We thank M. Boysen for comments on the manuscript. This project was funded by the Deutsche Forschungsgemeinschaft (SFB638 and MA 1278/4-1 to M.P.M., and Cluster of Excellence: CellNetworks EXC 81/1). M.P.M. is investigator of the Cluster of Excellence: CellNetworks.

1. Preparation of Buffers and Protein Samples
<ol>
	<li>Prepare H2O buffer. Use Hsp90 standard buffer (40 mM HEPES/KOH, pH 7.5, 50 mM KCl, 5 mM MgCl2, 10% glycerol) as H2O buffer. 
	Note: If the sample was dialyzed before analysis, use the dialysis buffer as H2O buffer. It is essential that the D2O buffer differs from the H2O buffer only in the hydrogen isotope. Volatile buffers like NH4CO3 or NH4-acetate or buffer components are not suitable!</li>
	<li>Prepare D2O buffer by lyophilization of Hsp90 standard buffer using a vacuum concentrator. After complete evaporation of H2O add pure D2O to the tube to reach the initial total volume (e.g. 1 ml buffer with 15% glycerol require the addition of 850 &#956;l D2O). Repeat the complete evaporation of the buffer and redissolve the buffer/salt components in D2O 2x.</li>
	<li>Prepare quench buffer (0.4 M KH2PO4/H3PO4 pH 2.2). 
	Note: To improve the efficiency of peptic digest of very stable proteins 4 M guanidine hydrochloride and 0.5 M Tris(2-carboxyethyl)phosphine (TCEP-HCl) can be added.</li>
	<li>Prepare 100% control sample (6 M guanidine hydrochloride, D2O). Add guanidine hydrochloride to an Hsp90 aliquot to reach a final concentration of 6 M. Completely evaporate H2O from the sample and add D2O to the tube to reach the initial total volume (e.g. 100 &#956;l sample with 10% glycerol require the addition of 90 &#956;l D2O). Repeat the complete evaporation of the buffer and redissolve the buffer/salt components in D2O. 
	Note: 20-100 pmol of sample are required for each injection. Prepare enough sample to have a 100% control for every day of HX-MS experiments.</li>
	<li>Prepare 50 pmol Hsp90 in 5 &#956;l Hsp90 standard buffer. 
	Note: An amount of 20-100 pmol of sample is required for each point of raw data. The volume of sample in the reaction is 1-5 &#956;l ideally. Adjust the concentration to fit these requirements. Any buffer can be used as long as it doesn&#39;t contain detergents or volatile components.</li>
</ol>
2. Preparation of Immobilized Pepsin on Aldehyde Activated Beads
<ol>
	<li>Dissolve 80 mg fresh pepsin in 2 ml 50 mM sodium citrate (pH 5).</li>
	<li>Dissolve 20 mg of sodium cyanoborohydride, handle with care (very toxic!) in 1 ml 2 M Na2SO4 and add to pepsin solution.</li>
	<li>Incubate mixture for 10 min at room temperature while gently agitating (e.g. overhead shaker).</li>
	<li>Add 600 mg beads with immobilized aldehyde groups to the mixture and incubate for 5-10 min at room temperature.</li>
	<li>Add 2.2 ml of 2 M Na2SO4 (pH 5) in 100 &#956;l aliquots every 3 min over one hour to slowly salt out the pepsin. Gently mix the sample between additions in an overhead shaker at room temperature.</li>
	<li>Incubate the pepsin beads at 4 &#176;C for 14-16 hr/overnight in an overhead shaker.</li>
	<li>Quench the reaction by adding of 1 ml 1 M ethanolamine and incubation at room temperature for 2 hr.</li>
	<li>Spin down the beads in a 50 ml falcon tube at 500 rpm, discard the supernatant and resuspend the beads in 0.1% formic acid. Repeat this step 2x. After the last centrifugation step, discard supernatant and estimate volume of beads. Add an equivalent volume of 0.1% formic acid and store at 4 &#176;C.</li>
</ol>
3. Preparation of Columns for Amide Hydrogen-exchange
<ol>
	<li>Use guard columns with an inner diameter of 1 mm for trapping columns and with 2 mm for pepsin columns.</li>
	<li>Unscrew one side of the guard column and remove the filter. Tightly screw packing funnel onto the open end of the column. Use a 1/16&#160;inch adapter and tubing to attach an empty syringe (5 ml) to the bottom outlet of the column. Make sure to fix it gas-tight to the guard column.</li>
	<li>Apply a few drops of slurry bead material on top of the funnel. Pull the plunger of the syringe to suck the slurry through the funnel into the guard column. Apply more slurry bead material onto the funnel and continue the procedure until the guard column is completely filled with bead material. Remove the funnel and place filter and filter ring onto the open end. Tightly screw the column cap onto the guard column and remove the syringe from the other side. Close both ends of the guard columns with plugs to avoid drying out of column material.</li>
</ol>
4. Setting up the System for Hydrogen Exchange Mass Spectrometry (HX-MS)
<ol>
	<li>Connect the trap column with the HPLC system (Figure 1). Equilibrate the column by setting the flow rate of pump A to 0.4 ml/min with 0.1% formic acid as solvent. Do not connect the pepsin column nor the analytical column yet.</li>
	<li>Calibrate the mass spectrometer and connect the outlet of the HPLC to the source of the mass spectrometer.</li>
</ol>
5. Determination of the Dynamic Range of Exchange
<ol>
	<li>Prepare ultrapure solvent A (0.1% formic acid in water) and ultrapure solvent B (0.1% formic acid in acetonitrile); ready mixed solvents are commercially available. Purge the HPLC pumps. Set up the chromatography and mass spectrometry methods in the control software by choosing a program with a step gradient after a short desalting step. For full-length Hsp90 use a 1-2 min desalting step followed by switching the 6-port valve from DESALT/LOAD to ELUTE and a step gradient from 90% solvent A/10% solvent B to 5% solvent A/95% solvent B. Before injection set the injection valve to LOAD and the 6-port valve to DESALT/LOADING position. 
	Note: Do not use a pepsin or analytical column during this experiment.</li>
	<li>Prepare 100-200 pmol of Hsp90 in 1-10 &#956;l H2O buffer and incubate for 10 min at 30 &#176;C. Add temperature adjusted D2O buffer to bring the sample volume up to 100 &#956;l and incubate exactly for a defined period of time (e.g. 10 sec, 100 sec, 1,000 sec). Add 100 &#956;l quench buffer and mix by pipetting up and down. Inject the 200 &#956;l sample into the injection port of the injection valve with a Hamilton syringe. Start the chromatography program and switch the injection valve into INJECT position. After 2 min switch the 6-port valve from DESALT/LOADING to ELUTE. Repeat this for at least three time-points.</li>
	<li>Repeat step 5.2 but add 2-3x excess of Sti1 to the Hsp90 sample prior to incubating for 10 min at 30 &#176;C.</li>
	<li>Determine the full-length protein masses by deconvolution of spectra in the mass spectrometry software. Calculate the number of incorporated deuterons by comparing the molecular weight of full-length Hsp90 with the observed mass after each run (e.g. 10 sec, 100 sec, 1,000 sec).</li>
	<li>Plot the incorporated deuterons for Hsp90 in absence and presence of Sti1 (y-axis) versus time (x-axis). Determine the time-point in the dynamic range where the difference between both curves is maximal. Use this value for the incubation time in D2O when identifying Hsp90-Sti1 interface and dynamics on peptide level.</li>
