The authors would like to thank Dr. Greg Martin (Cornell University) and his group for providing the DC3000 strain, tomato seeds, and advice. They would also like to thank Dr. Zhonglin Mou for help with the DAB protocol and the Proteomics Division at UF Interdisciplinary Center for Biotechnology Research for assistance in the method development. The protocol for protein extraction was modified from Hurkman and Tanaka16. The protocol on cysTMT labeling, steps 4 to 6 was adapted based on the original Thermo Pierce Fisher Scientific product manual17. This work was funded by National Science Foundation (MCB 0818051 to S Chen).

1. Growing Seedlings and Preparing Bacteria
<ol>
	<li>Seedlings are germinated in MetroMix 500 soil mixture (BWI Companies) in a growth chamber (160 &mu;mol photons m-2s-1 with a photoperiod of 8 hour light at 22 &deg;C and 16 hour dark at 20 &deg;C for 1 week. Relative humidity was set to 70%. Seedlings were watered with tap water as needed to keep the soil from becoming dry.</li>
	<li>Two seedlings of similar size are then transplanted to 4" diameter pots containing the MetroMix 500 soil and grown for an additional 3 weeks in the same conditions as 1.1.</li>
	<li>Pseudomonas syringae (Pst) is initially T-streaked onto Kings B Medium (KBM) plates (per liter: 20 g Proteose Peptone, 0.1975 g MgSO4, 1% Glycerol, 1.5 g K2HPO4, 18 g Agar) and grown for two days at 28 &deg;C. A single colony is collected, mixed into 300&mu;l liquid Kings B Medium (20 g Proteose peptone, 0.1975 g MgSO4, 10 mL Glycerol, 10 mL of 100x K2HPO4 stock (1.5 g of K2HPO4 in 10 mL of H2O ), and spread onto a Kings B Medium plate. This is cultured over night at 28 &deg;C.</li>
	<li>The bacteria inoculum is prepared by scraping the cultured bacteria from the KBM plate into 20 mL H2O. An OD600 of 0.03 (~106 cfu/ml) is prepared in 1 L of inoculation buffer (10 mM MgCl2 + 400 &mu;L Silwett-70). Inoculation buffer minus bacteria was prepared for the control solution.</li>
</ol>
2. Inoculation of Tomato with Pst and H2O2 Histochemical Stain
<ol>
	<li>Four week old plants were inoculated by dipping into the above inoculation solution for 30 seconds. Clear plastic bags were put over the plants immediately after inoculation.</li>
	<li>3'-3' diaminobenzidine (DAB) stain11 was prepared (1 mg/ml DAB in H2O, pH 3.8 (HCl)) three days before use to achieve complete solubilization. The solution was rocked to mix at room temperature.</li>
	<li>Twenty four hours after treatment, the fully expanded leaf was removed, placed in DAB stain epidermal side up, and vacuum infiltrated for 15 min. The leaves were then put in the dark at room temperature over night for staining.</li>
	<li>Leaves were boiled in 95% ethanol for 10 min, and then kept in 75% ethanol.</li>
</ol>
3. Protein Extraction
<ol>
	<li>After harvesting, tomato leaves were ground to fine powder with liquid nitrogen using mortar and pestle.</li>
	<li>For 0.5 g of tissue, add 1.25 mL Tris pH 8.8 buffered phenol and 1.25 mL / 0.5 g of extraction buffer (0.1 M Tris-HCl pH 8.8, 10 mM EDTA, 0.9 M sucrose) then continue grinding for a few more min in a fume hood.</li>
	<li>Transfer the extract to Oakridge centrifuge tube (Thermo Fisher Scientific Nalgene Company) and agitate for 2 hrs at room temperature.</li>
	<li>Centrifuge at 5,000x g and 15 &deg;C for 10 min. Transfer the top phenol phase to a new clean tube.</li>
	<li>Back-extract the aqueous phase with equal volume extracted phenol buffer; agitate on a shaker for 30 min. Centrifuge and transfer the extract to a new Falcon tube.</li>
	<li>Repeat the above step one more time and transfer the extract to new Oakridge tube.</li>
	<li>Precipitate phenol extracted proteins by adding 5 volumes of 0.1 M ammonium acetate in 100% methanol (stored at -20 &deg;C). Vortex and incubate at -20 &deg;C for overnight.</li>
	<li>Collect the protein pellet by centrifugation at 20,000x g, 4 &deg;C for 20 min .</li>
	<li>Wash the pellet 2 times with the cold 0.1M ammonium acetate in methanol and 2 times with 80% cold acetone. Add 1.5 mL cold 70% ethanol to pellet and suspend. Transfer to a 2 mL microcentrifuge tube (USA Scientific)and centrifuge at 14,000 rpm, 4 &deg;C for 20 min. Remove ethanol.</li>
	<li>Dry the pellet briefly in a SpeedVac concentrator and dissolve in a protein extraction buffer, e.g. ReadyPrep Protein Extraction Kit Reagent 3 (8 M Urea, 4% CHAPS, 40 mM Tris-base, 2 M Thiourea) (Bio-Rad). Centrifuge at 14,000 rpm and 20 &deg;C for 30 min to form a pellet. Collect the supernatant.</li>
	<li>Measure the protein concentration using a CB-X protein assay according to manufacturer's manual (Geno Technology).</li>
</ol>
4. Sample Preparation and Peptide Labeling with cysTMTs
<ol>
	<li>Prepare protein sample at a concentration of 2-5 &mu;g/&mu;L. Here we use 100 &mu;g of protein for each mass tag, 20 &mu;l-50 &mu;L samples. For this experiment all 6 tags were used.</li>
