The authors are thankful to Meytal Radzinski for her helpful discussions and critical reading of the article, and to Patrick Griffin and his lab members for their unlimited assistance while establishing the HDX analysis platform. The authors are grateful to the German-Israel Foundation (I-2332-1149.9/2012), the Binational Science Foundation (2015056), the Marie-Curie integration grant (618806), the Israel Science Foundation (1765/13 and 2629/16), and the Human Frontier Science Program (CDA00064/2014) for their financial support.

1. Preparation of Fully Reduced and Fully Oxidized Proteins
<ol>
	<li>Preparation of a fully reduced protein 
	Note: Here, we describe the reduction of a zinc-containing protein, and use a ZnCl2 solution to restore the Zn-incorporated, reduced protein state. The ZnCl2 solution can be replaced or discarded. Note that the time and temperature of the reduction process depends on the protein stability and function, and is thus specific per protein.
	<ol>
		<li>Thaw the protein sample on ice and spin it down to remove aggregates. Incubate the sample for at least 1.5&#160;h at 37 &#176;C with 5 mM DTT and 20 &#181;M ZnCl2 (up to 70% of protein concentration). 
		NOTE: The temperature at this step is protein-dependent and should be adjusted to the protein stability.</li>
		<li>Remove DTT using desalting columns. 
		NOTE: There are different desalting columns available. The protocol below describes the procedure using specific desalting columns (see Table of Materials). Before using other desalting columns, we recommend examining their efficiency of DTT removal and protein recovery, since some desalting columns may partially absorb the protein of interest.
		<ol>
			<li>Equilibrate the column with a potassium phosphate (KPi) buffer (40 mM, pH 7.5) by filling the column completely with the buffer and letting the buffer drip out. Repeat this process 2x.</li>
			<li>Remove the white disk filter in the column by gently pushing it down with tweezers and removing it; it is easiest to remove while the column is filled with the KPi buffer. Refill the column with KPi buffer and centrifuge it at 1,000 x g for 3 min.</li>
			<li>Transfer the column into a clean tube, add the protein sample slowly to the middle of the column and centrifuge it at 1,000 x g for 2 min. The DTT-free protein is now in the flow-through.</li>
		</ol>
		</li>
		<li>Check its concentrations (e.g., use a UV/Vis spectrometer) and measure its absorbance at 280 nm. Calculate the protein concentration using Beer&#39;s law.</li>
		<li>Distribute half of the protein samples into aliquots. Incubate the aliquots in anaerobic conditions (e.g., using an anaerobic chamber) for 20 min for a complete removal of oxygen. Seal the tubes with plastic film and store the samples at -20 &#176;C or -80 &#176;C, depending on the protein. 
		NOTE: Instead of using the anaerobic chamber, the tubes can also be flashed with argon gas to remove oxygen.</li>
	</ol>
	</li>
	<li>Preparation of a fully oxidized protein 
	NOTE: It is recommended to prepare oxidized protein samples from fully reduced proteins (described previously). This will reduce the heterogeneity in oxidation states of cysteines. It is possible to use different oxidizing reagents; here, we are focusing on hydrogen peroxide (H2O2). 
	CAUTION: Avoid over-oxidation as it can lead to undesirable intramolecular disulfide bonds and irreversible oxidation of different amino acids, including cysteine, methionine, tyrosine, and others.
	<ol>
		<li>To the remaining protein sample, add 5 mM H2O2 (freshly diluted) and incubate it for 3 h at 40 &#176;C while shaking. 
		NOTE: The temperature at this step is protein-dependent and should be adjusted to the protein stability.</li>
		<li>Equilibrate the column with KPi buffer (40 mM, pH 7.5) by filling the column completely with the buffer and letting the buffer drip out. Repeat this process 2x.</li>
		<li>Remove the white disk filter in the column by gently pushing it down with tweezers and removing it; it is easiest to remove while the column is filled with the KPi buffer.</li>
		<li>Refill the column with KPi buffer and centrifuge it at 1,000 x g for 3 min.</li>
		<li>Transfer the column into a clean tube, add the oxidized protein sample slowly to the middle of the column and centrifuge it at 1,000 x g for 2 min; the oxidized protein is now in the flow-through.</li>
		<li>Check the protein concentrations as in step 1.2.5, divide the oxidized proteins into aliquots, and store them at -20 &#176;C or -80 &#176;C, protein-dependent.</li>
	</ol>
	</li>
</ol>
2. Light Scattering Aggregation Assay
Note: All concentrations in this assay are chaperone- and substrate-specific, and should be calibrated. All buffers should be 0.22 &#181;m-filtered, as it is extremely important that the buffers are free of any particles or air bubbles and the cuvettes are clean and dust-free. It is very important to use a stirrer placed in the quartz cuvette. Check different stirrer sizes and shapes in order to ensure an efficient mixing of the entire solution without producing undesirable air bubbles. Moreover, there are different flouorospectrometers available in laboratories and facilities. Here, a specific fluorospectrometer (see Table of Materials) was used. Different instruments have a diverse sensitivity, measurement speed, and sampler parameters. Therefore, the exact measurement parameters (e.g., emission and excitation bandwidth, sensitivity, and others) should be optimized using a known aggregation-prone protein and its corresponding conditions. Using citrate synthase (CS) and/or luciferase as initial substrates in nanomolar concentrations is recommended.
<ol>
	<li>Chemical aggregation assay
	<ol>
		<li>Prepare the denatured substrate by incubating 12 &#181;M CS overnight in 40 mM HEPES (pH 7.5) and 4.5 M GdnCl. In order to preserve the pH, dissolve the GdnCl in the 40 mM HEPES (pH 7.5).</li>
		<li>Open the fluorospectrometer software and go to Time course measurement. Set the parameters to: Temperature: 25 &#176;C; &#955;em: 360; Em bandwidth: 5 nm; &#955;ex: 360; Ex bandwidth: 2.5 nm; and Data interval: 0.5 s.</li>