</ol>
6. Determination of Peptic Peptides Using MS/MS Spectra
<ol>
	<li>Connect the pepsin column and analytical column to the system.</li>
	<li>Set up the parameters for the chromatography and mass spectrometry in the control software by choosing gradient type and mass spectrometry method. Choose a long gradient (e.g. more than 90 min) to ensure good chromatographic resolution. Enable MS/MS spectra on the mass spectrometer. 
	Note: Good resolution on the HPLC and high mass accuracy have highest priority in this step.</li>
	<li>Prepare 100-200 pmol of Hsp90 in 100 &#956;l H2O buffer. Add 100 &#956;l quench buffer and mix by pipetting up and down. Inject the 200 &#956;l sample into the injection port of the injection valve with a Hamilton syringe. Start the chromatography program and switch the injection valve into INJECT position. After 2 min switch the 6-port valve from DESALT/LOADING to ELUTE position. 
	Note: If more than one protein is to be analyzed in HX-MS (i.e. protein-protein interaction interfaces) the peptides for each protein have to be determined individually.</li>
	<li>Identify peptic peptides of Hsp90 by searching a database (i.e. Mascot) for the resulting peptides. 
	Note: It is possible to use a custom database as the aim is to determine as many peptic peptides as possible and analysis is done with purified protein. Sample purity will have to be taken into account.</li>
	<li>Repeat this step without MS/MS and with the gradient that will be used for the actual HX experiment. For Hsp90 and Sti1 use a 10 min gradient from 90% solvent A/10% solvent B to 45% solvent A/55% solvent B. 
	Note: Gradients are normally between 5-15 min and highly depend on sample complexity and HX system specifications.</li>
	<li>Determine the retention times of the peptic peptides identified in 6.4 in the gradient used in 6.5 and create a list comprising 1.) peptide sequence 2.) peptide charge state and 3.) retention time. This will be used to identify each peptide after HX experiments. 
	Note: Be aware that peptides with small m/z differences, identical charge state and identical retention time can be a source of ambiguity.</li>
</ol>
7. Identification of Protein-protein Interaction Interfaces
<ol>
	<li>Set up the gradient and mass spectrometry method in the control software. Use a 10 min linear gradient from 90% solvent A/10% solvent B to 45% solvent A/55% solvent B. Load a mass spectrometry method that is optimized for detection of masses between 300-1,500 m/z, although most of the peptides will be below 1,000 m/z. Set the injection valve into LOAD position, the 6-port valve in LOADING/DESALTING Position. Set the flow rate of the loading pump to 0.4 ml/min. 
	Note: Length and type of gradient are sample dependent and may have to be optimized. Longer gradients improve the resolution of the chromatography but decrease the incorporation of deuterons into proteins due to back-exchange. The chosen methods need to be the same for all experiments to be compared.</li>
	<li>For the unexchanged reference prepare 20-100 pmol of Hsp90 in 100 &#956;l H2O buffer. Add 100 &#956;l ice-cold quench buffer, pipette up and down twice and inject sample into the injection valve of the HPLC. Immediately start the chromatography program and switch the injection valve into INJECT position. After 2 min switch the 6-port valve from DESALT/LOADING to ELUTE position. Do this for Hsp90 and Sti1 individually and for the mixture of proteins.</li>
	<li>Prepare 20-100 pmol of Hsp90 in a volume of 1-5 &#956;l. Add temperature adjusted D2O buffer to bring the sample volume up to 100 &#956;l and incubate exactly for a defined period of time (e.g. 30 sec; for conformational dynamics see Note). Add 100 &#956;l of ice-cold quench buffer, pipette up and down twice and quickly inject the 200 &#956;l into the injection valve of the HPLC. Immediately start the chromatography program and switch the injection valve into INJECT position. After 2 min switch the 6-port valve from DESALT/LOADING to ELUTE position. Do this for each protein individually, to determine the deuteron incorporation into each peptide in absence of the interacting protein. 
	Note: When studying the dynamics of conformational changes, repeat the experiment with different incubation times of Hsp90 in D2O buffer. Try to cover a wide timescale by choosing incubation times logarithmically (e.g. 10 sec, 30 sec, 100 sec, 300 sec, 1,000 sec, etc.). D2O buffer and sample buffer must be exactly identical except for the hydrogen isotope.</li>
	<li>Prepare an equal amount of 100% control sample (20-100 pmol) and add D2O buffer to bring the sample volume up to 100 &#956;l. Add 100 &#956;l of ice-cold quench buffer, pipette up and down twice and quickly inject the 200 &#956;l into the injection valve of the HPLC. Immediately start the chromatography program and switch the injection valve into INJECT position. After 2 min switch the 6-port valve from DESALT/LOADING to ELUTE position.</li>
	<li>To determine the interaction surface mix Hsp90 with an at least 2-fold excess of Sti1 to shift the equilibrium to the bound state (see Note) and incubate at the desired temperature until complex formation is at equilibrium. Add temperature adjusted D2O buffer to bring the sample volume up to 100 &#956;l and incubate exactly for a defined period of time (e.g. 30 sec). Add 100 &#956;l of ice-cold quench buffer, pipette up and down twice and quickly inject into the injection valve of the HPLC. After 2 min switch the 6-port valve from DESALT/LOADING to ELUTE position. Repeat this experiment with 20-100 pmol of Sti1 and excess of Hsp90. 
	Note: The absolute concentrations depend on the dissociation equilibrium constant of the interaction. Ideally, after dilution into D2O the concentration of the protein added in excess should be at least 10x&#160;the KD (corresponding to &#62;85% of the lower concentrated protein bound).</li>
	<li>Analyze the acquired data with suitable software and use the retention times determined in step&#160;6.5 to find each peptide in the analysis. Calculate the centroid of the isotope distribution for the unexchanged protein (step&#160;7.2) and for the HX experiments (step&#160;7.3). 
	Note: Although the isotope patterns of exchanged peptides look different from their unexchanged counterparts, both the knowledge about charge state of the peptide and the high mass accuracy of the mass spectrometer allows for easy determination of the centroids.</li>
	<li>Compare incorporation of deuterons of target protein alone and with excess of binding partner. This can be done automatically with commercial software or manually with a spreadsheet program. The 100% control sample values denote the maximal exchange for each peptide and can be used to determine the amount of D2O label lost during the experiment due to back exchange.</li>
</ol>

