Thiol-Redox Proteomics to Study Reversible Protein Thiol Oxidations in Bacteria

Thiol-redox proteomics methods are rapidly developing tools in redox biology. These are applied to identify and quantify proteins with reversible thiol oxidations that are formed under normal growth and oxidative stress conditions inside cells. The proteins with reversible thiol oxidations are usually prepared by alkylation of reduced thiols, subsequent reduction of disulﬁde bonds followed by a second differential alkylation of newly released thiols. Here, we describe two methods for detection of protein S -thiolations in Gram-positive bacteria using the direct shotgun approach and the ﬂuorescent-label thiol-redox proteomics method that have been successfully applied in our previous work.


Introduction
Bacteria are exposed to reactive oxygen species (ROS) during aerobic respiration or under infection conditions by the oxidative burst encountered by activated neutrophils [1,2]. ROS can damage all cellular macromolecules, but the most susceptible target is the thiol group of cysteine in proteins. The thiol group is subject to reversible oxidation by ROS to form intramolecular or intermolecular protein disulfides or mixed protein disulfides with low molecular weight (LMW) thiols (termed as S-thiolations). Highly reactive ROS, such as hydroxyl radicals are formed in the Fenton reaction and can lead to irreversible overoxidation of protein thiols to sulfinic or sulfonic acids leading to a loss of protein function [3] (Fig. 1).
Protein S-thiolations protect the thiol groups against overoxidation and function as thiol-redox switches to control protein activities. In eukaryotes and Gram-negative bacteria, the LMW thiol glutathione (GSH) is used for protein S-glutathionylation. Protein S-glutathionylation controls numerous cellular functions and is involved in many physiological and pathophysiological processes [4]. However, most Gram-positive bacteria lack the ability to synthesize GSH and alternative redox buffers are used for protein Sthiolations. Actinomycetes utilize the alternative LMW thiol mycothiol (MSH, Acetyl-Cys-GlcN-myoinositol) and Firmicutes produce bacillithiol (BSH, Cys-GlcN-malate) [5]. Thus far, the knowledge about the targets and the physiological role of protein S-thiolations in bacteria is limited. Recent advances in redox proteomics and mass spectrometry have facilitated the detection of various forms of protein S-thiolations in bacteria, including Scysteinylation, S-bacillithiolations, and S-mycothiolations [6][7][8][9][10]. Here, we provide two protocols that were applied in our lab to identify these different forms of protein S-thiolations in Gram-positive bacteria, such as Staphylococcus carnosus, Bacillus subtilis, and Corynebacterium glutamicum. We describe first the fluorescent-label based thiol-redox proteomics method that was developed to quantify and visualize reversibly oxidized protein thiols using the two-dimensional gel electrophoresis method (2D PAGE) [7,9]. In this approach, all reversibly oxidized proteins are reduced and labeled with the fluorescent dye BODIPY FL C 1 Fig. 1 Thiol chemistry of reactive oxygen species (ROS). Reversible thiol oxidation by ROS leads first to a Cys sulfenic acid intermediate (R-SOH) that is unstable and reacts further to form intramolecular and intermolecular disulfides or mixed disulfides with LMW thiols, such as glutathione, bacillithiol, cysteine, or CoASH, termed as S-thiolations. The Cys sulfenic acid can be also overoxidized to Cys sulfinic and sulfonic acids which are irreversible thiol oxidations (N-(4,4-difluoro-5,7-dimethyl-4-bora-3a,4a-diaza-s-indacene-3-l) methyl)iodoacetamide). The BODIPY-labeled protein extracts are separated by 2D-PAGE and scanned for BODIPY-fluorescence. The gels are subsequently stained for total protein amounts by Coomassie blue or a compatible fluorescent protein dye. Using the Decodon Delta 2D software an overlay image is generated of the fluorescent-labeled thiol-redox proteome and the Coomassie stained protein amount image. Finally, the level of oxidation is quantified in comparison to the protein amount and the oxidized proteins can be identified using MALDI-TOF mass spectrometry (Figs. 2b and 3).
