NS is funded by the University Council Diamond Jubilee Scholarship. JJP is supported by a UKRI Future Leaders Fellowship [Grant number: MR/T02223X/1].

1. Protein expression and purification of aSyn
<ol>
	<li>Prepare aSyn following a previously published report9.</li>
	<li>Dialyze into a safe storage buffer (e.g., Tris, pH 7.2 ).</li>
	<li>If required, concentrate the sample (e.g., spin filter microcentrifuge tubes using 3 kDa MWCO, 14,000 x g for approximately 10-30 min, see Table of Materials). 
	NOTE: It is advised not to concentrate excessively. The integrity of the monomer ensemble has not been verified beyond 25 &#181;M.</li>
	<li>Aliquot and store at &#8722;80 &#176;C 
	&#8203;NOTE: The aSyn monomer protein is stable for up to 1 year in these storage conditions.</li>
</ol>
2. HDX buffer preparation
NOTE: Since Tris has a high temperature coefficient, the pH measurement needs to be adjusted for the temperature at which the HDX reaction will be done, which is 20 &#176;C in this protocol.
<ol>
	<li>Prepare equilibrium buffer for State A and State B by weighing 0.002 mol of Tris into 100 mL of LC-MS grade water. For State B, add 29.8 mg of KCl, 14.2 mg of MgCl2, 36.8 g of CaCl2, and 836 mg of NaCl to the Tris buffer. Adjust the pH to 7.40 &#177; 0.05. 
	NOTE: The equilibrium buffer must contain the conditions at which aSyn is to be studied. In this case, it is 20 mM Tris at pH 7.4 +/&#8722; salts.</li>
	<li>Prepare labeling buffer for State A and State B by weighing 0.002 mol of Tris into 100 mL of deuterated water. For State B, add 29.8 mg of KCl, 14.2 mg of MgCl2, 36.8 g of CaCl2,&#160;and 836 mg of NaCl to the Tris labeling buffer. The pD of the labeling buffer corresponds to the pH of the equilibrium buffer. Since pH = pD - 0.41, adjust so that the pH meter reads 6.99 &#177; 0.0523,24. 
	NOTE: The labeling buffer needs to have the same components as the equilibrium buffer, except that it is prepared using deuterated water.</li>
	<li>Prepare quench buffer by weighing out 0.010 mol of Tris and 0.050 mol of urea and make up to 100 mL with LC-MS grade water. Adjust the pH to 2.50 &#177; 0.05 at 0.5 &#176;C. 
	NOTE: A quench buffer screen must be performed prior to HDX experiments to identify the best quench buffer for the protein of interest. Different concentrations and combinations of denaturants (e.g., urea and guanidinium hydrochloride) and reducing agents (e.g., tris(2-carboxyethyl)phosphine) are screened, along with physical parameters such as trapping volume and temperature, to effectively unfold and digest the quenched protein. A quench buffer comprising 100 mM Tris and 0.5 M urea at pH 2.50 is optimal for the present study.</li>
	<li>Prepare digestion column wash buffer by weighing 0.125 mol of guanidinium hydrochloride into a Duran glass bottle. Add 25 mL of methanol and 250 &#181;L of formic acid. Make up to 250 mL with LC-MS grade water. 
	NOTE: For the Enzymate BEH pepsin column (see Table of Materials), use a column wash buffer of 0.5 M guanidinium hydrochloride, 10% (v/v) methanol, and 0.1% (v/v) formic acid.</li>
	<li>Prepare the syringe weak wash by pipetting 0.5 &#181;L of formic acid into 249.5 mL of LC-MS grade water.</li>
	<li>Prepare the syringe strong wash by mixing equal parts of LC-MS grade water, methanol, acetonitrile, and isopropanol. Add formic acid to a final 2% (v/v) concentration. 
	&#8203;NOTE: To prevent cross-contamination between the various buffers and the protein and to enable cleaning of the injection port, it is critical that syringe wash solutions are prepared and that the flow path to the valve (often named the &#34;wash liner&#34;) is fully primed with liquid. The formic acid is optional for the syringe weak wash.</li>
</ol>
3. Peptide mapping procedure
<ol>
	<li>Prepare the sample following the step below.
	<ol>
		<li>Filter thawed aSyn protein stock from the &#8722;80 &#176;C freezer with 0.22 &#181;m syringe filters. Measure the absorbance of the filtered stock protein at 280 nm to determine the concentration by the Beer-Lambert law. Dilute the protein to a concentration of 5 &#181;M in the equilibrium buffer (step 2.1.). 
		NOTE: The Beer-Lambert law: A = &#949;cl, where A is absorbance, &#949; is the extinction coefficient of the protein at the measured wavelength (280 nm here) with units M&#8722;1cm&#8722;1, c is the protein concentration in M, and l is the path length in cm. For wild-type aSyn26, &#949; = 5960 M&#8722;1cm&#8722;1.</li>
	</ol>
	</li>
	<li>Set up the liquid chromatography method.
	<ol>
		<li>Create an inlet file with a loading/trapping time of 3 min at a pressure of 7000-9000 psi, followed by a gradient of 5% acetonitrile to 40% acetonitrile in 7 min, followed by repeated washing steps of 5%-95% acetonitrile-water for 10 min.</li>