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D., Konermann, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Conformational+dynamics+of+a+membrane+transport+protein+probed+by+h/d+exchange+and+covalent+labeling:+the+glycerol+facilitator.">Conformational dynamics of a membrane transport protein probed by h/d exchange and covalent labeling: the glycerol facilitator.</a> J. Mol. Biol. 416 (3), 400-413 (2012).</li><li>Hebling, C. M., Morgan, C. R., Stafford, D. W., Jorgenson, J. W., Rand, K. D., Engen, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Conformational+analysis+of+membrane+proteins+in+phospholipid+bilayer+nanodiscs+by+hydrogen+exchange+mass+spectrometry.">Conformational analysis of membrane proteins in phospholipid bilayer nanodiscs by hydrogen exchange mass spectrometry.</a> Anal chem. 82 (13), 5415-5419 (2010).</li><li>Abzalimov, R. R., Kaplan, D. A., Easterling, M. L., Kaltashov, I. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+conformations+can+be+probed+in+top-down+HDX+MS+experiments+utilizing+electron+transfer+dissociation+of+protein+ions+without+hydrogen+scrambling.">Protein conformations can be probed in top-down HDX MS experiments utilizing electron transfer dissociation of protein ions without hydrogen scrambling.</a> J. Am. Soc. Mass Spectrom. 20 (8), 1514-1517 (2009).</li></ol>

