HYDROGEN EXCHANGE:

What is it?
Hydrogen exchange combined with mass spectrometry is a method of analyzing proteins to learn things about their structure and their dynamics.  It promises to significantly aid our understanding of proteins and protein structure in the coming years.  Traditionally, hydrogen exchange has been used in conjunction with NMR.   However, now it is also possible to use mass spectrometry to measure hydrogen exchange.

The technique began in the early 1990s with the methods capable of introducing protein molecules into a mass spectrometer for mass analysis.  Methods development is ongoing and the technique is being applied to relevant biological problems, important proteins related to disease and to the vast number of proteins for which structural information is hard to obtain with other methods.

More history can be found here.

How does it work?
Some of the hydrogen atoms in proteins are capable of switching places with hydrogen atoms from the solvent molecules surrounding the protein.   If an isotope of hydrogen is used as the solvent, namely deuterium oxide (D2O), its heavier mass gets incorporated into the protein.  Because the protein now weighs more than normal, this change in mass can be monitored with high resolution mass spectrometers.

What is it used for?

Hydrogen exchange measurements can be used to sense changes in protein structure on a specific timescale (see FIGURE 1).  Some amide hydrogens, such as those at the surface of proteins, exchange very rapidly.  These hydrogens can be used to sense binding to other proteins and to analyze complexes.  Other hydrogens are buried in the hydrophobic core of the protein and may not exchange for hours, days, or even months.  Therefore the movements of proteins, and the rate of such movements can be studied.
  Some of the things you can use this technique for include:
  • Protein unfolding, either natural or induced by denaturants
  • Measurement of folding or unfolding rates
  • Protein folding, on timescales from milliseconds to days
  • Binding, binding constants and interacting surfaces
  • DNA-protein interactions

  Several advantages of the technique include:

  • Compared to other techniques, very little protein is required (~ 500 pmol)
  • There is limitation on the size of the protein -- even large protein complexes can be studied
  • Membrane proteins, nearly impossible with other techniques, can be studied

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FIGURE 1

How big can you go?
The size of protein or protein complex is primarily limited by how complex an experiment you want to do.   Modern mass spectrometers can measure proteins in the hundreds of thousands of daltons, even up to 1,000,000 Da.  By coupling UPLC to the mass spectrometer, complexes of many proteins can be analyzed.

The largest macromolecular complexes studied so far with hydrogen exchange are viral capsids, including the capsid of P22 (19.6 MDa)! Other large things that have been studied are the capsid of the brome mosaic virus (3.6 mDa) and the GroEL/ GroES complex.  All of these complexes are composed of one or two subunits, repeated many times.

The most complex, large macromolecular complex studied so far is the E coli ribosome.  While smaller than some capsids, the ribosome is composed of 54 unique proteins and 3 large RNAs for a total mass of 2.5 mDa.

Exchange theory
The exchange of hydrogens occurs at a specific rate, which is a function of the protein structure and solvent accessibility.  By measuring hydrogen exchange rates, we can draw conclusions about the dynamics of proteins.

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FIGURE 2

MOVIE

 There are three kinds of hydrogens in proteins (see FIGURE 2).   Hydrogens covalently bonded to carbon essentially to do not exchange.  The ones on the side chains exchange very fast and typically cannot be detected.  The ones at the backbone amide positions (yellow) exchange at rates that can be measured.  Each amino acid, except proline, has one amide hydrogen.  Therefore, hydrogen exchange rates can be measured along the entire length of the protein backbone.  Additionally, the backbone amide hydrogens are involved in formation of hydrogen bonds in secondary structural elements -- both alpha helicies and beta sheets; therefore, their exchange rates are a reflection of structure and structural stability.

The rate of HX depends on hydrogen bonding and solvent accessibility.  Folded proteins can have amino acids with HX rates as much as 1 billion times slower than the same amino acid that is not in a folded protein.  Protein folding and unfolding, whether in cells or in the test tube, represent large changes in protein structure, hydrogen bonding and solvent accessibility that can be investigated with HX MS.  Smaller structural changes critical for protein function can also be probed with HX MS.  Several comprehensive reviews have been published on the subject (see Publications page).

pH is the secret - Well, part of it
The secret to making hydrogen exchange measurements with mass spectrometry is in controlling the pH.  The rate of hydrogen exchange is very sensitive to pH -- a change in one pH unit equals a ten-fold change in the exchange rate!! (see FIGURE 3).

