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JoVE Journal Neuroscience

Neuroscience

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

Published: June 23, 2022 doi: 10.3791/64050
Automatic Translation
1, 1
1Living Systems Institute, Department of Biosciences, University of Exeter

Summary

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

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

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

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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Representative Results

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 in two different solution conditions: State A and State B8, as defined in steps 2.1.-2.2.

First, a mapping experiment on aSyn was performed, and a peptide coverage map was obtained, as shown in Figure 1. The map covers 100% of the protein sequence and has an average redundancy of 3.79. The 100% coverage value indicates that all the amino acids in the protein were found in the protein digests and will enable a comprehensive analysis of the exchange behavior of aSyn. The redundancy value indicates the number of overlapping peptides. A higher redundancy value increases the structural resolution of the final map, given subtractive flattening of the data for overlapping peptides32.

Using the fast-mixing quench-flow instrument prototype (see Table of Materials), high-quality, millisecond-timescale HDX-MS data on aSyn at pH 7.4 in State A and State B were collected (Figure 2). Following an isotopic assignment in DynamX, "crude" deuterium uptake curves were obtained, as shown in Figure 3A. It shows uptake curves for three peptides selected across each protein domain. The deuterium incorporation over time is displayed. The x-axis is on the millisecond timescale, which aligns with the very fast kinetics of aSyn at physiological conditions. The red shaded region shows the data typically obtained from conventional HDX instruments, with starting measurements from 30 s. Importantly, this cannot be further reduced by pH manipulation for so-called "time-window expansion"; that approach is invalid for the study of intrinsically disordered proteins/regions, as the pH shift will perturb the conformational ensemble of the weakly stable polypeptide. As can be seen here, most of aSyn is fully exchanged by 1 s (Figure 3C). This shows the importance of millisecond HDX measurements for monomeric aSyn as the full kinetic uptake curve for the exchange reaction is captured, which yields the most accurate measurement of the monomer conformations.

HDfleX performed back-exchange correction using the plateau deuterium incorporation. The data points were subsequently fitted according to Equation 1, providing an observed rate constant, kobs, indicative of the solvent accessibility and hydrogen bonding involvement of that particular peptide (Figure 3B).

Equation 1    Equation 1

where Dt is the deuterium incorporation at time t, nExp is the number of exponential phases, N is the maximum number of labile hydrogens, kobs is the observed exchange rate constant, and β is a stretching factor30,33.

Following curve fitting, the uptake area under the fitted curve can be calculated by integrating the fitted function describing the uptake curve within the experimental time window. Statistical significance analyses between the uptake area of the two states were performed. First, a global significance threshold for the uptake area was calculated in HDfleX at a confidence level of 95%. Uptake area difference plots were then generated, showing the difference between State A and State B at two levels of structural resolution: peptide resolution (Figure 4A) and amino acid resolution (Figure 4B). The peptide resolution difference plot shows the difference in uptake area between State A and State B for each individual peptide, while the amino acid resolution difference plot shows the difference in uptake area between State A and State B flattened across the entire amino acid sequence of aSyn30,34. Both plots indicate an overall greater deuterium uptake throughout the aSyn monomer in State B compared to State A. This finding can be justified by examining the deuterium uptake plots in Figure 3, where the State B uptake curve is always above the State A uptake curve. Furthermore, it can be seen that the magnitude of the uptake area difference is much higher at the C-terminus. Once again, this can be justified by tracing back to the original uptake curves, where the C-terminal peptides (peptides 124-140 shown in Figure 3) show a much bigger gap between the uptake curves than the rest of the protein. In conclusion, the solution conditions in State B cause an increase in solvent exposure or decrease in hydrogen-bonding network participation throughout the protein but more so at the C-terminus.

Figure 1
Figure 1: Peptide coverage map of wild-type aSyn with a total of 30 peptides and 100% sequence coverage. The three domains of aSyn are highlighted as follows: N-terminus (blue), non-amyloid beta component region (yellow), and C-terminus (red). Please click here to view a larger version of this figure.

Figure 2
Figure 2: Workflow for a millisecond HDX-MS experiment on aSyn. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Example uptake plots from three peptides selected across the three domains of aSyn for State A (yellow) and State B (blue). (A) Unfitted and non-back-exchange corrected uptake plot. (B) Fitted and back-exchange corrected uptake plots. The red shaded region represents data obtainable by conventional HDX-MS systems, typically starting from 30 s. Error bars correspond to the standard deviation of the three replicates. (C) Heatmap plot of the percentage deuterium uptake across amino acid sequence per timepoint. The color bar represents the percentage of deuterium uptake. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Uptake area difference plots at maximum curve plateau time (14,084 ms). (A) Peptide residue resolution plots show each peptide's uptake area difference between State A and State B. (B) Amino acid resolution plot showing the uptake area difference between State A and State B flattened across the amino acid sequence. Please click here to view a larger version of this figure.

