Download Hydrogen exchange mass spectrometry for the analysis of protein

HYDROGEN EXCHANGE MASS SPECTROMETRY FOR THE
ANALYSIS OF PROTEIN DYNAMICS
Thomas E. Wales and John R. Engen*
Department of Chemistry, University of New Mexico, Albuquerque,
New Mexico 87131
Received 5 April 2005; received (revised) 5 July 2005; accepted 6 July 2005
Published online 5 October 2005 in Wiley InterScience (www.interscience.wiley.com) DOI 10.1002/mas.20064
Hydrogen exchange coupled to mass spectrometry (MS) has
become a valuable analytical tool for the study of protein
dynamics. By combining information about protein dynamics
with more classical functional data, a more thorough understanding of protein function can be obtained. In many cases,
protein dynamics are directly related to specific protein
functions such as conformational changes during enzyme
activation or protein movements during binding. The method
is made possible because labile backbone hydrogens in a
protein will exchange with deuterium atoms when the protein is
placed in a D2O solution. The subsequent increase in protein
mass over time is measured with high-resolution MS. The
location of the deuterium incorporation is determined by
monitoring deuterium incorporation in peptic fragments that
are produced after the labeling reaction. In this review, we will
summarize the general principles of the method, discuss the
latest variations on the experimental protocol that probe
different types of protein movements, and review other recent
work and improvements in the field. # 2005 Wiley Periodicals,
Inc., Mass Spec Rev 25:158–170, 2006
Keywords: deuterium; electrospray; MALDI; protein folding;
conformation
I. INTRODUCTION
Proteins are not static structures in solution. They move and flex
naturally or in response to external stimuli. The movements of
proteins, collectively termed protein dynamics, can be extremely
important for protein function. To ascertain how dynamics play a
role in protein function, actual protein motions themselves need
to be investigated and understood. Currently, there are not a large
number of analytical tools capable of providing the necessary
resolution to connect specific protein movements with protein
function. The development of new tools and refinement of those
that are currently in use is therefore of great interest.
To this end, hydrogen exchange (HX) coupled with mass
spectrometry (MS) presents an opportunity for the analysis of
proteins and protein motions in ways that were not imagined even
5 years ago. Traditionally, hydrogen exchange methodology has
been used in conjunction with NMR analysis [see (Dyson &
————
Contract grant sponsor: The National Institutes of Health; Contract
grant numbers: R01-GM070590, R01-GM068901, R24-CA088339,
P20-RR016480.
*Correspondence to: John R. Engen, Clark Hall 242, MSC03-2060,
Department of Chemistry, University of New Mexico, Albuquerque,
NM 87131-0001. E-mail: [email protected]
Mass Spectrometry Reviews, 2006, 25, 158– 170
# 2005 by Wiley Periodicals, Inc.
Wright, 2004) for a review]. In comparison, hydrogen exchange
MS is a more recent development. The first demonstrated use of
HX MS came shortly after the development of electrospray
ionization (Chowdhury, Katta, & Chait, 1990; Katta & Chait,
1991). Important developments through the 1990s were led by
Prof. David Smith [reviewed in (Smith, Deng, & Zhang, 1997;
Engen & Smith, 2001)]. Hydrogen exchange MS has been the
subject of several comprehensive reviews (Kaltashov & Eyles,
2002a,b; Hoofnagle, Resing, & Ahn, 2003; Eyles & Kaltashov,
2004; Garcia, Pantazatos, & Villarreal, 2004). Recent developments in this field, which will be summarized in the following
sections, offer unparalleled limits of detection, low sample
consumption requirements, the promise of single amino acid
resolution, potential for automation and the ability to analyze
increasingly more complex mixtures. Future refinements that
could substantially improve the method will also be discussed.
II. OVERVIEW OF THE METHOD
An general scheme for hydrogen exchange MS experiments is
shown in Figure 1. The integration of deuterium into the
protein(s) of interest relies on the natural phenomena of hydrogen
exchange. The introduction of deuterium into a peptide or protein
can be accomplished in several ways and its incorporation can be
analyzed using several methods. The mass spectrometer is used
to monitor the increase in mass as hydrogen is exchanged for
deuterium. Throughout this review, we discuss several parts in
the scheme (Fig. 1, numbered circles) where recent improvements have been made and where future refinements are
anticipated.
III. HYDROGEN EXCHANGE FUNDAMENTALS
The details of hydrogen exchange mechanisms have been widely
reviewed (Hvidt & Nielsen, 1966; Woodward, Simon, &
Tüchsen, 1982; Englander & Kallenbach, 1984; Kim &
Woodward, 1993; Mayo & Baldwin, 1993; Bai et al., 1995;
Miller & Dill, 1995; Loh et al., 1996; Clarke, Itzhaki, & Fersht,
1997; Kaltashov & Eyles, 2002b; Hoofnagle, Resing, & Ahn,
2003; Eyles & Kaltashov, 2004; Krishna et al., 2004; Smith,
Deng, & Zhang, 1997). Here a short summary, collected from
these references, on the basics of hydrogen exchange and how it is
applied to the study of protein dynamics is presented. Hydrogens
that are located at peptide amide linkages (also referred to
as the backbone amide hydrogens) undergo replacement with
deuterons within 1–10 s when the peptides are incubated in D2O
PROTEIN DYNAMICS BY HYDROGEN EXCHANGE MASS SPECTROMETRY
at pD 7.0. In folded proteins, some backbone amide hydrogens
exchange quickly while others exchange only after months.
