The authors acknowledge support from the National Institutes of Health (NIH) under Grant R01 GM075092. The Thermo Orbitrap Fusion mass spectrometer used to acquire some of the data described here was acquired with funds from the National Institutes of Health grant S10OD010645.

1. Protein and reagent preparation
NOTE: This protocol includes an example workflow for labeling a protein with DEPC. Some conditions and reagent concentrations listed may vary based on the protein of choice.
<ol>
	<li>Prepare all reagent solutions in 1.5 mL microcentrifuge tubes.</li>
	<li>Prepare a protein solution of desired concentration, usually in the range of tens of &#181;M, in a 10 mM 3-(N-morpholino)propanesulfonic acid (MOPS) buffer at pH 7.4. Alternatively, prepare a buffer-exchange existing protein solution in 10 mM pH 7.4 MOPS if the sample contains a nucleophilic buffer that would be reactive with DEPC. Other buffers (e.g., phosphate buffered saline) can also be used, as long as they do not have nucleophilic functional groups.</li>
	<li>Prepare a 100 mM DEPC solution in dry acetonitrile (ACN) by pipetting 1.45 &#181;L of the stock 6.9 M DEPC solution into 98.55 &#181;L of the ACN. 
	NOTE: There is no one set of concentrations that will work with every protein, though the optimal concentrations can be estimated based on the number of His and Lys residues23. For example, with 50 &#181;L of a 50 &#181;M &#946;2-microglobulin solution, react the protein with 0.2 &#181;L of the 100 mM DEPC for a final DEPC concentration of 200 &#181;M (equal to 4x the protein concentration) to ensure the desired molar ratio using this general example (Table 1). DEPC labeling is a 2nd order reaction, so changing either the concentration of protein or DEPC in the reaction mixture will change the labeling rate.</li>
	<li>Prepare 1 M imidazole solution by weighing out 10 mg of imidazole and dissolving in 146.9 &#181;L of HPLC-grade water.</li>
</ol>
2. Covalent labeling of intact protein
<ol>
	<li>Set water bath temperature to 37 &#176;C and wait for the bath to reach a stable temperature. 
	NOTE: Reagent concentrations and volumes for an example labeling protocol can be found in Table 1.</li>
	<li>In a new microcentrifuge tube, mix MOPS buffer and protein solution in volumes listed in Table 1.</li>
	<li>To the protein and buffer add 0.2 &#181;L of the DEPC solution, making sure to properly mix the resulting solution, and then place the tube containing the reaction mixture into the 37&#160;&#176;C water bath for 1 minute. 
	NOTE: The volume of ACN added should not exceed 1% of the total reaction volume to avoid perturbation of the protein&#39;s structure during the labeling reaction. Reaction time is up to the user, although a 1 minute reaction under the example conditions minimizes overlabeling and the potential hydrolysis of DEPC14.</li>
	<li>After 1 minute, remove the tube containing the reaction mixture from the water bath and quench the reaction with 1 &#181;L of the 1 M imidazole solution to scavenge the remaining unreacted DEPC. 
	NOTE: The final concentration of imidazole in the reaction mixture should equal 50x the concentration of DEPC in the reaction mixture. This will ensure that remaining unreacted DEPC is scavenged.</li>
</ol>
3. Preparation of protein digest for bottom-up LC-MS
NOTE: Choose digestion conditions that are amenable to the protein of interest. Common steps involve unfolding the protein and reducing and alkylating any disulfide bonds.
<ol>
	<li>Unfold the protein by adding an appropriate unfolding reagent to the reaction mixture. 
	NOTE: Common unfolding agents include ACN, urea, and guanidine hydrochloride (GuHCl).</li>
	<li>Prepare solutions of Tris(2-carboxyethyl)phosphine (TCEP) and iodoacetamide (IAM) by weighing out 5 mg of each and dissolving them in new microcentrifuge tubes in 174.4 and 270.3 &#181;L of 10 mM pH 7.4 MOPS buffer, respectively, for the reduction and alkylation steps.</li>
	<li>Reduce disulfide bonds by adding 2 &#181;L of the 100 mM TCEP (final concentration of 2 mM in reaction mixture) solution to the reaction mixture and reacting for 3 minutes at room temperature. 
	NOTE: The final concentration of TCEP should equal 40x the protein concentration per disulfide bond present in the solution.</li>
	<li>Alkylate reduced cysteines with 4 &#181;L of the 100 mM IAM solution (final concentration of 4 mM in reaction mixture) for 30 minutes in the dark. IAM is light-sensitive and will decompose under direct light. 
	NOTE: The final concentration of IAM in solution should be twice the concentration used for TCEP, or 80x the protein concentration per disulfide bond.</li>
	<li>Digest the protein with an appropriate enzyme such as trypsin or chymotrypsin. A 10:1 protein:enzyme ratio for a 3-hour digestion at 37&#160;&#176;C with immobilized enzyme at a shaking rate of 300 strokes/min is typically sufficient for DEPC-labeled proteins. See Discussion.</li>
	<li>After digestion, separate the immobilized enzyme from the digested peptides by centrifugation at 12,000 rpm for 5 minutes.</li>
	<li>Analyze the sample immediately by LC-MS/MS or flash-freeze the sample with liquid nitrogen to minimize sample degradation and label loss. Store the flash-frozen samples at &#60; -20 &#176;C until ready for LC-MS/MS analysis.</li>
</ol>
4. LC-MS/MS Analysis
NOTE: Standard LC-MS/MS parameters for bottom-up proteomics can be used to identify labeled sites on the proteolytic peptide fragments. A general example is outlined below.
<ol>
	<li>Separate the DEPC-labeled peptides using a reversed-phase C18 stationary phase. Use a typical LC mobile phase of two solvents: (A) water + 0.1% formic acid and (B) ACN + 0.1% formic acid using a gradient (e.g., Figure 2) to achieve the best separation of peptides. 
	NOTE: The separation time can be optimized based on sample complexity, and mobile phase flow rate depends on whether capillary or nano LC is used.</li>
	<li>Use a mass spectrometer capable of doing on-line LC-MS and MS/MS to identify DEPC modification sites on the peptide. In our experiments, we have successfully used several types of mass spectrometers. Any mass spectrometer capable of automatically performing MS/MS of many peptides during the course of an LC-MS analysis should be suitable. Relevant MS parameters include: ESI source voltage = -4000 V for regular ESI; -2000 V for nanospray; Orbitrap resolution = 60,000; Dynamic exclusion duration = 30 s; MS/MS activation type: CID, ETD, or both; Mass scan range = 200-2,000; Automatic Gain Control = 4.0E5 (MS1 in Orbitrap) and 5.0E4 (MS2 in linear quadrupole ion trap).</li>
	<li>Load and inject the digested, labeled protein sample into the LC system and start the LC-MS/MS acquisition. If the sample has been flash-frozen, thaw before analysis. Divert the LC effluent to waste for the first 5 minutes to avoid excessive salts from getting into the ESI source. 
	&#8203;NOTE: A 5 &#181;L injection loop is generally utilized, allowing for injection of approximately 2.5 &#181;g of protein to the LC-MS/MS. This is dependent on the loading conditions of the LC as to not clog the sample injector.</li>
</ol>
5. Data analysis
<ol>
	<li>Identify DEPC label sites and quantify peptide peak areas using appropriate software for the mass spectrometer that is used.</li>
	<li>Include DEPC addition (72.02 Da) and carbamidomethylation (57.02 Da) as variable modifications. Additional search parameters for the MS/MS analysis are as follows: Maximum missed cleavages = 3; Fragment ion types = b and y; Precursor m/z tolerance = 10 ppm (this value should be higher if a quadrupole ion trap mass spectrometer is used); Fragment m/z tolerance = 0.5 Da (this value should be lower if a high-resolution mass spectrometer is used for a product ion scan); Precursor charge = 1-4. 
	NOTE: Different database search algorithms have different scoring systems, and many can have difficulty identifying DEPC-modified peptides because modification levels can be low. Adjusting the score cutoff may be necessary to identify more labeled peptides. If so, then manual interrogation of the MS/MS data should be used to verify low-scoring peptides. The product of the hydrolysis of the DEPC label is not included in the data searching because the hydrolyzed DEPC is no longer reactive toward nucleophilic side chains.</li>
	<li>Determine residue-level modification percentages using the chromatographic peak areas of the modified and unmodified versions of the peptides. 
	NOTE: Any peptide containing the modified residue of interest must be considered and all charge states that are included must be present in all the measured samples. Peptides having different ionization efficiencies and eluting at different times causes this value to be a relative rather than absolute measure of the modification of a specific site. 
	<img alt="Equation 1" src="/files/ftp_upload/61983/61983eq01.jpg" /> 
	where Ai,z represents peak area of any given peptide (i) that contains the residue of interest and considers all detected charge states (z).</li>
	<li>Determine if a labeling change between a control and experimental sample is significant using statistical evaluation. Three replicate measurements for each sample is typical, and t-tests are most commonly utilized with 95 or 99% confidence intervals.</li>
</ol>