	<li>Add equal volume of alkylation buffer (100 mM Tris-HCl pH7.5, 200 mM iodoacetamide) to the protein sample to block the free thiol group. Buffer should be prepared fresh. Alkylation is performed at 37 &deg;C for 1 hour.</li>
	<li>Precipitate the protein by adding 1 mL of cold 80% acetone at -20&deg;C for overnight. Pellet the protein by centrifuging at 14,000 rpm, 4 &deg;C for 20 min. Wash the pellet 3 times with 80% Acetone vortexing each time. Remove the supernatant and allow to air dry while on ice.</li>
	<li>Resuspend the pellet in 50 &mu;L lysis buffer (6 M Urea, 50 mM Tris base, 1 mM EDTA). 0.5 &mu;L of 100 mM Tris(2-Carboxyethyl) Phosphine (TCEP) is then added to reduce the disulfide bonds by incubating for one hour at room temperature.</li>
	<li>Prepare tags by adding 20 &mu;L of acetonitrile to the reagent tubes to solubilize the tags. Vortex and spin down to the bottom.</li>
	<li>Wash out extra TCEP using a Microcon 3KD centrifugal filter device (Millipore). Add 50 &mu;L sample to Microcon 3KD column, and 50 &mu;L lysis buffer to the column. Centrifuge for 15 min at 10,000x g and 4 &deg;C. Repeat this step for a total of three times. Remove column and invert into a clean tube. Centrifuge for 5 min at 1,000x g and 4 &deg;C.</li>
	<li>Check the pH. If needed, adjust the pH to 7.0-8.0 with 1 M HCl.</li>
	<li>Add 5 &mu;L of the cysTMT reagent to each sample (The cysTMT reagent kit provides 20 &mu;L tags for up to 500 &mu;g protein. This method uses 100 &mu;g protein, therefore we use one-fifth of the tag.) . Allow the reaction to proceed for 2 hr at room temperature.</li>
	<li>Samples were combined with Laemmli Sample Buffer (Bio-Rad) in a 1:1 ratio.</li>
</ol>
5. Removal of Non-reacted Tag Sample Fractionation
<ol>
	<li>Samples were boiled for 5 min and separated on a pre-cast 12% polyacrylamide gel (Bio-Rad). The gel was rinsed three times in 5 minute intervals with filtered H2O and stained with Coomassie Blue (Bio-Rad) for 1 hour. The gel was de-stained overnight with H2O.</li>
	<li>Twelve fractions were collected from each gel lane. Fractions were determined by protein band concentrations. Fractions were chopped into 1mm pieces and collected into a 1.5 mL tube.</li>
	<li>Gel pieces were de-stained using 50% acetonitrile and 0.1 M ammonium bicarbonate in H2O. This step must be repeated until blue color is removed from gel pieces (~3-4 times, 15 min each time).</li>
	<li>De-stained gel pieces were dried using 100% acetonitrile (cover gel pieces) for 15 min, supernatant removed, and the pieces dried using a SpeedVac.</li>
	<li>Dissolve trypsin (Promega) at 1:10 (trypsin: protein) ratio in the same volume (as pooled labeled sample) of 25 mM ammonium bicarbonate. For example, 600 &mu;g protein sample with a volume of 500 &mu;L needs 60 &mu;g trypsin dissolved in 500 &mu;L of 25 mM ammonium bicarbonate.</li>
	<li>Add the trypsin solution to the gel pieces and set on ice to rehydrate. If pieces are not covered add 50 mM ammonium bicarbonate buffer. After rehydration, incubate at 37 &deg;C for 12-16 hr.</li>
	<li>Remove the liquid protein extract from the incubated samples.</li>
	<li>Stop the reaction and remove any remaining protein digest by covering gel pieces with 5% formic acid, 50% acetonitrile.. Shake gel pieces for 20 min at room temperature. Transfer the supernatant to the extracted protein sample tubes from 5.7. Repeat the extraction procedure twice. Dry the extracted peptides using the SpeedVac.</li>
</ol>
6. Sample Enrichment of cysTMT Labeled-peptides
<ol>
	<li>Add a gradient of anti-TMT resin to 0.5 mL tubes for a 50% slurry. (Gradient was determined from band concentration in fractionation gel. If a fraction collection consists of a dark band, more resin is needed. Fractions with weaker bands require less resin as there is less protein.)</li>
	<li>The resin is then washed three times with one column volume of 1x Tris-buffered saline (TBS) (25 mM Tris, 0.15 M NaCl, pH 7.2) (Thermo Scientific Pierce Protein Research Products).</li>
	<li>Add 200 &mu;L 1xTBS to each sample. Samples are then added to anti-TMT resin (Thermo Scientific Pierce Protein Research Products) and agitated at room temperature for 2 hr followed by rocking overnight at 4 &deg;C.</li>
	<li>Add sample to column (Thermo Scientific Pierce Protein Research Products).</li>
	<li>Wash each column three times with 1x 200 &mu;L TBS. This is then followed with washing three times with 0.05% CHAPs (dissolved in 1x TBS).</li>
	<li>The column is then washed three times with 4 M urea in 1x TBS. Two hundred microliters of H2O is then used to wash the column three times.</li>
	<li>Each sample is eluted three times with 200 &mu;L 50% elution buffer (50% acetonitrile, 0.4% TFA).</li>
	<li>Samples are then dried in a vacuum concentrator.</li>
</ol>
