx1 implies that they contain cysteine residues prone to oxidation, probably by the ROS endogenously produced by E. coli (87). This prompted us to test if exposure of cells to an exogenous more severe oxidative stress would lead to the identification of additional Trx substrates. To that end, we expressed the Trx1_WCGPA variant in ΔtrxAΔtrxC cells grown in LB. When the culture reached an A₆₀₀ of 0.5, HOCl was added. HOCl is a fast acting oxidant released by neutrophils to kill invading bacteria and reacting most rapidly with cysteine and methionine residues (88, 89), causing oxidative protein unfolding and the formation of incorrect disulfide bonds. The complexes involving Trx1_WCGPA were purified by affinity chromatography and subjected to diagonal gel electrophoresis (Fig. 3A), as described above. Following MS/MS analysis, we identified 213 proteins that were found in a mixed-disulfide complex with Trx1_WCGPA (they were off the diagonal) in two independent experiments (supplemental Table S9, constructed from supplemental Tables S5 and S6). Eighty-two of the 213 identified proteins had already been found covalently-bound to Trx1_WCGPA in the experiment performed without HOCl (Fig. 3B), which validates the data shown in Table I. The 131 remaining proteins were only found in complex with Trx1_WCGPA when an oxidative stress was applied. Interestingly, the new Trx substrates include several proteins involved in the defense mechanisms against oxidative stress, such as the peroxiredoxin OsmC (90, 91), the transcription factor OxyR (92–94), and the methionine sulfoxide reductase MsrA (95, 96), a well known Trx substrate. Of note, 112 of the newly identified Trx1 partners had never been reported to interact with thioredoxin in any living organism. These proteins are involved in a variety of cellular functions including energy metabolism, amino acid synthesis, cell division, lipopolysaccharide synthesis, protein homeostasis, and transcription (see supplemental Table S9). These data highlight the important role played by Trx1 in rescuing a large number of proteins damaged by oxidative stress.

Fig. 3.

Trapping of Trx1 substrates in a ΔtrxAΔtrxC E. coli strain in the presence of HOCl and in a in a ΔtrxAΔgshA E. coli strain in vivo. A, To identify Trx1 substrates upon oxidative stress, ΔtrxAΔtrxC cells expressing Trx1_WCGPA were exposed to HOCl for 10 min. The mixed-disulfide complexes were then purified by affinity chromatography using Ni-NTA agarose. After concentration, the proteins trapped by Trx1_WCGPA were separated by SDS-PAGE (nonreducing dimension). The complexes were then separated in a second, reducing dimension. Proteins were identified by mass spectrometry. The spots analyzed by mass spectrometry were numbered as indicated (see supplemental Table S9). The molecular mass markers (in kDa) are indicated on the left. The inset shows the reducing pathway that was genetically modified. Here, by deleting the trxA and trxC genes, it is the Trx pathway that was impaired. Trx, thioredoxin; TR, thioredoxin reductase; Grx, glutaredoxin; gshA encodes the gamma-glutamylcysteine synthetase; gshB encodes the glutathione synthetase; GR, glutaredoxin reductase. B, Venn diagram representing the numbers of proteins identified in the ΔtrxAΔtrxC strain with or without HOCl stress (from Tables I and supplemental Table S9). C, To identify the Trx1 substrates that may have been missed because of the activity of the Grx pathway, Trx1_WCGPA was expressed in a ΔtrxAΔgshA mutant. The mixed-disulfide complexes were then purified, concentrated and separated as described in A. Proteins were identified by mass spectrometry. The spots analyzed by mass spectrometry were numbered as indicated (see supplemental Table S10). The molecular mass markers (in kDa) are indicated on the left. The inset shows the reducing pathways that were genetically modified. Here, by deleting the trxA and gshA genes, both the Trx and the Grx pathways were impaired. Legend is the same as in A. D, Venn diagram representing the numbers of proteins identified in the ΔtrxAΔtrxC (with HOCl stress or not) and the ΔtrxAΔgshA strains (from Tables I, S9 and S10).