The authors have nothing to disclose.

Critical Steps 
Several points regarding experimental design should be considered to ensure reliable labeling results. First, to maximize protein labeling, it is necessary to avoid buffers with strongly nucleophilic groups (e.g., Tris) because they can react with DEPC and lower the extent of labeling. It is also conceivable that such buffers could react with labeled residues, causing the removal of the label and therefore loss of structural information. We recommend MOPS as a buffer, but phosphate buffered saline works as well. Second, dithiothreitol should be avoided for reduction of disulfide bonds because the free thiols in this reagent can react with labeled residues and remove the DEPC modification. This same chemistry is why it is essential to alkylate reduced disulfides because free Cys can cause intra-protein label scrambling33. Third, the quench step with imidazole (Protocol step 2.4) is critical for stopping the labeling reaction so any remaining DEPC is unable to continue to react with the protein.
It is also important to appreciate that the DEPC labeling reaction is 2nd order, meaning that changing either the protein or DEPC concentration will affect the extent of labeling. We have empirically found that a 4:1 DEPC:protein concentration ratio is a reasonable value when the protein has a concentration in the 10&#39;s of &#181;M range, as it tends to avoid any labeling-induced structural changes14. However, the optimum DEPC:protein concentration ratio is protein-dependent as well as concentration-dependent, and some proteins require higher ratios to achieve sufficient labeling. The optimal DEPC concentration to minimize the structural perturbation of labeling a given protein can be estimated from the number of solvent accessible His and Lys residues23.
To avoid variability in labeling extents due to reagent degradation over time, the control and experimental reactions should ideally be performed on the same day. DEPC undergoes rapid hydrolysis when exposed to water and will degrade during storage as well, so good lab practice and reagent handling are key. When not in use, the DEPC stock bottle should be stored in a desiccator.
Modifications and Troubleshooting 
Because DEPC CL is a kinetically controlled reaction, the labeling rate, and thus the modification level, are controlled by protein and reagent concentrations, labeling time, and temperature. As indicated above, often DEPC CL is performed for 1 minute at 37 &#176;C with a DEPC to protein molar ratio of 4 to 1. At this molar ratio (4x), proteins can be safely labeled without structural perturbations14. DEPC:protein molar ratios that are greater than 4 can be used for some proteins, and not surprisingly higher DEPC concentrations result in more extensive labeling and more structural information. The safest way to use higher DEPC concentrations without perturbing structure is to generate dose-response plots in which the reactivity of a protein&#39;s proteolytic fragments are measured as a function of DEPC concentration (see Figure 4B)14,24. However, this approach can be time-consuming, as it requires multiple measurements. Recently, we demonstrated that the optimum DEPC concentration can be estimated for a given protein from the number and SASA of His and Lys residues in that protein23. In recent work, we have also suggested alternate ways to assess that a protein&#39;s structure is not perturbed during CL9,24.
Proteolytic digestion is also an important step in DEPC CL-MS, as the peptides that are generated allow one to localize structural information after LC-MS/MS analysis. In general, serine proteases such as trypsin and chymotrypsin are effective for protein digestion. Trypsin is more commonly used due to its cleavage efficiency and specificity at the C-terminal side of Lys and Arg residues. However, trypsin usually does not cleave after DEPC-labeled Lys residues, causing missed cleavage at labeled Lys residues that can sometimes complicate data analysis. The activity of chymotrypsin, which cleaves after bulky hydrophobic residues, is typically not affected by DEPC labeling, but this enzyme has lower cleavage efficiency and specificity than trypsin. In addition, for proteins with numerous hydrophobic residues, chymotrypsin can generate several short peptide fragments that can be difficult separate and detect under standard LC-MS conditions.
LC-ESI-MS/MS analyses of protein digests requires a stable electrospray to obtain reliable semiquantitative results. Capillary and nano LC are commonly used separation platforms where efficient separation can be obtained with small amounts of protein. However, in our experience, capillary LC provides more reliable quantitative information (i.e., modification levels) than nano LC because of the higher sample amounts that are used. For large proteins that are digested into a large number of labeled and unlabeled peptides, coelution of peptides with similar hydrophobicity may happen. In these cases, longer LC gradients should be used for better separations.
Fast and efficient MS/MS is important for good sequence coverage and label site identification. Mass spectrometers with quadrupole ion traps are excellent instruments for this purpose. Generally, CID is a common and highly effective method for peptide dissociation; however, we have observed cases of DEPC label scrambling during CID of labeled peptide ions, resulting in ambiguity in labeling site identity34. In these somewhat rare cases, ETD provides information that is more reliable. Consequently, if possible, alternating between both dissociation techniques can be helpful. Because DEPC can label many different residue types, it is common that isomers of a given peptide fragment are generated that differ in the side chain that is modified. Many times, these peptide isomers can be separated by LC, although they tend to elute at very similar times. This likelihood should be considered when setting MS/MS exclusion times during automated LC-MS/MS analyses. Typically, shorter than usual exclusion times should be used.
Limitations 
While DEPC CL-MS is tremendously beneficial for studying protein structure and interactions, and significant progress has been made to address a wide variety of protein systems, some limitations of this technique remain. If labeling conditions are not well optimized (e.g., using too high a concentration of DEPC), over labeling can perturb the protein structure and result in inaccurate structural information14,23,24. In addition, the accuracy and precision of the measured modification levels is affected by several factors, including label loss if samples sit too long before LC-MS analysis and errors in the chemical labeling steps. Consistency in each experimental step is important for reliable results. One of the more challenging issues currently associated with DEPC-based CL-MS is data analysis software. Readily available data analysis programs are not optimized to successfully identify peptides with low modification percentages (e.g., &#60; 1%). Consequently, we have developed and used specialized software to enable peptides with low modification levels to be readily identified18.
Even though DEPC CL-MS can provide moderate protein structural resolution, a detailed 3D structure cannot be obtained from the labeling technique. Recently, some structural prediction tools based on labeling data have been developed35,36. However, more developments are required to optimally incorporate DEPC labeling results with computational modeling to predict protein structure.
Significance Compared to Alternative Methods 
The structural resolution possible with DEPC-based CL-MS is moderate when compared to techniques like X-ray crystallography or NMR, so it is most appropriate to compare the technique to HDX-MS, XL-MS, and other CL-MS, or footprinting, approaches. As was mentioned in the introduction, CL-MS is complementary to HDX-MS because it provides information about the side chains of residues in a protein, whereas HDX-MS reports on backbone structure and dynamics. This complementarity allows the two techniques to be used together to obtain more structural information, as has been shown by several groups37,38,39,40,41. An emerging idea relevant to DEPC-based CL-MS is the synergistic information that is available when it is used in conjunction with HDX-MS22. Because of the labeling timeframes associated with the two techniques, DEPC-based CL-MS can often clarify ambiguity with protein regions that undergo decreased HDX. Decreased HDX can arise from either reduced solvent accessibility due to binding or decreased protein dynamics. DEPC reactions are inherently 2-3 orders of magnitude slower than HDX reactions, so DEPC labeling is largely unaffected by changes in protein dynamics as long as they are not accompanied by changes in solvent accessibility.
CL-MS has some advantages over HDX-MS experiments in that label scrambling and label loss are minimal when appropriate precautions are taken. In HDX experiments, label loss or back-exchange is a standard feature of the technique, and fast digestions and LC separations at low pH are attempts to minimize this problem. It is important to recognize, though, that the back exchange issue is counterbalanced by the higher number of labeled sites that are typically measured in HDX-MS. Labeling scrambling is a larger problem in HDX-MS because incorrect information would then be obtained, so analysis conditions must be optimized to minimize this concern.
XL-MS and CL-MS have similar advantages with regard to label loss and scrambling, as both involve the formation of a covalent bond that is robust enough to remain during digestion and LC-MS analyses. Sequencing and identification of cross-linked peptides, however, is more challenging than for covalently labeled peptides, requiring the use of specialized software. One key advantage that XL-MS has over CL-MS is its ability to provide distance constraints, which can be valuable when predicting or narrowing down possible protein structures. CL-MS data has been used to facilitate protein structure prediction36,42,43, but this area requires more work to fully reach its potential.
When compared to other CL-MS methods, including those that use specific or non-specific labeling reagents, DEPC has some advantages and disadvantages. Like methods that use amino acid specific reagents, DEPC labeling is straightforward; the reagent only needs to be added to the sample of interest. This simplicity contrasts with methods such as fast photochemical oxidation of proteins (FPOP) or synchrotron based HRF that require sophisticated light sources to produce hydroxyl radicals. Unlike methods that use specific reagents, DEPC can label six different amino acids and the N-terminus, enabling it to provide higher structural resolution. However, the number of residues that can be labeled by DEPC are fewer than the number that can be oxidized by hydroxyl radicals, so the structural resolution obtainable by DEPC-based CL-MS is lower. One practical ramification of fewer residues being labeled by DEPC is that using the reagent sometimes does not provide all desired structural information, and so it can benefit from being used in conjunction with other labeling reagents. We have recently demonstrated the value of using DEPC together with the reagent pair 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC)/glycine ethyl ether (GEE), which can label glutamate and aspartate residues19.