7. Mass Spectrometry Analysis
<ol>
	<li>Resuspend the samples in 12 &mu;L of 3% acetonitrile with 0.1% formic acid and inject 5 &mu;L directly onto an Eksigent NanoLC-1D high pressure liquid chromatography column (AB SCIEX, USA).</li>
	<li>Peptides will be separated on a Proteopep II C18 column 75 &mu;m ID x 20 cm (New Objective, USA) using a 4-60% gradient (A: 3% acetonitrile, 0.1% formic acid; B: 97% acetonitrile, 0.1% formic acid) at 3 &mu;L/min over 60 min.</li>
	<li>A Thermo Scientific LTQ Orbitrap XL mass spectrometer was used to detect peptides using a top 2 x 3 experiment consisting of single stage MS followed by acquisition of 3 MS/MS spectra with higher-energy C-trap dissociation (HCD) fragmentation followed by 3 MS/MS with collision induced dissociation (CID) for protein identification. Parameters in this mode were: Isolation width: 3.0 m/z; Collision energy: 50% (10% with two steps). Only doubly- and triply-charged peptides were selected for fragmentation. Dynamic exclusion parameters were set to: Repeat count = 1; Repeat Duration = 60; Exclusion list size = 500; Exclusion duration = 28. Target values are as follows: MS = 5 e5; MS/MS (HCD) = 1 e5. Ion transfer times were set to 500 for FTMS and 300 for MS/MS (HCD). Two microscans were required for HCD spectra.</li>
</ol>
8. Database Searching and Quantitation
<ol>
	<li>The acquired CID and HCD data were analyzed using Thermo Scientific Proteome Discoverer software 1.2 (Thermo Scientific Pierce Protein Research Products) through a branched workflow which implemented a Reporter Ion Quantizer (20 ppm mass tolerance of fragment ions) to quantify the ratio. A separate segment processed the MS2 spectra through the Spectrum Selector, Spectrum Normalizer, and Spectrum Grouper nodes.</li>
	<li>Data were then searched against the SEQUEST search engine. A 20 ppm mass tolerance was used. Static modifications were set to the cysTMT reagent (304.18 Da) and dynamic modifications included phosphorylation and methionine oxidation (15.99 Da). A custom database for Solanum lycopersicum was composed using RNA data (with about 350,000 entries from Harvard University)13 was used in both cases.</li>
</ol>
9. Representative Results
A representative image of a control tomato plant leaf and a Pseudomonas inoculated leaf is shown in Figure 1. A difference between control treated and Pseudomonas treated leaves is observed. After the leaves are removed and stained using DAB, the de-staining process allows for the histochemical stain to show signs of ROS in the leaf tissue (Figure 2). Figure 2A is representative of a control leaf with no staining. Figure 2B is representative of a leaf treated with Pseudomonas and positive staining for H2O2 production. An example of Proteome Discover data output of a differentially redox regulated protein is shown in Figure 3. This protein is a known redox regulated protein ferredoxin-114 and has been shown to play a role in defense against Pseudomonas syringae pv tomato15. Peak intensity between control and inoculated samples is used to obtain relative quantification, which showed significant changes in ferredoxin-1 redox regulation (p&lt; 0.05). High intensity peaks suggest that this protein is oxidized in response to the pathogen treatment. Figure 4 is an example of Proteome Discover data output of a protein that has similar redox regulation of a protein between a control and inoculated sample. Peaks of similar intensity suggest the presence of disulfide bonds not regulated by a change in treatment. The method will revolutionize how scientists detect redox responsive cysteines and disulfides10.
<img alt="Figure 1" src="/files/ftp_upload/3766/3766fig1.jpg" /> 
Figure 1. A representative image of inoculated tomato leaves with control solution (A) and Pseudomonas (B).
<img alt="Figure 2" src="/files/ftp_upload/3766/3766fig2.jpg" /> 
Figure 2. A representative image of DAB staining of inoculated tomato leaves with control solution (A) and Pseudomonas (B). Leaves were stained using 3'-3' diaminobenzidine. Chlorophyll was removed from leaves by boiling in 95% ethanol. Dark staining indicates the presence of H2O2. Only leaves inoculated with bacteria culture showed dark staining.
<img alt="Figure 3" src="/files/ftp_upload/3766/3766fig3.jpg" /> 
Figure 3. An example of Proteome Discover data output of ferredoxin-1, differentially redox regulated protein14. Peak intensity above each peak is used for absolute quantification. Peak intensity between control and inoculated samples is used to perform relative quantification.
<img alt="Figure 4" src="/files/ftp_upload/3766/3766fig4.jpg" /> 
Figure 4. An example of Proteome Discover data output of a protein that has similar peak intensity between a control and inoculated sample. Peaks of similar intensity suggest the presence of disulfide bonds not regulated by a change in treatment.

