The Engen Laboratory
       
 
Labeling methods
 
Deuterium Introduction
For more information see the publication below as well as the Publications page:
 
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 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.
 

Dilution
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
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 pulsed-labeling or continuous labeling (see Figure 1)

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Figure 1
 

Pulsed-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.
 

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 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 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) pulsed-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.
Figure 2
 
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 in Figure 3:

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Figure 3
 
Which ionization method to chose?
 

Currently, there are two primary ionization methods for doing hydrogen exchange analysis:

Electrospray (ESI)
MALDI

In the past, FAB was also used.

Electrospray

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

 MALDI

  • Potential for higher throughput
  • Deuterium losses can be harder to control
  • Generally easier to do than ESI
  • All data are aquired in a single spectra
  • Sample salts and impurities are much more tolerated
  • Lack of multiple charge states make spectra simple

People using MALDI:

Komives Lab, UCSD

Use of MALDI to look for binding surfaces
(PNAS 95, 14705-14710)

Fitzgerald Lab, Duke

SUPREX technique to analyze protein stability
(PNAS 97, 8296-8301)
 
The basic experiment
 
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 deg. C.  Its VERY important that that the pH change is precisely controled.  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 ensure that the pH will drop to 2.5 and stay there.  See below "Controls & Calculations".

To do electrospray (see the HX MS background page as well as above) mass spectral analysis, the sample must be desalted before it enters into the instrument.  Desalting serves a second, and very important purpose - it washes away the deuterium from amino acid sidechains (see Figure 4).  The exchange rate of the amide hydrogens in side chains (for example in arginine, or histidine) is very-very fast compared to the exchange rate of the amide hydrogens in the backbone.  When desalting is performed with standard HPLC buffers (which have a pH of 2.5 and are made of H2O), the hydrogen in the water of the HPLC 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.

Figure 5 shows an HPLC column, injectors and associated tubing in a bath of ice water to maintain the temperature at 0 deg. C.  Using this set-up reduces the losses of deuterium during analysis, whether it be from a whole protein or from peptides.

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 deg. C.  When the sample is ready for analysis, it is digested for 5 minutes at 0 deg. C with pepsin or it can be digestion inline immediately prior to RP-HPLC separation (see Figure 5).  After digestion, the peptides are separated quickly at 0 deg. C in 5-6 minutes.  The column, solvent lines and injector are kept in an ice bath (see Figure 5).  Figure 1 panel B illustrates a brief flow-chart of a continuous labeling experiment.
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Figure 4

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Figure 5
 
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 Background/Theory page).  Without doing so, the results of such analyses can be meaningless.

It is also critical to USE PROPER CONTROL samples.   The details of this are discussed in the section titled "Controls and Calculations" below.
 
Controls and calculations
 
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):
a 0% deuterium control
and
a 100% deuterium control
 
Control samples representing 0% and 100% exchanged protein are analyzed with each set of samples. 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.  The adjustment for loss of deuterium is explained with the help of Figure 6.
 
Figure 6 -- Example of data processing steps for adjusting the value of incorporated deuterium. (a). The centroid value of each isotopic distribution representing a given peptide is determined and adjusted with the equation in (b) for loss of deuterium during analysis. See text for definition of terms in (b).  Panel (c) compares raw data for a segment of protein. Observed: measured amount of deuterium taken-up at the given time; Adjusted: amount of deuterium in the peptide after adjusting the data with the equation in (b); Calculated: amount of deuterium in an unstructured peptide with the sequence of this segment of protein under identical exchange-in conditions.
 
Panel a is an example of data for a segment of protein, displaying the 0% and 100% references. The equation used to apply the adjustment is shown in Figure 6b where D is the adjusted deuterium level, m is the experimentally observed mass, m0% is the 0% or undeuterated control, m100% is the totally deuterated control and N is the total number of amide hydrogens in the fragment.
 
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).
 
Figure 6c shows the deuterium level plotted versus exchange-in time for a segment of protein before and after the adjustment for back-exchange was considered. The absolute number of deuterium is affected, but the profile of the curve is not. For comparison, a curve for exchange of the same sequence in an unstructured peptide [calculated from the parameters in (Bai et al., 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 is not required as long as the analysis conditions are identical.
 
 
Calculation of rate constants
 
From exchange-in time-course data, one can estimate the distribution of rate constants describing isotope exchange at each linkage within a peptide.  An example of fitting the data is shown in Figure 7.
 

Figure 7 -- Example of the data processing steps involved in determining the distribution of amide hydrogens within a peptide fragment and the rate constants for these amide hydrogens. (a). An example of curve-fit data from a protein. It is derived by fitting the data with the equation in (b). Details of the terms in (b) are explained in the text. (c). Final form of the equation in (b) for the example data in (a). Amide hydrogens have been grouped (according to similar exchange rates) into the catagories a, b, c [also shown in (a)]. The rates of exchange of each of these groups, (m1, m2, m3, respectively) are also shown.
 
Real data for a fragment of a protein are shown in panel a. A smooth line corresponding to the equation that best fits the experimental data was calculated using the equation in panel b. Rate constants are fit to the experimental data with a series of first order rate expressions where D is the number of deuterium present in a peptide, N is the number of peptide amide linkages in a segment and ki are the hydrogen-deuterium exchange rate constants for each peptide linkage (Zhang et al., 1996).
 
Although single amino acid resolution is not generally possible, exchange rates describing groups of hydrogens with similar exchange rates can be calculated. Therefore, a three-term exponential equation was used where the amide hydrogens are divided into fast, medium and slow categories according to their exchange rates
 
Figure 7c shows the actual equation used for the data in Fig 7a. The number of amide hydrogens that exchange at similar exchange rates is calculated for each term of the equation. At the same time the average rate constant for each group is calculated. The results for fitting the experimental data are shown in the bottom of panel c where the number of amide hydrogens that exchange at a given rate are tabulated along with the average rate constant calculated for each group
 
Software packages such as Kaleidograph (Synergy Software) or SigmaPlot (Jandel) can be used for fitting.
 

References
 
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. Biochemistry 35, 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.
 
 
Updated on 04.Nov.2006