Engineering E. coli to Optimize Trapping

The fact that PAPS reductase and the catalytic subunit of RNR (subunit alpha), two major Trx substrates (4, 6), were not found in complex with Trx1_WCGPA, even when an exogenous oxidative stress was applied, indicated that certain Trx targets had managed to escape the hunt. We reasoned that this could be because of the activity of the glutathione/glutaredoxin (Grx) pathway, a reducing system functioning in parallel to the Trx pathway in the E. coli cytosol. In the Grx pathway, electrons flow from NADPH to glutathione via glutathione oxidoreductase and then from glutathione to three dithiol glutaredoxins (6, 97, 98), some of which share a partially redundant substrate specificity with Trx1. Therefore, in order to identify Trx1 substrates that may have been missed because of their reduction by the Grx pathway, we decided to proceed with the trapping experiment in cells where both reducing pathways are impaired. Although E. coli, like many other bacteria, needs either the Trx or Grx pathway for survival (99, 100), strains with partially functioning pathways can be engineered (100). For instance, cells lacking trxA and gshA, encoding an enzyme required for glutathione biosynthesis, are blocked for the Grx pathway and do not express Trx1 but remain viable because of the presence of Trx2. We decided to use these cells to search for additional Trx1 substrates. Indeed, as Trx2 has a limited substrate specificity, which we showed using a Trx2_WCGPA mutant (supplemental Fig. S2), ΔtrxAΔgshA cells are likely to accumulate most Trx1 substrates in the oxidized state, even those that can otherwise be reduced by the Grx pathway, making these cells ideally suited for this search.

The Trx1_WCGPA variant was expressed in ΔtrxAΔgshA cells, purified by affinity chromatography and subjected to diagonal gel electrophoresis (Fig. 3C) as described before. MS/MS analysis led to the identification of 170 proteins that were found in a mixed-disulfide complex with Trx1_WCGPA in two independent experiments (supplemental Table S10, constructed from supplemental Tables S7 and S8). Almost half of the identified proteins (71 out of 170) had already been found covalently-bound to Trx1_WCGPA when this protein was expressed in the ΔtrxAΔtrxC strain (Fig. 3D), further validating the data shown in Table I. Remarkably, expressing Trx1_WCGPA in ΔtrxAΔgshA cells led to the identification of 99 additional proteins that were released from Trx1_WCGPA in the second dimension, 82 of which had never been reported to interact with thioredoxin in any living organism. The new putative Trx1 substrates are involved in a variety of cellular functions ranging from energy metabolism, amino acid synthesis, protein homeostasis to transcription (see supplemental Table S10). They also included the enzymes RNR (catalytic subunit alpha) and PAPS reductase, in agreement with our starting hypothesis, and strongly suggesting that some of the new Trx1 substrates identified in ΔtrxAΔgshA cells possess oxidation-prone cysteine residue(s) that can be reduced by both cytoplasmic reducing pathways. It is also interesting to note that 53 out of the 99 substrates identified in the ΔtrxAΔgshA mutant were also found when ΔtrxAΔtrxC cells were exposed to HOCl (Fig. 3D). This probably results from the fact that impairing both the Trx and Grx systems causes a more severe endogenous oxidative stress in the ΔtrxAΔgshA mutant.

Identification of PtsI as a Novel Redox-regulated Enzyme

To further investigate the interaction between Trx1 and the newly identified proteins, we decided to focus on PtsI. This enzyme was selected because it plays an essential role in the coupled translocation/phosphorylation of several carbohydrates including glucose in bacteria (101, 102) and its activity can be easily monitored in vivo. First, we confirmed that Trx1 (12 kDa) and PtsI (64 kDa) form a mixed-disulfide complex by detecting a DTT-sensitive 76 kDa band in the fraction containing purified Trx1_WCGPA bound to its substrates, using a specific anti-PtsI antibody (Fig. 4A). PtsI has four cysteine residues, of which only one (Cys502) is strictly conserved (supplemental Fig. S3). Before dissecting the Trx1-PtsI interaction, we decided to determine the importance of the various cysteine residues for PtsI activity. As shown in Fig. 4B, mutation of Cys502 to serine prevented E. coli to ferment glucose, confirming the importance of this active site residue for activity (103, 104). In contrast, replacement of the three other cysteines, either alone (not shown) or together, had no impact, indicating that they are dispensable for the function of PtsI. Note that all mutant proteins were expressed at physiological levels from the ptsI chromosomal locus. Given the importance of Cys502 for PtsI activity, we postulated that Trx1 might be involved in maintaining this residue reduced in order to keep PtsI active. To test if Trx1 specifically interacts with Cys502, we expressed Trx1_WCGPA in a ΔtrxAΔgshA strain producing PtsI_AMCS from the ptsI locus (all three nonessential cysteine residues are mutated in PtsI_AMCS). As shown in Fig. 4C, we observed the formation of a DTT-sensitive complex between Trx1_WCGPA and PtsI_AMCS that was recognized by the anti-PtsI antibody. This result implies that Cys502 becomes a Trx1 target following its oxidation, most likely to a sulfenic acid. Because of their high reactivity, sulfenic acids are often difficult to identify. To test if Cys502 of PtsI can be oxidized to a sulfenic acid, PtsI_AMCS was incubated with H₂O₂ and dimedone, a sulfenic acid specific reagent. The protein was then digested with trypsin and the peptide mixture analyzed by LC-MS/MS. As shown in Fig. 4D, the dimedone-modified peptide, corresponding to the sulfenic acid form of PtsI_AMCS, could be detected, indicating that Cys502 of PtsI can be oxidized to the sulfenic state. PtsI having three additional cysteines, it is possible that the sulfenic acid rearranges in vivo into an intramolecular disulfide involving one of these residues. Regardless of whether this rearrangement occurs or not, our data indicate that Trx1 recognizes oxidized PtsI, suggesting that it plays a role in protecting PtsI from oxidative inactivation. It is interesting to note that, in the course of PtsI purification, we observed a noncovalent complex between Trx1 and PtsI that remained remarkably stable following successive chromatography steps (not shown), further supporting the functional link between these two proteins.