We have nothing to disclose.

Binding of an interaction partner to a protein inevitably causes changes in solvent accessibility on the binding site. Additionally, many proteins undergo dynamic conformational changes upon binding, which affect other regions than the actual binding interface. HX-MS is a robust method to monitor these changes and is even capable of revealing conformational changes in proteins on timescales that other methods cannot cover.
To successfully perform HX-MS three points are critical: 1) an optimal ...

The number of crystal structures of proteins and protein complexes increased rapidly in recent years. They present invaluable snapshots of the structural organization of these proteins and provide a basis for structure-function analysis. However, the dynamics of proteins and the conformational changes, which are essential for their functions, are rarely revealed by X-ray crystallography. Cryo-electronmicroscopy, on the other hand, is able to capture protein and protein complexes in different conformations but generally cannot resolve conformational changes down to secondary structure level1. Conformational dynamics of proteins in solution at atomic details can only be resolved by NMR, but this method is still restricted to proteins of relatively small sizes (generally &#8804; 30 kDa) and needs high concentrations of proteins (&#8805; 100 &#956;M), which hampers experiments with oligomerization or aggregation prone proteins2. One method that is able to bridge between high-resolution X-ray crystallography and cryo-electronmicroscopy and which is not limited by protein size or concentration is amide hydrogen-1H/2H-exchange (HX) in combination with mass spectrometry (MS). In recent years this method has developed to a valuable analytical tool for the analysis of protein dynamics, protein folding, protein stability and conformational changes3-5. The molecular basis of this method is the labile nature of backbone amide hydrogens in proteins, which will exchange with deuterium atoms when the protein is placed in a D2O solution. The subsequent increase in protein mass over time is measured with high-resolution MS.
In short unstructured peptides HX only depends on temperature, catalyst concentration (OH-, H3O+ i.e. pH, see Figure 3) and amino acid side chains of adjacent residues due to inductive, catalytic and steric effects. These effects on the intrinsic chemical exchange rate kch have been elegantly quantified by Bai et al.6 and a program is available (courtesy Z. Zhang), which calculates kch for each amino acid within a polypeptide dependent on pH and temperature. At neutral pH and ambient temperatures kch is in the order of 101-103&#160;sec-1. In folded proteins HX can be 2-9 orders of magnitude slower mainly due to hydrogen bonding in the secondary structure and to a minor degree due to restricted access of hydrated OH- ions to the interior of a tightly folded protein. HX in native proteins therefore implicates&#160;partial or global unfolding, chemical exchange and refolding to the native state according to equation (1) and the observed exchange rates kobs depend on the opening rate kop, the closing rate kcl and the intrinsic chemical exchange rate kch according to equation (2).
<img alt="Equations 1-2" src="/files/ftp_upload/50839/50839eq1-2.jpg" /> 
Under native state conditions kop is much smaller than kch and can be neglected in the denominator. There are two extreme exchange regimes called EX1 and EX2. If the kcl is much smaller than kch (EX1) the observed rate is practically equal to the opening rate and HX allows immediate observation of the unfolding of a structural element. Such an exchange regime, where all amide protons exchange at once upon opening of the structural element, is readily observable in MS by a bimodal distribution of the isotope peaks7. If kcl is much greater than kch (EX2) the observed rate is proportional to kch whereby the proportionality constant is equal to the folding-unfolding equilibriums constant Ku = kop/kcl. Under these conditions, many opening and closing events are necessary before all amide protons exchange for deuterons, leading to a gradual increase in average mass while the isotopic distribution remains roughly the same. The EX2 regime allows the determination of the free energy of unfolding &#916;Gu and therefore the stability of a structural element. Under native state condition the EX2 regime is most common. Increase of pH and addition of chaotropic agents can shift the exchange mechanism to EX1. Therefore, HX-MS can be used to explore thermodynamic as well as kinetic parameters of protein folding and conformational changes.
As mentioned above HX is intrinsically pH and temperature dependent and the exchange half-life of a completely solvent exposed proton of the backbone amide group is between 5-400 msec at physiological pH (pH 7.6) and 30 &#176;C, but 10 min to &#62;15 hr with an average of &#62;2 hr at pH 2.9 and 0 &#176;C (except for the proton of the first backbone amide bond of a polypeptide, which exchanges with a half-life of ca. 1-2 min). Under such slow exchanging conditions it is possible to digest the sample using proteases (e.g. pepsin) that are active under these conditions, with out losing all the information contained in the incorporated deuterons. Since the introduction of peptic digestion under slow exchanging conditions, not only the overall HX kinetics of full-length proteins can be analyzed but HX can be localized to specific regions8,9. Spatial resolution is currently limited to the size of the peptic fragments generated, which is in general between 10-30 residues. However, overlapping fragments created due to the nonspecific nature of cleavage by pepsin could lead to an increase in spatial resolution. In addition, several other proteases were found to be active under quench conditions, however, much less efficient than pepsin10. Further increase of spatial resolution can be reached by fragmentation of peptides in the gas phase by methods that preserved the deuteration pattern such as electron capture dissociation (ECD), electron transfer dissociation (ETD) and infrared multiphoton dissociation (IRMPD)11-13. These techniques prevent the loss of spatial resolution due to intramolecular proton migration (&#34;scrambling&#34;), which is observed by collision-induced dissociation (CID) the most commonly used fragmentation technique. However, these methods require optimization for every individual peptide and is thus still quite challenging.
HX-MS has been used to analyze protein-ligand and protein-protein interactions including viral capsid assembly14-17. Protein unfolding and refolding as well as temperature induced conformational changes were investigated7,18,19. Phosphorylation and single amino acid mutation-related conformational changes16,20 and nucleotide-induced changes were analyzed21,22. Therefore, this method seems ideally suitable to analyze assembly and dynamics of molecular machines. One attractive candidate, whose mechanism is of great general interest, is the Hsp90 chaperone complex.