		<li>Prepare the sample by adding 1,600 &#181;L of 40 mM HEPES to a quartz cuvette. Insert the cuvette into the sample holder and let the sample reach the desired temperature.</li>
		<li>Set the stirring to 600 rpm and begin the measurement until a baseline is established. Keep the stirring on for the entire measurement.</li>
		<li>To measure the CS aggregation in the absence of a chaperone, at 120 s into the measurement, add (gently, but quickly) 10 &#181;L of denatured CS (final concentration of 75 nM). Continue the measurement for 1,200 s.</li>
		<li>To measure the CS aggregation in the presence of Hsp33, at 60 s into the measurement, add Hsp33 (final concentration of 300 nM). After an additional 60 s, add 10 &#181;L of denatured CS (final concentration of 75 nM). Continue the measurement for 1,200 s. 
		NOTE: When adding the substrate or chaperone, in order to avoid insertion of bubbles, use a 10-&#181;L pipette.</li>
	</ol>
	</li>
	<li>Thermal aggregation assay 
	NOTE: The temperature for the thermal aggregation assay depends on the protein stability and should be adjusted to each protein independently.
	<ol>
		<li>Open the fluorospectrometer software and go to Time course measurement. Set the parameters as follows: Temperature: 43 &#176;C; &#955;em: 360; Emission bandwidth: 5 nm; &#955;ex: 360; Excitation bandwidth: 2.5 nm; and Data interval: 0.5 s.</li>
		<li>Prepare the sample by adding 1,600 &#181;L of pre-warmed 40 mM HEPES to a quartz cuvette. Insert the cuvette into the sample holder and let the sample reach 43 &#176;C.</li>
		<li>Set the stirring to 600 rpm and begin the measurement until a baseline is established. Keep the stirring on for the entire measurement.</li>
		<li>To measure the CS aggregation in the absence of a chaperone, at 120 s into the measurement, gently add CS (final concentration of 125 nM) and continue measuring for 1,200 s.</li>
		<li>To measure the CS aggregation in the presence of Hsp33, at 60 s into the measurement, add Hsp33 (final concentration of 600 nM). After an additional 60 s, add CS (final concentration of 125 nM). Continue the measurement for 1,200 s. 
		NOTE: When adding the substrate or chaperone, avoid the insertion of bubbles.</li>
	</ol>
	</li>
	<li>Data analysis and noise removal by Kfits 
	Note: Kfits is available at use of Kfits has previously been described in Rimon et al.45
	<ol>
		<li>Upload the data file. 
		NOTE: The input is the raw results from light scattering measurements in a textual comma-separated or tab-delimited format.</li>
		<li>To remove any noise, choose analysis parameters. Use Automatic best model (recommended), mark the Noise is always above signal flag.</li>
		<li>Manually remove obvious outliers using the green and red adjustable lines; this step will filter the noise above the green line and below the red line.</li>
		<li>Set the baseline and the fit curve. After applying the noise threshold, download the processed data.</li>
	</ol>
	</li>
</ol>
3. Hydrogen-deuterium Exchange Mass Spectrometry
<ol>
	<li>Preparation of buffers and protein samples (Hsp33 and Hsp33-CS complex)
	<ol>
		<li>Dilute the protein samples to a final concentration of 1 mg/mL in 25 mM Tris-HCl buffer at pH 7.5 and transfer them to 1.5-mL vials.</li>
		<li>Prepare protein-substrate complex samples by incubating Hsp33 with CS at a ratio of 1:1.5 at 43 &#176;C.
		<ol>
			<li>Add CS in a step-wise manner to avoid any rapid aggregation and ensure a fruitful binding between the Hsp33 and the thermally unfolded CS.</li>
			<li>Use at least four steps (i.e., each time add a quarter of the final volume) and incubate the samples for 15 min after every addition to allow CS protection by Hsp33.</li>
		</ol>
		</li>
		<li>Remove any aggregates using a 30-min centrifugation at 16,000 x g at 4 &#176;C. 
		NOTE: New protein-substrate complexes must be prepared fresh. Substrate addition should be made gradually; otherwise, aggregation will occur. Temperature, substrate, and substrate concentrations are protein-specific.</li>
		<li>Prepare a buffer H (25 mM Tris-HCl, pH 7.5), which serves as deuteration control, and a buffer D (25 mM Tris-DCl, pH 7.09), which is the deuteration buffer. Also prepare a fresh quenching buffer (150 mM TCEP, 3 M GdnCl, 0.1% formic acid). 
		NOTE: The quenching buffer should be optimized for the protein of interest in order to obtain its maximal sequence coverage after the pepsin digestion.</li>
		<li>Transfer all buffers and samples into vials, and place them in proper trays. Hold buffers H and D at 25 &#176;C (tray 25 &#176;C); on the other hand, hold the samples and quenching buffer at 0 - 2 &#176;C (tray 0 &#176;C). Place the samples in 150-&#181;L glass inserts (see the Table of Materials) first, and then transfer them into vials.</li>
	</ol>
	</li>
	<li>Preparation of the instrument 
	NOTE: The mass spectrometer comes with two accompanying software programs: one controls the pumps, and the other controls the mass spectrometer (refer to the Table of Materials). The next steps will be described using these two software programs.
	<ol>
		<li>Manually turn on all cooling units. Once all cooling systems reach their target temperatures, manually switch on both the high-performance liquid chromatography (HPLC) and loading pumps. 
		NOTE: In our case, these consist of two cooling baths and a cooling box (which contains the trap and analytical columns). One cooling unit cools the tray at 0 &#176;C, while the other keeps the tray at 25 &#176;C; the temperature of the cooling box is 1 - 2 &#176;C.</li>
		<li>Open the software which controls both pumps, as well as the software which controls the mass spectrometer; make sure that MS is on Standby.</li>
		<li>Disconnect the HPLC outlet valve from the MS source and wash the system first with a &#34;triple cleaner&#34; solution (1% formic acid, 33% acetonitrile, 33% isopropanol, 33% methanol), then with buffer B (80% acetonitrile, 0.1% formic acid), and finally with buffer A (0.1% formic acid, pH 2.25), and then insert the pepsin column into the system. If changing buffers, make sure to purge both pumps before proceeding.</li>