Fig. 4.

Identification of PtsI as a novel redox-regulated enzyme. A, Trx1_WCGPA was expressed in a ΔtrxAΔgshA mutant, purified and concentrated. The elution fraction was then analyzed by Western blotting using an anti-PtsI antibody. The bands corresponding to PtsI alone or in complex with Trx1_WCGPA are indicated. Upon DTT addition, the band corresponding to the complex between PtsI and Trx1 disappeared whereas the PtsI band increased, indicating that Trx1_WCGPA is released from PtsI under reducing conditions. The molecular mass markers (in kDa) are indicated on the left. The fact that PtsI was observed in the nonreduced sample indicates that a portion of this protein co-purified with Trx1_WCGPA on the Ni-NTA agarose column. B, Strains expressing different versions of PtsI were streaked on MacConkey plates (containing 0.5% glucose) and glucose fermentation was analyzed. Red colonies indicate glucose fermentation whereas white colonies show no fermentation. Strains are as follows: 1. Wild type, 2. ΔptsI, 3. ptsI::ptsI_AMCS, 4. ptsI::ptsI_C502S. Cells with a nonfunctional PtsI cannot ferment glucose because of an impaired glucose uptake. C, The complex formed between Trx1_WCGPA and PtsI_AMCS was analyzed as in A, in a ΔtrxAΔgshA mutant expressing PtsI_AMCS from the chromosome (instead of wild-type PtsI). The molecular mass markers (in kDa) are indicated on the left. D, Identification of dimedone on Cys502 of PtsI. The LC-MS/MS spectrum shows data obtained from a +2 parent ion of [M+2H]²⁺ = 789.9. “m” in the peptide sequence corresponds to a methionine sulfoxide. The Cx residue corresponds to a sulfenic acid modified by dimedone which produces a +138Da mass increment; the y- and b- series of ions allow exact localization of the modified Cys.

CONCLUSIONS

E. coli Trx1 has been identified in 1964 (4). Since then, it has been the focus of intense scrutiny and has become one of the best-characterized members of a large family of oxidoreductases present in all domains of life. However, despite a wealth of information on Trx1, a comprehensive search for proteins depending on Trx1 for reduction had never been carried out. In our study, by expressing a Trx1 mutant with a catalytic site engineered for substrate trapping in cells with altered redox properties, we identified a total of 201 new putative Trx1 targets, considerably expanding the number of intracellular proteins depending on Trx1 for reduction. In addition, we identified proteins that become Trx1 substrates following exposure of the cells to an exogenous oxidative stress as well as proteins that most likely depend on both the Trx and Grx pathways for reduction. Although it is possible that Trx1 reduces disulfide bonds that could form in some of these proteins as a result of a catalytic activity, it is more likely that the majority of the newly identified proteins depend on Trx1 for the reduction of sensitive cysteine residues prone to oxidation by ROS. Determining the identity of the cysteine residues involved in the formation of mixed-disulfide intermediates with Trx1 and identifying the nature of the oxidative modifications affecting the Trx1 substrates will be fields of future research. Furthermore, it will be particularly interesting to elucidate if the oxidation of some of the newly identified Trx1 substrates is part of a regulation mechanism controlling their activity under more oxidizing redox conditions, which could lead to the discovery of new redox-regulated pathways.