The direct identification of the different forms of protein Sthiolations is based on a bottom-up proteomics approach using shotgun LC-MS/MS analysis of alkylated proteins samples under non-reducing conditions [6,7,10]. The reduced thiols in the protein extracts are blocked using iodoacetamide (IAM) or N-ethylmaleimide (NEM) during cell disruption and the extracts The MS/MS spectrum shows as example the S-bacillithiolated MetE-Cys730 peptide. Bacillithiol (BSH) is composed of cysteine, glucosamine and malate. The abundant neutral loss precursor ion is characteristic for the loss of malate (À134 Da) and serves as indicator for the S-bacillithiolated peptide since malate of BSH is lost during fragmentation. (b) For the thiol redox proteome analysis, reduced protein thiols are alkylated with IAM and reversibly oxidized proteins (-S-SR) are reduced with TCEP and labeled with the fluorescent dye BODIPY FL C 1 -IA. Proteins are separated using 2D PAGE, scanned for BODIPY fluorescence and stained with Colloidal Coomassie blue are directly applied for tryptic in-gel digestion and mass spectrometry to identify for example S-bacillithiolated or S-mycothiolated peptides (Fig. 2a). The 2D gel-based method and gel-free shotgun approach use similar protocols for thiol-trapping by IAM or NEM during sample preparation leading to a significant overlap between the identified reversibly oxidized proteins with those identified with S-thiolations [7][8][9]. However, even in combination both methods cover only a small fraction of protein S-thiolations and have several limitations. For example, false-positive redox-regulated proteins may be identified using the 2D gel-based approach and S-thiolated proteins cannot be enriched using both methods. Hence, these 2D gel-based method and shotgun-approach should be combined with more sensitive mass spectrometry-based methods of redox  Fig. 3 Fluorescence-label thiol redox proteomics to visualize reversibly oxidized proteins in B. subtilis before (control) and after diamide stress. Diamide is a thiol-reactive azo compound that causes disulfide stress in cells as shown by the chemistry above. For the thiol redox proteome analysis, cells were harvested at control and diamide stress conditions and prepared as described in Fig. 2b. The overlay image shows the BODIPY fluorescence image (redox proteome, red image) compared with the Coomassie-stained protein amount image (green image). Quantitative image analysis is performed using the DECODON Delta 2D software (http:/ / www.decodon.com) to calculate fluorescence (redox) ratios versus protein amounts. Proteins which appear in red in the overlay images are highly oxidized proteins with low protein levels, yellow spots are abundant oxidized proteins and green proteins are reduced proteins. The strongly increased reversible protein thiol oxidation after diamide stress is visualized which includes also protein S-thiolations proteomics, such as OxICAT or the NEM-Biotin-switch assay for comprehensive quantification of protein S-thiolations [11][12][13][14]. These mass spectrometry-based thiol-trapping assays could be even further developed to use purified bacilliredoxins or mycoredoxins for selective reduction of S-bacillithiolations and Smycothiolations, respectively. This would facilitate the enrichment and more comprehensive identification and quantification of the various protein S-thiolation forms in bacteria. Similarly, the glutaredoxin-coupled NEM-Biotin-switch assay has been successfully applied for global identification of S-glutathionylations in different eukaryotic cells [11,13]. Further advances in mass spectrometry and the design of novel chemical probes will provide leads for a more comprehensive description of the dynamics of the complete S-thiolome in various bacterial cells.

Materials
Centrifugation of protein samples in Eppendorf tubes is performed using a microfuge. For concentration of protein samples, a vacuum concentrator (SpeedVac) is used. Cells are disrupted using a Precellys-homogenizer (Peqlab). The rehydration chamber is required for rehydration of the IPG-strips. The Multiphor electrophoresis instrument including a power supply (e.g., EPS3500-XL from Pharmacia) is used for the IEF run. The 2D electrophoresis unit and a power supply are used for the subsequent second dimension (SDS-PAGE). In addition, casting trays must be provided for casting the 1D and 2D gels. Scanning of the 2D redox gels is performed using the Typhoon Scanner.