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+stimulation+of+an+oxidative+burst+during+elicitation+of+cultured+plant+cells:+role+in+defense+and+signal+transduction.">Rapid stimulation of an oxidative burst during elicitation of cultured plant cells: role in defense and signal transduction.</a> Plant Physiol. 90, 109-116 (1989).</li><li>Orozco-Cardenas, M., Ryan, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hydrogen+peroxide+is+generated+systemically+in+plant+leaves+by+wounding+and+systemin+via+the+octadecanoid+pathway.">Hydrogen peroxide is generated systemically in plant leaves by wounding and systemin via the octadecanoid pathway.</a> Proc. Natl. Acad. Sci. U.S.A. 96, 6553-6557 (1999).</li><li>Tonks, N. 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No conflicts of interest declared.

This protocol provides information on performing DAB staining as well as cysTMT labeled redox cysteine quantification. These procedures are beneficial in examining production of ROS as well as the effect on protein regulation when Solanum lycopersicum is inoculated with Pseudomonas syringae. The methods presented in this protocol provide a way to examine ROS in whole leaf samples in a way that causes the least amount of damage to leaf tissue. The labeling procedure provides a way to examine potentially redox regulated proteins by utilizing a cysteine labeling method. This is beneficial when examining an early stage of stress response.
Methods such as isotope-coded affinity tag (ICAT) and cysTMT can be used in examining potential redox regulated proteins in biological samples. ICAT allows labeling and comparison of two samples12. Both methods label free cysteines and can be used for protein quantification10,12. However, the cysTMT method allows for a decrease in experimental variation as well as multiplexing10. The number of tags available allows researchers to include replicates or multiple samples in their experimental design. Having more samples provides the potential for a higher number of proteins identified. A major disadvantage of the cysTMT technique is that it compromises the overall quality of protein identification because of the selective enrichment steps for cysTMT labeled-peptides (6.5-6.6). The number of peptides for the protein identification depends largely on the number of cysteine residues in the protein sequence. This problem can be overcome by submitting part of the tryptic sample before enrichment for mass spectrometry protein identification.
Due to the nature of the experimental design as well as the labeling mechanism that the cysTMT method utilizes, certain steps are critical. While performing protein precipitation and pellet washings (3.9) it is important to keep samples chilled on ice to reduce protein degradation. During cysTMT labeling, removal of the reducing reagent (4.6) is important because samples may undergo reverse labeling. Reverse labeling is possible if reducing reagent remains in the sample. If samples are reduced after labeling, the cysTMT tag can be removed. Once the labels are added to the samples, the pH level must be checked (4.7) in order to have optimal labeling efficiency. In addition, data analysis is dependent on what is required of the researcher and the ultimate goal in using the protocol. It is also dependent on the software being used as each software has different algorithms.
This experiment utilizes pathogen as an elicitor for enhanced production of reactive oxidative species in tomato; however, other redox regulated responses may be measured accordingly. This experimental design is adaptable to other plant and animal systems.

<table border="1">
<tbody>
 <tr>
 <td>Name 			of the reagent</td>
 <td>Company</td>
 <td>Catalogue 			number</td>
 </tr>
 <tr>
 <td>MetroMix 			500</td>
 <td>BWI 			Companies</td>
 <td>TX-500</td>
 </tr>
 <tr>
 <td>3,3'-Diaminobenzidine</td>
 <td>Sigma-Aldrich</td>
 <td>D8001</td>
 </tr>
 <tr>
 <td>ReadyPrep 			Sequential Extraction kit Reagent 3</td>
 <td>Bio-Rad</td>
 <td>163-2104</td>
 </tr>
 <tr>
 <td>CB-X 			protein assay</td>
 <td>Geno 			Technology</td>
 <td>786-12x</td>
 </tr>
 <tr>
 <td>cysTMT 			reagents</td>
 <td>Thermo 			Scientific Pierce Protein Research Products</td>
 <td>90071</td>
 </tr>
 <tr>
 <td>Laemmli 			Sample Buffer</td>
 <td>Bio-Rad</td>
 <td>161-0737</td>
 </tr>
 <tr>
 <td>Bio-Safe 			Comassie (G-250 stain)</td>
 <td>Bio-Rad</td>
 <td>161-0786</td>
 </tr>
 <tr>
 <td>Microcon 			3KD column</td>
 <td>Millipore</td>
 <td>42403</td>
 </tr>
 <tr>
 <td>Immobilized 			Anti-TMT resin</td>
 <td>Thermo 			Scientific Pierce Protein Research Products</td>
 <td>90076</td>
 </tr>
 <tr>
 <td>Centrifuge 			column</td>
 <td>Thermo 			Scientific Pierce Protein Research Products</td>
 <td>89896</td>
 </tr>
 <tr>
 <td>Proteopep 			II C18 column</td>
 <td>New 			Objective</td>
 <td>PFC7515-PP2-10</td>
 </tr>
 <tr>
 <td>NanoLC-1D 			HPLC</td>
 <td>AB 			Sciex</td>
 <td>90389</td>
 </tr>
 <tr>
 <td>LTQ 			Orbitrap XL</td>
 <td>Thermo 			Scientific</td>
 <td>0020137580</td>
 </tr>
 <tr>
 <td>SpeedVac</td>
 <td>Labconco</td>
 <td>7812013</td>
 </tr>
 <tr>
 <td>Proteome 			Discoverer 1.2 software</td>
 <td>Thermo 			Scientific Pierce Protein Research Products</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Trypsin</td>
 <td>Promega</td>
 <td>V5111</td>
 </tr>
 <tr>
 <td>Oakridge 			Centrifuge Tube</td>
 <td>Thermo 			Scientific Nalgene Company</td>
 <td>3139-0050</td>
 </tr>
 <tr>
 <td>Microcentrifuge 			tube (2ml)</td>
 <td>USA 			Scientific</td>
 <td>1620-2700</td>
 </tr>
 <tr>
 <td>12% 			Mini-PROTEAN 			TGX 			Precast Gel</td>
 <td>Bio-Rad</td>
 <td>456-1043</td>
 </tr>
 <tr>
 <td> Top of Form 
&gt;Bio-Safe 			Coomassie StainBottom 			of Form
 </td>
 <td>Bio-Rad</td>
 <td>161-0786</td>
 </tr>
 <tr>
 <td>TMT 			enrichment kit</td>
 <td>Thermo 			Scientific Pierce Protein Research Products</td>
 <td>90077</td>
 </tr>
 </tbody>
</table>