The rates of the most slowly exchanging amide hydrogens may
be reduced by as much as 108 of their rates in unfolded forms of
the same protein (Englander & Kallenbach, 1984). Nearly all
peptide amide hydrogens in folded proteins are hydrogen
bonded, either intramolecularly to another part of the protein or
to water. The large reduction of amide hydrogen exchange rates
in folded proteins is primarily due to restricted access of solvent
to the interior of the protein and to intramolecular hydrogen
bonding. It is not possible to differentiate between these two
contributing parameters as they occur concomitantly. At physiological pH, base-catalyzed exchange is the dominant mechanism
for hydrogen exchange. Base-catalyzed isotope exchange can
occur only when a hydrogen bond is severed in the presence of the
catalyst (hydroxide) and the source of the new hydrogen (water).
The rate constant for isotope exchange at each individual
amide linkage in a normally folded protein, kex, can be described
by Equation 1
kex ¼ kf þ ku ¼ ðb þ Kunf Þk2
ð1Þ
where kex is expressed as the sum of the contributions of exchange
from folded (kf) and unfolded (ku) forms of the protein (Kim &
Woodward, 1993). The mechanisms described by Equation 1 are
illustrated graphically in Figure 2. Exchange from the folded
form likely dominates for amide hydrogens that are not
participating in intramolecular hydrogen bonding and that are
located near the surface. Exchange in unfolded forms requires
substantial movement of the backbone. Unfolding to expose
backbone amide sites to deuterium can be isolated to small
regions (localized unfolding) or may involve the entire protein
(global unfolding).
The rate constant for exchange from the folded state, kf, is
described by Equation 2
kf ¼ bk2
ð2Þ
where b is a probability factor for exchange from folded forms
and k2 is the rate constant for HX at each amide linkage in an
unstructured peptide, a value that can be calculated (Bai et al.,
1993). The value for b ranges from 0 to 1 and is a function of
several parameters including solvent accessibility and intramolecular hydrogen bonding. When b is closer to 1, there is a
higher probability that a particular amide hydrogen is exposed
to water and catalyst at the same time that it is also exchange
competent.
Kunf in Equation 1 is the equilibrium constant describing the
unfolding process. HX NMR studies have used denaturants to
distinguish between b and Kunf (Bai et al., 1994; Itzhaki, Neira, &
Fersht, 1997; Chamberlain & Marqusee, 1998). The rate constant
for exchange from unfolded forms of proteins depends on the rate
constant for exchange from an unfolded peptide (k2) as well as the
unfolding dynamics described by k1 and k1 as shown in
Equation 3 where F and U are the folded and unfolded forms,
respectively.
k1
k2
k1
k1
D2 O
k1
FH Ð UH ! UD Ð FD
ð3Þ
&
When k2 >> k1 (termed EX1 kinetics), the rate constant for
exchange from unfolded forms is given by the unfolding rate
constant k1 (Eq. 4).
ku ¼ k1
ð4Þ
However, under physiological conditions it is more common
for k1 >> k2. In this case (EX2 kinetics), the rate constant
for exchange from unfolded forms is given by Equation 5 where
Kunf is the equilibrium constant describing the unfolding
process.
ku ¼
k1
k2 ¼ Kunf k2
k1
ð5Þ
While only a few proteins undergo EX1 kinetics naturally, all
proteins undergo EX2 exchange kinetics under physiological
conditions. EX2 kinetics may be envisioned as involving many
rapid and random visits to a state capable of exchange. However,
the probability of exchange during a single visit is small. EX1
kinetics are described as a cooperative unfolding event involving
several residues, all of which exchange before refolding (k1)
occurs (Eq. 3). Proteins can be induced to exhibit EX1 kinetics
with denaturant (Deng & Smith, 1998) or by increasing pH
(Swint-Kruse & Robertson, 1996). Some proteins may contain
regions that undergo EX1 and EX2 kinetics simultaneously.
Regions in which exchange occurs by either EX1 or EX2 kinetics
can be identified by characteristic isotope patterns in mass spectra
(Miranker et al., 1993).
Structural changes required for HX described by kf and ku
differ in the magnitude of atomic displacement(s) required for
isotope exchange. Due to the highly compact nature of proteins in
their native state, relative to their denatured states, exchange at
individual sites is believed to involve small atomic movements,
probably less than an angstrom, but sufficient to allow diffusion
of OD and D2O to the exchange site (Kim & Woodward, 1993).
In parallel with this highly localized motion, short segments, as
well as the entire backbone of a protein, can exchange through
unfolding processes. Molecular motions associated with unfolding of large segments of the backbone require displacing many
atoms several angstroms from their equilibrium positions in the
native structure and global unfolding requires gross movement of
the entire backbone. Thus, exchange from the folded form (kf) of
a protein involves primarily low amplitude motions (small
displacement) while exchange from unfolded forms (ku) requires
much larger amplitude motions. Results of a theoretical study by
Miller and Dill suggest that large structural changes with little
changes in free energy are possible, but uncommon (Miller &
Dill, 1995).
IV. DEUTERIUM INTRODUCTION: DEUTERIUM,
MEET PROTEIN; PROTEIN, MEET DEUTERIUM
To measure the incorporation of deuterium into a protein, the
protein must, of course, be exposed to deuterium. The
introduction of deuterium sounds like a relatively simple process.