		<li>Ensure that the flow-rate for both pumps is 0.1 mL/min and the pressure is stable.</li>
		<li>After washing the system with a steady flow and steady pressure, insert the HPLC outlet into the MS source and turn the MS on.</li>
	</ol>
	</li>
	<li>Mass spectrometer parameters
	<ol>
		<li>Set the peptide ionization to electrospray ionization (ESI) at 175 &#176;C, the sheath gas flow at 17, the aux gas glow at 2, and the spray voltage at 4.5 kV (Supplementary Figure 1).</li>
		<li>Set up the parameters as follows for the non-deuterated samples.
		<ol>
			<li>Set the parameters: Scan Range m/z to 300 - 1500; Resolution to 70,000; Automatic gain control target (AGC) to 106; and the Maximum injection time (IT) at 100 ms. 
			NOTE: The 5 most intense ions with charge states between +2 and +6 (their intensity higher than 1.3 x 104) and a dynamic window of 30 in will be isolated.</li>
			<li>Perform the ion fragmentation by higher-energy collisional dissociation (HCD) in the multiple-collision cell with normalized collision energy (NCE) equal to 28. Detect fragment ions by tandem MS at a resolution of 35,000, AGC target of 105, maximum IT at 60 ms, and an isolation window of 2.0 m/z.</li>
		</ol>
		</li>
		<li>For the deuterated samples, employ MS1 only with a higher resolution of 140,000 and with otherwise similar parameters.</li>
	</ol>
	</li>
	<li>Running the experiment
	<ol>
		<li>Set the trays in the following manner.
		<ol>
			<li>Place buffer H and D in Tray 25 &#176;C, then place the samples and quenching buffer in the 0 &#176;C tray.</li>
			<li>Place X empty vials, aligned in both trays, where X = the number of samples x the number of time points. 
			NOTE: In the 25 &#176;C tray, the empty vials serve as reaction vials, and in the tray 0 &#176;C, they serve as quenching vials.</li>
			<li>Each of the empty vials contains an empty glass insert; discard and replace said insert between experiments. 
			NOTE: In order to run the HDX experiment in the most efficient and least time-consuming way, a software which allows samples to be run simultaneously (Supplementary Figure 2) was used. The next steps will be described using this software.</li>
		</ol>
		</li>
		<li>Open the software. Set up the program parameters to perform the following.
		<ol>
			<li>Incubate each sample (5 &#181;L) with 45 &#181;L of buffer D for several minutes, depending on the time points selected (e.g., 1, 3, 18, 40, and 100 min) at 25 &#176;C. Incubate the non-deuterated sample with 45 &#181;L of buffer H instead.</li>
			<li>Immediately after the incubation, mix 50 &#181;L of deuterated protein into 50 &#181;L of ice-cold quenching buffer and inject them into an immobilized pepsin column (20 mm in length, 2.0 mm in diameter).</li>
			<li>Elute any peptides from the pepsin column into the pre-column by washing it with buffer A for 6 min at a rate of 50 &#181;L/min. Keep buffer A in the cooling box at 0 &#176;C.</li>
			<li>Elute the peptides out of the pre-column into the C18 analytical column (C18 Column, 130 &#197;, 1.7 &#181;m, 2.1 mm x 50 mm, kept at 0 &#176;C) and separate them by applying a linear gradient of acetonitrile at 100 &#181;L/min. Run the acetonitrile gradient using acetonitrile 100% as buffer B: 7 min at 2%, 7 min at 10 - 30%, 2.5 min at 30 - 90%, 1.5 min at 90%, rapid gradient for 1 min at 90 - 8%, and finally equilibrate the column for 4 min at 2%.</li>
		</ol>
		</li>
		<li>Build a running sequence (see Supplementary Figure 2): enter all time points, sample names, and locations in the trays, as well as the buffer names and locations. 
		NOTE: The software allows the alignment of all the different running times into a program that runs several samples at once, efficiently decreasing running times.</li>
	</ol>
	</li>
	<li>Data analysis 
	NOTE: We work with several software programs in the process of data analysis, including Proteome Discoverer and MaxQuant (free software) for the peptide identification and analysis of sequence coverage, as well as HDX workbench, a free software that allows the analysis of the deuterium incorporation in proteins, and a comparison of the HDX rates between different sets of experiments. The next steps will be described using these software programs.
	<ol>
		<li>Analyze the peptide coverage of the sample by running the non-deuterated control samples through the software (Supplementary Figure 3). Set up the software using the following parameters.
		<ol>
			<li>Use the Sequest HT method with a No-enzyme (unspecific) cleave. Search for peptides of 4 - 144 amino acids with a mass tolerance of 7 ppm and 0.5 Da for the precursor&#39;s ion and fragment. Allow for a dynamic methionine oxidation.</li>
			<li>Create a database for the protein, through which the software can determine the peptide coverage. Export the identified peptides in textual format.</li>
		</ol>
		</li>
		<li>Analyze the deuterated results using the HDX workbench free software58.
		<ol>
			<li>Open the protein editor and define the protein by inserting the protein sequence and peptide coverage file.</li>
			<li>Next, open the Setup experiment wizard editor and enter all the inputs requested. 
			NOTE: The program requires the number of samples, the number of time points, the number of replicates, the pH value of the buffers, and the temperature.</li>
			<li>Finally, input the parameters regarding the mass spectrometer used, after which the program generates a list of detected peptides. 
			NOTE: The software detects the &#34;HDX&#34; peptides, inspects isotope clusters, and demonstrates a clear visualization of deuteration in the samples.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Data presentation
	<ol>
		<li>Label the regions with a deferential deuteration uptake on the structure using the PyMol software or any other visualization software.</li>
		<li>Introduce the deuterium uptake levels instead of the B-factor values. 
		Note: A basic tutorial can be found at https://pymolwiki.org/index.php/Practical_Pymol_for_Beginners.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

