Method Article

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

DOI:

10.3791/64050

⸱

June 23rd, 2022

 , 
1Living Systems Institute, Department of Biosciences, University of Exeter

In This Article

Summary

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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.

Abstract

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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.

Introduction

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Parkinson'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'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'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 "bottom-up" 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.

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Protocol

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1. Protein expression and purification of aSyn

  1. Prepare aSyn following a previously published report9.
  2. Dialyze into a safe storage buffer (e.g., Tris, pH 7.2 ).
  3. 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 µM.
  4. Aliquot and store at −80 °C
    ​NOTE: The aSyn monomer protein is stable for up to 1 year in these storage conditions.

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 °C in this protocol.

  1. 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 ± 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 +/− salts.
  2. 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, 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 ± 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.
  3. 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 ± 0.05 at 0.5 °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.
  4. 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 µL 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.
  5. Prepare the syringe weak wash by pipetting 0.5 µL of formic acid into 249.5 mL of LC-MS grade water.
  6. 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.
    ​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 "wash liner") is fully primed with liquid. The formic acid is optional for the syringe weak wash.

3. Peptide mapping procedure

  1. Prepare the sample following the step below.
    1. Filter thawed aSyn protein stock from the −80 °C freezer with 0.22 µm 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 µM in the equilibrium buffer (step 2.1.).
      NOTE: The Beer-Lambert law: A = εcl, where A is absorbance, ε is the extinction coefficient of the protein at the measured wavelength (280 nm here) with units M−1cm−1, c is the protein concentration in M, and l is the path length in cm. For wild-type aSyn26, ε = 5960 M−1cm−1.
  2. Set up the liquid chromatography method.
    1. 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.
    2. 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.
  3. 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.
    1. 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.
    2. 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.
  4. Set up the autosampler robot (see Table of Materials).
    1. Add 50 µL of the 5 µM 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 °C.
    2. 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 °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.
    3. 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.
    4. 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).
  5. Process the mapping data.
    1. Identify peptides from the mapping experiment files using appropriate software (see Table of Materials).
    2. 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.
    3. 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.
    4. Manually curate isotopic assignments to obtain the final peptide coverage map of aSyn in DynamX.

4. Millisecond hydrogen/deuterium exchange study

  1. Clean the FastHDX prototype instrument (see Table of Materials) before starting HDX experiments.
    1. Open the compatible HDX software GUI and allow the system to initialize.
    2. Enter the Sample Chamber temperature as 20 °C and the Quench Chamber as 0.5 °C. Click on Set to apply the new temperatures.
    3. Set up the centrifuge tubes with LC-MS grade water at all inlets.
    4. 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.
    5. 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.
    6. Repeat step 4.1.5. 1x more.
    7. 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.
  2. Set up the FastHDX prototype instrument for HDX experiments.
    1. Add 500 uL of filtered 5 µM aSyn in a total recovery vial and place inside a tabletop fridge to prevent temperature-induced oligomerization and aggregation.
    2. 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.
    3. Add 50 mL of column wash buffer (2. HDX buffer preparation step 4) to the pepsin wash inlet.
    4. 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 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.
    5. Repeat steps 4.1.5. 1x.
    6. In the Manual Quench Flow tab, enter the required settings, explained in steps 4.2.7.-4.2.10.
    7. 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.
    8. Set Trap Time (mins) to be the length of the trapping time. Here, it is 3.00.
    9. 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.
    10. 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.
    11. 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.
      ​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.

5. Data processing

  1. 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).
  2. Import raw files into the preferred mass measurement software (e.g., DynamX, HDExaminer, etc.). Open the "Data" menu and click on MS Files. Click on New State to create states for each protein condition studied.
    1. Click 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.
  3. Automatically assign isotopes first (this is automatic following step 5.2. above), then manually curate the isotopic assignment to ensure high data quality.
  4. 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.
  5. Open the Data menu in mass measurement software and click on Export Cluster Data.