Mapping Energy Level Ramp Voltage (V)
Low 20-40
Medium 25-45
High 30-50
Very high 35-55

Table 1: Mapping energy levels and corresponding transfer ramp voltages.

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Discussion

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 only around 2 h. Given the fully-automated instrumentation used here, a complete dataset can be acquired in 1 day. However, when handling samples and preparing buffers, care must be taken to ensure that measurements are derived from weakly stable protein (or protein regions) in the desired state. Importantly, a previous study showed that different storage conditions, such as freezing and lyophilizing, resulted in different aSyn monomer conformers and that it is important to characterize the potential impact of sample handling on the aSyn monomer conformational ensemble10. Indeed, HDX-MS is a highly sensitive measure of such conformational perturbations, with a dynamic range from microseconds to at least months. In addition, if strictly studying only the aSyn monomer, filtration is strongly advised to remove unwanted oligomers and fibrils that might have formed in the sample upon storage or handling. Furthermore, the HDX-MS buffers need to be tightly controlled within 0.05 of the desired pH or pD, as any discrepancies will significantly affect the intrinsic rate of exchange and lead to unwanted errors. It is also important to note that comparisons between solution conditions for any protein that differs in pH, temperature, or salt composition will alter the intrinsic rate. Therefore, these data will require further corrections, such as applying a pH adjustment factor35 or an empirical correction factor8,30.

In terms of instrumentation, there are no commercially available systems that allow the acquisition of millisecond HDX-MS data. Several research groups have developed their own systems, from quench-flow systems13,15,36 to microfluidic chips37,38,39 to capture the rapid exchange kinetics of certain proteins. Another method that has been used to achieve millisecond timescale HDX-MS data is known as the time expansion method40,41, whereby the pH of the buffers is reduced to slow down the exchange kinetics. However, this method does not apply to aSyn (or to any weakly stable protein features) as (1) the lowering of the pH drastically changes the charge density of the protein and increases the rate of aggregation8,42, and (2) the aSyn conformers are only meta-stable and are likely to be perturbed by these pH alterations. For these reasons, it is recommended to maintain a consistent pH in the HDX-MS buffers when studying monomeric aSyn conformations, unless physiologically relevant, and employ a millisecond labeling instrument.

Most of the aSyn monomer fully exchanges within 1 s, and at most, it takes approximately 15 s to fully exchange with deuterium (Figure 3) at a physiologically relevant pH of 7.4 (reflective of intracellular cytosolic conditions at the presynapse). Using conventional HDX-MS systems, starting at 30 s is not appropriate as the HDX-MS data would correspond to the plateau of the exchange reaction, which does not provide any useful conformational information. However, the lower limit of measurement of the millisecond HDX instrument (corresponding to a "dead-time" of 50 ms) enables monitoring of the exchange reaction from ~25% completion for the aSyn monomer at pH 7.4. This allowed us to capture the majority of the kinetic uptake curve. Fitting the deuterium uptake curve to Equation 1 provides important kinetic information; it corresponds to an estimate of the observed rate constant, kobs. While not covered here, it is possible to carry out aggregation kinetics experiments and examine fibril morphologies of aSyn under the same solution conditions as the HDX-MS experiments since HDX-MS is highly tolerant of a wide range of buffers8. Thus, for example, the kobs from the HDX-MS experiment can be correlated with the results from the aggregation experiments to gain insight into which conformations are most prone to certain aggregation behaviors and fibril morphologies.

For the simple case of differential HDX-MS experiments, where two or more conditions or protein variants are to be compared, the area under the fitted uptake curve can be integrated for each state and compared to each other. In this study, the uptake areas for State A and State B were compared at two different levels of structural resolution: peptide resolution and amino acid resolution, both of which have distinct strengths and challenges. For instance, the peptide resolution data reflect the raw spectral data more closely and have undergone the least processing. However, the "flattened" amino acid resolution data allow both peptide and soft-fragmentation information to be combined into a single output rather than separate unmergeable outputs and, ultimately, present the data at the highest structural resolution. One limitation of the mass spectrometry detection of HDX labeling is the challenge of obtaining amino acid resolution. While "soft-fragmentation" techniques, such as electron transfer dissociation (ETD), electron capture dissociation (ECD), and ultraviolet photodissociation (UVPD), have been proven to be effective at generating higher resolutions, they remain challenging, unpredictable, and inefficient30,43,44,45,46,47,48.

Compared to other structural techniques, millisecond HDX-MS has the unique advantage of capturing the conformational dynamics of monomeric aSyn at high structural and temporal resolutions. As the fast exchange kinetics of the monomer are no longer a limiting factor, further studies can be performed on monomeric aSyn with different mutations, post-translational modifications, salt components and concentrations, and binding partners at physiological pH. Correlating the HDX-MS results with functional studies, such as aggregation kinetics and fibril morphologies, can provide insight into conformers that either promote normal cellular function or are disease-prone. Ultimately, it is anticipated that such millisecond HDX-MS may be crucial for discovering targeted drugs that stabilize specific physiologically tolerated conformers.