Proteins are essential biomolecules in virtually every physiological process. The variety of functions that proteins perform are possible because of the structures they adopt and the interactions that they have with other biomolecules. To understand protein function at a deeper level, biochemical and biophysical tools are needed to elucidate these important structural features and interactions. Traditionally, X-ray crystallography, cryogenic electron microscopy, and nuclear magnetic resonance (NMR) spectroscopy have provided the desired atomic-level detail to reveal protein structure. However, numerous protein systems cannot be interrogated by these techniques because of poor crystallization behavior, limited protein availability, excessive sample heterogeneity, or molecular weight limitations. Consequently, newer analysis methods have emerged that overcome these limitations. Among the emerging techniques that can provide protein structural information is mass spectrometry.
Mass spectrometry (MS) measures a molecule&#39;s mass-to-charge (m/z) ratio, so protein higher-order structural information must be obtained by encoding the desired structural information into the mass of the protein. Several approaches to encode this information have been developed, including hydrogen-deuterium exchange (HDX)1,2,3,4, chemical crosslinking (XL)5,6, and covalent labeling (CL)7,8,9,10. In HDX, backbone amide hydrogens are exchanged by slightly more massive deuteriums at rates that depend on solvent accessibility and H-bonding extent. The extent of HDX can be localized by rapidly digesting the protein into peptide fragments before separating and measuring these fragments by the mass spectrometer or by dissociating the protein in a top-down experiment. Determining the rate of exchange provides further insight into protein dynamics. HDX has proven to be a valuable tool for characterizing protein structure despite challenges associated with back exchange and the need for specialized equipment to maximize reproducibility. In XL-MS, proteins are reacted with bi-functional reagents that covalently link adjacent residue side chains within a given protein or between two proteins. In doing so, XL-MS can provide distance constraints that can be used to characterize protein structure. The regions of the protein that are cross-linked can be identified by proteolytic digestion followed by liquid chromatography (LC)-MS analysis. While XL-MS is a versatile tool that has been used to study a variety of protein complexes, including inside cells, identification of the XL products is challenging and requires specialized software.
CL-MS has emerged recently as a complementary and sometimes alternative MS-based tool to study protein structure and interactions. In CL-MS, a protein or protein complex is covalently modified with a mono-functional reagent that can react with solvent-exposed side chains (Figure 1). By comparing the modification extents of a protein or protein complex under different conditions, conformation changes, binding sites, and protein-protein interfaces can be revealed. After the CL reaction, site-specific information, often at the single amino-acid level, can be obtained using typical bottom-up proteomics workflows in which proteins are proteolytically digested, peptide fragments are separated by LC, and modified sites are identified using tandem MS (MS/MS). The rich history of bioconjugate chemistry has made numerous reagents available for CL-MS experiments. CL reagents fall into two general categories: (i) specific and (ii) non-specific. Specific reagents react with a single functional group (e.g., free amines)8,10 and are easy to implement, but they tend to provide limited structural information. Non-specific reagents react with a wide range of side chains, but often require specialized equipment such as lasers or synchrotron sources to produce these highly reactive species. Hydroxyl radicals are the most commonly used non-specific reagent, having been applied in hydroxyl radical footprinting (HRF)7,11,12,13 experiments to study a wide range of proteins and protein complexes under a variety of conditions.
Our research group has successfully used another relatively non-specific reagent called diethylpyrocarbonate (DEPC) to study protein structure and interactions in the context of CL-MS experiments14,15,16,17,18,19,20,21,22,23,24,25. DEPC offers the simplicity of specific labeling reagents (i.e., no specialized equipment is necessary to perform the reactions), while reacting with up to 30% of amino acids in the average protein. As a highly electrophilic reagent, DEPC is capable of reacting with the N-terminus and the nucleophilic side chains of cysteine, histidine, lysine, tyrosine, serine, and threonine residues. Typically, a single product of these reactions is generated, resulting in a mass increase of 72.02 Da. This single type of product contrasts with the up to 55 different products that can be produced when proteins react with hydroxyl radicals7. Such simple chemistry facilitates identification of labeled sites.
Here, we provide protocols for using DEPC-based CL-MS to study protein structure and interactions. Details associated with reagent preparation, DEPC-protein reactions, protein digestion conditions, LC-MS and MS/MS parameters, and data analysis are described. We also demonstrate the utility of DEPC labeling by providing example results from protein-metal, protein-ligand, and protein-protein interactions as well as proteins undergoing structural changes upon heating.

<table><tbody><tr><td>1.5 mL microcentrifuge tube</td><td>Thermo Fisher Scientific</td><td>3448</td><td></td></tr><tr><td>3-(N-morpholino)propanesulfonic acid</td><td>Millipore Sigma</td><td>M1254</td><td></td></tr><tr><td>3-(N-morpholino)propanesulfonic acid sodium salt</td><td>Millipore Sigma</td><td>M9381</td><td></td></tr><tr><td>Acclaim PepMap RSLC C18 Column</td><td>Thermo Scientific</td><td>164537</td><td>300 &amp;mu;m x 15 cm, C18, 2 &amp;mu;m, 100 A</td></tr><tr><td>Acetonitrile</td><td>Fisher Scientific</td><td>A998-1</td><td></td></tr><tr><td>Diethylpyrocarbonate</td><td>Millipore Sigma</td><td>D5758</td><td></td></tr><tr><td>HPLC-grade water</td><td>Fisher Scientific</td><td>W5-1</td><td></td></tr><tr><td>Imidazole</td><td>Millipore Sigma</td><td>I5513</td><td></td></tr><tr><td>Immobilized chymotrypsin</td><td>ProteoChem</td><td>g4105</td><td></td></tr><tr><td>Immobilized trypsin, TPCK Treated</td><td>Thermo Fisher Scientific</td><td>20230</td><td></td></tr><tr><td>Iodoacetamide</td><td>Millipore Sigma</td><td>I1149</td><td></td></tr><tr><td>Tris(2-carboxyethyl)phosphine</td><td>Millipore Sigma</td><td>C4706</td><td></td></tr></tbody></table>

covalent labeling with diethylpyrocarbonate for studying protein higher-order structure by mass spectrometry

Characterizing a protein&#39;s higher-order structure is essential for understanding its function. Mass spectrometry (MS) has emerged as a powerful tool for this purpose, especially for protein systems that are difficult to study by traditional methods. To study a protein&#39;s structure by MS, specific chemical reactions are performed in solution that encode a protein&#39;s structural information into its mass. One particularly effective approach is to use reagents that covalently modify solvent accessible amino acid side chains. These reactions lead to mass increases that can be localized with residue-level resolution when combined with proteolytic digestion and tandem mass spectrometry. Here, we describe the protocols associated with use of diethylpyrocarbonate (DEPC) as a covalent labeling reagent together with MS detection. DEPC is a highly electrophilic molecule capable of labeling up to 30% of the residues in the average protein, thereby providing excellent structural resolution. DEPC has been successfully used together with MS to obtain structural information for small single-domain proteins, such as &#946;2-microglobulin, to large multi-domain proteins, such as monoclonal antibodies.