In this paper, we provided protocols for the analysis of redox-dependent chaperone activity and the characterization of structural changes upon the binding of a client protein. These are complementary methodologies to define potential chaperone-substrate complexes and analyze potential interaction sites.
Here, we applied these protocols for the characterization of a complex between the redox-regulated chaperone Hsp33 with a well-studied chaperone substrate CS. We presented two different types ...

Cells frequently encounter an accumulation of reactive oxygen species (ROS) produced as byproducts of respiration1,2, protein and lipid oxidation3,4, and additional processes5,6,7. Despite ROS&#39; beneficial role in diverse biological processes such as cellular signaling8,9 and immune response10, an imbalance between ROS production and its detoxification might occur, leading to oxidative stress7. The biological targets of ROS are proteins, lipids, and nucleic acids, the oxidation of which affect their structure and function. Therefore, the accumulation of cellular oxidants is strongly linked to a diverse range of pathologies including cancer9,11, inflammation12,13, and aging14,15, and have been found to be involved in the onset and progression of neurodegenerative disorders such as Alzheimer&#39;s, Parkinson&#39;s, and ALS disease16,17,18.
Both newly synthesized and mature proteins are highly sensitive to oxidation due to the potentially harmful modifications of their side chains, which shape protein structure and function19,20. Therefore, oxidative stress usually leads to a widespread protein inactivation, misfolding and aggregation, eventually leading to cell death. One of the elegant cellular strategies to cope with the potential damage of protein oxidation is to utilize redox-dependent chaperones, which inhibit the widespread protein aggregation, instead of forming stable complexes with misfolded client proteins21,22,23. These first-line defense chaperones are rapidly activated by a site-specific oxidation (usually on cysteine residues) that converts them into potent anti-aggregation molecules24. Since oxidative stress results in the inhibition of respiration and in decreases in the cellular ATP levels25, canonical ATP-dependent chaperones are less effective during oxidative stress conditions25,26,27. Therefore, redox-activated ATP-independent chaperones play a vital role in maintaining protein homeostasis upon the accumulation of oxidants in bacteria and eukaryotes (e.g., Hsp3328 and RidA29 in bacteria, Get330 in yeast, peroxiredoxins31 in eukaryotes). The activity of these chaperones strongly depends on reversible structural conformational changes induced by a site-specific oxidation that uncovers hydrophobic regions involved in the recognition of misfolded client proteins.
Research of the anti-aggregation mechanism and the principles governing the recognition of the client proteins by chaperones is not easy due to the dynamic and heterogenic nature of chaperone-substrate interactions32,33,34,35,36,37. However, stress-regulated chaperones have an opportunity to advance our understanding of the anti-aggregation function due to their ability to: 1) obtain two different forms of the chaperone, active (e.g., oxidized) and inactive (e.g., reduced), with the introduction or removal of a stress condition easily switching between them (e.g., oxidant and reducing agent), 2) have a broad range of substrates, 3) form highly stable complexes with the client proteins that may be evaluated by different structural methodologies, and 4) focus solely on the substrate recognition and release, mediated by redox-dependent conformational changes, as the majority of these chaperones lacks the folding capability.
Here, we analyze the bacterial redox-regulated chaperone Hsp33&#39;s anti-aggregation activity, a vital component of the bacterial defense system against oxidation-induced protein aggregation28. When reduced, Hsp33 is a tightly folded zinc-binding protein with no chaperone activity; however, when exposed to oxidative stress, Hsp33 undergoes extensive conformational changes which expose its substrate binding regions38,39. Upon oxidation, the zinc ion that is strongly bound to four highly conserved cysteine residues of the C-terminal domain is released40. This results in the formation of two disulfide bonds, an unfolding of the C-terminal domain, and a destabilization of the adjacent linker region41. The C-terminal and linker regions are highly flexible and are defined as intrinsically or partially disordered. Upon return to non-stress conditions, the cysteines become reduced and the chaperone returns to its native folded state with no anti-aggregation activity. The refolding of the chaperone leads to a further unfolding and destabilization of the bound client protein, which triggers its transfer to the canonical chaperone system, DnaK/J, for refolding38. Analysis of Hsp33&#39;s interaction sites suggests that Hsp33 uses both its charged disordered regions as well as the hydrophobic regions on the linker and N-terminal domain to capture misfolded client proteins and prevent their aggregation38,42. In the folded state, these regions are hidden by the folded linker and C-terminal domain. Interestingly, the linker region serves as a gatekeeper of Hsp33&#39;s folded and inactive state, &#34;sensing&#34; the folding status of its adjacent C-terminal domain34. Once destabilized by mutagenesis (either by a point mutation or a full sequence perturbation), Hsp33 is converted to a constitutively active chaperone regardless the redox state of its redox-sensitive cysteines43.
The protocols presented here allow monitoring of Hsp33&#39;s redox-dependent chaperone activity, as well as mapping conformational changes upon the activation and binding of client proteins. This methodology can be adapted to research other chaperone-client recognition models as well as non-chaperone protein-protein interactions. Moreover, we present protocols for the preparation of fully reduced and oxidized chaperones that can be used in studies of other redox-switch proteins, to reveal potential roles of protein oxidation on the protein activity.
Specifically, we describe a procedure to monitor chaperone activity in vitro and define its substrate specificity under different types of protein aggregation (chemically or thermally induced) using light scattering (LS) measured by a fluorospectrometer44. During aggregation, light scattering at 360 nm increases rapidly due to the increasing turbidity. Thus, aggregation can be monitored in a time-dependent manner at this wavelength. LS is a fast and sensitive method for testing protein aggregation and thus the anti-aggregation activity of a protein of interest using nanomolar concentrations, enabling the characterization of protein aggregation-related kinetic parameters under different conditions. Moreover, the LS protocol described here does not require expensive instrumentation, and can be easily established in any laboratory.
Nevertheless, it is quite challenging to obtain &#34;clean&#34; kinetic curves and to derive a protein&#39;s kinetic parameters from such light scattering experiments, due to noise and the large number of outliers generated by air bubbles and large aggregates. To overcome this obstacle, we present a novel graphical tool, Kfits45, used for reducing noise levels in different kinetic measurements, specifically fitted for protein aggregation kinetic data. This software provides preliminary kinetic parameters for an early assessment of the results and allows the user to &#34;clean&#34; large quantities of data quickly without affecting its kinetic properties. Kfits is implemented in Python and available in open source at 45.
One of the challenging questions in the field relates to mapping interaction sites between chaperones and their client proteins and understanding how chaperones recognize a wide range of misfolded substrates. This question is further complicated when studying highly dynamic protein complexes which involve intrinsically disordered chaperones and aggregation-prone substrates. Fortunately, structural mass spectrometry has dramatically advanced over the last decade and has successfully provided helpful approaches and tools to analyze the structural plasticity and map residues involved in protein recognition46,47,48,49. Here, we present one such technique-hydrogen-deuterium exchange mass spectrometry (HDX-MS)-which allows the mapping of residue-level changes in a structural conformation upon protein modification or protein/Ligand binding35,50,51,52,53,54,55. HDX-MS uses the continuous exchange of backbone hydrogens by deuterium, the rate of which is affected by the chemical environment, accessibility, and covalent and non-covalent bonds56. HDX-MS tracks these exchange processes using a deuterated solvent, commonly heavy water (D2O), and allows measurement based on the change in molecular weight following the hydrogen to deuterium exchange. Slower rates of hydrogen-deuterium exchange can result from hydrogens participating in hydrogen bonds or, simply, from steric hindrance, which indicates local changes in structure57. Changes upon a ligand binding or post-translational modifications can also lead to differences in the hydrogen environment, with a binding resulting in differences in the hydrogen-deuterium exchange (HDX) rates46,53.
We applied this technology to 1) map Hsp33 regions which rapidly unfold upon oxidation, leading to the activation of Hsp33, and 2) define the potential binding interface of Hsp33 with its full-length misfolded substrate, citrate synthase (CS)38.
The methods described in this manuscript can be applied to study redox-dependent functions of proteins in vitro, defining anti-aggregation activity and the role of structural changes (if any) in protein function. These methodologies can be easily adapted to diverse biological systems and applied in the laboratory.