		<li>Ensure lock spray (e.g., leucine enkephalin, see Table of Materials) is flowing at 2000 psi and connected to the source lock spray probe of the mass spectrometer.</li>
	</ol>
	</li>
	<li>Set up the mass spectrometry MSE methods. 
	NOTE: MSE is a broadband data-independent acquisition method with no precursor mass isolation. Therefore, all ions within the selected m/z range are fragmented further using collision-induced dissociation (CID)27.
	<ol>
		<li>In the MS method file, choose MSE Continuum and set up an acquisition time between 2-10 min, electrospray source, and positive resolution mode. Acquire MSE over 50-2000 Da, scanning every 0.3 s.</li>
		<li>For function 1 (low energy), set up the trap and transfer collision energies to be 4 V. For function 2 (high energy), set up the ramp transfer collision energy to be constant at 4V and the trap collision energy to be as stated in Table 1 for each mapping energy level. 
		NOTE: Ion-mobility MSE methods can also be used. Alternative mapping methods (e.g., data-dependent acquisition or DDA) can be used at user discretion.</li>
	</ol>
	</li>
	<li>Set up the autosampler robot (see Table of Materials).
	<ol>
		<li>Add 50 &#181;L of the 5 &#181;M protein to a total recovery vial. Place the vial in a sample position in the HDX right chamber. Ensure this chamber is at 0.5 &#176;C.</li>
		<li>Add one reagent vial of equilibrium buffer and two reagent vials of labeling buffer to reagent positions one, two, and three in the HDX left chamber. Ensure this chamber is at 20 &#176;C by setting the Peltier temperature controller (see Table of Materials). Add one reagent vial of quench buffer to reagent position one in the HDX right chamber.</li>
		<li>Add eight total recovery vials in the reaction positions of the HDX left chamber and eight maximum recovery vials in the reaction positions of the HDX right chamber. 
		NOTE: To ensure full cleaning and maximum reproducibility of dispensed volumes, it is advised to perform a syringe wash and prime wash on the autosampler syringes. For example, execute a sequence of (1) weak wash, (2) strong wash, (3) weak wash prior to starting the mapping experiments. This sequence can be repeated extensively, and it is recommended to do it up to 20x or until the syringe is fully wetted.</li>
		<li>Set up a sample list with appropriate LC and MS methods in the scheduling software and start the schedule. 
		NOTE: For the present study, Chronos is used as the scheduling software (see Table of Materials).</li>
	</ol>
	</li>
	<li>Process the mapping data.
	<ol>
		<li>Identify peptides from the mapping experiment files using appropriate software (see Table of Materials).</li>
		<li>Import peptide identification data into DynamX (see Table of Materials) using the following peptide threshold parameters: minimum intensity = 5000, minimum sequence length = 0, maximum sequence length = 40, minimum products = 1, minimum products per amino acid = 0.25, minimum consecutive products = 2, minimum sum intensity for products = 0, minimum score = 0, and maximum MH+ Error (ppm) = 0. 
		NOTE: Alternatively, derive the optimized settings according to a recommended workflow28.</li>
		<li>Select Data from the menu and click on Import PLGS Results. Click on Add to choose the relevant data files for the spectral assignment. When all have been added, click on Next and enter the above filter settings. Then, click on Finish.</li>
		<li>Manually curate isotopic assignments to obtain the final peptide coverage map of aSyn in DynamX.</li>
	</ol>
	</li>
</ol>
4. Millisecond hydrogen/deuterium exchange study
<ol>
	<li>Clean the FastHDX prototype instrument (see Table of Materials) before starting HDX experiments.
	<ol>
		<li>Open the compatible HDX software GUI and allow the system to initialize.</li>
		<li>Enter the Sample Chamber temperature as 20 &#176;C and the Quench Chamber as 0.5 &#176;C. Click on Set to apply the new temperatures.</li>
		<li>Set up the centrifuge tubes with LC-MS grade water at all inlets.</li>
		<li>In the Titrator Plumbing Delivery tab, check both left and right syringes and click on Prime to remove any latent air bubbles in the tubes. Repeat until all bubbles are gone.</li>
		<li>In the Macros tab, check all the boxes for the syringes. Click on Calibrate Syringes Home Position. Click on Wash Syringe Load Loop. Click on Wash All Mixing Loop Volumes.</li>
		<li>Repeat step 4.1.5. 1x more.</li>
		<li>If any bubbles appear in the buffer syringes, degas by disconnecting the syringe and ejecting the bubble vertically. Replace the syringe and recalibrate to the zero position.</li>
	</ol>
	</li>