However, different methods of deuterium introduction allow one
to probe different aspects of protein dynamics. In addition, the
sample quantity requirements of the mass spectrometer,
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WALES AND ENGEN
especially in the case of analysis of protein complexes, may
dictate D2O introduction of a specific type.
The primary method for introducing deuterium into a protein
sample is by dilution. Typically, a solution of protein in a protiated
buffer is diluted with a deuterated buffer that has a deuterium
content of 99% or more. Dilutions of 15-fold or greater will
produce final deuterium concentrations of >95%. This serves to
force the labeling reaction (k2) in one direction (see Eq. 3).
However, with this labeling method the original protein sample is
diluted. Such a dilution may not be compatible with the sample
quantity requirements of the mass spectrometer and such diluted
samples may therefore require an additional experimental step to
concentrate the protein sample prior to analysis. Concentration
can be accomplished by rapidly trapping the protein with online
HPLC at higher flow rates (usually >100 mL/min). The
concentration step works best when analyses are performed with
online HPLC-ESI as too much deuterium would be lost if this step
were done prior to MALDI mass analyses (see below). An
alternative to the dilution technique is to carry out a rapid buffer
switch with small gel filtration spin columns (Engen & Smith,
2000). Although the buffer switch technique takes a bit longer than
the dilution method, the protein is not nearly as diluted.
There are two kinds of labeling experiments: continuous
labeling and pulse labeling [see also (Deng, Pan, & Smith,
1999a)]. In continuous labeling experiments (Fig. 1, top right),
protein is exposed to D2O while the populations of folded
and unfolded species are in flux. The populations may be in flux
as a result of natural protein motions that result from some
population-altering force (i.e. the addition of denaturant, change
in pH or temperature, in response to protein function, ligand
binding, or protein–protein complex formation, etc.) designed to
cause a shift in the population of folded versus unfolded (or the
reverse depending on the experiment). Once a protein has made a
transition from a folded to unfolded state, it becomes labeled with
D2O and the mass increases. As the D2O concentration is very
high, once a molecule is labeled, it is not able to revert to a
protiated species (see Eq. 3). In other words, the transition from a
protiated species to one that is deuterated is unidirectional. The
deuterium level in the protein sample at any point in the course of
the labeling experiment integrates the number of molecules in the
sample that had unfolded (or folded) up to that point (Miranker
et al., 1993). With continuous labeling, it is possible to sample the
population of and potentially observe the transition through
various intermediate states whose numbers are very small at any
given moment and therefore may go undetected using conventional spectroscopic methods. Continuous labeling is most useful
for monitoring slow unfolding transitions [i.e. (Engen et al.,
1997)], the majority of unfolding events in proteins. Given
enough time, all proteins should become totally deuterated
during a continuous labeling experiment as a result of protein
motions and protein breathing. Because the transition is very
slow for many proteins, the addition of denaturants may be used
to induce unfolding.
Most HX MS experiments involve continuous labeling
simply because they are technically simpler to perform. Far fewer
experiments are of the pulse labeling variety. Generally in pulse
labeling experiments (Fig. 1, top left), a population of protein
molecules is either induced to undergo some kind of conformational change by addition of a perturbing agent or it is already in
the process of changing its structure through protein folding. The
perturbing agent is most often a chemical denaturant, although
heat, pH or binding to substrates can also be used. The sample is
then exposed to deuterium for a very brief time (the pulse). Only
those molecules that are unfolded when the sample is pulse
labeled will undergo isotopic exchange; the remainder of the
population remains unlabeled. The resulting deuterium levels
then indicate the instantaneous population of folded and unfolded
molecules. Pulsed labeling has recently been used to identify
protein folding mechanisms [see (Wu & Engen, 2004) for a more
detailed description] as well as to probe significantly populated
kinetic intermediate states in a folding reaction (either on or off
pathway intermediates) (Deng & Smith, 1999; Chen et al., 2001;
Wintrode et al., 2003; Mazon et al., 2004; Pan & Smith, 2004; Pan
et al., 2004; Rojsajjakul et al., 2004). With pulse labeling, it is
FIGURE 1. Overall scheme for hydrogen exchange mass spectrometry experiments. A: Pulse labeling. After
a protein has been exposed to a perturbant (chemical denaturant, heat, pH, binding, complex formation,
pressure, etc.), unfolded regions (gray) become labeled with deuterium (red) during a quick pulse of D2O
(typically 10 s). Deuterium exchange is quenched by reducing the pH and temperature. B: Continuous
labeling. D2O buffer is added to a protein (in H2O buffer) such that the final D concentration is >95%. After a
set period of time, an aliquot of the labeled protein is removed from the original tube and mixed with quench
buffer to reduce the pH and temperature. Aliquot removal is repeated for subsequent labeling times. The
protein concentration and solution volume are controlled such that all the aliquots are identical upon quench
except for the amount of time the protein was exposed to D2O. C: Localized exchange information. Quenched
samples (from part A, part B, or both) are digested with pepsin or another acid protease. The resulting peptides
are analyzed with online HPLC-ESI-MS or with MALDI-MS. The resulting data analysis provides
information on deuterium exchange in short fragments of the peptide backbone. D: Global exchange
information. Quenched samples (from part A, part B, or both) are directly analyzed with HPLC-ESI-MS or
MALDI-MS. The data provide a global picture of how the protein behaves in D2O. It is often recommended
that Part D be performed prior to Part C. Areas of recent and future improvement have been marked with
numbers. 1—Robotic automation of mixing, buffer addition, etc.; 2—Rapid-mixing techniques such as
quench-flow analyses (see text for details); 3—Analysis based upon relative deuterium levels instead of
absolute levels (see text for details); 4—Use of acid proteases other than porcine pepsin. Alternative proteases
include protease type XIII from Aspergillus saitoi and protease type XVIII from Rhizhopus species (Cravello,
Lascoux, & Forest, 2003). 5—CID, ECD, and ETD as fragmentation techniques to provide single amino acid
resolution of hydrogen exchange information; 6—Solvent-free MALDI; 7—Nano-ESI-MS and other
miniaturization; 8—chromatographic and labeling techniques to accomplish HX MS of single proteins in
complexes and mixtures; 9—automation of data analysis and increase interpretation speed.