Methods
Fluorescent-label thiol-redox proteomics to visualize reversibly oxidized proteins in bacteria in response to oxidative stress.

Fluorescent-Label Thiol-Redox Proteomics: Growth, Harvesting of Cells, and Blocking of Reduced Thiols by IAM
Fluorescent-label thiol-redox proteomics is carried out to visualize reversibly oxidized proteins in bacteria in response to oxidative stress.
1. Inoculate an overnight culture of bacterial cells into two flasks each containing 100 mL defined minimal medium to an optical density at 500 nm (OD 500 ) of 0.05.
2. Grow bacterial cells until the mid-exponential phase (e.g., OD 500 of 0.4). Flask 1 is used as control and flask 2 is exposed to sublethal NaOCl concentration which allows the cells to recover from growth arrest. The sublethal NaOCl concentration has to be determined for the specific bacterium before the stress experiment.
3. Harvest bacterial cultures in flask 1 before the stress (control) and that in flask 2 after 30 min of NaOCl stress exposure in 2 Â 50 mL Falcon-like tubes on ice by rapid centrifugation (ca 9500 Â g, 5 min, 4 C).
4. Wash cell pellets immediately in 1-2 mL TE-IAM buffer to remove the oxidant, transfer the cells to 2 mL Eppendorf-like tubes, repeat centrifugation using a microfuge (19,400 Â g, 5 min, 4 C).
5. Resuspend washed cell pellet on ice immediately in 400 μL UCE buffer with 100 mM IAM. Do not vortex to avoid air bubbles and resuspend the cell pellet slowly using a pipette tip (see Note 2).
6. Break cells in UCE-IAM buffer immediately using a homogenizer in the presence of glass beads and remove glass beads by short centrifugation in a microfuge (19,400 Â g, 5 min, 4 C).
Thiol alkylation is performed for 15 min in the dark (see Note 2).
7. Add four parts pure acetone to one part of protein extract in 2 mL Eppendorf-like tubes (e.g., add 0.4 mL protein extract to 1.6 mL acetone), precipitate proteins for 1 h on ice and centrifuge protein pellet in a microfuge (19,400 Â g, 20 min, room temperature (RT)).
8. Wash protein pellet with 80% (v/v) acetone by mixing up the pellet mechanically with a pipette tip to ensure the proteins are suspended and any trace of IAM is removed. Remove supernatant after centrifugation using a microfuge (19,400 Â g, 20 min, 20 C) and repeat washing and centrifugation with 80% (v/v) acetone four times. Dry the washed protein pellet in a vacuum centrifuge and store proteins frozen at À20 C until reduction and BODIPY labeling (see Note 3).

5.
Place the IPG strips in the isoelectric focusing chamber. On top at both ends of the IPG strips place electrode strips that are soaked with distilled water. Place the electrodes on top of these electrode strips and connect these with the cathode and anode.
Overlay the IPG strips with mineral oil, start running the IEF using the power supply EPS3500-XL according to the IEF following parameters (see Note 7):

5.
Step: 3000 V 1 mA 5 W 57,000 Vh 19 h 6. Remove IPG strips from the chamber and soak the mineral oil from the surface of the strips by a filter paper. Store IPG strips at À20 C until separation by SDS-PAGE.

Second
Dimension of 2D SDS-PAGE of Fluorescent-Labeled Proteins 1. For 12.5% acrylamide-bisacrylamide 2D gels prepare the separating gel solution as described in Subheading 2.1.2, item 19. Immediately after initiating the polymerization process cast the separating gel solution into the funnel of the multicasting chamber. Remove bubbles by pressing the flexible tube which connects the funnel and the multicasting chamber. Open the valve to cast slowly the gel solution through the flexible tube into the multicasting chamber. Make sure to avoid air bubbles in the chamber. Cover the separating gels with butanol to ensure a homogeneous polymerization of the gel. Acrylamide polymerization is finished within 2 h.