profiling thiol redox proteome using isotope tagging mass spectrometry

Pseudomonas syringae pv. tomato strain DC3000 not only causes bacterial speck disease in Solanum lycopersicum but also on Brassica species, as well as on Arabidopsis thaliana, a genetically tractable host plant1,2. The accumulation of reactive oxygen species (ROS) in cotyledons inoculated with DC3000 indicates a role of ROS in modulating necrotic cell death during bacterial speck disease of tomato3. Hydrogen peroxide, a component of ROS, is produced after inoculation of tomato plants with Pseudomonas3. Hydrogen peroxide can be detected using a histochemical stain 3'-3' diaminobenzidine (DAB)4. DAB staining reacts with hydrogen peroxide to produce a brown stain on the leaf tissue4. ROS has a regulatory role of the cellular redox environment, which can change the redox status of certain proteins5. Cysteine is an important amino acid sensitive to redox changes. Under mild oxidation, reversible oxidation of cysteine sulfhydryl groups serves as redox sensors and signal transducers that regulate a variety of physiological processes6,7. Tandem mass tag (TMT) reagents enable concurrent identification and multiplexed quantitation of proteins in different samples using tandem mass spectrometry8,9. The cysteine-reactive TMT (cysTMT) reagents enable selective labeling and relative quantitation of cysteine-containing peptides from up to six biological samples. Each isobaric cysTMT tag has the same nominal parent mass and is composed of a sulfhydryl-reactive group, a MS-neutral spacer arm and an MS/MS reporter10. After labeling, the samples were subject to protease digestion. The cysteine-labeled peptides were enriched using a resin containing anti-TMT antibody. During MS/MS analysis, a series of reporter ions (i.e., 126-131 Da) emerge in the low mass region, providing information on relative quantitation. The workflow is effective for reducing sample complexity, improving dynamic range and studying cysteine modifications. Here we present redox proteomic analysis of the Pst DC3000 treated tomato (Rio Grande) leaves using cysTMT technology. This high-throughput method has the potential to be applied to studying other redox-regulated physiological processes.

Watch this Scientific Journal Video about Profiling Thiol Redox Proteome Using Isotope Tagging Mass Spectrometry at JoVE.com

Profiling Thiol Redox Proteome Using Isotope Tagging Mass Spectrometry

Reactive oxygen species level is elevated when cells encounter stress conditions. Here we show the example of 3'-3' diaminobenzidine staining as well as cysTMT labeling and mass spectrometry to profile the redox proteome in Pseudomonas syringae treated tomato leaves.

Pseudomonas syringae pv. tomato strain DC3000 not only causes bacterial speck disease in Solanum lycopersicum but also on Brassica species, as well as on ...

profiling-thiol-redox-proteome-using-isotope-tagging-mass-spectrometry

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

16.1K Views.  University of Florida. Reactive oxygen species level is elevated when cells encounter stress conditions. Here we show the example of 3'-3' diaminobenzidine staining as well as cysTMT labeling and mass spectrometry to profile the redox proteome in Pseudomonas syringae treated tomato leaves.

Video: Profiling Thiol Redox Proteome Using Isotope Tagging Mass Spectrometry

MALDI Sample Preparation: the Ultra Thin Layer Method

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS) 1,2. The ultra-thin layer method involves the production of a substrate layer of matrix crystals (alpha-cyano-4-hydroxycinnamic acid) on the sample plate, which serves as a seeding ground for subsequent crystallization of a matrix/analyte mixture. Advantages of the ultra-thin layer method over other sample deposition approaches (e.g. dried droplet) are that it provides (i) greater tolerance to impurities such as salts and detergents, (ii) better resolution, and (iii) higher spatial uniformity. This method is especially useful for the accurate mass determination of proteins. The protocol was initially developed and optimized for the analysis of membrane proteins and used to successfully analyze ion channels, metabolite transporters, and receptors, containing between 2 and 12 transmembrane domains 2. Since the original publication, it has also shown to be equally useful for the analysis of soluble proteins. Indeed, we have used it for a large number of proteins having a wide range of properties, including those with molecular masses as high as 380 kDa 3. It is currently our method of choice for the molecular mass analysis of all proteins. The described procedure consistently produces high-quality spectra, and it is sensitive, robust, and easy to implement.

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS).

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption ...

A Quantitative Assessment of The Yeast Lipidome using Electrospray Ionization Mass Spectrometry

Lipids are one of the major classes of biomolecules and play important roles membrane dynamics, energy storage, and signalling1-4. The budding yeast Saccharomyces cerevisiae, a genetically and biochemically manipulable unicellular eukaryote with annotated genome and very simple lipidome, is a valuable model for studying biological functions of various lipid species in multicellular eukaryotes2,3,5. S. cerevisiae has 10 major classes of lipids with chain lengths mainly of 16 or 18 carbon atoms and either zero or one degree of unsaturation6,7. Existing methods for lipid identification and quantification - such as high performance liquid chromatography, thin-layer chromatography, fluorescence microscopy, and gas chromatography followed by MS - are well established but have low sensitivity, insufficiently separate various molecular forms of lipids, require lipid derivitization prior to analysis, or can be quite time consuming. Here we present a detailed description of our experimental approach to solve these inherent limitations by using survey-scan ESI/MS for the identification and quantification of the entire complement of lipids in yeast cells. The described method does not require chromatographic separation of complex lipid mixtures recovered from yeast cells, thereby greatly accelerating the process of data acquisition. This method enables lipid identification and quantification at the concentrations as low as g/ml and has been successfully applied to assessing lipidomes of whole yeast cells and their purified organelles. Lipids extraction from whole yeast cells for using this method of lipid analysis takes two to three hours. It takes only five to ten minutes to run each sample of extracted and dried lipids on a Q-TOF mass spectrometer equipped with a nano-electrospray source.

We describe a new quantitative lipidomics method for identifying numerous lipid species in yeast using survey-scan electrospray ionization mass spectrometry (ESI/MS). This method exceeds currently available methods for lipid identification and quantification in the ability to resolve various molecular forms of lipids, sensitivity, and speed.

Lipids are one of the major classes of biomolecules and play important roles membrane dynamics, energy storage, and signalling1-4. The budding yeast ...