	<li>Set up the FastHDX prototype instrument for HDX experiments.
	<ol>
		<li>Add 500 uL of filtered 5 &#181;M aSyn in a total recovery vial and place inside a tabletop fridge to prevent temperature-induced oligomerization and aggregation.</li>
		<li>Add 50 mL of equilibrium, labeling, and quench buffers to each buffer (2. HDX buffer preparation steps 1-3) inlet in the left and right chambers.</li>
		<li>Add 50 mL of column wash buffer (2. HDX buffer preparation step 4) to the pepsin wash inlet.</li>
		<li>To prime the protein and column wash lines 1x, check both left and right syringes in the Titrator Plumbing Delivery tab and click on Prime&#160;once. 
		NOTE: Any subsequent click on Prime will cause additional repeats of the prime process, resulting in the consumption of large amounts of protein sample. Care is advised to avoid this, even when there is a delay in the software after a button click has been attempted.</li>
		<li>Repeat steps 4.1.5. 1x.</li>
		<li>In the Manual Quench Flow tab, enter the required settings, explained in steps 4.2.7.-4.2.10.</li>
		<li>For a time-course experiment, enter the times in milliseconds using the Symbolic dots button. If replicates are required, add the same timepoint multiple times, e.g., for a triplicate of 50 ms, 50 50 50 needs to be entered. The sample list in the mass spectrometer software (MassLynx is used here, see Table of Materials) needs to match those timepoints exactly. 
		NOTE: The mass spectrometer sample list file names and/or sample text should be used to ensure a permanent record of the HDX labeling times corresponding to those entered in the FastHDX software GUI. Labeling times for each sample run will not be stored anywhere else.</li>
		<li>Set Trap Time (mins) to be the length of the trapping time. Here, it is 3.00.</li>
		<li>Set Wait for HPLC (mins) = (trap time + run time + 1.5 min). 
		NOTE: For example, for an experiment with 3 min trapping and 17 min gradient, this will be 21.50 min.</li>
		<li>Click on the Run Blank box only if running blank experiments between sample runs. If yes, ensure an entry (i.e., a valid row on the sample list) in the software for the blank run after each sample run.</li>
		<li>Once the sample list is ready on the software, highlight the appropriate entries and start the run in the software by clicking on the Play button and FastHDX in the software. 
		&#8203;NOTE: Due to hydrogen/deuterium scrambling, MS-only methods or soft-fragmentation techniques (electron transfer dissociation, electron capture dissociation, and ultraviolet photo-dissociation) can be used only29. For aSyn at a physiological pH of 7.40, timepoints ranging from 50 ms to 300 s are most applicable as these cover the entire deuterium uptake curve8.</li>
	</ol>
	</li>
</ol>
5. Data processing
<ol>
	<li>Load the file of spectrally assigned peptides from the peptide mapping experiments. Open the File menu and click on Open in the DynamX software (see Table of Materials).</li>
	<li>Import raw files into the preferred mass measurement software (e.g., DynamX, HDExaminer, etc.). Open the &#34;Data&#34; menu and click on MS Files. Click on New State to create states for each protein condition studied.
	<ol>
		<li>Click&#160;on New Exposure to add each HDX timepoint. Click on New RAW to import the .raw files. Drag each .raw file to the correct position. Click on OK when done.</li>
	</ol>
	</li>
	<li>Automatically assign isotopes first (this is automatic following step 5.2. above), then manually curate the isotopic assignment to ensure high data quality.</li>
	<li>Export the cluster data into a .csv file with columns in the following order: protein name, sequence start number, sequence end number, sequence, modification, fragment, maximum possible uptake, mass of monoisotopic species, state name, exposure time, file name, charge, retention time, intensity, and centroid.</li>
	<li>Open the Data menu in mass measurement software and click on Export Cluster Data.</li>
</ol>
6. Data analysis
<ol>
	<li>Load the exported cluster data into the preferred HDX analysis software. Here, HDfleX is used30 (see Table of Materials).</li>
	<li>Fit the experimental data for all peptides and states, selecting the appropriate back-exchange correction methods to get observed rate constants for the HDX reaction.</li>
	<li>Calculate a global significance threshold by the preferred method (HDfleX supports several options for this) and perform hybrid significance testing to determine the significant differences across the states compared31,32. 
	NOTE: If the difference observed is greater than the global threshold and the p-value is less than the chosen confidence level (e.g., 95%), the difference is considered significant.</li>
</ol>

