Our research laboratory has written extensively on the methodology of HX MS and some of our recent reviews and articles that describe how to do this method are:

Wales & Engen (2006). Hydrogen exchange mass spectrometry for the analysis of protein dynamics.
Mass Spectrom. Rev. 25, 158.  DOI: 10.1002/mas.20064. Pubmed: 16208684.

Morgan & Engen (2009). Investigating Solution-Phase Protein Structure and Dynamics by Hydrogen Exchange Mass Spectrometry. Curr Protoc Protein Sci. Chapter 17:Unit17.6. DOI: 10.1002/0471140864.ps1706s58. Pubmed: 19937720. 

Marcsisin & Engen (2010). Hydrogen Exchange Mass Spectrometry: What is it and what can it tell us?
Anal Bioanal Chem. 397(3), 967.  DOI: 10.1007/s00216-010-3556-4. Pubmed: 20195578. 

Wales TE, Eggertson MJ & Engen (2013). Considerations in the analysis of hydrogen exchange mass spectrometry data.  Methods Mol Biol. 1007:263-288. DOI: 10.1007/978-1-62703-392-3_11. Pubmed: 23666730.

Labeling methods Which ionization method? The basic experiment Controls & Processing


Deuterium introduction
Deuterium can be introduced to the protein sample in a number of ways.  Two primary methods are dilution and gel filtration.  It is always recommended that deuterium is introduced into a protein sample that is already in solution.  Addition of D2O to lyophilized protein complicates the data at early exchange-in times due to solvation effects.

Engen JR, Smith DL. (2000).  Investigating the higher order structure of proteins. Hydrogen exchange, proteolytic fragmentation, and mass spectrometry.  Methods Mol Biol. 146, 95. Pubmed: 10948498

With the protein in a buffer of H2O, the sample is diluted by adding an excess of the same buffer but made with D2O, in a dilution factor of 15-20 fold by volume.  This effectively raises the deuterium level to greater than 95%.  Dilution is an easy method but requires more protein since the concentration is significantly reduced.

Gel filtration
This method works by introducing the protein in a buffer of H2O into a small spin column that has been equilibrated with the same buffer made with D2O.  The spin column is filled with G-25 gel filtration media.   After a brief spin in a table-top centrifuge, the protein elutes out the column into a receiving tube.  The protein is now in D2O and the H2O is trapped in the gel filtration media.  Using this technique requires precise timing of how long it takes to elute the protein from a specific bed volume at a specific centrifuge speed.  A simple test can be done with a visible protein like cytochrome c, myoglobin, or green fluorescent protein to determine the optimal parameters.

Pulsed vs. Continuous-labeling
The information below was summarized from this publication:

Deng, Y, Zhang, Z. and Smith, D.L. (1999). Comparison of continuous and pulsed labeling amide hydrogen exchange/mass spectrometry for studies of protein dynamics. J. Am. Soc. Mass Spectrom. 10, 675.

The actual labeling step, when the protein is in deuterium, can also be done in several ways.  Two primary methods are pulsed-labeling or continuous labeling.

Pulse Labeling
The protein sits under specific conditions for a period of time (i.e. in denaturant for 30 minutes).  Then a short pulse of deuterium is introduced and the reaction is quenched.  A good example of pulse labeling is for the study of protein folding, as shown at the right.

Continuous Labeling
This is the most common type of HX MS experiment.  The protein is exposed to deuterium at time=0 and left in deuterium.  At a various times the reaction is quenched by moving an aliquot of sample to quench buffer.

The primary difference between the two methods:
In continuous labeling, a cumulative summary of the populations of the molecules is evident in the spectra.  In pulse labeling, an instantaneous snapshot of the protein populations is evident.  Continuous labeling is useful for looking at populations of protein molecules under conditions where the folded state is favored since the minor contribution of the unfolded states is integrated over time.  Pulse labeling is useful for measuring the unfolding and refolding rate constants in conditions where the unfolded state has been artificially increased by addition of chemical denaturant, heat or pH changes.

Two spectra are shown at the left.  They are from the same peptide of a larger protein that was labeled by either (A) continuous or (B) pulse labeling.

In (A), the protein hydrogens were available for exchange with deuterium for 30 minutes before being analyzed.  In (B), the protein was equilibrated in 3M urea for 30 minutes, was pulsed for 10 seconds with deuterium at pH 7 and then analyzed.

From the spectra in (A), the cumulative population of molecules that have unfolded is evident, since once the molecule unfolds, it becomes labeled (a process that cannot be reversed since the concentration of deuterium is >95%).  In (B), there are 2 populations of molecules, indicated by the two isotope distributions.  The higher mass distribution represents the percent of the population that is unfolded after 30 minutes in 3M urea.

Quench-flow and rapid mixing
Rapid mixing systems, such as the Biologic quench flow system or other such systems can be used to investigate events that are as fast as 10 milliseconds.  Things under this category can include rapid exchange of surface amide hydrogens or protein folding events.  Protein folding can be studied with a type of pulsed-labeling.  A denatured protein is allowed to refold by dilution of the denaturant.  After certain refolding times, the protein is pulsed with deuterium and the reaction quenched.  Analyses of this type can be automated with quench-flow devices.  An example of a set-up that can be used is shown on the right.

Labeling methods Which ionization method? The basic experiment Controls & Processing


Currently, there are two primary ionization methods for doing hydrogen exchange analysis: Electrospray and MALDI.  In the past, FAB was also used.


