Hydrogen–deuterium exchange (also called H–D or H/D exchange) is a chemical reaction in which a covalently bonded hydrogen atom is replaced by a deuterium atom, or vice versa. Usually the examined protons are the amides in the backbone of a protein. The method gives information about the solvent accessibility of various parts of the molecule, and thus the tertiary structure of the protein. Hydrogen exchange was first shown and explored by Kaj Ulrik Linderstrøm-Lang.
In solution, amide hydrogens in the peptide bonds of proteins exchange protons with the solvent. By changing the solvent from H2O to D2O, deuterons will be incorporated in the amide positions and the exchange reaction can be followed. Most often, deuterium is added to a protein in H2O by diluting the H2O solution with D2O (e.g. tenfold). Usually exchange is performed at physiological pH (7.0–8.0) where proteins are in their most native ensemble of conformational states. See also .
Because the exchange reaction can be either acid or base catalyzed, it is strongly pH dependent. For the backbone amide hydrogens, the minimum exchange rate occurs at approximately pH 2.6, on average. By performing the exchange at neutral pH and then rapidly changing the pH, the exchange rates of the backbone amide hydrogens can be dramatically slowed, or quenched. The pH at which the reaction is quenched depends on the analysis method. For detection by NMR, the pH may be moved to around 4.0–4.5. For detection by mass spectrometry, the pH is dropped to the minimum of the exchange curve, pH 2.6. In the most basic experiment, the reaction is allowed to take place for a set time before it is quenched.
The deuteration pattern of a quenched protein can be stably maintained in aprotic environments. However, analysis of the deuteration is usually performed in an aqueous solution, which means that exchange will continue at a slow rate even after the reaction is quenched. Reversion of deuterated positions after the quench step is referred to as back-exchange and various methods have been devised to correct for this.
H–D exchange was measured originally by the father of hydrogen exchange Kaj Ulrik Linderstrøm-Lang using density gradient tubes. In modern times, H–D exchange has primarily been monitored by the methods: NMR spectroscopy and mass spectrometry. Each of these methods have their advantages and drawbacks.
Hydrogen and deuterium nuclei are grossly different in their magnetic properties. Thus it is possible to distinguish between them by NMR spectroscopy. Typically HSQC spectra are recorded at a series of timepoints while the hydrogen is exchanging with the deuterium. Since the HSQC experiment is specific for hydrogen, the signal will decay exponentially as the hydrogen exchanges. It is then possible to fit an exponential function to the data, and obtain the exchange constant. This method gives residue-specific information for all the residues in the protein simultaneously. The major drawback is that it requires a prior assignment of the spectrum for the protein in question. This can be very labor intensive, and usually limits the method to proteins smaller than 25 kDa. Because it takes minutes to hours to record a HSQC spectrum, amides that exchange quickly must be measured using other pulse sequences.
Mass spectrometry has several advantages over NMR with respect to analysis of H–D exchange reactions: Much less material is needed, the concentration of protein can be very low (as low as 0.1 uM), the size limit is much greater, and data can usually be collected and interpreted much more quickly.
The deuterium nucleus is twice as heavy as the hydrogen nucleus because it contains a neutron as well as a proton. Thus a protein that contains some deuterium will be heavier than one that contains all hydrogen. As a protein is increasingly deuterated, the molecular mass increases correspondingly. Detecting the change in the mass of a protein upon deuteration was made possible by modern protein mass spectrometry, first reported in 1991 by Katta and Chait .
The location and relative amount of deuterium exchange along the peptide backbone can be determined roughly by subjecting the protein to proteolysis after the exchange reaction has been quenched. Individual peptides are then analyzed for overall deuteration of each peptide fragment. Using this technique the resolution of deuterium exchange is determined by the size of the peptides produced during digestion . Pepsin, an acid protease, is commonly used for proteolysis, as the quench pH must be maintained during the proteolytic reaction. To minimize the back-exchange, proteolysis and subsequent mass spectrometry analysis must be done as quickly as possible. HPLC separation of the peptic digest is often carried out at low temperature just prior to electrospray mass spectrometry to minimize back-exchange. More recently, UPLC has been used due to its superior separation capabilities .
It was proposed in 1999 that it might be possible to achieve single-residue resolution by using collision-induced dissociation fragmentation of deuterated peptides in conjunction with tandem mass spectrometry. It was soon discovered that CID causes "scrambling" of the deuterium position within the peptides . However, fragmentation produced by electron capture dissociation and electron transfer dissociation proceed with little or no scrambling under the correct experimental conditions . This suggests that eventually it may be possible to obtain single-residue resolution of H/D exchange reactions on a routine basis.
It is not possible to determine the structure of a protein with H/D exchange nor is it possible to define secondary structural elements. The reasons for this are related to the way in which protein structure slows exchange. Exchange rates are a function of two parameters: solvent accessibility and hydrogen bonding. Thus an amide which is part of an intramolecular hydrogen bond will exchange slowly if at all, while an amide on the surface of protein hydrogen bonded to water will exchange rapidly. Amides buried from the solvent but not hydrogen bonded may also have very slow exchange rates. Because both solvent accessibility and hydrogen bonding contribute to the rate of exchange, it becomes difficult to attribute a given exchange rate to a structural element without x-ray crystallography or NMR structural data.
H–D exchange has been used to characterize the folding pathway of proteins, by refolding the protein under exchange conditions. The parts of the structure that form rapidly, will be protected quickly, and thus not exchanged, whereas areas that fold late in the pathway will be exposed to the exchange for longer periods of time. Thus H/D exchange can be used to determine the sequence of various folding events. The critical factor determining the time resolution of this approach is the time required for quenching.
H–D exchange has been used to characterize protein–protein interactions. The exchange reaction needs to be carried out with the isolated proteins and with the complex. The exchanging regions are then compared. If a region is buried by the binding, the amides in this region may be protected in the complex and exchange slowly. However, one must bear in mind that H–D exchange cannot be used to locate binding interfaces for all protein-protein interactions. Some protein-protein interactions are driven by electrostatic forces of side chains and are unlikely to change the exchange rate of backbone amide hydrogens, particularly if the amide hydrogens are located in stable structural elements such as alpha helicies.
Lastly, H–D exchange can be used to monitor conformational changes in proteins as they relate to protein function. If conformation is altered as result of post-translational modification, enzyme activation, drug binding or other functional events, there will likely be a change to H/D exchange that can be detected.
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