Identifying DEPC modification sites and modification percentages 
Mass addition due to covalent labeling can be measured at the (a) intact protein and (b) peptide levels8,9. At the intact level, a distribution of protein species with different numbers of labels can be obtained from direct analysis or LC-MS of labeled protein samples. To obtain higher resolution structural information (i.e., site-specific labeling data), measurements must be performed at the peptide level. After the labeling and quenching steps, the labeled proteins are subjected to bottom-up proteomic analysis (i.e., disulfide reduction, alkylation, proteolytic digestion, and LC-MS/MS). Figure 3 shows how the DEPC labeled sites are identified and their modification levels are calculated for a DEPC CL-MS experiment on the protein &#946;-2-microglobulin (&#946;2m)21. MS/MS is used to sequence the labeled peptides and pinpoint their DEPC labeled sites, while modification percentages are calculated from their relative peak areas in extracted ion chromatograms (See Figure 3A).
Protein surface mapping using DEPC CL-MS 
Because of the relationship between protein topology and labeling rate, DEPC has been used to study changes in protein higher-order structure (HOS) and identify protein interaction sites8,9. An example is the effect of Cu(II) binding on the structure of &#946;2m. DEPC modification rate coefficients of peptide fragments from unbound &#946;2m and Cu(II)-bound &#946;2m (b2m-Cu) can be measured (Table 2) by varying DEPC concentrations and generating second-order kinetic plots14,24.&#160;The DEPC reactivity of residues in &#946;2m-Cu reveals significant labeling rate changes for His31, Ser33, and Thr4, while other sites, including different His residues, are statistically unchanged. These data are consistent with the fact that His31, but not other His residues in &#946;2m, binds Cu causing structural changes near His31 (i.e., Ser33) and near the N-terminus (i.e., Thr4) (Figure 4)14,26,27.
DEPC CL-MS for identifying changes in HOS 
DEPC labeling with MS detection is also a valuable tool for characterizing HOS changes to proteins, which has important implications for protein therapeutics, which are currently the fastest growing segment of the pharmaceutical market28. DEPC CL can identify specific protein regions that undergo structural changes upon thermal and oxidative stress18. After &#946;2m is exposed to heat stress, many residues that undergo significant decreases in labeling extents (N-terminus, Ser28, His31, Ser33, Ser55, Ser57, Lys58) are clustered on one side of the protein, suggesting that this region of the protein undergoes a conformational change or possibly mediates aggregation (Figure 5A). In addition to these residues, after the protein is exposed to oxidative stress, other residues with a decrease in labeling (Ser11, His13, Lys19, Lys41, Lys94) form a cluster on another face of the protein, indicating that the oxidation-induced conformational changes occur elsewhere (Figure 5B). Other work from our group has also shown that DEPC CL-MS can detect and identify sites of conformational changes in heat-stressed monoclonal antibody therapeutics20.
DEPC CL-MS to study protein-protein interactions 
Insight into protein-protein interaction sites and aggregation interfaces for amyloid-forming proteins can be obtained using DEPC labeling as well9. Decreases in the modification levels of residues upon oligomer formation can reveal the binding interfaces. DEPC CL-MS was used to characterize the pre-amyloid oligomers of &#946;2m, which is the protein that forms amyloids in dialysis-related amyloidosis29, after initiating amyloid formation with Cu(II)15,16,26. Comparing the DEPC reactivity of the &#946;2m monomer with the pre-amyloid &#946;2m dimer that is formed 2 h after adding Cu(II) shows that nine residues undergo decreases in labeling while six residues undergo no change or slight increases in labeling (Figure 6A). Upon mapping these changes on the &#946;2m, it is immediately apparent that the dimer interface involves the ABED &#946;-sheets of &#946;2m monomers (Figure 6B)15.
DEPC CL-MS to study protein-ligand binding 
Ligand binding to proteins leads to decreases in the solvent accessibility of residues that interact with the ligand, and DEPC-based CL-MS can be used to identify ligand-binding sites on proteins. For example, the binding sites of epigallocatechin-3-gallate (EGCG) on &#946;2m under amyloid-forming conditions can be identified by significant decreases in DEPC labeling at the residues buried by EGCG binding. In the presence of ECGC, Lys6 and Lys91 have lower DEPC modification percentages (Figure 7), and these residues are clustered in one region of the protein, indicating residue protection due to ligand binding. The N-terminus, Thr4, and His31, meanwhile, undergo increases in labeling extents (Figure 7), which are indicative of ECGC-induced structural changes and suggest that the Cu(II) binding sites on &#946;2m (N-terminus and His31)14,30,31 may be disrupted as a result of ECGC binding32. In addition, the binding sites of the other two small-molecule inhibitors of &#946;2m amyloid formation, rifamycin SV and doxycycline, have been identified using CL-MS19. In this study, however, results from DEPC labeling alone were not enough to map the binding sites with sufficient structural resolution. Results from three different CL reagents, namely DEPC, BD, and 1-ethyl-3-(3-(dimethylamino)propyl)carbodiimide - glycine ethyl ester (EDC/GEE) pair were necessary to better pinpoint the binding sites19.
Improving structural resolution and reducing label scrambling 
Even though DEPC labeling causes the formation of a covalent bond, label loss due to hydrolysis can occur, especially for Ser, Thr, and Tyr residues (Figure 8). Label loss can be minimized by decreasing the time between DEPC CL reaction and LC-MS analysis using short proteolytic digestions (e.g., 2 h digestion with immobilized enzymes) rather than overnight digestions. Fast digestions result in more modified residues being measured, increasing the amount of protein structural information. For example, a 2 h digestion with immobilized chymotrypsin consistently leads to the identification of more DEPC-modified residues in &#946;2m (Figure 9)17. With an overnight digestion about 75% of the Ser, Thr, Tyr, His, and Lys residues in &#946;2m are detected, but when the time between labeling and LC-MS is reduced by using a 2 h digestion, 95% of these residues in &#946;2m are detected. Most of the newly detected modified residues are Tyr and Thr residues that are prone to hydrolysis.
Over the course of our work with DEPC, we have noticed that in some proteins, Cys residues that from disulfide bonds are modified by DEPC, even though the disulfide bonds are intact during the DEPC labeling reactions. Such labeling of Cys residues occurs because free thiols can react with other modified residues in solution (Figure 10), leading to so-called &#34;label scrambling&#34; as the carbethoxy group is transferred to the Cys residue. Label scrambling decreases the modification levels at other residues and provides incorrect protein structural information. To avoid label scrambling, the free Cys thiols must be fully alkylated right after disulfide reduction33.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61983/61983fig01.jpg" /> 