<table><tbody><tr><td>&lt;strong&gt;Chemicals, Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Acetonitrile HPLC plus</td><td>Sigma Aldrich</td><td>34998-2.5L</td><td>solvent</td></tr><tr><td>Formic acid Optima LC/MS</td><td>Fisher Chemicals</td><td>A117-50</td><td>solvent supplement</td></tr><tr><td>Isopropyl alcohol, HPLC grade</td><td>Fisher Chemicals</td><td>P750717</td><td>solvent</td></tr><tr><td>Methanol</td><td>Fisher Chemicals</td><td>A456-212</td><td>solvent</td></tr><tr><td>Tris(hydroxymethyl)aminomethane</td><td>Sigma Aldrich</td><td>252859</td><td>buffer</td></tr><tr><td>Trifluoroacetic acid</td><td>Sigma Aldrich</td><td>76-05-1</td><td>solvent</td></tr><tr><td>Water for HPLC</td><td>Sigma Aldrich</td><td>270733-2.5L-M</td><td>solvent</td></tr><tr><td>ZnCl2, Zinc Chloride</td><td>Merck</td><td>B0755416 308</td><td>reagent</td></tr><tr><td>DTT</td><td>goldbio</td><td>27565-41-9</td><td>reducing agent</td></tr><tr><td>PD mini trap G-25 columns GE healthcare</td><td>GE healthcare</td><td>29-9180-07</td><td>desalting column</td></tr><tr><td>Potassium Phosphate</td><td>United states Biochemical Corporation</td><td>20274</td><td>buffer</td></tr><tr><td>Hydrogen peroxide 30%</td><td>Merck</td><td>K46809910526</td><td>oxidizing agent</td></tr><tr><td>citrate synthase</td><td>sigma aldrich</td><td>C3260</td><td>substrate</td></tr><tr><td>HEPES acid free</td><td>sigma aldrich</td><td>7365-45-9</td><td>buffer</td></tr><tr><td>Gndcl</td><td>sigma aldrich</td><td>G3272-500G</td><td>denaturant</td></tr><tr><td>Deuterium Chloride Solution</td><td>sigma aldrich</td><td>543047-10G</td><td>buffer</td></tr><tr><td>Deuterium Oxide 99%</td><td>sigma aldrich</td><td>151882-100G</td><td>solvent</td></tr><tr><td>TCEP</td><td>bioworld</td><td>42000058-2</td><td>reducing agent</td></tr><tr><td>150uL Micro-Insert with Mandrel Interior &amp;amp; Polymer Feet, 29*5mm</td><td>La-Pha-Pack -Thermo Fischer Scientific</td><td></td><td></td></tr><tr><td>1.5mL Clear Short Thread Vial 9mm Thread, 11.6*32mm</td><td>La-Pha-Pack -Thermo Fischer Scientific</td><td></td><td></td></tr><tr><td>quartz cuvette</td><td>Hellma 101-QS</td><td></td><td></td></tr><tr><td>&lt;strong&gt;Instruments&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Jasco FP-8500 Fluorospectrometer</td><td>Jasco</td><td></td><td></td></tr><tr><td>Thermomixer Comfort</td><td>Eppendorf</td><td>13058/0</td><td></td></tr><tr><td>Heraeus Megafuge 16R, bench topCentrifuge</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>pH meter , PB-11 sartorius</td><td>Sartorius</td><td>13119/0</td><td></td></tr><tr><td>AffiPro Immobilized Pepsin column (20mm length, 2.0mm diameter).</td><td>AffiPro</td><td></td><td></td></tr><tr><td>Waters Pre-column (ACQUITY UPLC BEH C18 VanGuard 130 &amp;Aring;, 1.7um, 2.1mmx5mm)</td><td>Waters</td><td></td><td></td></tr><tr><td>C18 analytical column (ACQUITY UPLC Peptide BEH c18 Column, 130 &amp;Aring;, 1.7um, 2.1mmx50mm)</td><td></td><td></td><td></td></tr><tr><td>Vinyl Anaerobic chamber with Airlock door</td><td>COY</td><td></td><td></td></tr><tr><td>Q-exactive-orbitrap mass spectrometer</td><td>Thermo-Fischer Scientific</td><td></td><td></td></tr><tr><td>PAL system LHX - robotic system for handling HDX samples</td><td>PAL system</td><td>https://www.palsystem.com/index.php?id=840</td><td></td></tr><tr><td>Dionex Ultimate 3000, XRS pump</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>Dionex AXP-MS auxiliary pump</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>&lt;strong&gt;Software, Software Tools, Database search&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Kfits: Fit aggregation Data</td><td>http://kfits.reichmannlab.com/fitter/</td><td></td><td></td></tr><tr><td>Thermo Scientific Xcalibur software</td><td>https://www.thermofisher.com/order/catalog/product/OPTON-30487</td><td></td><td></td></tr><tr><td>Q Exactive MS Series Tune Interface (Tune)</td><td>https://tools.thermofisher.com/content/sfs/brochures/WS-MS-Q-Exactive-Calibration-Maintenance-iQuan2016-EN.pdf</td><td></td><td></td></tr><tr><td>Chronos software (Axel Semrau)</td><td>http://www.axel-semrau.de/en/Software/Software+Solutions/Chronos-p-966.html</td><td></td><td></td></tr><tr><td>Proteome Discoverer V1.4 software</td><td>https://www.thermofisher.com/order/catalog/product/OPTON-30795</td><td></td><td></td></tr><tr><td>HDX workbench software</td><td>http://hdx.florida.scripps.edu/hdx_workbench/Home.html</td><td></td><td></td></tr></tbody></table>