The authors declare no competing interests.

In the present article, the following procedures are described: (1) performing peptide mapping experiments on monomeric aSyn to obtain the highest sequence coverage, (2) acquiring millisecond HDX-MS data on monomeric aSyn under physiological conditions, and (3) performing data analysis and interpretation of the resulting HDX-MS data. The provided procedures are generally simple to execute, each labeling experiment typically lasts only around 8 h for three replicates and eight timepoints, and the mapping experiment lasts ...

Parkinson&#39;s disease (PD) is a neurodegenerative illness affecting millions of people worldwide1. It is characterized by the formation of cytoplasmic inclusions known as Lewy bodies and Lewy neurites in the brain&#39;s substantia nigra pars compacta region. These cytoplasmic inclusions have been found to contain aggregates of the intrinsically disordered protein aSyn2. In PD and other synucleinopathies, aSyn transforms from a soluble disordered state into an insoluble, highly structured diseased state. In its native form, monomeric aSyn adopts a wide range of conformations stabilized by long-range electrostatic interactions between its N- and C-termini and hydrophobic interactions between its C-terminus and non-amyloid beta component (NAC) region3,4,5,6. Any disruptions in those stabilizing interactions, such as mutations, post-translational modifications, and changes in the local environment, can lead to the misfolding of the monomer, thus triggering the process of aggregation7.
While a vast amount of research exists on the oligomeric and fibrillar forms of aSyn8,9,10,11, there is a crucial need to study the monomeric form of the protein and better understand which conformers are functional (and how) and which are prone to aggregate8,9,10,11. Being intrinsically disordered, only 14 kDa in size, and difficult to crystallize, the aSyn monomer is not amenable to most structural biological techniques. However, one technique capable of measuring the conformational dynamics of monomeric aSyn is millisecond HDX-MS, which has recently generated important structural observations that would be challenging or impossible to obtain otherwise12,13,14. Millisecond HDX-MS sensitively measures the average of the protein conformational ensemble by monitoring the isotopic exchange at amide hydrogens, indicating solvent accessibility and hydrogen-bonding network participation of a particular protein region on the millisecond timescale. It is necessary to stress the millisecond aspect of the HDX-MS as, due to its natively unfolded, meta-stable nature, aSyn exhibits very fast hydrogen-exchange kinetics that manifest well below the lower limit of conventional HDX-MS systems. For example, most of the aSyn molecule has completely exchanged hydrogen for deuterium under intracellular conditions in less than 1 s. Several laboratories have now built fast-mixing instrumentation; in this case, a prototype fast-mixing quench-flow instrument capable of performing HDX-MS with a dead-time of 50 ms and a temporal resolution of 1 ms is used15. While millisecond HDX-MS has recently been acutely important in the study of aSyn, it stands to be valuable in studying intrinsically disordered proteins/regions more widely and a large number of proteins with loops/regions that are only weakly stable. For example, peptide drugs (e.g., insulin; GLP-1/glucagon; tirzepatide) and peptide-fusion proteins (e.g., the HIV inhibitor FN3-L35-T1144) are major drug formats where solution-phase structural and stability information can be a critical input for drug development decisions, and, yet, the peptide moiety is often only weakly stable and intractable by HDX-MS at the seconds timescale16,17,18,19,20. Emergent HDX-MS methods with labeling in the seconds/minutes domains have been shown to derive structural information for DNA G-quadruplexes, but it should be possible to extend this to more diverse oligonucleotide structures by the application of millisecond HDX-MS21.
HDX-MS experiments can be performed at three different levels: (1) bottom-up (whereby the labeled protein is digested proteolytically), (2) middle-down (whereby the labeled protein is digested proteolytically, and the resulting peptides are fragmented further by soft-fragmentation techniques), and (3) top-down (whereby soft-fragmentation techniques directly fragment the labeled protein)22. Thus, sub-molecular HDX-MS data allow us to localize the exchange behavior to specific regions of a protein, making it critical to have adequate sequence coverage for such experiments. The structural resolution of any HDX-MS experiment relies on the number of proteolytic peptides or fragments derived from the protein upon digestion or soft-fragmentation, respectively. In each of the three experiment types outlined above, the change in amide exchange at each peptide/fragment is mapped back onto the protein&#39;s primary structure to indicate the behavior of localized regions of the protein. While the highest structural resolution is achieved through soft-fragmentation, the description of these experiments is out of the scope of the current study, which focuses on the measurement of aSyn monomer conformations. Excellent results can be obtained with the commonly applied &#34;bottom-up&#34; workflow described here.
Here, procedures are provided on (1) how to prepare and handle aSyn samples and HDX-MS buffers, (2) how to perform peptide mapping for a bottom-up HDX-MS experiment, (3) how to acquire HDX-MS data on monomeric aSyn under physiological conditions, specifically in the millisecond time domain (using a custom-built instrument; alternative instruments for millisecond labeling have also been described), and (4) how to process and analyze the HDX-MS data. Methods using monomeric aSyn at physiological pH (7.40) in two solution conditions are exemplified here. While critically useful in the study of aSyn, these procedures can be applied to any protein and are not limited to intrinsically disordered proteins.