Figure 1: Covalent labeling-mass spectrometry (CL-MS). A monofunctional reagent is used to modify solvent accessible amino acids and can be used to provide site-specific information about conformational changes or interaction interfaces in proteins (top vs. bottom image). The modified protein is proteolytically digested, and the resulting peptides are analyzed by liquid chromatography (LC) in conjunction with MS and tandem MS (MS/MS). Figure has been adapted from Limpikirati, P., Liu, T., Vachet, R. W. Covalent Labeling-Mass Spectrometry for Studying Protein Structure and Interactions. Methods&#160;144, 79-93 (2018). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61983/61983fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Volume Used (&#956;L)</td>
			<td>Concentration in Solution (&#956;M)</td>
		</tr>
		<tr>
			<td>Protein (100 &#956;M, MOPS)</td>
			<td>50</td>
			<td>50</td>
		</tr>
		<tr>
			<td>MOPS (10 mM, pH 7.4)</td>
			<td>48.8</td>
			<td>n/a</td>
		</tr>
		<tr>
			<td>DEPC (100 mM)</td>
			<td>0.2</td>
			<td>200 (=4x[protein])</td>
		</tr>
		<tr>
			<td>Imidazole (1 M)</td>
			<td>1</td>
			<td>10,000 (=50x[DEPC])</td>
		</tr>
		<tr>
			<td>Total Volume</td>
			<td>100</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 1. General DEPC labeling protocol.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61983/61983fig02.jpg" /> 
Figure 2. Example LC gradient for separation of peptides. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61983/61983fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61983/61983fig03.jpg" /> 
Figure 3 Illustration of how DEPC labeled sites are identified and their modification levels are calculated. After DEPC labeling and proteolytic digestion, (A) LC-MS analysis of the digested protein is performed. Peak areas of (B) unlabeled and (C) labeled peptides in a chromatogram are used to calculate the labeling percentage. During LC-MS, peptides are subjected to CID MS/MS. Tandem mass spectra of (D) unlabeled and (E) &#38; (F) labeled peptides obtained at specific retention times are used for peptide sequencing and identification of DEPC labeled sites. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61983/61983fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="0" cellpadding="0" cellspacing="0" style="width:700px;" width="698">
	<tbody>
		<tr>
			<td>Residue</td>
			<td>b2m</td>
			<td>b2m-Cu</td>
		</tr>
		<tr>
			<td>Thr4</td>
			<td>0.082 &#177; 0.004</td>
			<td>0.052 &#177; 0.009</td>
		</tr>
		<tr>
			<td>His13</td>
			<td>0.041 &#177; 0.003</td>
			<td>0.033 &#177; 0.005</td>
		</tr>
		<tr>
			<td>His31</td>
			<td>0.010 &#177; 0.001</td>
			<td>0.003 &#177; 0.001</td>
		</tr>
		<tr>
			<td>Ser33</td>
			<td>0.010 &#177; 0.002</td>
			<td>0.004 &#177; 0.001</td>
		</tr>
		<tr>
			<td>His51</td>
			<td>0.036 &#177; 0.003</td>
			<td>0.030 &#177; 0.004</td>
		</tr>
		<tr>
			<td>Ser88</td>
			<td>0.029 &#177; 0.007</td>
			<td>0.020 &#177; 0.006</td>
		</tr>
	</tbody>
</table>
Table 2 Site-specific DEPC modification rate coefficients (k, M-1s-1) for b2m in the absence and presence of Cu(II).
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61983/61983fig04.jpg" /> 
Figure 4 Using DEPC CL-MS to determine the effect of Cu(II) binding on the structure of &#946;2m: (A) Protein surface mapping of the residues with significant decreases in DEPC labeling rates (blue), indicating changes in their solvent accessibility upon Cu(II) binding, and those with no significant changes in labeling rate (magenta) (PDB accession code 1JNJ). (B) Example site-specific second-order kinetic plots of the reaction of &#946;2m with different concentrations of DEPC in the presence and absence of Cu(II). The labeling rate coefficient (k) can be obtained from the slope of the kinetic plot. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61983/61983fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61983/61983fig05.jpg" /> 
Figure 5 Covalent labeling results for stressed vs. native &#946;2m: (A) Heat stress - heating at 75 &#176;C for 24 h and (B) oxidative stress - with 3% H2O2 for 24 h. Significant changes in modification percentages are shown in blue (decrease in labeling) and red (increase in labeling) while residues with no significant labeling changes are shown in pale green (PDB accession code 1JNJ). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61983/61983fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61983/61983fig06.jpg" /> 
Figure 6&#160;Using DEPC CL-MS to determine the dimer interface for the pre-amyloid dimer of &#946;2m: (A) Summary of DEPC modification level changes for modified residues in the monomer (i.e., t = 0) and dimer 2 h after adding Cu(II). Residues with significant decreases in labeling extent are denoted by an asterisk (*). (B) Covalent labeling results mapped on the &#946;2m structure. Residues with decreased labeling are shown in blue and residues with no changes or slight increases in labeling are shown in red (PDB accession code 1LDS). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61983/61983fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61983/61983fig07.jpg" /> 
Figure 7 Covalent labeling results for EGCG-bound vs. unbound b2m. The DEPC modification percentages are shown for the displayed residues. Statistically significant differences in the covalent labeling percentage are denoted by an asterisk (*). Increases in labeling upon ECGC binding are shown in red while decreases are shown in blue. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61983/61983fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/61983/61983fig08.jpg" /> 
Figure 8&#160;DEPC covalent labeling reactions (nucleophilic acyl substitution) and label loss (hydrolysis of carbethoxylated residues) <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61983/61983fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/61983/61983fig09.jpg" /> 
Figure 9 Mapping of the measured modification sites on &#946;2m. (A) Labeled sites after conventional overnight digestion, and (B) labeled sites after a 2 h digestion with immobilized chymotrypsin. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61983/61983fig09large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" class="xfigimg" src="/files/ftp_upload/61983/61983fig10.jpg" /> 
Figure 10&#160;Hypothesized mechanism of DEPC label scrambling (Cys capture of carbethoxylated His) <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61983/61983fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Covalent Labeling with Diethylpyrocarbonate for Studying Protein Higher-Order Structure by Mass Spectrometry at JoVE.com