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		<tr>
			<td>Reagent/Equipment</td>
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			<td>Catalogue number</td>
			<td>Comments (optional)</td>
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			<td>maXis QTOF</td>
			<td>Bruker</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>nanoAcquity UPLC</td>
			<td>Waters Corp.</td>
			<td></td>
			<td></td>
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			<td>Shimadzu 10AD-VP</td>
			<td>Shimadzu</td>
			<td></td>
			<td></td>
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		<tr>
			<td>6-port Valve EPC6W with microelectric actuator</td>
			<td>Valco</td>
			<td>#EPC6W</td>
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		</tr>
		<tr>
			<td>Injection valve (manual)</td>
			<td>Rheodyne</td>
			<td>#7725</td>
			<td></td>
		</tr>
		<tr>
			<td>Poros AL20 media</td>
			<td>Applied Biosystems</td>
			<td>#1-6029-06</td>
			<td></td>
		</tr>
		<tr>
			<td>Poros R2</td>
			<td>Applied Biosystems</td>
			<td>#1-1118-02</td>
			<td></td>
		</tr>
		<tr>
			<td>Pepsin</td>
			<td>Sigma</td>
			<td>#P6887</td>
			<td>use fresh pepsin</td>
		</tr>
		<tr>
			<td>Microbore (1 mm)</td>
			<td>IDEX</td>
			<td>#C-128</td>
			<td></td>
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			<td>Microbore (2 mm)</td>
			<td>IDEX</td>
			<td>#C-130B</td>
			<td></td>
		</tr>
		<tr>
			<td>Acquity UPLC BEH C8 Column</td>
			<td>Waters Corp.</td>
			<td>#186002876</td>
			<td></td>
		</tr>
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			<td>Thermomixer</td>
			<td>Eppendorf</td>
			<td>#5355000.011</td>
			<td></td>
		</tr>
		<tr>
			<td>Tubing (various diameters)</td>
			<td>IDEX</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fittings</td>
			<td>IDEX</td>
			<td>#PK-110 with PK-100</td>
			<td></td>
		</tr>
	</tbody>
</table>

analyzing protein dynamics using hydrogen exchange mass spectrometry

All cellular processes depend on the functionality of proteins. Although the functionality of a given protein is the direct consequence of its unique amino acid sequence, it is only realized by the folding of the polypeptide chain into a single defined three-dimensional arrangement or more commonly into an ensemble of interconverting conformations. Investigating the connection between protein conformation and its function is therefore essential for a complete understanding of how proteins are able to fulfill their great variety of tasks. One possibility to study conformational changes a protein undergoes while progressing through its functional cycle is hydrogen-1H/2H-exchange in combination with high-resolution mass spectrometry (HX-MS). HX-MS is a versatile and robust method that adds a new dimension to structural information obtained by e.g. crystallography. It is used to study protein folding and unfolding, binding of small molecule ligands, protein-protein interactions, conformational changes linked to enzyme catalysis, and allostery. In addition, HX-MS is often used when the amount of protein is very limited or crystallization of the protein is not feasible. Here we provide a general protocol for studying protein dynamics with HX-MS and describe as an example how to reveal the interaction interface of two proteins in a complex. &#160;&#160;