defining hsp33's redox-regulated chaperone activity and mapping conformational changes on hsp33 using hydrogen-deuterium exchange mass spectrometry

Living organisms regularly need to cope with fluctuating environments during their life cycle, including changes in temperature, pH, the accumulation of reactive oxygen species, and more. These fluctuations can lead to a widespread protein unfolding, aggregation, and cell death. Therefore, cells have evolved a dynamic and stress-specific network of molecular chaperones, which maintain a "healthy" proteome during stress conditions. ATP-independent chaperones constitute one major class of molecular chaperones, which serve as first-line defense molecules, protecting against protein aggregation in a stress-dependent manner. One feature these chaperones have in common is their ability to utilize structural plasticity for their stress-specific activation, recognition, and release of the misfolded client. 
In this paper, we focus on the functional and structural analysis of one such intrinsically disordered chaperone, the bacterial redox-regulated Hsp33, which protects proteins against aggregation during oxidative stress. Here, we present a toolbox of diverse techniques for studying redox-regulated chaperone activity, as well as for mapping conformational changes of the chaperone, underlying its activity. Specifically, we describe a workflow which includes the preparation of fully reduced and fully oxidized proteins, followed by an analysis of the chaperone anti-aggregation activity in vitro using light-scattering, focusing on the degree of the anti-aggregation activity and its kinetics. To overcome frequent outliers accumulated during aggregation assays, we describe the usage of Kfits, a novel graphical tool which allows easy processing of kinetic measurements. This tool can be easily applied to other types of kinetic measurements for removing outliers and fitting kinetic parameters. To correlate the function with the protein structure, we describe the setup and workflow of a structural mass spectrometry technique, hydrogen-deuterium exchange mass spectrometry, that allows the mapping of conformational changes on the chaperone and substrate during different stages of Hsp33 activity. The same methodology can be applied to other protein-protein and protein-ligand interactions.

The two methods presented make it possible to follow the kinetic activity and dynamics of protein interactions between a chaperone and its substrate. Moreover, the reduction-oxidation protocol allows the preparation of a fully reduced and fully oxidized chaperone, giving a more in-depth understanding of the activation mechanism of redox-dependent disordered chaperones.
First, we used light scattering in order to examine the redo...

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Defining Hsp33's Redox-regulated Chaperone Activity and Mapping Conformational Changes on Hsp33 Using Hydrogen-deuterium Exchange Mass Spectrometry

One of the most challenging stress conditions that organisms encounter during their lifetime involves the accumulation of oxidants. During oxidative stress, cells heavily rely on molecular chaperones. Here, we present methods used to investigate the redox-regulated anti-aggregation activity, as well as to monitor structural changes governing the chaperone function using HDX-MS.

Living organisms regularly need to cope with fluctuating environments during their life cycle, including changes in temperature, pH, the accumulation of ...

defining-hsp33-s-redox-regulated-chaperone-activity-mapping

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8.6K Views.  The Hebrew University of Jerusalem. Een van de meest uitdagende stress voorwaarden die organismen tijdens hun leven tegenkomen gaat de accumulatie van oxidanten. Tijdens oxidatieve stress afhankelijk cellen sterk zijn van moleculaire chaperones. Hier presenteren we methoden voor het onderzoeken van de redox-gereglementeerde anti-aggregatie activiteit, evenals het monitoren van de structurele veranderingen in het bestuur van de Chaperon-functie met behulp van HDX-MS.