Covalent Labeling with Diethylpyrocarbonate for Studying Protein Higher-Order Structure by Mass Spectrometry

The experimental procedures for performing diethylpyrocarbonate-based covalent labeling with mass spectrometric detection are described. Diethylpyrocarbonate is simply mixed with the protein or protein complex of interest, leading to the modification of solvent accessible amino acid residues. The modified residues can be identified after proteolytic digestion and liquid chromatography/mass spectrometry analysis.

Characterizing a protein&#39;s higher-order structure is essential for understanding its function. Mass spectrometry (MS) has emerged as a powerful tool ...

covalent-labeling-with-diethylpyrocarbonate-for-studying-protein

Nanyang Technological University

Research

JoVE Journal

Chemistry

5.3K Views.  University of Massachusetts Amherst. The experimental procedures for performing diethylpyrocarbonate-based covalent labeling with mass spectrometric detection are described. Diethylpyrocarbonate is simply mixed with the protein or protein complex of interest, leading to the modification of solvent accessible amino acid residues. The modified residues can be identified after proteolytic digestion and liquid chromatography/mass spectrometry analysis.

Video: Covalent Labeling with Diethylpyrocarbonate for Studying Protein Higher-Order Structure by Mass Spectrometry

Formation of Ordered Biomolecular Structures by the Self-assembly of Short Peptides

In nature, complex functional structures are formed by the self-assembly of biomolecules under mild conditions. Understanding the forces that control self-assembly and mimicking this process in vitro will bring about major advances in the areas of materials science and nanotechnology. Among the available biological building blocks, peptides have several advantages as they present substantial diversity, their synthesis in large scale is straightforward, and they can easily be modified with biological and chemical entities1,2. Several classes of designed peptides such as cyclic peptides, amphiphile peptides and peptide-conjugates self-assemble into ordered structures in solution. Homoaromatic dipeptides, are a class of short self-assembled peptides that contain all the molecular information needed to form ordered structures such as nanotubes, spheres and fibrils3-8. A large variety of these peptides is commercially available.
This paper presents a procedure that leads to the formation of ordered structures by the self-assembly of homoaromatic peptides. The protocol requires only commercial reagents and basic laboratory equipment. In addition, the paper describes some of the methods available for the characterization of peptide-based assemblies. These methods include electron and atomic force microscopy and Fourier-Transform Infrared Spectroscopy (FT-IR). Moreover, the manuscript demonstrates the blending of peptides (coassembly) and the formation of a &#34;beads on a string&#34;-like structure by this process.9 The protocols presented here can be adapted to other classes of peptides or biological building blocks and can potentially lead to the discovery of new peptide-based structures and to better control of their assembly.