Hsp90 is a molecular chaperone in yeast and member of the Hsp90 chaperone family. By going through a complex ATPase cycle it assists late folding steps of many protein clients. Efficient folding requires transfer of clients from Hsp70 and interaction of the co-chaperone Sti1/Hop. Sti1 directly binds to Hsp90 and facilitates client binding by inhibition of Hsp90&#39;s ATPase activity. Interaction of Hsp90 with Sti1 was recently studied using HX-MS23. Here we present representative results of the underlying expe...

Watch this Scientific Journal Video about Analyzing Protein Dynamics Using Hydrogen Exchange Mass Spectrometry at JoVE.com

Analyzing Protein Dynamics Using Hydrogen Exchange Mass Spectrometry

Protein conformation and dynamics are key to understanding the relationship between protein structure and function. Hydrogen exchange coupled with high-resolution mass spectrometry is a versatile method to study the conformational dynamics of proteins as well as characterizing protein-ligand and protein-protein interactions, including contact interfaces and allosteric effects.

All cellular processes depend on the functionality of proteins. Although the functionality of a given protein is the direct consequence of its unique ...

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18.3K Views.  University of Heidelberg. Protein conformation and dynamics are key to understanding the relationship between protein structure and function. Hydrogen exchange coupled with high-resolution mass spectrometry is a versatile method to study the conformational dynamics of proteins as well as characterizing protein-ligand and protein-protein interactions, including contact interfaces and allosteric effects.

Video: Analyzing Protein Dynamics Using Hydrogen Exchange Mass Spectrometry

Real-time Breath Analysis by Using Secondary Nanoelectrospray Ionization Coupled to High Resolution Mass Spectrometry

Exhaled volatile organic compounds (VOCs) have aroused considerable interest, since they can serve as biomarkers for disease diagnosis and environmental exposure in a non-invasive manner. In this work, we present a protocol to characterize the exhaled VOCs in real time by using secondary nanoelectrospray ionization coupled to high resolution mass spectrometry (Sec-nanoESI-HRMS). The homemade Sec-nanoESI source was readily set up based on a commercial nanoESI source. Hundreds of peaks were observed in the background-subtracted mass spectra of exhaled breath, and the mass accuracy values are -4.0-13.5 ppm and -20.3-1.3 ppm in the positive and negative ion detection modes, respectively. The peaks were assigned with accurate elemental composition according to the accurate mass and isotopic pattern. Less than 30 s is used for one exhalation measurement, and it takes approximately 7 min for six replicated measurements.

A protocol for characterizing chemical composition of exhaled breath in real time by using secondary nanoelectrospray ionization coupled to high resolution mass spectrometry is demonstrated.

Exhaled volatile organic compounds (VOCs) have aroused considerable interest, since they can serve as biomarkers for disease diagnosis and environmental ...

Synthesis and Mass Spectrometry Analysis of Oligo-peptoids

Peptoids are sequence-controlled peptide-mimicking oligomers consisting of N-alkylated glycine units. Among many potential applications, peptoids have been thought of as a type of molecular information storage. Mass spectrometry analysis has been considered the method of choice for sequencing peptoids. Peptoids can be synthesized via solid phase chemistry using a repeating two-step reaction cycle. Here we present a method to manually synthesize oligo-peptoids and to analyze the sequence of the peptoids using tandem mass spectrometry (MS/MS) techniques. The sample peptoid is a nonamer consisting of alternating N-(2-methyloxyethyl)glycine (Nme) and N-(2-phenylethyl)glycine (Npe), as well as an N-(2-aminoethyl)glycine (Nae) at the N-terminus. The sequence formula of the peptoid is Ac-Nae-(Npe-Nme)4-NH2, where Ac is the acetyl group. The synthesis takes place in a commercially available solid-phase reaction vessel. The rink amide resin is used as the solid support to yield the peptoid with an amide group at the C-terminus. The resulting peptoid product is subjected to sequence analysis using a triple-quadrupole mass spectrometer coupled to an electrospray ionization source. The MS/MS measurement produces a spectrum of fragment ions resulting from the dissociation of charged peptoid. The fragment ions are sorted out based on the values of their mass-to-charge ratio (m/z). The m/z values of the fragment ions are compared against the nominal masses of theoretically predicted fragment ions, according to the scheme of peptoid fragmentation. The analysis generates a fragmentation pattern of the charged peptoid. The fragmentation pattern is correlated to the monomer sequence of the neutral peptoid. In this regard, MS analysis reads out the sequence information of the peptoids.