6. Data analysis

  1. Load the exported cluster data into the preferred HDX analysis software. Here, HDfleX is used30 (see Table of Materials).
  2. 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.
  3. 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.

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Results

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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 ...

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Discussion

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

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 ...

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Disclosures

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

The authors declare no competing interests.

Acknowledgements

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

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
1 × 100 mm ACQUITY BEH 1.7 μm C18 column Waters Corporation186002346Analytical column
Acetonitrile HPLC grade >99.9% HiPerSolvVWR20060.420For LC mobile phases
CaCl2Sigma AldrichC5670Salt for HDX buffers
ChronosAxel Semrau (Purchased from Waters Corporation)667006090Scheduling software to enable multiple HDX-MS sample injections automatically. Alternative software is available from other vendors e.g. HDXDirector or LEAP Shell
Deuterium chlorideGoss Scientific (Cambridge Isotope Laboratories)DLM-2-50For HDX labelling buffers
Deuterium oxide (99.9% D2O)Goss Scientific (Cambridge Isotope Laboratories)DLM-4Deuterated water
DynamX 3.0Waters Corporation176016027Isotopic assignment and deuterium incorporation calculation
Enzymate BEH Pepsin ColumnWaters Corporation186007233Pepsin digestion column
Formic Acid, 99.0% LC/MS GradeFisher Scientific10596814For LC mobile phases
Guanidinium hydrochlorideSigma AldrichRDD001-500GChaotrope/Denaturant
HDfleXUniversity of ExeterN/Ahttps://ore.exeter.ac.uk/repository/handle/10871/127982
KClSigma AldrichP3911Salt for HDX buffers
LEAP HDX-2 CTC PAL sampling robotWaters Corporation725000637Autosampler robot
Leucine enkephalinWaters Corporation186006013For mass spectrometry lockspray calibration.
MassLynxWaters Corporation667004007Software controlling inlet methods and mass spectrometer
Maximum recovery vialsWaters Corporation600000670CV100 pack including caps - used for quench tray in LEAP HDX-2
MgCl2Sigma AldrichM8266Salt for HDX buffers
Millipore 0.22 µm syringe filtersMilliporeN9CA7069BSyringe filters
ms2minApplied Photophysics LtdN/Afast-mix quench-flow millisecond hdx instrument
NaClSigma AldrichS9888Salt for HDX buffers
Peltier temperature controllerLEAP Technologies Inc.HP115-COOL/DPeltier controller to set precise temperature of chambers in the LEAP robot.
ProteinLynx Global Server 3.0Waters Corporation715001030Peptide identification software. Alternative software is available from other vendors.
Reagent pot capsWaters Corporation186004632100 pack
Reagent pots for LEAP HDX-2Waters Corporation186001420100 pack excluding caps - used for buffers in LEAP HDX-2
Sodium deuteroxide (99.5% in D2O)Goss Scientific (Cambridge Isotope Laboratories)DLM-57For HDX labelling buffers
Spin filter microcentrifuge tubes (3 kDa MWCO)Amicon (Merck Sigma Aldrich)UFC5003Micro centrifuge tubes to concentrate protein. This facilitates buffer exchange and accurate sample loading for HDX-MS experiments.
Synapt G2-Si mass spectrometerWaters Corporation176850035Mass spectrometer
Total recovery vialsWaters Corporation600000671CV100 pack including caps - used for labelling tray in LEAP HDX-2
Tris-HClSigma AldrichT3253-250GBuffer
Trizma baseSigma AldrichT60040-B2005Buffer
UreaSigma AldrichU5378-1KGChaotrope/Denaturant
VanGuard 2.1 x 5 mm ACQUITY BEH C18 column Waters Corporation186004623Trap desalting column

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Intrinsically Disordered ProteinsPeptide MappingProtein AggregationConformational DynamicsDeuterium UptakeStatistical Significance Testing

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