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with Hydrogen Exchange and Mass Spectrometry





For more information see the publication:

Engen, J.R. & Smith, D.L. (2000). Investigating the higher order structure of proteins: Hydrogen exchange, proteolytic fragmentation & mass spectrometry. In "Protein and Peptide Analysis: New Mass Spectrometric Applications" (J. Chapmann, ed.). Meth. Mol. Biol. Vol. 146.

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 to protein that are already in solution.   Addition of D2O to lyophilized protein complicates the data at early exchange-in times due to solvation effects.


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 it become diluted.

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.




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, 675-684.

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

Continuous labeling

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 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.


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A flow chart of the differences between continuous and pulsed-labeling is shown in Figure 1.  In this example, urea is used to denature the protein prior to pulsed-labeling.


The primary difference between the two methods:
in continuous labeling, a cumulative summary of the populations of the molecules if evident in the spectra.  In pulsed-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.  Pulsed-labeling is useful for measuring the unfolding and refolding rate constants in conditions where the unfolded state has been artificially increased by addition of denaturant, heat or pH changes.


Two spectra are shown at the left.   They are from the same peptide of a larger proteins that was labeled with (A) continuous or (B) pulsed-labeling.

In (A), the protein sat in deuterium for 30 minutes before being analyzed.  In (B), the protein sat 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.
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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 below:

quench1.jpg (23282 bytes)



For more reading, see the following publications:

Yang, H., and Smith, D.L. (1997). Kinetics of cytochrome c folding examined by hydrogen exchange and mass spectrometry. Biochemistry 36, 14992-14999.

Dharmasiri, K., and Smith, D.L. (1996). Mass spectrometric determination of isotopic exchange rates of amide hydrogens located on the surfaces of proteins. Anal. Chem. 68, 2340-2344.