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Disclosures

The authors declare no competing interests.

Acknowledgments

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

Materials

Name Company Catalog Number Comments
1 × 100 mm ACQUITY BEH 1.7 μm C18 column  Waters Corporation 186002346 Analytical column
Acetonitrile HPLC grade >99.9% HiPerSolv VWR 20060.420 For LC mobile phases
CaCl2 Sigma Aldrich C5670 Salt for HDX buffers
Chronos Axel Semrau (Purchased from Waters Corporation) 667006090 Scheduling software to enable multiple HDX-MS sample injections automatically. Alternative software is available from other vendors e.g. HDXDirector or LEAP Shell
Deuterium chloride Goss Scientific (Cambridge Isotope Laboratories) DLM-2-50 For HDX labelling buffers
Deuterium oxide (99.9% D2O) Goss Scientific (Cambridge Isotope Laboratories) DLM-4 Deuterated water
DynamX 3.0 Waters Corporation 176016027 Isotopic assignment and deuterium incorporation calculation
Enzymate BEH Pepsin Column Waters Corporation 186007233 Pepsin digestion column
Formic Acid, 99.0% LC/MS Grade Fisher Scientific 10596814 For LC mobile phases
Guanidinium hydrochloride Sigma Aldrich RDD001-500G Chaotrope/Denaturant
HDfleX University of Exeter N/A https://ore.exeter.ac.uk/repository/handle/10871/127982
KCl Sigma Aldrich P3911 Salt for HDX buffers
LEAP HDX-2 CTC PAL sampling robot Waters Corporation 725000637 Autosampler robot
Leucine enkephalin Waters Corporation 186006013 For mass spectrometry lockspray calibration.
MassLynx Waters Corporation 667004007 Software controlling inlet methods and mass spectrometer
Maximum recovery vials Waters Corporation 600000670CV 100 pack including caps - used for quench tray in LEAP HDX-2
MgCl2 Sigma Aldrich M8266 Salt for HDX buffers
Millipore 0.22 µm syringe filters Millipore N9CA7069B Syringe filters
ms2min Applied Photophysics Ltd N/A fast-mix quench-flow millisecond hdx instrument
NaCl Sigma Aldrich S9888 Salt for HDX buffers
Peltier temperature controller LEAP Technologies Inc. HP115-COOL/D Peltier controller to set precise temperature of chambers in the LEAP robot.
ProteinLynx Global Server 3.0 Waters Corporation 715001030 Peptide identification software. Alternative software is available from other vendors.
Reagent pot caps Waters Corporation 186004632 100 pack
Reagent pots for LEAP HDX-2 Waters Corporation 186001420 100 pack excluding caps - used for buffers in LEAP HDX-2
Sodium deuteroxide (99.5% in D2O) Goss Scientific (Cambridge Isotope Laboratories) DLM-57 For HDX labelling buffers
Spin filter microcentrifuge tubes (3 kDa MWCO) Amicon (Merck Sigma Aldrich) UFC5003 Micro centrifuge tubes to concentrate protein. This facilitates buffer exchange and accurate sample loading for HDX-MS experiments.
Synapt G2-Si mass spectrometer Waters Corporation 176850035 Mass spectrometer
Total recovery vials Waters Corporation 600000671CV 100 pack including caps - used for labelling tray in LEAP HDX-2
Tris-HCl Sigma Aldrich T3253-250G Buffer
Trizma base Sigma Aldrich T60040-B2005 Buffer
Urea Sigma Aldrich U5378-1KG Chaotrope/Denaturant
VanGuard 2.1 x 5 mm ACQUITY BEH C18 column  Waters Corporation 186004623 Trap desalting column

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References

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Tags

Millisecond Hydrogen/deuterium-exchange Mass Spectrometry Alpha-synuclein Structural Dynamics Proteome Disorder Fully Automated Analytical Technique Peptide Mapping Sample Preparation Protein Concentration Equilibrium Buffer Autosampler Robot HDX Robotics Mass Spectrometer
Millisecond Hydrogen/Deuterium-Exchange Mass Spectrometry for the Study of Alpha-Synuclein Structural Dynamics Under Physiological Conditions
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Seetaloo, N., Phillips, J. J.… More

Seetaloo, N., Phillips, J. J. Millisecond Hydrogen/Deuterium-Exchange Mass Spectrometry for the Study of Alpha-Synuclein Structural Dynamics Under Physiological Conditions. J. Vis. Exp. (184), e64050, doi:10.3791/64050 (2022).

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