<table><tbody><tr><td>1 &amp;times; 100 mm ACQUITY BEH 1.7 &amp;mu;m C18 column&amp;nbsp;</td><td>Waters Corporation</td><td>186002346</td><td>Analytical column</td></tr><tr><td>Acetonitrile HPLC grade &gt;99.9% HiPerSolv</td><td>VWR</td><td>20060.420</td><td>For LC mobile phases</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma Aldrich</td><td>C5670</td><td>Salt for HDX buffers</td></tr><tr><td>Chronos</td><td>Axel Semrau (Purchased from Waters Corporation)</td><td>667006090</td><td>Scheduling software to enable multiple HDX-MS sample injections automatically. Alternative software is available from other vendors e.g. HDXDirector or LEAP Shell</td></tr><tr><td>Deuterium chloride</td><td>Goss Scientific (Cambridge Isotope Laboratories)</td><td>DLM-2-50</td><td>For HDX labelling buffers</td></tr><tr><td>Deuterium oxide (99.9% D&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Goss Scientific (Cambridge Isotope Laboratories)</td><td>DLM-4</td><td>Deuterated water</td></tr><tr><td>DynamX 3.0</td><td>Waters Corporation</td><td>176016027</td><td>Isotopic assignment and deuterium incorporation calculation</td></tr><tr><td>Enzymate BEH Pepsin Column</td><td>Waters Corporation</td><td>186007233</td><td>Pepsin digestion column</td></tr><tr><td>Formic Acid, 99.0% LC/MS Grade</td><td>Fisher Scientific</td><td>10596814</td><td>For LC mobile phases</td></tr><tr><td>Guanidinium hydrochloride</td><td>Sigma Aldrich</td><td>RDD001-500G</td><td>Chaotrope/Denaturant</td></tr><tr><td>HDfleX</td><td>University of Exeter</td><td>N/A</td><td>https://ore.exeter.ac.uk/repository/handle/10871/127982</td></tr><tr><td>KCl</td><td>Sigma Aldrich</td><td>P3911</td><td>Salt for HDX buffers</td></tr><tr><td>LEAP HDX-2 CTC PAL sampling robot</td><td>Waters Corporation</td><td>725000637</td><td>Autosampler robot</td></tr><tr><td>Leucine enkephalin</td><td>Waters Corporation</td><td>186006013</td><td>For mass spectrometry lockspray calibration.</td></tr><tr><td>MassLynx</td><td>Waters Corporation</td><td>667004007</td><td>Software controlling inlet methods and mass spectrometer</td></tr><tr><td>Maximum recovery vials</td><td>Waters Corporation</td><td>600000670CV</td><td>100 pack including caps - used for quench tray in LEAP HDX-2</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma Aldrich</td><td>M8266</td><td>Salt for HDX buffers</td></tr><tr><td>Millipore 0.22 &amp;micro;m syringe filters</td><td>Millipore</td><td>N9CA7069B</td><td>Syringe filters</td></tr><tr><td>ms2min</td><td>Applied Photophysics Ltd</td><td>N/A</td><td>fast-mix quench-flow millisecond hdx instrument</td></tr><tr><td>NaCl</td><td>Sigma Aldrich</td><td>S9888</td><td>Salt for HDX buffers</td></tr><tr><td>Peltier temperature controller</td><td>LEAP Technologies Inc.</td><td>HP115-COOL/D</td><td>Peltier controller to set precise temperature of chambers in the LEAP robot.</td></tr><tr><td>ProteinLynx Global Server 3.0</td><td>Waters Corporation</td><td>715001030</td><td>Peptide identification software. Alternative software is available from other vendors.</td></tr><tr><td>Reagent pot caps</td><td>Waters Corporation</td><td>186004632</td><td>100 pack</td></tr><tr><td>Reagent pots for LEAP HDX-2</td><td>Waters Corporation</td><td>186001420</td><td>100 pack excluding caps - used for buffers in LEAP HDX-2</td></tr><tr><td>Sodium deuteroxide (99.5% in D&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Goss Scientific (Cambridge Isotope Laboratories)</td><td>DLM-57</td><td>For HDX labelling buffers</td></tr><tr><td>Spin filter microcentrifuge tubes (3 kDa MWCO)</td><td>Amicon (Merck Sigma Aldrich)</td><td>UFC5003</td><td>Micro centrifuge tubes to concentrate protein. This facilitates buffer exchange and accurate sample loading for HDX-MS experiments.</td></tr><tr><td>Synapt G2-Si mass spectrometer</td><td>Waters Corporation</td><td>176850035</td><td>Mass spectrometer</td></tr><tr><td>Total recovery vials</td><td>Waters Corporation</td><td>600000671CV</td><td>100 pack including caps - used for labelling tray in LEAP HDX-2</td></tr><tr><td>Tris-HCl</td><td>Sigma Aldrich</td><td>T3253-250G</td><td>Buffer</td></tr><tr><td>Trizma base</td><td>Sigma Aldrich</td><td>T60040-B2005</td><td>Buffer</td></tr><tr><td>Urea</td><td>Sigma Aldrich</td><td>U5378-1KG</td><td>Chaotrope/Denaturant</td></tr><tr><td>VanGuard 2.1 x 5 mm ACQUITY BEH C18 column&amp;nbsp;</td><td>Waters Corporation</td><td>186004623</td><td>Trap desalting column</td></tr></tbody></table>

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.