Video: Defining Hsp33's Redox-regulated Chaperone Activity and Mapping Conformational Changes on Hsp33 Using Hydrogen-deuterium Exchange Mass Spectrometry

Detection of the pH-dependent Activity of Escherichia coli Chaperone HdeB In Vitro and In Vivo

Bacteria are frequently exposed to environmental changes, such as alterations in pH, temperature, redox status, light exposure or mechanical force. Many of these conditions cause protein unfolding in the cell and have detrimental impact on the survival of the organism. A group of unrelated, stress-specific molecular chaperones have been shown to play essential roles in the survival of these stress conditions. While fully folded and chaperone-inactive before stress, these proteins rapidly unfold and become chaperone-active under specific stress conditions. Once activated, these conditionally disordered chaperones bind to a large number of different aggregation-prone proteins, prevent their aggregation and either directly or indirectly facilitate protein refolding upon return to non-stress conditions. The primary approach for gaining a more detailed understanding about the mechanism of their activation and client recognition involves the purification and subsequent characterization of these proteins using in vitro chaperone assays. Follow-up in vivo stress assays are absolutely essential to independently confirm the obtained in vitro results.
This protocol describes in vitro and in vivo methods to characterize the chaperone activity of E. coli HdeB, an acid-activated chaperone. Light scattering measurements were used as a convenient read-out for HdeB&#39;s capacity to prevent acid-induced aggregation of an established model client protein, MDH, in vitro. Analytical ultracentrifugation experiments were applied to reveal complex formation between HdeB and its client protein LDH, to shed light into the fate of client proteins upon their return to non-stress conditions. Enzymatic activity assays of the client proteins were conducted to monitor the effects of HdeB on pH-induced client inactivation and reactivation. Finally, survival studies were used to monitor the influence of HdeB&#39;s chaperone function in vivo.

Detection of the pH-dependent Activity of Escherichia coli Chaperone HdeB In Vitro and In Vivo

This study describes biophysical, biochemical and molecular techniques to characterize the chaperone activity of Escherichia coli HdeB under acidic pH conditions. These methods have been successfully applied for other acid-protective chaperones such as HdeA and can be modified to work for other chaperones and stress conditions.

Bacteria are frequently exposed to environmental changes, such as alterations in pH, temperature, redox status, light exposure or mechanical force. Many ...

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;

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 ...

Chemistry

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. ...

Resolving Affinity Purified Protein Complexes by Blue Native PAGE and Protein Correlation Profiling

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological processes. Affinity purification coupled to mass spectrometry (AP-MS) has become the method of choice for identifying protein-protein interactions. However, conventional AP-MS studies provide information on protein interactions, but the organizational information is lost. To address this issue, we developed a strategy to unravel the distinct functional assemblies a protein might be involved in, by resolving affinity-purified protein complexes prior to their characterization by mass spectrometry. Protein complexes isolated through affinity purification of a bait protein using an epitope tag and competitive elution are separated through blue native electrophoresis. Comparison of protein migration profiles through correlation profiling using quantitative mass spectrometry allows assignment of interacting proteins to distinct molecular entities. This method is able to resolve protein complexes of close molecular weights that might not be resolved by traditional chromatographic techniques such as gel filtration. With little more work than conventional AP-geLC-MS/MS, we demonstrate this strategy may in many cases be adequate for obtaining protein complex topological information concomitantly to identifying protein interactions.

Here we present protocols for affinity purification of protein complexes and their separation by blue native PAGE, followed by protein correlation profiling using label free quantitative mass spectrometry. This method is useful to resolve interactomes into distinct protein complexes.

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological ...

Lipid Droplet Isolation for Quantitative Mass Spectrometry Analysis

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of virion morphogenesis. Quantitative lipid droplet proteome analysis can be used to identify proteins that localize to or are displaced from lipid droplets under conditions such as virus infections. Here, we describe a protocol that has been successfully used to characterize the changes in the lipid droplet proteome following infection with HCV. We use Stable Isotope Labeling with Amino Acids in Cell Culture (SILAC) and thus label the complete proteome of one population of cells with &#34;heavy&#34; amino acids to quantitate the proteins by mass spectrometry. For lipid droplet isolation, the two cell populations (i.e.&#160;HCV-infected/&#34;light&#34; amino acids and uninfected control/&#34;heavy&#34; amino acids) are mixed 1:1 and lysed mechanically in hypotonic buffer. After removing the nuclei and cell debris by low speed centrifugation, lipid droplet-associated proteins are enriched by two subsequent ultracentrifugation steps followed by three washing steps in isotonic buffer. The purity of the lipid droplet fractions is analyzed by western blotting with antibodies recognizing different subcellular compartments. Lipid droplet-associated proteins are then separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) followed by Coomassie staining. After tryptic digest, the peptides are quantified by liquid chromatography-electrospray ionization-tandem mass spectrometry (LC-ESI-MS/MS). Using this method, we identified proteins recruited to lipid droplets upon HCV infection that might represent pro- or antiviral host factors. Our method can be applied to a variety of different cells and culture conditions, such as infection with pathogens, environmental stress, or drug treatment.

Lipid droplets are important organelles for the replication of several pathogens, including the Hepatitis C Virus (HCV). We describe a method to isolate lipid droplets for quantitative mass spectrometry of associated proteins; it can be used under a variety of conditions, such as virus infection, environmental stress, or drug treatment.

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of ...

Using Three-color Single-molecule FRET to Study the Correlation of Protein Interactions

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For many molecular machines in a cell proteins have to act together&#160;with interaction partners in a functional cycle to fulfill their task. The extension of two-color to multi-color smFRET makes it possible to simultaneously probe more than one interaction or conformational change. This not only adds a new dimension to smFRET experiments but it also offers the unique possibility to directly study the sequence of events and to detect correlated interactions when using an immobilized sample and a total internal reflection fluorescence microscope (TIRFM). Therefore, multi-color smFRET is a versatile tool for studying biomolecular complexes in a quantitative manner and in a previously unachievable detail.
Here, we demonstrate how to overcome the special challenges of multi-color smFRET experiments on proteins. We present detailed protocols for obtaining the data and for extracting kinetic information. This includes trace selection criteria, state separation, and the recovery of state trajectories from the noisy data using a 3D ensemble Hidden Markov Model (HMM). Compared to other methods, the kinetic information is not recovered from dwell time histograms but directly from the HMM. The maximum likelihood framework allows us to critically evaluate the kinetic model and to provide meaningful uncertainties for the rates.
By applying our method to the heat shock protein 90 (Hsp90), we are able to disentangle the nucleotide binding and the global conformational changes of the protein. This allows us to directly observe the cooperativity between the two nucleotide binding pockets of the Hsp90 dimer.

Here, we present a protocol to obtain three-color smFRET data and its analysis with a 3D ensemble Hidden Markov Model. With this approach, scientists can extract kinetic information from complex protein systems, including cooperativity or correlated interactions.