A protocol is described for the manual synthesis of oligo-peptoids followed by sequence analysis by mass spectrometry.

Peptoids are sequence-controlled peptide-mimicking oligomers consisting of N-alkylated glycine units. Among many potential applications, peptoids have ...

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 ...

Large-scale Top-down Proteomics Using Capillary Zone Electrophoresis Tandem Mass Spectrometry

Capillary zone electrophoresis-electrospray ionization-tandem mass spectrometry (CZE-ESI-MS/MS) has been recognized as a useful tool for top-down proteomics that aims to characterize proteoforms in complex proteomes. However, the application of CZE-MS/MS for large-scale top-down proteomics has been impeded by the low sample-loading capacity and narrow separation window of CZE. Here, a protocol is described using CZE-MS/MS with a microliter-scale sample-loading volume and a 90-min separation window for large-scale top-down proteomics. The CZE-MS/MS platform is based on a linear polyacrylamide (LPA)-coated separation capillary with extremely low electroosmotic flow, a dynamic pH-junction-based online sample concentration method with a high efficiency for protein stacking, an electro-kinetically pumped sheath flow CE-MS interface with extremely high sensitivity, and an ion trap mass spectrometer with high mass resolution and scan speed. The platform can be used for the high-resolution characterization of simple intact protein samples and the large-scale characterization of proteoforms in various complex proteomes. As an example, a highly efficient separation of a standard protein mixture and a highly sensitive detection of many impurities using the platform is demonstrated. As another example, this platform can produce over 500 proteoform and 190 protein identifications from an Escherichia coli proteome in a single CZE-MS/MS run.

A detailed protocol is described for the separation, identification, and characterization of proteoforms in protein samples using capillary zone electrophoresis-electrospray ionization-tandem mass spectrometry (CZE-ESI-MS/MS). The protocol can be used for the high-resolution characterization of proteoforms in simple protein samples and the large-scale identification of proteoforms in complex proteome samples.

Capillary zone electrophoresis-electrospray ionization-tandem mass spectrometry (CZE-ESI-MS/MS) has been recognized as a useful tool for top-down ...

High-throughput and Comprehensive Drug Surveillance Using Multisegment Injection-Capillary Electrophoresis-Mass Spectrometry

New analytical methods are urgently needed to enable high-throughput, yet comprehensive drug screening, given an alarming opioid and prescription drug crisis in public health. Conventional urine drug testing based on a two-tier immunoassay screen followed by a gas chromatography-tandem mass spectrometry (GC-MS/MS) or liquid chromatography-tandem mass spectrometry (LC-MS/MS) method are expensive and prone to bias while being limited to targeted panels of known drugs of abuse (DoA). Herein, we outline an improved method for drug surveillance that allows for the resolution and detection of an expanded panel of DoA and their metabolites when using multisegment injection-capillary electrophoresis-mass spectrometry (MSI-CE-MS). Multiplexed separations of ten urine samples with a quality control by CE (&lt; 3 min/sample) in conjunction with full-scan data acquisition using a time-of-flight mass spectrometer (TOF-MS) under positive ion mode detection allows for the identification and quantification of DoA above recommended cut-off levels. An excellent resolution of drug isomers and isobars, including background interferences, are achieved when using MSI-CE-MS with an electrokinetic spacer between sample segments, where accurate mass/molecular formula together with the comigration of a matching deuterated internal standard and the detection of one or more bio-transformed metabolites facilitate DoA identification over a wider detection window. Additionally, urine samples can be analyzed directly without enzyme deconjugation for the rapid screening without complicated sample workup. MSI-CE-MS enables the surveillance of a broad spectrum of DoA that is required for the treatment monitoring of high-risk patients, including confirming prescribed drug adherence, revealing illicit drug use/substitution, and evaluating optimal dosage regimes as required for new advances in precision medicine.

Here we describe a high-throughput method for comprehensive drug surveillance that allows for the improved resolution and detection of large panels of drugs of abuse and their metabolites, with quality control based on multisegment injection-capillary electrophoresis-mass spectrometry.

New analytical methods are urgently needed to enable high-throughput, yet comprehensive drug screening, given an alarming opioid and prescription drug ...