Analyzing Large Protein Complexes by Structural Mass Spectrometry

Living cells control and regulate their biological processes through the coordinated action of a large number of proteins that assemble themselves into an array of dynamic, multi-protein complexes1. To gain a mechanistic understanding of the various cellular processes, it is crucial to determine the structure of such protein complexes, and reveal how their structural organization dictates their function. Many aspects of multi-protein complexes are, however, difficult to characterize, due to their heterogeneous nature, asymmetric structure, and dynamics. Therefore, new approaches are required for the study of the tertiary levels of protein organization.
One of the emerging structural biology tools for analyzing macromolecular complexes is mass spectrometry (MS)2-5. This method yields information on the complex protein composition, subunit stoichiometry, and structural topology. The power of MS derives from its high sensitivity and, as a consequence, low sample requirement, which enables examination of protein complexes expressed at endogenous levels. Another advantage is the speed of analysis, which allows monitoring of reactions in real time. Moreover, the technique can simultaneously measure the characteristics of separate populations co-existing in a mixture. 
Here, we describe a detailed protocol for the application of structural MS to the analysis of large protein assemblies. The procedure begins with the preparation of gold-coated capillaries for nanoflow electrospray ionization (nESI). It then continues with sample preparation, emphasizing the buffer conditions which should be compatible with nESI on the one hand, and enable to maintain complexes intact on the other. We then explain, step-by-step, how to optimize the experimental conditions for high mass measurements and acquire MS and tandem MS spectra. Finally, we chart the data processing and analyses that follow. Rather than attempting to characterize every aspect of protein assemblies, this protocol introduces basic MS procedures, enabling the performance of MS and MS/MS experiments on non-covalent complexes. Overall, our goal is to provide researchers unacquainted with the field of structural MS, with knowledge of the principal experimental tools.

Mass spectrometry has proven to be a valuable tool for analyzing large protein complexes. This method enables insights into the composition, stoichiometry and overall architecture of multi-subunit assemblies. Here, we describe, step-by-step, how to perform a structural mass spectrometry analysis, and characterize macromolecular structures.

Living cells control and regulate their biological processes through the coordinated action of a large number of proteins that assemble themselves into an ...

OLIgo Mass Profiling (OLIMP) of Extracellular Polysaccharides

The direct contact of cells to the environment is mediated in many organisms by an extracellular matrix. One common aspect of extracellular matrices is that they contain complex sugar moieties in form of glycoproteins, proteoglycans, and/or polysaccharides. Examples include the extracellular matrix of humans and animal cells consisting mainly of fibrillar proteins and proteoglycans or the polysaccharide based cell walls of plants and fungi, and the proteoglycan/glycolipid based cell walls of bacteria. All these glycostructures play vital roles in cell-to-cell and cell-to-environment communication and signalling.
An extraordinary complex example of an extracellular matrix is present in the walls of higher plant cells. Their wall is made almost entirely of sugars, up to 75% dry weight, and consists of the most abundant biopolymers present on this planet. Therefore, research is conducted how to utilize these materials best as a carbon-neutral renewable resource to replace petrochemicals derived from fossil fuel. The main challenge for fuel conversion remains the recalcitrance of walls to enzymatic or chemical degradation due to the unique glycostructures present in this unique biocomposite.
Here, we present a method for the rapid and sensitive analysis of plant cell wall glycostructures. This method OLIgo Mass Profiling (OLIMP) is based the enzymatic release of oligosaccharides from wall materials facilitating specific glycosylhydrolases and subsequent analysis of the solubilized oligosaccharide mixtures using matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF/MS)1 (Figure 1). OLIMP requires walls of only 5000 cells for a complete analysis, can be performed on the tissue itself2, and is amenable to high-throughput analyses3. While the absolute amount of the solubilized oligosaccharides cannot be determined by OLIMP the relative abundance of the various oligosaccharide ions can be delineated from the mass spectra giving insights about the substitution-pattern of the native polysaccharide present in the wall.
OLIMP can be used to analyze a wide variety of wall polymers, limited only by the availability of specific enzymes4. For example, for the analysis of polymers present in the plant cell wall enzymes are available to analyse the hemicelluloses xyloglucan using a xyloglucanase5, 11, 12, 13, xylan using an endo-&beta;-(1-4)-xylanase 6,7, or for pectic polysaccharides using a combination of a polygalacturonase and a methylesterase 8. Furthermore, using the same principles of OLIMP glycosylhydrolase and even glycosyltransferase activities can be monitored and determined 9.

OLIgo Mass Profiling OLIMP of Extracellular Polysaccharides

A rapid way is described to gain insights into the structure of polysaccharides in an extracellular matrix. The method takes advantage of the specificity of glycosylhydrolases and the sensitivity of mass spectrometry allowing minute amounts of materials to be analyzed. This technique is adaptable to be used directly on tissue itself.

The direct contact of cells to the environment is mediated in many organisms by an extracellular matrix. One common aspect of extracellular matrices is ...

Direct Analysis of Single Cells by Mass Spectrometry at Atmospheric Pressure

Analysis of biochemicals in single cells is important for understanding cell metabolism, cell cycle, adaptation, disease states, etc. Even the same cell types exhibit heterogeneous biochemical makeup depending on their physiological conditions and interactions with the environment. Conventional methods of mass spectrometry (MS) used for the analysis of biomolecules in single cells rely on extensive sample preparation. Removing the cells from their natural environment and extensive sample processing could lead to changes in the cellular composition. Ambient ionization methods enable the analysis of samples in their native environment and without extensive sample preparation.1 The techniques based on the mid infrared (mid-IR) laser ablation of biological materials at 2.94 &mu;m wavelength utilize the sudden excitation of water that results in phase explosion.2 Ambient ionization techniques based on mid-IR laser radiation, such as laser ablation electrospray ionization (LAESI) and atmospheric pressure infrared matrix-assisted laser desorption ionization (AP IR-MALDI), have successfully demonstrated the ability to directly analyze water-rich tissues and biofluids at atmospheric pressure.3-11 In LAESI the mid-IR laser ablation plume that mostly consists of neutral particulate matter from the sample coalesces with highly charged electrospray droplets to produce ions. Recently, mid-IR ablation of single cells was performed by delivering the mid-IR radiation through an etched fiber. The plume generated from this ablation was postionized by an electrospray enabling the analysis of diverse metabolites in single cells by LAESI-MS.12 This article describes the detailed protocol for single cell analysis using LAESI-MS. The presented video demonstrates the analysis of a single epidermal cell from the skin of an Allium cepa bulb. The schematic of the system is shown in Figure 1. A representative example of single cell ablation and a LAESI mass spectrum from the cell are provided in Figure 2.