  • HPLC step washes away all the labile hydrogens at side chain positions
  • Temperature and pH are easily controlled
  • When coupled to HPLC or UPLC, large, complex digests can be analyzed
  • Samples are desalted and made free of impurities
  • Multiple charge states provides redundancy


  • Deuterium losses can be harder to control
  • All data are acquired in a single spectra - limits upper protein size limit
  • Sample salts and impurities are much more tolerated
  • Lack of multiple charge states make spectra simple

Labeling methods Which ionization method? The basic experiment Controls & Processing


For a continuous labeling experiment, a protein is labeled with deuterium at pH=7.0.   The reaction is quenched by lowering the pH to 2.5 and the temperature to 0 degrees C.  Its VERY important that that the pH change is precisely controlled.  The best way to do that is to use a weak (25 mM) phosphate buffer for the labeling step at pH =7.0 and then use a strong (100 mM) phosphate buffer for the quench step.  This ensures that the pH will drop to 2.5 and stay there.  See below "Controls & Calculations".

To do electrospray (see the Tutorial: theory page as well as above) mass spectral analysis, the sample must be desalted before it enters the instrument. Desalting serves a second, and very important purpose - it washes away the deuterium from amino acid sidechains. The exchange rate of the amide hydrogens in side chains (for example in lysine or glutamic acid) is very-very fast compared to the exchange rate of the amide hydrogens in the backbone. When desalting is performed with standard LC buffers (which have a pH of 2.5 and are made of H2O), the hydrogen in the water of the LC buffers replaces the deuterium that exchanged into the side chains. This leaves only deuterium label at the backbone amide positions. Having only these amide hydrogens to analyze is a convenient situation:

  1. it gives an individual exchange-rate sensor for every amino acid (expect proline).
  2. it eliminates messy details of exchange into side chains, which can be influenced by things other than secondary structure and solvent shielding.

To maintain the quench conditions, the HPLC/UPLC column, injectors and associated tubing etc. were put into a bath of ice water to ensure the temperature is 0 degrees C. Using this set-up reduces the losses of deuterium during analysis, whether it be from a whole protein or from peptides.  Modern systems use refrigeration to provide the low temperature, rather than ice baths.

Whole proteins can be analyzed, or digested into smaller pieces with an enzyme. The enzyme that seems to work the best is porcine pepsin. It likes working at pH 2.5 and can tolerate being at 0 degrees C. When the sample is ready for analysis, it is digested in a test tube for 5 minutes at 0 degrees C with a pepsin solution or it can be digestion online immediately prior to LC separation. After digestion, the peptides are separated quickly in 5-6 minutes. The column, solvent lines and injector are kept cold. 

Other experiments
There are many variations one can do to the general type of experiment explained above. Whether analysis if done with MALDI rather than electrospray, whether a pH pulse is used, whether denaturants are thrown in.... all experiment must take into account the pH factor and the temperature factor (see Tutorial: Theory page). Without doing so, the results of such analyses can be meaningless.

Labeling methods Which ionization method? The basic experiment Controls & Processing


Relative versus absolute
The amount of deuterium that is incorporated can be measured in a relative sense or an absolute sense. Relative measurements are easier because no back-exchange controls are required.  For most types of comparisons experiments, relative measurements are the best. 

Absolute measurements: adjustment for deuterium loss
Although some loss of deuterium label from the peptic fragments during peptic digestion and HPLC is unavoidable in these experiments, simple adjustments can be made for these losses.  These adjustments are based on analysis of appropriate controls (Zhang, 1995; Zhang & Smith, 1993): 0% control sample and 100% control sample.

The 0% control is protein that had never been exposed to deuterium and the 100% control is protein in which all exchangeable hydrogens have been replaced with deuterium.  An equation is used to adjust the levels of deuterium at each labeling time point (Zhang & Smith, 1993). Since the adjustment is based on the assumption that loss of deuterium from a totally deuterated fragment is proportional to the loss from the same fragment when partially deuterated, the accuracy of the adjustment depends on the sequence of the peptide (Zhang & Smith, 1993).  The average error that results from the equation in Fig 6a is about 5%, with 92% of fragments having an error <10%, 3% having an error >15% and 0.5% having an error >25% (Zhang & Smith, 1993). Adjustment is most useful when determining the actual amount of deuterium incorporated in a fragment. However, when the levels of deuterium in the same fragment are compared (such as comparing the amount of deuterium in a given region in the free or ligand bound state) the correction and a relative measurment is more appropriate.

Data processing
The amount of deuterium in each peptic peptide is determined and then plotted against the exchange-in time.  From graphs of deuterium uptake, comparisons can be made for different forms of the protein.  The many details of data processing have recently been described in:

Wales TE, Eggertson MJ & Engen (2013). Considerations in the analysis of hydrogen exchange mass spectrometry data.  Methods Mol Biol. 1007, 263.
DOI: 10.1007/978-1-62703-392-3_11. Pubmed: 23666730.


Bai, Y., Milne, J. S., Mayne, L., & Englander, S. W. (1993). Primary structure effects on peptide group hydrogen exchange. Proteins: Struct. Funct. Genet. 17, 75-86.

Zhang, Z., Post, C. B., & Smith, D. L. (1996). Amide hydrogen exchange determined by mass spectrometry: application to rabbit muscle aldolase. Biochemistry35, 779-791.

Zhang, Z., & Smith, D. L. (1993). Determination of amide hydrogen exchange by mass spectrometry: A new tool for protein structure elucidation. Protein Sci.2, 522-531.