3D Depth Profile Reconstruction of Segregated Impurities Using Secondary Ion Mass Spectrometry

The presented protocol combines excellent detection limits (1 ppm to 1 ppb) using secondary ion mass spectrometry (SIMS) with reasonable spatial resolution (~1 &#181;m). Furthermore, it describes how to obtain realistic three-dimensional (3D) distributions of segregated impurities/dopants in solid state materials. Direct 3D depth profile reconstruction is often difficult to achieve due to SIMS-related measurement artifacts. Presented here is a method to identify and solve this challenge. Three major issues are discussed, including the i) nonuniformity of the detector being compensated by flat-field correction; ii) vacuum background contribution (parasitic oxygen counts from residual gases present in the analysis chamber) being estimated and subtracted; and iii)&#160;performance of all steps within a stable timespan of the primary ion source. Wet chemical etching is used to reveal the position and types of dislocation in a material, then the SIMS result is superimposed on images obtained via scanning electron microscopy (SEM). Thus, the position of agglomerated impurities can be related to the position of certain defects. The method is fast and does not require sophisticated sample preparation stage; however, it requires a high-quality, stable ion source, and the entire measurement must be performed quickly to avoid deterioration of the primary beam parameters.

The presented method describes how to identify and solve measurement artifacts related to secondary ion mass spectrometry as well as obtain realistic 3D distributions of impurities/dopants in solid state materials.

The presented protocol combines excellent detection limits (1 ppm to 1 ppb) using secondary ion mass spectrometry (SIMS) with reasonable spatial ...

Capillary Electrophoresis Mass Spectrometry Approaches for Characterization of the Protein and Metabolite Corona Acquired by Nanomaterials

The adsorption of biomolecules from surrounding biological matrices to the surface of nanomaterials (NMs) to form the corona has been of interest for the past decade. Interest in the bio-nano interface arises from the fact that the biomolecular corona confers a biological identity to NMs and thus causes the body to identify them as &#8220;self&#8221;. For example, previous studies have demonstrated that the proteins in the corona are capable of interacting with membrane receptors to influence cellular uptake and established that the corona is responsible for cellular trafficking of NMs and their eventual toxicity. To date, most research has focused upon the protein corona and overlooked the possible impacts of the metabolites included in the corona or synergistic effects between components in the complete biomolecular corona. As such, this work demonstrates methodologies to characterize both the protein and metabolite components of the biomolecular corona using bottom-up proteomics and metabolomics approaches in parallel. This includes an on-particle digest of the protein corona with a surfactant used to increase protein recovery, and a passive characterization of the metabolite corona by analyzing metabolite matrices before and after NM exposures. This work introduces capillary electrophoresis &#8211; mass spectrometry (CESI-MS) as a new technique for NM corona characterization. The protocols outlined here demonstrate how CESI-MS can be used for the reliable characterization of both the protein and metabolite corona acquired by NMs. The move to CESI-MS greatly decreases the volume of sample required (compared to traditional liquid chromatography &#8211; mass spectrometry (LC-MS) approaches) with multiple injections possible from as little as 5 &#181;L of sample, making it ideal for volume limited samples. Furthermore, the environmental consequences of analysis are reduced with respect to LC-MS due to the low flow rates (&#60;20 nL/min) in CESI-MS, and the use of aqueous electrolytes which eliminates the need for organic solvents.

Here we present a protocol to characterize the complete biomolecular corona, proteins, and metabolites, acquired by nanomaterials from biofluids using a capillary electrophoresis &#8211; mass spectrometry approach.

The adsorption of biomolecules from surrounding biological matrices to the surface of nanomaterials (NMs) to form the corona has been of interest for the ...

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.

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 ...

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 ...

Organic Solvent-Based Protein Precipitation for Robust Proteome Purification Ahead of Mass Spectrometry

While multiple advances in mass spectrometry (MS) instruments have improved qualitative and quantitative proteome analysis, more reliable front-end approaches to isolate, enrich, and process proteins ahead of MS are critical for successful proteome characterization. Low, inconsistent protein recovery and residual impurities such as surfactants are detrimental to MS analysis. Protein precipitation is often considered unreliable, time-consuming, and technically challenging to perform compared to other sample preparation strategies. These concerns are overcome by employing optimal protein precipitation protocols. For acetone precipitation, the combination of specific salts, temperature control, solvent composition, and precipitation time is critical, while the efficiency of chloroform/methanol/water precipitation depends on proper pipetting and vial manipulation. Alternatively, these precipitation protocols are streamlined and semi-automated within a disposable spin cartridge. The expected outcomes of solvent-based protein precipitation in the conventional format and using a disposable, two-stage filtration and extraction cartridge are illustrated in this work. This includes the detailed characterization of proteomic mixtures by bottom-up LC-MS/MS analysis. The superior performance of SDS-based workflows is also demonstrated relative to non-contaminated protein.

The present protocol describes solvent-based protein precipitation under controlled conditions for robust and rapid recovery and purification of proteome samples prior to mass spectrometry.

While multiple advances in mass spectrometry (MS) instruments have improved qualitative and quantitative proteome analysis, more reliable front-end ...