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FIGURE 3 		Consider FIGURES 3 and 4, for example:
if the exchange rate of an average amide hydrogen in a completely unstructured peptide at pH=7 equals 10 per second, the same average amide hydrogen in the same unstructured peptide at pH=5 would have an exchange rate of 0.1 per second.  The actual time involved to make the exchange in this example is far less than 1 second (see half-life in Figure 3).
So in the simplest case scenario, one would want to do the exchange reaction at a higher pH and then reduce the pH to stop the exchange from occurring.  This is essential to do a proper mass spectrometry analysis, since it takes some time to carry out the mass spectral analysis. 		click for a
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FIGURE 4

And don't forget, proteins are not just unstructured peptides! Their exchange rates will be slowed by:
    1.  Hydrogen bonding that creates the secondary structure, primarily alpha helices and beta sheets.
    2.  Protection from the solvent, primarily due to being buried in the hydrophobic core of the protein.
    3.  Hydrogen bonding to water in the solvent, a much smaller effect than #'s 1 and 2.

The pH effect can be used in a number of ways in addition to the simplest one mentioned above.  For example, one could do protein folding at pH 7 and then pulse quickly to pH=10 to "flash-label" any amide hydrogen not involved in hydrogen bonds with other parts of the protein.

The rest of the secret
Another problem remains! Considering the example in Figure 3, even if you do an experiment at pH 7.0 and reduce the pH to 2.5 (the minimum of the curve in Figure 2), there will only be about 11 minutes (half-life) for you to do the mass spectral analysis before you loose half the deuterium label!. To further slow the exchange rates, the temperature must be lowered to zero degrees celsius (see Figure 3). At pH=2.5 and temp=0, the average half-life of exchange for the average amide hydrogen is 30-60 minutes. These conditions give enough time to analyze the sample.

AND A MINOR COMPLICATION....
Even in unstructured peptides, every sequence will not exchange at the same rate. That means that the loss of deuterium from a labeled protein will not be the same. There are influences from the neighboring amino acids, which depends of course on sequence. The amino acid at the N-terminus will also exchange its backbone amide hydrogen much faster than ones further down the line. These issues have been addressed in the literature. To read more, see:

For sequence effects:
Connelly GP, Bai Y, Jeng MF, Englander SW. (1993). Isotope effects in peptide group hydrogen exchange. Proteins 17(1), 87-92.
Bai Y, Milne JS, Mayne L, Englander SW. (1993). Primary structure effects on peptide group hydrogen exchange. Proteins 17(1), 75-86.

For calculating the average deuterium loss for an average peptide:
Zhang, Z., and Smith, D.L. (1993). Determination of amide hydrogen exchange by mass spectrometry: A new tool for protein structure elucidation. Protein Sci. 2, 522-531.

MASS SPECTROMETRY:

Introduction
Mass spectrometry is invaluable for measuring the molecular weight of proteins. Typical analysis can identify the molecular weight of a 20,000 kD protein to within +/- 2.0 daltons while analysis of peptides can be even more accurate. It is for these reasons that mass spectrometry is used in the Engen laboratory to investigate the incorporation of hydrogen into proteins during hydrogen exchange.

Because deuterium weights 2 Da and hydrogen weighs 1 Da, deuterated and non-deuterated proteins and/or peptides can easily be distinguished

This is a simple diagram of a mass spectrometer. It can be divided into three parts: ionization, mass analyzer, and detector.

General mass spectrometry geometry

Ionization
Proteins and peptides must first be converted to the gas phase via ionization.   Ionization of proteins/peptides can be done in several ways. MALDI (matrix assisted laser desorption mass spectrometry) uses a laser beam to zap crystallized protein/matrix mixtures into the gas phase.  ESI (electrospray ionization) converts a liquid-protein solution into fine droplets.  The solvent is evaporated from the fine droplets and charge is deposited on the protein molecules.  ESI can be coupled to HPLC.  As peptides/proteins elute from an HPLC column they are sent directly into the mass spectrometer where they are ionized.

Mass analyzer
Once proteins/peptides are ionized, they must be separated according to their molecular weight. There are several types of mass analyzers: magnetic sector, quadrupole, ion trap, time-of flight.

All mass analysis is done in a vacuum so that molecules do not collide with each other. Sometimes, molecular collisions are desirable and can be used to fragment larger molecules into smaller ones. This procedure is called MS/MS.

Detector
After a collection of proteins/peptides have been separated according to mass, they must be detected.  Ions can be detected with electron multipliers or with diode array detectors.

We use mass spectrometers from the Waters Corporation.  Take a tour of our lab and the instruments in the James L. Waters Mass Spectrometry Facility.