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.
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.
CONTINUOUS VS. PULSED-LABELING
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,
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
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.
|click for a
|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
|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.
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
An example of a set-up that can be used is shown below:
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
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.