160
FIGURE 1.
161
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WALES AND ENGEN
FIGURE 2. Models for hydrogen exchange into the folded form (A) and
into unfolded forms (B) of proteins. The unfolding and refolding rate
constants are described by k1 and k1, respectively. k2 is the rate constant
for exchange from an unfolded peptide, a value that can be calculated
(Bai et al., 1993). See text and Equation 3 for further details.
possible to complement data obtained using conventional
spectroscopic and NMR experiments (Nishimura, Wright, &
Dyson, 2003).
Pulse labeling experiments may also be performed using a
quench-flow scheme originally described for HX MS by (Yang &
Smith, 1997) and more recently discussed by Konerman &
Simmons, 2003; Wintrode et al., 2003. The addition of deuterium
and other steps in the preparation of pulse or continuous labeled
samples has been automated (see section on Automation later in
this review). With automation, the reproducibility of sample
preparation is improved. It is anticipated that fully automated
sample preparation devices will become commonplace (Fig. 1,
circles 1,2).
during the HPLC step and for MALDI analyses, losses may occur
during the sample preparation process. However, an adjustment
can be made to compensate for back-exchange. An adjustment
calculation was first described by Zhang & Smith, 1993. In the
appendix to their 1993 publication, they describe the derivation,
accuracy, and proper use of the equation used for the calculation.
While other adjustment methods have been described (Resing,
Hoofnagle, & Ahn, 1999; Hoofnagle, Resing, & Ahn, 2004), they
do not significantly improve upon this original comprehensive
description.
When properly controlled, the back-exchange in most
modern and well tuned ESI mass spectrometers is on the order
of 1–3%. An additional 10–20% of the deuterium label may be
lost during in-solution digestion and HPLC separation depending
on the length of time that it takes to perform each step. However,
peptide and protein recovery differ widely [discussed in (Pan &
Smith, 2004)]. More losses can be expected when MALDI is used
for the analysis [see below and (Kipping & Schierhorn, 2003)].
However, even when 15% of the label reverts back to hydrogen,
less than 1 in 6 deuterium is lost. The end result is that if two
curves indicating changes in deuterium levels in a given protein
or peptide are obtained and show a difference of 2–3 deuterium, a
correction for back-exchange will only change the difference
between the two curves by 0.3–0.4 Da. Therefore the comparison
of exchange curves with and without the correction yields almost
no new information except for a closer approximation of the
number of deuterium that were incorporated into the protein or
peptide of interest. An example of this is shown in Figure 3. Here,
the back-exchange correction was applied to raw data where D2O
losses were abnormally high (35%). After correction for the
V. IMPROVING MASS ANALYSIS
A. Retaining the Label
Once the incorporation of the isotope label is complete, the task
becomes the identification of which amides have been deuterated. To maximize the amount of retained label, the deuterium
back-exchange to hydrogen must be controlled. Back-exchange
is the undesirable exchange of the deuterium for hydrogen and
results in loss of some of the label. To minimize back-exchange,
sample analysis must be as rapid as possible and be done at 08C.
The majority of back-exchange occurs because proteolytic
digestion and the subsequent analysis of the deuterium levels
are done with protiated solvents. For ESI analyses, losses occur
162
FIGURE 3. Back-exchange correction. A: Back-exchange adjustment
equation as described by (Zhang & Smith, 1993). 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 exchangeable amide hydrogens in the sequence of
interest. B: Example of the use of the back-exchange correction in part A.
The average deuterium loss for this example peptide was 35%. Closed
circles: raw data; Open squares: raw data adjusted using the equation in
(A); Dashed line: the calculated amount (Bai et al., 1993) of deuterium in
an unstructured peptide with the same sequence as the example peptide
examined under identical conditions (pH 7.0 and 228C).
PROTEIN DYNAMICS BY HYDROGEN EXCHANGE MASS SPECTROMETRY
&
samples and all other variables related to back-exchange cancel
out.
B. Localizing the Label
FIGURE 4. Model peptide showing the fragmentation sites for b/y and
c/z ions [see Roepstorff & Fohlman, 1984].
35% D2O loss, the curve is shifted upward, with the most
significant deviation from the raw, uncorrected data occurring at
larger deuterium levels. This is an extreme example, as many
peptides do not loose 35% of their label during a well controlled
experiment.
The back-exchange correction is only necessary when one
wishes to know the exact number of deuterium atoms in a given
protein or peptide fragment. In many cases where biological
functional information is being probed, the actual number of
deuterons that have exchanged is not as important as where the
exchange has occurred. For example, if deuterium levels are
being compared in peptic fragments of a folded and denatured
version of the same protein, the location of the unfolding is of
primary interest. The same protein is being analyzed, but under
different experimental conditions. In such cases, a relative
deuterium level can be used rather than making a back-exchange
correction to obtain the absolute number of deuterons for each
exchange time-point (Fig. 1, circle 3).