Single cell analysis is performed by mass spectrometry on plant and animal cells at atmospheric pressure by using a sharpened optical fiber to sample the cells for laser ablation electrospray ionization (LAESI) mass spectrometry.

Analysis of biochemicals in single cells is important for understanding cell metabolism, cell cycle, adaptation, disease states, etc. Even the same cell ...

Profiling of Methyltransferases and Other S-adenosyl-L-homocysteine-binding Proteins by Capture Compound Mass Spectrometry (CCMS)

There is a variety of approaches to reduce the complexity of the proteome on the basis of functional small molecule-protein interactions such as affinity chromatography 1 or Activity Based Protein Profiling 2. Trifunctional Capture Compounds (CCs, Figure 1A) 3 are the basis for a generic approach, in which the initial equilibrium-driven interaction between a small molecule probe (the selectivity function, here S-adenosyl-L-homocysteine, SAH, Figure 1A) and target proteins is irreversibly fixed upon photo-crosslinking between an independent photo-activable reactivity function (here a phenylazide) of the CC and the surface of the target proteins. The sorting function (here biotin) serves to isolate the CC - protein conjugates from complex biological mixtures with the help of a solid phase (here streptavidin magnetic beads). Two configurations of the experiments are possible: "off-bead" 4 or the presently described "on-bead" configuration (Figure 1B). The selectivity function may be virtually any small molecule of interest (substrates, inhibitors, drug molecules).
S-Adenosyl-L-methionine (SAM, Figure 1A) is probably, second to ATP, the most widely used cofactor in nature 5, 6. It is used as the major methyl group donor in all living organisms with the chemical reaction being catalyzed by SAM-dependent methyltransferases (MTases), which methylate DNA 7, RNA 8, proteins 9, or small molecules 10. Given the crucial role of methylation reactions in diverse physiological scenarios (gene regulation, epigenetics, metabolism), the profiling of MTases can be expected to become of similar importance in functional proteomics as the profiling of kinases. Analytical tools for their profiling, however, have not been available. We recently introduced a CC with SAH as selectivity group to fill this technological gap (Figure 1A).
SAH, the product of SAM after methyl transfer, is a known general MTase product inhibitor 11. For this reason and because the natural cofactor SAM is used by further enzymes transferring other parts of the cofactor or initiating radical reactions as well as because of its chemical instability 12, SAH is an ideal selectivity function for a CC to target MTases. Here, we report the utility of the SAH-CC and CCMS by profiling MTases and other SAH-binding proteins from the strain DH5&alpha; of Escherichia coli (E. coli), one of the best-characterized prokaryotes, which has served as the preferred model organism in countless biochemical, biological, and biotechnological studies. Photo-activated crosslinking enhances yield and sensitivity of the experiment, and the specificity can be readily tested for in competition experiments using an excess of free SAH.

Profiling of Methyltransferases and Other S-adenosyl-L-homocysteine-binding Proteins by Capture Compound Mass Spectrometry CCMS

Capture Compounds are trifunctional small molecules to reduce the complexity of the proteome by functional reversible small molecule-protein interaction followed by photo-crosslinking and purification. Here we use a Capture Compound with S-adenosyl-L-homocysteine-binding as selectivity function to isolate methyltransferases from an Escherichia coli whole cell lysate and identify them by MS.

There is a variety of approaches to reduce the complexity of the proteome on the basis of functional small molecule-protein interactions such as affinity ...

Quantification of Proteins Using Peptide Immunoaffinity Enrichment Coupled with Mass Spectrometry

There is a great need for quantitative assays in measuring proteins. Traditional sandwich immunoassays, largely considered the gold standard in quantitation, are associated with a high cost, long lead time, and are fraught with drawbacks (e.g. heterophilic antibodies, autoantibody interference, 'hook-effect').1 An alternative technique is affinity enrichment of peptides coupled with quantitative mass spectrometry, commonly referred to as SISCAPA (Stable Isotope Standards and Capture by Anti-Peptide Antibodies).2 In this technique, affinity enrichment of peptides with stable isotope dilution and detection by selected/multiple reaction monitoring mass spectrometry (SRM/MRM-MS) provides quantitative measurement of peptides as surrogates for their respective proteins. SRM/MRM-MS is well established for accurate quantitation of small molecules 3, 4 and more recently has been adapted to measure the concentrations of proteins in plasma and cell lysates.5-7 To achieve quantitation of proteins, these larger molecules are digested to component peptides using an enzyme such as trypsin. One or more selected peptides whose sequence is unique to the target protein in that species (i.e. "proteotypic" peptides) are then enriched from the sample using anti-peptide antibodies and measured as quantitative stoichiometric surrogates for protein concentration in the sample. Hence, coupled to stable isotope dilution (SID) methods (i.e. a spiked-in stable isotope labeled peptide standard), SRM/MRM can be used to measure concentrations of proteotypic peptides as surrogates for quantification of proteins in complex biological matrices. The assays have several advantages compared to traditional immunoassays. The reagents are relatively less expensive to generate, the specificity for the analyte is excellent, the assays can be highly multiplexed, enrichment can be performed from neat plasma (no depletion required), and the technique is amenable to a wide array of proteins or modifications of interest.8-13 In this video we demonstrate the basic protocol as adapted to a magnetic bead platform.

Stable Isotope Standards and Capture by Anti-Peptide Antibodies (SISCAPA) couples affinity enrichment of peptides with stable isotope dilution mass spectrometry (MRM-MS) to provide quantitative measurement of peptides as surrogates for their respective proteins. Here we describe the protocol using magnetic particles in a partially automated format.