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For ...

The Determination of Protease Specificity in Mouse Tissue Extracts by MALDI-TOF Mass Spectrometry: Manipulating PH to Cause Specificity Changes

Proteases have several biological functions, including protein activation/inactivation and food digestion. Identifying protease specificity is important for revealing protease function. The method proposed in this study determines protease specificity by measuring the molecular weight of unique substrates using Matrix-assisted Laser Desorption/Ionization Time-of-Flight (MALDI-TOF) mass spectrometry. The substrates contain iminobiotin, while the cleaved site consists of amino acids, and the spacer consists of polyethylene glycol. The cleaved substrate will generate a unique molecular weight using a cleaved amino acid. One of the merits of this method is that it may be carried out in one pot using crude samples, and it is also suitable for assessing multiple samples. In this article, we describe a simple experimental method optimized with samples extracted from mouse lung tissue, including tissue extraction, placement of digestive substrates into samples, purification of digestive substrates under different pH conditions, and measurement of the substrates&#39; molecular weight using MALDI-TOF mass spectrometry. In summary, this technique allows for the identification of protease specificity in crude samples derived from tissue extracts using MALDI-TOF mass spectrometry, which may easily be scaled up for multiple sample processing.

Here, we present a protocol to determine protease specificity in crude mouse tissue extracts using MALDI-TOF spectrometry.

Proteases have several biological functions, including protein activation/inactivation and food digestion. Identifying protease specificity is important ...

A Hydrogen-Deuterium Exchange Mass Spectrometry (HDX-MS) Platform for Investigating Peptide Biosynthetic Enzymes

Hydrogen-deuterium exchange mass spectrometry (HDX-MS) is a powerful method for the biophysical characterization of enzyme conformational changes and enzyme-substrate interactions. Among its many benefits, HDX-MS consumes only small amounts of material, can be performed under near native conditions without the need for enzyme/substrate labeling, and can provide spatially resolved information on enzyme conformational dynamics&#8722;even for large enzymes and multiprotein complexes. The method is initiated by the dilution of the enzyme of interest into buffer prepared in D2O. This triggers the exchange of protium in peptide bond amides (N-H) with deuterium (N-D). At the desired exchange time points, reaction aliquots are quenched, the enzyme is proteolyzed into peptides, the peptides are separated by ultra-performance liquid chromatography (UPLC), and the change in mass of each peptide (due to the exchange of hydrogen for deuterium) is recorded by MS. The amount of deuterium uptake by each peptide is strongly dependent on the local hydrogen bonding environment of that peptide. Peptides present in very dynamic regions of the enzyme exchange deuterium very rapidly, whereas peptides derived from well-ordered regions undergo exchange much more slowly. In this manner, the HDX rate reports on local enzyme conformational dynamics. Perturbations to deuterium uptake levels in the presence of different ligands can then be used to map ligand binding sites, identify allosteric networks, and to understand the role of conformational dynamics in enzyme function. Here, we illustrate how we have used HDX-MS to better understand the biosynthesis of a type of peptide natural products called lanthipeptides. Lanthipeptides are genetically encoded peptides that are post-translationally modified by large, multifunctional, conformationally dynamic enzymes that are difficult to study with traditional structural biology approaches. HDX-MS provides an ideal and adaptable platform for investigating the mechanistic properties of these types of enzymes.

A Hydrogen-Deuterium Exchange Mass Spectrometry HDX-MS Platform for Investigating Peptide Biosynthetic Enzymes

Lanthipeptide synthetases catalyze multistep reactions during the biosynthesis of peptide natural products. Here, we describe a continuous, bottom-up, hydrogen-deuterium exchange mass spectrometry (HDX-MS) workflow that can be employed to study the conformational dynamics of lanthipeptide synthetases, as well as other similar enzymes involved in peptide natural product biosynthesis.

Hydrogen-deuterium exchange mass spectrometry (HDX-MS) is a powerful method for the biophysical characterization of enzyme conformational changes and ...

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 ...

Millisecond Hydrogen/Deuterium-Exchange Mass Spectrometry for the Study of Alpha-Synuclein Structural Dynamics Under Physiological Conditions

Alpha-synuclein (aSyn) is an intrinsically disordered protein whose fibrillar aggregates are abundant in Lewy bodies and neurites, which are the hallmarks of Parkinson's disease. Yet, much of its biological activity, as well as its aggregation, centrally involves the soluble monomer form of the protein. Elucidation of the molecular mechanisms of aSyn biology and pathophysiology requires structurally highly resolved methods and is sensitive to biological conditions. Its natively unfolded, meta-stable structures make monomeric aSyn intractable to many structural biology techniques. Here, the application of one such approach is described: hydrogen/deuterium-exchange mass spectrometry (HDX-MS) on the millisecond timescale for the study of proteins with low thermodynamic stability and weak protection factors, such as aSyn. At the millisecond timescale, HDX-MS data contain information on the solvent accessibility and hydrogen-bonded structure of aSyn, which are lost at longer labeling times, ultimately yielding structural resolution up to the amino acid level. Therefore, HDX-MS can provide information at high structural and temporal resolutions on conformational dynamics and thermodynamics, intra- and inter-molecular interactions, and the structural impact of mutations or alterations to environmental conditions. While broadly applicable, it is demonstrated how to acquire, analyze, and interpret millisecond HDX-MS measurements in monomeric aSyn.

The structural ensemble of monomeric alpha-synuclein affects its physiological function and physicochemical properties. The present protocol describes how to perform millisecond hydrogen/deuterium-exchange mass spectrometry and subsequent data analyses to determine conformational information on the monomer of this intrinsically disordered protein under physiological conditions.

Alpha-synuclein (aSyn) is an intrinsically disordered protein whose fibrillar aggregates are abundant in Lewy bodies and neurites, which are the hallmarks ...

Defining Hsp33's Redox-regulated Chaperone Activity and Mapping Conformational Changes on Hsp33 Using Hydrogen-deuterium Exchange Mass Spectrometry

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