There are several advantages of using relative levels rather
than absolute deuterium levels. First, as proper back-exchange
correction relies on analysis of a totally deuterated form of the
molecule of interest, a totally deuterated form must be prepared.
It can be difficult to prepare such a control, especially for larger
proteins. As one is never really certain that the totally deuterated
control sample is really 100% deuterated at all backbone amide
positions, one can never really be sure of the validity of a
correction that assumes 100% deuteration. Using relative
deuterium levels does not require the preparation or analysis of
a totally deuterated sample. Second, because back-exchange is a
complex process that depends to some extent on the sequence
(Zhang & Smith, 1993), recoveries differ widely between
different proteins and peptides [discussed in (Pan & Smith,
2004)]. Sequence variation is no longer a factor when using the
relative method because the same sequence is being compared to
itself. The only variable that has changed is the conformation of
the protein(s). An added benefit of relative results is that other
variables such as slight changes in buffer pH, temperature,
concentration, etc. all cancel out. Ordinarily these types of
variables, in addition to back-exchange variability, are significant
enough to prevent the use of relative deuterium levels to compare
samples obtained at different times. To take advantage of the
relative method, all experiments that one wishes to compare must
be completed together under identical experimental conditions.
For experiments performed at the same time and under identical
conditions, back-exchange is statistically the same for all
Determination of deuterium levels in whole proteins does not
provide the type of local information desired. In other words, the
spatial resolution is very low. To increase the resolution,
proteolytic digestion is primarily used [first described by Zhang
& Smith, 1993]. Because digestion of the deuterium labeled
protein(s) must occur under quench conditions, acid proteases
must be used. To date, the best acid-protease for these purposes is
pepsin. A few complications exist however. As pepsin is a nonspecific protease (it generally cleaves at hydrophobic residues),
the sites of backbone cleavage cannot be predicted from the
amino acid sequence. However, pepsin will cut in the same place
given the same conditions, so reproducibility is not a problem. To
be certain of the identity of each peptic peptide, it becomes
necessary to sequence those that are generated. Most commonly,
this is accomplished using tandem MS techniques. A further
complication is that peptic digestion of large proteins may
produce undesirably long peptides (>15 residues long) rather
than the more desirable short ones (5–10 residues).
Eric Forest and colleagues recently demonstrated the use of
acid proteases other than pepsin for HX MS analyses (Cravello,
Lascoux, & Forest, 2003) (Fig. 1, circle 4). However, pepsin was
still the most efficient protease. The use of multiple enzymes
produced many overlapping fragments. Overlapping peptides are
highly desirable (Garcia, Pantazatos, & Villarreal, 2004) as they
increase the spatial resolution. Multiple shorter peptides were
also obtained from regions where pepsin cleavage produced
single, long peptide fragments. A form of pepsin that cleaves with
a greater specificity would significantly improve the digestion
step for HX MS. Enzyme engineering should be able to generate
recombinant analogues of pepsin that have increased specificity,
as seen with trypsin or other specific proteases.
In addition to the complications already mentioned, another
significant problem with using pepsin that is often overlooked is
the interpretation of small deuterium changes in the long peptic
fragments. For example, a change of 2 deuterium in a peptide of
20 amino acids cannot be attributed to changes in dynamics
within the whole peptide. While curve fitting (Zhang & Smith,
1993) serves to classify the hydrogens into categories, it cannot
identify them. Careful data interpretation is necessary when large
fragments are involved. A method to further improve the spatial
resolution is needed.
C. New Fragmentation Methods
It would be most desirable to be able to monitor deuterium
exchange at individual amino acids with MS. Attempts have been
made to use CID to fragment peptic peptides into shorter pieces
(b/y ions, see Figure 4) (Deng, Pan, & Smith, 1999b; Demmers
et al., 2000, 2002; Kim et al., 2001; Hoerner et al., 2004). It was
originally observed that b ions from high-energy CID with argon
as the collision gas yielded deuterium levels that were consistent
with NMR measured deuterium levels (Deng, Pan, & Smith,
1999b; Kim et al., 2001). However, deuterium was apparently
‘‘scrambled’’ in most y ions via migration during the fragmenta163
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WALES AND ENGEN
tion process. The scrambling process seemed to depend on the
sequence (Demmers et al., 2002) and other multiple factors
(Hoerner et al., 2004). Other work has shown that scrambling
appears to be 100% in both b and y ions (Jorgensen et al., 2005).
Other methods of fragmentation may offer a solution to the
scrambling issue. Kaltashov and Eyles have extensively
discussed the use of FTMS and electron capture dissociation
(ECD) methodology towards this end (Kaltashov & Eyles,
2002a,b; Eyles & Kaltashov, 2004). It has been shown that the
fragmentation of ions in ECD occurs before there is an
opportunity for the energy to be randomized to a more probable
bond-cleavage site (Turecek & McLafferty, 1984; Zubarev,
Kelleher, & McLafferty, 1998; Horn, Ge, & McLafferty, 2000).