This paper describes the formation of highly ordered peptide-based structures by the spontaneous process of self-assembly. The method utilizes commercially available peptides and common lab equipment. This technique can be applied to a large variety of peptides and may lead to the discovery of new peptide-based assemblies.

In nature, complex functional structures are formed by the self-assembly of biomolecules under mild conditions. Understanding the forces that control ...

Time-resolved ElectroSpray Ionization Hydrogen-deuterium Exchange Mass Spectrometry for Studying Protein Structure and Dynamics

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. Hydrogen-Deuterium Exchange (HDX) measured at rapid time scales is uniquely suited to detect structures and hydrogen bonding networks that are briefly populated, allowing for the characterization of transient conformers in native ensembles. Coupling of HDX to mass spectrometry offers several key advantages, including high sensitivity, low sample consumption and no restriction on protein size. This technique has advanced greatly in the last several decades, including the ability to monitor HDX labeling times on the millisecond time scale. In addition, by incorporating the HDX workflow onto a microfluidic platform housing an acidic protease microreactor, we are able to localize dynamic properties at the peptide level. In this study, Time-Resolved ElectroSpray Ionization Mass Spectrometry (TRESI-MS) coupled to HDX was used to provide a detailed picture of residual structure in the tau protein, as well as the conformational shifts induced upon hyperphosphorylation.

Conformational flexibility plays a critical role in protein function. Herein, we describe the use of time-resolved electrospray ionization mass spectrometry coupled to hydrogen-deuterium exchange for probing the rapid structural changes that drive function in ordered and disordered proteins.

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. ...

Biochemistry

Mass Spectrometric Approaches to Study Protein Structure and Interactions in Lyophilized Powders

Amide hydrogen/deuterium exchange (ssHDX-MS) and side-chain photolytic labeling (ssPL-MS) followed by mass spectrometric analysis can be valuable for characterizing lyophilized formulations of protein therapeutics. Labeling followed by suitable proteolytic digestion allows the protein structure and interactions to be mapped with peptide-level resolution. Since the protein structural elements are stabilized by a network of chemical bonds from the main-chains and side-chains of amino acids, specific labeling of atoms in the amino acid residues provides insight into the structure and conformation of the protein. In contrast to routine methods used to study proteins in lyophilized solids (e.g., FTIR), ssHDX-MS and ssPL-MS provide quantitative and site-specific information. The extent of deuterium incorporation and kinetic parameters can be related to rapidly and slowly exchanging amide pools (Nfast, Nslow) and directly reflects the degree of protein folding and structure in lyophilized formulations. Stable photolytic labeling does not undergo back-exchange, an advantage over ssHDX-MS. Here, we provide detailed protocols for both ssHDX-MS and ssPL-MS, using myoglobin (Mb) as a model protein in lyophilized formulations containing either trehalose or sorbitol.

Here, we present detailed protocols for solid-state amide hydrogen/deuterium exchange mass spectrometry (ssHDX-MS) and solid-state photolytic labeling mass spectrometry (ssPL-MS) for proteins in solid powders. The methods provide high-resolution information on protein conformation and interactions in the amorphous solid-state, which may be useful in formulation design.

Amide hydrogen/deuterium exchange (ssHDX-MS) and side-chain photolytic labeling (ssPL-MS) followed by mass spectrometric analysis can be valuable for ...

Atomic Scale Structural Studies of Macromolecular Assemblies by Solid-state Nuclear Magnetic Resonance Spectroscopy

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the propagation of neurodegenerative disorders. Such assemblies consist in multiple protein subunits organized in a non-covalent way to form large macromolecular objects that can execute a variety of cellular functions or cause detrimental consequences. Atomic insights into the assembly mechanisms and the functioning of those macromolecular assemblies remain often scarce since their inherent insolubility and non-crystallinity often drastically reduces the quality of the data obtained from most techniques used in structural biology, such as X-ray crystallography and solution Nuclear Magnetic Resonance (NMR). We here present magic-angle spinning solid-state NMR spectroscopy (SSNMR) as a powerful method to investigate structures of macromolecular assemblies at atomic resolution. SSNMR can reveal atomic details on the assembled complex without size and solubility limitations. The protocol presented here describes the essential steps from the production of 13C/15N isotope-labeled macromolecular protein assemblies to the acquisition of standard SSNMR spectra and their analysis and interpretation. As an example, we show the pipeline of a SSNMR structural analysis of a filamentous protein assembly.

Structures of supramolecular protein assemblies at atomic resolution are of high relevance because of their crucial roles in a variety of biological phenomena. Herein, we present a protocol to perform high-resolution structural studies on insoluble and non-crystalline macromolecular protein assemblies by magic-angle spinning solid-state nuclear magnetic resonance spectroscopy (MAS SSNMR).

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the ...

Deep Proteome Profiling by Isobaric Labeling, Extensive Liquid Chromatography, Mass Spectrometry, and Software-assisted Quantification

Many exceptional advances have been made in mass spectrometry (MS)-based proteomics, with particular technical progress in liquid chromatography (LC) coupled to tandem mass spectrometry (LC-MS/MS) and isobaric labeling multiplexing capacity. Here, we introduce a deep-proteomics profiling protocol that combines 10-plex tandem mass tag (TMT) labeling with an extensive LC/LC-MS/MS platform, and post-MS computational interference correction to accurately quantitate whole proteomes. This protocol includes the following main steps: protein extraction and digestion, TMT labeling, 2-dimensional (2D) LC, high-resolution mass spectrometry, and computational data processing. Quality control steps are included for troubleshooting and evaluating experimental variation. More than 10,000 proteins in mammalian samples can be confidently quantitated with this protocol. This protocol can also be applied to the quantitation of post translational modifications with minor changes. This multiplexed, robust method provides a powerful tool for proteomic analysis in a variety of complex samples, including cell culture, animal tissues, and human clinical specimens.

We present a protocol to accurately quantitate proteins with isobaric labelling, extensive fractionation, bioinformatics tools, and quality control steps in combination with liquid chromatography interfaced to a high-resolution mass spectrometer.

Many exceptional advances have been made in mass spectrometry (MS)-based proteomics, with particular technical progress in liquid chromatography (LC) ...

Analyzing Protein Architectures and Protein-Ligand Complexes by Integrative Structural Mass Spectrometry

Proteins are an important class of biological macromolecules that play many key roles in cellular functions including gene expression, catalyzing metabolic reactions, DNA repair and replication. Therefore, a detailed understanding of these processes provides critical information on how cells function. Integrative structural MS methods offer structural and dynamical information on protein complex assembly, complex connectivity, subunit stoichiometry, protein oligomerization and ligand binding. Recent advances in integrative structural MS have allowed for the characterization of challenging biological systems including large DNA binding proteins and membrane proteins. This protocol describes how to integrate diverse MS data such as native MS and ion mobility-mass spectrometry (IM-MS) with molecular dynamics simulations to gain insights into a helicase-nuclease DNA repair protein complex. The resulting approach provides a framework for detailed studies of ligand binding to other protein complexes involved in important biological processes.