Due to its intrinsically disordered nature, it is difficult to capture the intricate structural changes in aSyn at physiological pH. HDX-MS monitors isotopic exchange at backbone amide hydrogens, probing the protein conformational dynamics and interactions. It is one of the few techniques to acquire this information at high structural and temporal resolutions. This protocol is broadly applicable to a wide range of proteins and buffer conditions, and this is exemplified by the measurement of the exchange kinetics of aSyn ...

Watch this Scientific Journal Video about Millisecond Hydrogen/Deuterium-Exchange Mass Spectrometry for the Study of Alpha-Synuclein Structural Dynamics Under Physiological Conditions at JoVE.com

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

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 ...

millisecond-hydrogendeuterium-exchange-mass-spectrometry-for-study

Research

JoVE Journal

Neuroscience

2.8K Views.  University of Exeter. Alle organismen hebben een wanorde in hun proteoom, geschat tot 33% Deze regio's zijn waarneembaar door waterstof / deuterium uitwisseling massaspectrometrie, maar alleen in het milliseconde tijdsbereik. Ons protocol laat u zien hoe u dit met succes kunt doen. Dit is een volledig geautomatiseerde analysetechniek.Je stelt gewoon alles in, programmeert de gewenste tijdmetingen, drukt op go en loopt weg. Deze techniek opent de opwindende mogelijkheid om te screenen op verbindingen die spe...

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

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. ...

Biochemistry

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

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

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

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

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 ...

Utilizing Time-Resolved Protein-Induced Fluorescence Enhancement to Identify Stable Local Conformations One &#945;-Synuclein Monomer at a Time

Using spectroscopic rulers to track multiple conformations of single biomolecules and their dynamics have revolutionized the understanding of structural dynamics and its contributions to biology. While the FRET-based ruler reports on inter-dye distances in the 3-10 nm range, other spectroscopic techniques, such as protein-induced fluorescence enhancement (PIFE), report on the proximity between a dye and a protein surface in the shorter 0-3 nm range. Regardless of the method of choice, its use in measuring freely-diffusing biomolecules one at a time retrieves histograms of the experimental parameter yielding separate centrally-distributed sub-populations of biomolecules, where each sub-population represents either a single conformation that stayed unchanged within milliseconds, or multiple conformations that interconvert much faster than milliseconds, and hence an averaged-out sub-population. In single-molecule FRET, where the reported parameter in histograms is the inter-dye FRET efficiency, an intrinsically disordered protein, such as the &#945;-Synuclein monomer in buffer, was previously reported as exhibiting a single averaged-out sub-population of multiple conformations interconverting rapidly. While these past findings depend on the 3-10 nm range of the FRET-based ruler, we sought to put this protein to the test using single-molecule PIFE, where we track the fluorescence lifetime of site-specific sCy3-labeled &#945;-Synuclein proteins one at a time. Interestingly, using this shorter range spectroscopic proximity sensor, sCy3-labeled &#945;-Synuclein exhibits several lifetime sub-populations with distinctly different mean lifetimes that interconvert in 10-100 ms. These results show that while &#945;-Synuclein might be disordered globally, it nonetheless attains stable local structures. In summary, in this work we highlight the advantage of using different spectroscopic proximity sensors that track local or global structural changes one biomolecule at a time.

Time-resolved single-molecule protein-induced fluorescence enhancement is a useful fluorescence spectroscopic proximity sensor sensitive to local structural changes in proteins. Here we show it can be used to uncover stable local conformations in &#945;-Synuclein, which is otherwise known as globularly unstructured and unstable when measured using the longer range FRET ruler.

Using spectroscopic rulers to track multiple conformations of single biomolecules and their dynamics have revolutionized the understanding of structural ...