This rapid and effective activation method may therefore result in
a decrease in the hydrogen scrambling as opposed to the slower
CID activation of ions. A report on the use of ECD to fragment
intact proteins after HX labeling has recently appeared
(Charlebois, Patrie, & Kelleher, 2003). Fragments produced by
ECD of whole proteins (c/z ions, see Figure 4) may still result in
incomplete sequence coverage, especially for larger proteins. It
may still be advisable to digest larger proteins and protein
complexes with pepsin and perform ECD on the peptides, thereby
providing single amino acid resolution (Fig. 1, circle 5). Other
instrumental methods have also reported on fragmentation of
proteins after HX (Akashi, Naito, & Takio, 1999; Lanman et al.,
2003).
Unfortunately, the ECD methods described to date are
limited to Fourier transform instruments for technical reasons
and are therefore cost prohibitive for many research laboratories.
Recent descriptions of ECD and ETD (electron transfer
dissociation) in ion traps may bring this technology into the
hands of researchers using standard mass analyzers (Baba et al.,
2004; Syka et al., 2004). ECD/ETD technology should facilitate
the eventual analysis of deuterium exchange at individual amino
acids without concern for potential label scrambling. Such
experiments would be a major improvement to the HX MS
method (Fig. 1, circle 5).
VI. WHICH KIND OF MASS SPECTROMETRY?
ESI VERSUS MALDI
The use of ESI-MS for the analysis of hydrogen/deuterium
exchange experiments has become very popular since the mid
1990s. Approximately 80% of the articles published with respect
to HX MS in the past 5 years have employed electrospray as the
ionization source. ESI-MS has become the most commonly used
ionization method for HX MS partially because the quenched
sample is introduced directly via HPLC into the electrospray
source. The advantages of using electrospray MS are described
below.
MALDI-MS can also be used for HX MS protein dynamics
studies. E. A. Komives reported the first use of MALDI for the
analysis of hydrogen exchange content in the mapping of a
protein–protein interface (Mandell, Falick, & Komives, 1998a).
She has used MALDI HX MS to identify several binding sites in
the cyclic-AMP-dependent protein kinase (PKA) complex with
the kinase inhibitor and ATP (Mandell, Falick, & Komives,
1998a) and to determine the antibody–antigen recognition site
164
for a monoclonal antibody raised against human thrombin
(Baerga-Ortiz et al., 2002). Others have reported on the use of
MALDI to probe the folding and assembly of viral capsids (Tuma
et al., 2001) and to investigate protein conformational changes as
a result of polyvaline and polyleucine a-helix aggregation
(Hosia, Johansson, & Griffiths, 2002). Topological information
about yeast F1-ATPase, a supramolecular hetero-oligomeric
protein complex of 370 kDa, was obtained using MALDI as the
ionization method (Nazabal et al., 2003).
The advantages of MALDI are that it is, in principle and
generally in practice, easier than ESI for non-experienced users.
All ions are singly charged and this simplifies data interpretation.
There is no HPLC separation and desalting. However, the
disadvantages seem to outweigh the advantages: only 20 % of
all HX MS publications have used MALDI as the ionization
technique. Deuterium losses are significantly higher in MALDI
than in ESI, although attempts have been made to reduce these
loses (Kipping & Schierhorn, 2003). While all of the peptides are
present in one spectrum and no HPLC steps are required
(seemingly a simplification over ESI methods), for large and
complex systems, the spectra become too crowded to interpret the
data. Simple HPLC separation is necessary to temporally resolve
a large number of peptides. Coupling HPLC to ESI has other
advantages: the HPLC step washes away deuterium label present
in the amino acid side-chains (Zhang & Smith, 1993), it is
compatible with all buffers and denaturants, it can afford rapid
concentration of very dilute samples (although ZipTips and the
like can do some concentration in MALDI analyses) and ESI can
handle some buffers and matrices that would be deleterious to
MALDI sample ionization.
The development of new solvent-free sample preparation
methods (Trimpin et al., 2001, 2002) may eliminate some of the
aforementioned complications for MALDI analysis of HX
samples. Eluent from a chromatographic source can be
continuously deposited onto a MALDI sample stage (that has
been pre-coated with matrix) by spraying the eluent at elevated
temperatures (Falkenhagen et al., 2003; Falkenhagen & Weidner,
2004). The advantages of this over the conventional dried droplet
method and the solvent-free grinding method are improved
compatibility of analyte and matrix as well as greater sensitivity.
It remains to be seen if coupling of chromatography to solventfree MALDI spotting will make the use of MALDI more
attractive for HX MS analyses (Fig. 1, circle 6).
VII. THINKING SMALLER
Although an obvious extension of the method, the use of nanoHX MS has not yet become widespread. Off-line nano-ESI-MS
analyses of continuous labeled samples have been used to probe
lipid interactions with transmembrane peptides (Demmers et al.,
2000). The a-helical structures in the molten globule state of wild
type human a-lactalbumin and several proline analogues were
also investigated with off-line nano-ESI-MS (Last et al., 2001).
Complete online digestion, separation and ESI-MS analysis on a
nano scale have also been described (Wang & Smith, 2003) and a
100-fold increase in sensitivity was reported. Nano-HX MS can
be particularly valuable for proteins that are hard to obtain in
large quantities or those that aggregate at higher concentrations.
PROTEIN DYNAMICS BY HYDROGEN EXCHANGE MASS SPECTROMETRY
Many rare, disease-relevant signaling proteins fit into this
category. It is anticipated that more and more proteins and
protein systems will require the use of nano-HX MS in the
coming years (Fig. 1, circle 7).
VIII. MORE COMPLEX WITH COMPLEXES
IN THE MIX
While analysis of proteins in vitro is providing much information
about them, the ‘‘Holy Grail’’ will be to analyze proteins in vivo.