Mass spectrometry (MS) has emerged as an important tool for the investigation of structure and dynamics of macromolecular assemblies. Here, we integrate MS-based approaches to interrogate protein complex formation and ligand binding.

Proteins are an important class of biological macromolecules that play many key roles in cellular functions including gene expression, catalyzing ...

Resin-Assisted Capture Coupled with Isobaric Tandem Mass Tag Labeling for Multiplexed Quantification of Protein Thiol Oxidation

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed procedure of a quantitative redox proteomics method that utilizes resin-assisted capture (RAC) in combination with tandem mass tag (TMT) isobaric labeling and liquid chromatography-tandem mass spectrometry (LC-MS/MS) to allow multiplexed stochiometric quantification of oxidized protein thiols at the proteome level. The site-specific quantitative information on oxidized cysteine residues provides additional insight into the functional impacts of such modifications. 
The workflow is adaptable across many sample types, including cultured cells (e.g., mammalian, prokaryotic) and whole tissues (e.g., heart, lung, muscle), which are initially lysed/homogenized and with free thiols being alkylated to prevent artificial oxidation. The oxidized protein thiols are then reduced and captured by a thiol-affinity resin, which streamlines and simplifies the workflow steps by allowing the proceeding digestion, labeling, and washing procedures to be performed without additional transfer of proteins/peptides. Finally, the labeled peptides are eluted and analyzed by LC-MS/MS to reveal comprehensive stoichiometric changes related to thiol oxidation across the entire proteome. This method greatly improves the understanding of the role of redox-dependent regulation under physiological and pathophysiological states related to protein thiol oxidation.

Protein thiol oxidation has significant implications under normal physiological and pathophysiological conditions. We describe the details of a quantitative redox proteomics method, which utilizes resin-assisted capture, isobaric labeling, and mass spectrometry, enabling site-specific identification and quantification of reversibly oxidized cysteine residues of proteins.

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed ...

Capillary Electrophoresis-based Hydrogen/Deuterium Exchange for Conformational Characterization of Proteins with Top-down Mass Spectrometry

Resolving conformational heterogeneity of multiple protein states that coexist in solution remains one of the main obstacles in the characterization of protein therapeutics and the determination of the conformational transition pathways critical for biological functions, ranging from molecular recognition to enzymatic catalysis. Hydrogen/deuterium exchange (HDX) reaction coupled with top-down mass spectrometric (MS) analysis provides a means to characterize protein higher-order structures and dynamics in a conformer-specific manner. The conformational resolving power of this technique is highly dependent on the efficiencies of separating protein states at the intact protein level and minimizing the residual non-deuterated protic content during the HDX reactions. 
Here we describe a capillary electrophoresis (CE)-based variant of the HDX MS approach that aims to improve the conformational resolution. In this approach, proteins undergo HDX reactions while migrating through a deuterated background electrolyte solution (BGE) during the capillary electrophoretic separation. Different protein states or proteoforms that coexist in solution can be efficiently separated based on their differing charge-to-size ratios. The difference in electrophoretic mobility between proteins and protic solvent molecules minimizes the residual non-deuterated solvent, resulting in a nearly complete deuterating environment during the HDX process. The flow-through microvial CE-MS interface allows efficient electrospray ionization of the eluted protein species following a rapid mixing with the quenching and denaturing modifier solution at the outlet of the sprayer. The online top-down MS analysis measures the global deuteration level of the eluted intact protein species, and subsequently, the deuteration of their gas-phase fragments. This paper demonstrates this approach in differential HDX for systems, including the natural protein variants coexisting in milk.

Presented here is a protocol for a capillary electrophoresis-based hydrogen/deuterium exchange (HDX) approach coupled with top-down mass spectrometry. This approach characterizes the difference in higher-order structures between different protein species, including proteins in different states and different proteoforms, by conducting concurrent differential HDX and electrophoretic separation.

Resolving conformational heterogeneity of multiple protein states that coexist in solution remains one of the main obstacles in the characterization of ...

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 ...

Biology

Metabolic Pathway Confirmation and Discovery Through 13C-labeling of Proteinogenic Amino Acids

Microbes have complex metabolic pathways that can be investigated using biochemistry and functional genomics methods. One important technique to examine cell central metabolism and discover new enzymes is 13C-assisted metabolism analysis 1. This technique is based on isotopic labeling, whereby microbes are fed with a 13C labeled substrates. By tracing the atom transition paths between metabolites in the biochemical network, we can determine functional pathways and discover new enzymes. 
As a complementary method to transcriptomics and proteomics, approaches for isotopomer-assisted analysis of metabolic pathways contain three major steps 2. First, we grow cells with 13C labeled substrates. In this step, the composition of the medium and the selection of labeled substrates are two key factors. To avoid measurement noises from non-labeled carbon in nutrient supplements, a minimal medium with a sole carbon source is required. Further, the choice of a labeled substrate is based on how effectively it will elucidate the pathway being analyzed. Because novel enzymes often involve different reaction stereochemistry or intermediate products, in general, singly labeled carbon substrates are more informative for detection of novel pathways than uniformly labeled ones for detection of novel pathways3, 4. Second, we analyze amino acid labeling patterns using GC-MS. Amino acids are abundant in protein and thus can be obtained from biomass hydrolysis. Amino acids can be derivatized by N-(tert-butyldimethylsilyl)-N-methyltrifluoroacetamide (TBDMS) before GC separation. TBDMS derivatized amino acids can be fragmented by MS and result in different arrays of fragments. Based on the mass to charge (m/z) ratio of fragmented and unfragmented amino acids, we can deduce the possible labeled patterns of the central metabolites that are precursors of the amino acids. Third, we trace 13C carbon transitions in the proposed pathways and, based on the isotopomer data, confirm whether these pathways are active 2. Measurement of amino acids provides isotopic labeling information about eight crucial precursor metabolites in the central metabolism. These metabolic key nodes can reflect the functions of associated central pathways.
13C-assisted metabolism analysis via proteinogenic amino acids can be widely used for functional characterization of poorly-characterized microbial metabolism1. In this protocol, we will use Cyanothece 51142 as the model strain to demonstrate the use of labeled carbon substrates for discovering new enzymatic functions.

Metabolic Pathway Confirmation and Discovery Through 13C-labeling of Proteinogenic Amino Acids

13C-isotope labeling is a useful technique for determining the cell central metabolism for various types of microorganisms. After cells have been cultured with a specific labeled substrate, GC-MS measurement can reveal functional metabolic pathways based on unique labeling patterns in proteinogenic amino acids.

Microbes have complex metabolic pathways that can be investigated using biochemistry and functional genomics methods. One important technique to examine ...