There is a great need for quantitative assays in measuring proteins. Traditional sandwich immunoassays, largely considered the gold standard in ...

Selected Reaction Monitoring Mass Spectrometry for Absolute Protein Quantification

Absolute quantification of target proteins within complex biological samples is critical to a wide range of research and clinical applications. This protocol provides step-by-step instructions for the development and application of quantitative assays using selected reaction monitoring (SRM) mass spectrometry (MS). First, likely quantotypic target peptides are identified based on numerous criteria. This includes identifying proteotypic peptides, avoiding sites of posttranslational modification, and analyzing the uniqueness of the target peptide to the target protein. Next, crude external peptide standards are synthesized and used to develop SRM assays, and the resulting assays are used to perform qualitative analyses of the biological samples. Finally, purified, quantified, heavy isotope labeled internal peptide standards are prepared and used to perform isotope dilution series SRM assays. Analysis of all of the resulting MS data is presented. This protocol was used to accurately assay the absolute abundance of proteins of the chemotaxis signaling pathway within RAW 264.7 cells (a mouse monocyte/macrophage cell line). The quantification of Gi2 (a heterotrimeric G-protein &#945;-subunit) is described in detail.

This protocol describes how to perform absolute quantification assays of target proteins within complex biological samples using selected reaction monitoring. It was used to accurately quantify proteins of the mouse macrophage chemotaxis signaling pathway. Target peptide selection, assay development, and qualitative and quantitative assays are described in detail.

Absolute quantification of target proteins within complex biological samples is critical to a wide range of research and clinical applications. This ...

Identification of MyoD Interactome Using Tandem Affinity Purification Coupled to Mass Spectrometry

Skeletal muscle terminal differentiation starts with the commitment of pluripotent mesodermal precursor cells to myoblasts. These cells have still the ability to proliferate or they can differentiate and fuse into multinucleated myotubes, which maturate further to form myofibers. Skeletal muscle terminal differentiation is orchestrated by the coordinated action of various transcription factors, in particular the members of the Muscle Regulatory Factors or MRFs (MyoD, Myogenin, Myf5, and MRF4), also called the myogenic bHLH transcription factors family. These factors cooperate with chromatin-remodeling complexes within elaborate transcriptional regulatory network to achieve skeletal myogenesis. In this, MyoD is considered the master myogenic transcription factor in triggering muscle terminal differentiation. This notion is strengthened by the ability of MyoD to convert non-muscle cells into skeletal muscle cells. Here we describe an approach used to identify MyoD protein partners in an exhaustive manner in order to elucidate the different factors involved in skeletal muscle terminal differentiation. The long-term aim is to understand the epigenetic mechanisms involved in the regulation of skeletal muscle genes, i.e., MyoD targets. MyoD partners are identified by using Tandem Affinity Purification (TAP-Tag) from a heterologous system coupled to mass spectrometry (MS) characterization, followed by validation of the role of relevant partners during skeletal muscle terminal differentiation. Aberrant forms of myogenic factors, or their aberrant regulation, are associated with a number of muscle disorders: congenital myasthenia, myotonic dystrophy, rhabdomyosarcoma and defects in muscle regeneration. As such, myogenic factors provide a pool of potential therapeutic targets in muscle disorders, both with regard to mechanisms that cause disease itself and regenerative mechanisms that can improve disease treatment. Thus, the detailed understanding of the intermolecular interactions and the genetic programs controlled by the myogenic factors is essential for the rational design of efficient therapies.

MyoD is a myogenic transcription factor with a strong capacity to induce myogenic transdifferentiation of many fully differentiated non-muscle cell lines. The epigenetic mechanisms involved in this transdifferentiation are largely unknown. Here we describe a double-affinity purification method followed by mass spectrometry to exhaustively characterize MyoD partners.

Skeletal muscle terminal differentiation starts with the commitment of pluripotent mesodermal precursor cells to myoblasts. These cells have still the ...

Cellular Redox Profiling Using High-content Microscopy

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, excessive ROS levels induce a state of oxidative stress, which is accompanied by irreversible oxidative damage to DNA, lipids and proteins. Thus, quantification of ROS provides a direct proxy for cellular health condition. Since mitochondria are among the major cellular sources and targets of ROS, joint analysis of mitochondrial function and ROS production in the same cells is crucial for better understanding the interconnection in pathophysiological conditions. Therefore, a high-content microscopy-based strategy was developed for simultaneous quantification of intracellular ROS levels, mitochondrial membrane potential (&#916;&#936;m) and mitochondrial morphology. It is based on automated widefield fluorescence microscopy and image analysis of living adherent cells, grown in multi-well plates, and stained with the cell-permeable fluorescent reporter molecules CM-H2DCFDA (ROS) and TMRM (&#916;&#936;m and mitochondrial morphology). In contrast with fluorimetry or flow-cytometry, this strategy allows quantification of subcellular parameters at the level of the individual cell with high spatiotemporal resolution, both before and after experimental stimulation. Importantly, the image-based nature of the method allows extracting morphological parameters in addition to signal intensities. The combined feature set is used for explorative and statistical multivariate data analysis to detect differences between subpopulations, cell types and/or treatments. Here, a detailed description of the assay is provided, along with an example experiment that proves its potential for unambiguous discrimination between cellular states after chemical perturbation.

This paper presents a high-content microscopy workflow for simultaneous quantification of intracellular ROS levels, as well as mitochondrial membrane potential and morphology &#8211; jointly referred to as mitochondrial morphofunction &#8211; in living adherent cells using the cell-permeant fluorescent reporter molecules 5-(and-6)-chloromethyl-2&#39;,7&#39;-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA) and tetramethylrhodamine methylester (TMRM).

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, ...