High-Resolution Neutron Spectroscopy to Study Picosecond-Nanosecond Dynamics of Proteins and Hydration Water

Neutron scattering offers the possibility to probe the dynamics within samples for a wide range of energies in a nondestructive manner and without labeling other than deuterium. In particular, neutron backscattering spectroscopy records the scattering signals at multiple scattering angles simultaneously and is well suited to study the dynamics of biological systems on the ps-ns timescale. By employing D2O-and possibly deuterated buffer components-the method allows monitoring of both center-of-mass diffusion and backbone and side-chain motions (internal dynamics) of proteins in liquid state.
Additionally, hydration water dynamics can be studied by employing powders of perdeuterated proteins hydrated with H2O. This paper presents the workflow employed on the instrument IN16B at the Institut Laue-Langevin (ILL) to investigate protein and hydration water dynamics. The preparation of solution samples and hydrated protein powder samples using vapor exchange is explained. The data analysis procedure for both protein and hydration water dynamics is described for different types of datasets (quasielastic spectra or fixed-window scans) that can be obtained on a neutron backscattering spectrometer.
The method is illustrated with two studies involving amyloid proteins. The aggregation of lysozyme into &#181;m sized spherical aggregates-denoted particulates-is shown to occur in a one-step process on the space and time range probed on IN16B, while the internal dynamics remains unchanged. Further, the dynamics of hydration water of tau was studied on hydrated powders of perdeuterated protein. It is shown that translational motions of water are activated upon the formation of amyloid fibers. Finally, critical steps in the protocol are discussed as to how neutron scattering is positioned regarding the study of dynamics with respect to other experimental biophysical methods.

Neutron backscattering spectroscopy offers a nondestructive and label-free access to the ps-ns dynamics of proteins and their hydration water. The workflow is presented with two studies on amyloid proteins: on the time-resolved dynamics of lysozyme during aggregation and on the hydration water dynamics of tau upon fiber formation.

Neutron scattering offers the possibility to probe the dynamics within samples for a wide range of energies in a nondestructive manner and without ...

Detection of Disease-associated &#945;-synuclein by Enhanced ELISA in the Brain of Transgenic Mice Overexpressing Human A53T Mutated &#945;-synuclein

In addition to established methods like Western blot, new methods are needed to quickly and easily quantify disease-associated &#945;-synuclein (&#945;SD) in experimental models of synucleopathies. A transgenic mouse line (M83) over-expressing the human A53T &#945;S and spontaneously developing a dramatic clinical phenotype between eight and 22 months of age, characterized by symptoms including weight loss, prostration, and severe motor impairment, was used in this study. For molecular analyses of &#945;SD (disease-associated &#945;S) in these mice, an ELISA was designed to specifically quantify &#945;SD in sick mice. Analysis of the central nervous system in this mouse model showed the presence of &#945;SD mainly in the caudal brain regions and the spinal cord. There were no differences in &#945;SD distribution between different experimental conditions leading to clinical disease, i.e., in uninoculated and normally aging transgenic mice and in mice inoculated with brain extracts from sick mice. The specific detection of &#945;SD immunoreactivity using an antibody against Ser129 phosphorylated &#945;S by ELISA essentially correlated with that obtained by Western blot and immunohistochemistry. Unexpectedly, similar results were observed with several other antibodies against the C-terminal part of &#945;S. The propagation of &#945;SD, suggesting the involvement of a &#8220;prion-like&#8221; mechanism, can thus be easily monitored and quantified in this mouse model using an ELISA approach.

An ELISA offering a novel quantitative approach is described. It specifically detects disease-associated &#945;-synuclein (&#945;SD) in a transgenic mouse model (M83) of synucleinopathy using several antibodies against either the Ser129 phosphorylated &#945;S form or the C-terminal part of the protein.

In addition to established methods like Western blot, new methods are needed to quickly and easily quantify disease-associated &#945;-synuclein (&#945;SD) ...

Medicine

Sequential Extraction of Soluble and Insoluble Alpha-Synuclein from Parkinsonian Brains

Alpha-synuclein (&#945;-syn) protein is abundantly expressed mainly within neurons, and exists in a number of different forms - monomers, tetramers, oligomers and fibrils. During disease, &#945;-syn undergoes conformational changes to form oligomers and high molecular weight aggregates that tend to make the protein more insoluble. Abnormally aggregated &#945;-syn is a neuropathological feature of Parkinson&#39;s disease (PD), dementia with Lewy bodies (DLB) and multiple system atrophy (MSA). Biochemical characterization and analysis of insoluble &#945;-syn using buffers with increasing detergent strength and high-speed ultracentrifugation provides a powerful tool to determine the development of &#945;-syn pathology associated with disease progression. This protocol describes the isolation of increasingly insoluble/aggregated &#945;-syn from post-mortem human brain tissue. This methodology can be adapted with modifications to studies of normal and abnormal &#945;-syn biology in transgenic animal models harbouring different &#945;-syn mutations as well as in other neurodegenerative diseases that feature aberrant fibrillar deposits of proteins related to their respective pathologies.

Here, we present a protocol for the isolation of increasingly insoluble/aggregated alpha-synuclein (&#945;-syn) from post-mortem human brain tissue. Through the utilization of buffers with increasing detergent strength and high-speed ultracentrifugation techniques, the variable properties of &#945;-syn aggregation in diseased and non-diseased tissue can be examined.