1.
Inoculate three flasks each containing 100 mL minimal medium with overnight cultures to an optical density of OD 500 of 0.07 and grow cells until the mid-exponential growth phase (OD 500 of 0.5-1.0). Flask 1 is used as untreated control. Flask 2 is exposed to diamide, and flask 3 is treated with NaOCl at sublethal concentrations each for 30 min. These sublethal NaOCl and diamide concentrations should reduce the growth rate and have to be determined before in detailed growth analyses.
2. Harvest the three bacterial cultures from untreated cells (control) and cells exposed for 30 min to diamide and NaOCl stress in 50 mL Falcon tubes on ice by rapid centrifugation (9500 Â g, 10 min, 4 C).
3. Wash bacterial cell pellets in TE-IAM buffer, transfer to Eppendorf tubes, centrifuge again (17,900 Â g, 5 min, 4 C) and resuspend the cell pellets in 1 mL UCE-IAM buffer on ice.
4. Disrupt cells using the homogenizer on ice and centrifuge cell extracts twice for 30 min at 4 C to remove the cell debris. Alkylate the proteins containing free thiol-groups by incubation of the protein extract for 15 min at RT in the dark. 2. Cell sampling, washing, and blocking of reduced thiols by IAM for the fluorescent-label Thiol-Redox proteomics method needs to be done at the same day until the acetoneprecipitation step is completed and the IAM-alkylated protein extracts is dried (step 8). If the cell pellet is not quickly dissolved and broken in UCE-IAM buffer, artificial oxidation can occur since reduced thiols are not blocked by IAM. The acetone precipitated protein pellet can be stored for several days at À20 C.
3. Washing of the IAM-alkylated protein extract in acetone must be done carefully by using a pipette tip to mechanically disturb the protein pellet. Traces of remaining IAM must be avoided since these prevent the BODIPY-IAM labeling.
4. For BODIPY-IAM labeling, we recommend to spin down the BODIPY-IAM stock solution for removal of precipitated fluorescent dye which affects the thiol labeling.
5. Equilibrate the Biorad spin columns directly before the removal of excess BODIPY-IAM dye from the samples.
6. Reswelling of the IPG strips, IEF and the SDS-PAGE should be performed in the dark. The IPG strips should be subjected to SDS-PAGE directly after the IEF run is finished to avoid loss of the fluorescence label.
7. The IEF running parameters are used according to the instructions as recommended for IPG BlueStrips from SERVA (http://www.scie-plas.com/documents/SCIE-PLAS/distributors/445.pdf.) 8. The scanning of the fluorescence images is performed using a Typhoon scanner and requires to wet the surface of the scanner with distilled water. After the gel run, the 2D gel is removed from the glass plates and placed immediately to the scanner surface in the water by avoiding air bubbles under the gel. For gel scanning, the PMT voltage needs to be adjusted according to the fluorescence intensity of the spots. We further recommend for the fluorescence quantification to run all gels of one experiment (e.g., gels of controls and stress samples) at the same day in one 2D gel run.
9. For preparation of protein extracts for direct identifications of S-thiolations using Orbitrap LC-MS/MS analysis, the reduced thiols need to be IAM-alkylated directly after cell harvesting and alkylated proteins are subsequently used for fractionation by non-reducing SDS-PAGE. Protein sample processing at the same day avoids precipitation of IAM-alkylated proteins which often occurs after freezing the IAM-alkylated protein extracts.
To maintain the thiol-modifications, it is required that cells need to be broken, alkylated, and processed by SDS-PAGE at the same day. However, it is possible to store cells frozen at À70 C before preparation of protein extracts for SDS-PAGE and mass spectrometry.