Further, the analysis of protein complexes and protein machines
(Gavin et al., 2002) is becoming more and more commonplace.
Techniques that can work with large protein assemblies and
complex matrices such as cell lysates or organelle preparations
should make this dream a reality. HX MS seems positioned to
make this advance and with proper method development, it may
soon be possible to investigate the dynamics of a single protein
from a large, multi-protein complex or a single protein in a very
complex mixture like cytoplasm.
In vivo hydrogen exchange has already been reported
(Ghaemmaghami & Oas, 2001). Other studies investigating
proteins in cell lysates have also appeared (Engen, Bradbury, &
Chen, 2002). The problem in all these experiments is the
separation or isolation of the protein of interest from other
cellular components. When such complex systems are involved,
mass alone is not sufficient to distinguish the protein of interest
from the components of the matrix. Systems in which protein
fragmentation occurs after initial mass separation (ECD as
described above, or several tandem stages of MS and HPLC) may
help alleviate this problem. Other methods to get around the
complexity problem include tagging the protein of interest so that
it stands out against the background of the matrix (Engen,
Bradbury, & Chen, 2002). These methods, although demonstrated in principle, have yet to be used practically. It would also work
to isolate the protein by affinity purification just prior to mass
analysis. However, the pH requirements for HX MS hinder this
possibility, although other chromatographic methods (IEX) could
potentially be better suited for this purpose. HX MS methods
development is moving in this direction and future investigators
will no doubt solve these technical problems (Fig. 1, circle 8).
IX. AUTOMATION
In an ideal automated system, an operator would select a time
point for analysis and automated sample preparation and analysis
instrumentation would do the rest. The end result would be a
graph of hydrogen exchange for a given region of a protein, also
automatically represented in a 3-dimensional modeling program.
While the plumbing and sample introduction (Fig. 1, circles 1, 2)
parts of this ideal scenario have been automated fairly easily
(Woods, 1997, 2001a,b,c), the rest has not come so easily. Data
processing at the end of data acquisition is substantial and mostly
has been done manually. Automation of the data analysis
[mentioned in (Garcia, Pantazatos, & Villarreal, 2004)] aids in
processing but is not yet ideal. Commercial versions of the
software are also not available. The coming years should see
more development in this area (Fig. 1, circle 9).
&
X. RECENT EXAMPLES OF WORK USING HXMS
There are many examples of analyses performed with HX MS
over the past 5 years. We would like to highlight several different
types of analyses with specific examples from the literature to
demonstrate the range of possibilities.
A. Protein Assemblies
MS has opened the door to investigating the dynamics of large
protein assemblies. The HIV-1 capsid assembly is one such
system that has been studied recently with HX MS (Lanman et al.,
2003, 2004). HX MS experiments were performed on unassembled and assembled capsid tubes to determine the putative N
domain to C domain interactions in the assembled capsid tubes.
Interactions between helices I and II of the N domain were
identified. In addition, a previously unrecognized inter-subunit N
domain–C domain interaction was observed (Lanman et al.,
2003). Further experiments on both the immature and mature
virion revealed that helix I and helix II were involved in intersubunit interactions in both the mature and immature virion
(Lanman et al., 2004). Together, these results illustrate the use of
HX MS methodologies to identify possible therapeutic targets
that cause disruption of viral capsid assembly. Other groups have
also investigated viral capsids with HX MS (Wang, Lane, &
Smith, 2001).
B. Protein Dynamics
Conformational changes may occur as a result of phosphorylation or by means of single amino acid mutation. Two recent
studies used HX MS to describe the effects of both phosphorylation and single amino acid mutation on the COOH-terminal Src
kinase (Csk), an enzyme that regulates signaling by the Srcfamily of tyrosine kinases. To provide a structural framework for
understanding phosphorylation-driven protein conformational
changes, HX MS was used to monitor the effects of nucleotide
binding on the solution conformation of Csk in the presence of
ADP and AMPPNP (a non-hydrolysable ATP analogue) (Lanman et al., 2003). The results implied that phosphorylation of Csk
results in conformational changes that may influence regulatory
motions in the catalytic pathway. The HX MS data also showed
that the conformational states of the protein are different
depending on whether substrate or product is bound (Lanman
et al., 2003).
To probe the conformational consequences of a single amino
acid substitution at amino acid position F183 in Csk, believed to
be of significant importance in the communication between the
SH2 and kinase domains of Csk, HX MS experiments for three
substitution analogues at this specific position were performed.
Three substitutions were explored: F ! G, F ! Y, F !A. HX
MS experiments revealed that compared to the wild-type protein,
glycine substitution at F183 reduced flexibility in several peptide
regions in Csk, tyrosine mutations increased flexibility, and
alanine mutations showed mixed results. In addition, the data
suggested that each mutant was well folded, since major regions
in all 3 domains had exchange patterns that were indistinguishable from those for the wild-type protein (Lanman et al., 2004).
165
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WALES AND ENGEN
Another example of the use of HX MS to probe protein
conformational dynamics involved the heat shock transcription
factor s32 (Rist et al., 2003). HX MS proved particularly valuable
for this task as previous CD and fluorescence measurements were
only able to detect global and local changes, respectively. As this
protein tended to aggregate at high concentrations, NMR was not
an available tool to study the protein solution conformation. HX
MS experiments were designed to probe whether or not s32 acted
as a thermosensor. The folded states of s32 were investigated at
two separate temperatures: 37 and 428C, optimal growth
conditions, and heat-stress conditions, respectively. The results
indicated that there was a high degree of protein flexibility at
378C, and that there was reversible unfolding of a small structural
motif at 428C. The location of this unfolded region was identified
by analyzing the HX into pepsin fragments of the deuterium
labeled samples. From these data, a map of the region that
unfolded at elevated temperatures was produced.