Alpha-synuclein (&#945;-syn) protein is abundantly expressed mainly within neurons, and exists in a number of different forms - monomers, tetramers, ...

Exogenous Administration of Microsomes-associated Alpha-synuclein Aggregates to Primary Neurons As a Powerful Cell Model of Fibrils Formation

For years, the inability of replicating formation of insoluble alpha-synuclein (&#945;S) inclusions in cell cultures has been a great limitation in the study of &#945;S aggregation in Parkinson&#39;s Disease (PD). Recently, the development of new animal models through the exogenous inoculation of brain extracts from diseased &#945;S transgenic mice or PD patients has given new hopes to the possibility of creating more adequate cell models of &#945;S aggregation. Unfortunately, when it comes to cells in cultures, administration of raw brain extracts has not proven as successful as in mice and the source of choice of exogenous aggregates is still in vitro preformed &#945;S fibrils.
We have developed a method to induce the formation of intracellular &#945;S inclusions in primary neurons through the exogenous administration of native microsomes-associated &#945;S aggregates, a highly toxic &#945;S species isolated from diseased areas of transgenic mice. This fraction of &#945;S aggregates that is associated with the microsomes vesicles, is efficiently internalized and induces the formation of intracellular inclusions positive for aggregated and phosphorylated &#945;S. Compared to in vitro-preformed fibrils which are made from recombinant &#945;S, our method is faster and guarantees that the pathogenic seeding is made with authentic &#945;S aggregates extracted from diseased animal models of PD, mimicking more closely the type of inclusions obtained in vivo. As a result, availability of tissues rich in &#945;S inclusions is mandatory.
We believe that this method will provide a versatile cell-based model to study the microscopic aspects of &#945;S aggregation and the related cellular pathophysiology in vivo and will be a starting point for the creation of more accurate and sophisticated cell paradigm of PD.

The goal of this protocol is to provide a cell-based system that replicates the formation of alpha-synuclein aggregates in vivo. Intracellular alpha-synuclein inclusions are seeded in primary neurons by the internalization and propagation of exogenous administered native microsomes-associated alpha-synuclein aggregates isolated from diseased alpha-synuclein transgenic mice.

For years, the inability of replicating formation of insoluble alpha-synuclein (&#945;S) inclusions in cell cultures has been a great limitation in the ...

Studying Pre-formed Fibril Induced &#945;-Synuclein Accumulation in Primary Embryonic Mouse Midbrain Dopamine Neurons

The goal of this protocol is to establish a robust and reproducible model of &#945;-synuclein accumulation in primary dopamine neurons. Combined with immunostaining and unbiased automated image analysis, this model allows for the analysis of the effects of drugs and genetic manipulations on &#945;-synuclein aggregation in neuronal cultures. Primary midbrain cultures provide a reliable source of bona fide embryonic dopamine neurons. In this protocol, the hallmark histopathology of Parkinson&#8217;s disease, Lewy bodies (LB), is mimicked by the addition of &#945;-synuclein pre-formed fibrils (PFFs) directly to neuronal culture media. Accumulation of endogenous phosphorylated &#945;-synuclein in the soma of dopamine neurons is detected by immunostaining already at 7 days after the PFF addition. In vitro cell culture conditions are also suitable for the application and evaluation of treatments preventing &#945;-synuclein accumulation, such as small molecule drugs and neurotrophic factors, as well as lentivirus vectors for genetic manipulation (e.g., with CRISPR/Cas9). Culturing the neurons in 96 well plates increases the robustness and power of the experimental setups. At the end of the experiment, the cells are fixed with paraformaldehyde for immunocytochemistry and fluorescence microscopy imaging. Multispectral fluorescence images are obtained via automated microscopy of 96 well plates. These data are quantified (e.g., counting the number of phospho-&#945;-synuclein-containing dopamine neurons per well) with the use of free software that provides a platform for unbiased high-content phenotype analysis. PFF-induced modeling of phosphorylated &#945;-synuclein accumulation in primary dopamine neurons provides a reliable tool to study the underlying mechanisms mediating formation and elimination of &#945;-synuclein inclusions, with the opportunity for high-throughput drug screening and cellular phenotype analysis.

Here, we present a detailed protocol to study neuronal &#945;-synuclein accumulation in primary mouse dopamine neurons. Phosphorylated &#945;-synuclein aggregates in neurons are induced with pre-formed &#945;-synuclein fibrils. Automated imaging of immunofluorescently labeled cells and unbiased image analysis make this robust protocol suitable for medium-to-high throughput screening of drugs that inhibit &#945;-synuclein accumulation.

The goal of this protocol is to establish a robust and reproducible model of &#945;-synuclein accumulation in primary dopamine neurons. Combined with ...

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

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