C. Protein Unfolding/Refolding
Studies of protein folding/unfolding have traditionally been
performed using the more conventional spectroscopic based
techniques: NMR, IR, CD, and Fluorescence. Hydrogen
exchange coupled to MS is also a useful tool for the study of
protein folding/unfolding reactions. To illustrate this, some
recent examples using pulse labeling HX MS are presented
below.
In studies of rabbit muscle triosephosphate isomerase (TIM)
(Pan et al., 2004), pulse labeling denaturation experiments have
revealed a bimodal isotope pattern of labeled protein in the raw
MS data, which is evidence for two-state unfolding behavior.
However, during the renaturation experiments there were three
envelopes of isotope peaks present, suggesting that there was an
intermediate in the refolding pathway. To obtain information
about this intermediate, peptic digestion was performed after the
pulse labeling experiment. The data showed that the intermediate
was a form in which the C-terminal half was folded while the Nterminal half was not. Further, TIM folding fit a 4 þ 4 model of
folding for (ba)8-proteins. In this set of experiments, HX MS
provided location information regarding the folding intermediates as well as following both the refolding and unfolding
properties.
In another example, pulse labeling HX MS was employed to
better characterize the molten globule intermediate state during
the unfolding of the multidomain dimeric protein MM-CK
(Mazon et al., 2004). In 0.8 M GdmCl, where the molten globule
state of this protein was maximally populated, MM-CK exhibited
a highly fluctuating structure that allowed for total deuteration.
The study also used intrinsic fluorescence, ANS binding, and farUV CD to probe the molten globule state of the protein. With HX
MS, only two species were detected during GdmCl denaturation
as opposed to more intermediate states detected by the
conventional methods.
The a-subunit of tryptophan synthase is a 29 kDa, single
domain protein that unfolds according to a four-state equilibrium
unfolding model. There are two intermediate states, I1 and I2.
The intermediate state I1 contains a significant amount of
secondary structure while I2 contains no detectable secondary
166
structure and mostly resembles the denatured state. Both of these
intermediates are on-pathway kinetic species. HX MS was used
to map the stable secondary structure in the I1 equilibrium
intermediate (Rojsajjakul et al., 2004). Such information
provided insights into the relationship between sequence,
structure, and folding in the a-subunit. The identification of
protected regions in the I1 intermediate was accomplished. The
identified regions (most of the N-terminal region 12–128 with 5
peptides regions of exception three of which are exposed loops)
represented a contiguous domain (a-helices 1–3 and b-strands
1–4). It was also shown that the C-terminus, residues 128–268,
was either unfolded or weakly folded in this intermediate. A
refolding study of the urea denatured a-subunit of tryptophan
synthase using pulse-quench HX MS has shown that there is an on
pathway kinetic folding intermediate that shares a similar folded
protein core with the I1 equilibrium intermediate (Wintrode et al.,
2005). Together the data obtained for the equilibrium and kinetic
intermediates show that the latter stages of the folding reaction
for the a-subunit of tryptophan synthase are under thermodynamic control (Wintrode et al., 2005).
When studying protein folding/unfolding reactions, it is not
uncommon to see HX MS technologies combined with other
methods. One such recent example is the integration of HX MS
with a cyanylation-based methodology (Li et al., 2004) to better
understand the conformation of disulfide-bonded proteins and
intermediates during refolding reactions. In the cyanylationbased methodology, a protein containing disulfide bonds is
trapped and the disulfide structure of a given cystinyl protein
folding intermediate is identified and preserved. HX MS is then
used to assess the other conformational features of the
intermediate. These technologies were used to trap a 1-disulfide
bond (early folding) intermediate and a 2-disulfide bond (later
forming) intermediate of long Arg3 insulin-like growth factor-I
(LR3IGF-I) (Li et al., 2004). HX MS data showed an increasing
degree of protection from exchange as a function of disulfide
bond formation. There was significantly more secondary
structure after formation of the second disulfide bond (specifically in helix 3).
It is clear that HX MS technology is even more valuable
when combined with conventional methods for the analysis of
protein folding and unfolding reactions. As spectroscopic
methods such as CD, IR, and Raman spectroscopy monitor
globally averaged changes to protein secondary structures, and
fluorescence monitors the exposure of aromatic residues to
solvent, there is the possibility that they might not be sensitive to
specific, local changes in protein conformation and dynamics.
One can see from the above examples that HX MS can be an
important complement to the traditional biophysical techniques
to probe protein folding/unfolding reactions. HX MS may reveal
a wealth of additional information that would otherwise go
undetected.
D. Binding Experiments
As a result of their significance to protein folding, stability,
association and function, binding interactions are under intense
investigation. These interactions include protein:small molecule,
protein:polypeptide, protein:lipid, protein:nucleic acid, and
PROTEIN DYNAMICS BY HYDROGEN EXCHANGE MASS SPECTROMETRY
protein:protein binding. Much information about the conformational changes that a protein undergoes during ligand binding
have been determined with high-resolution X-ray crystallography or NMR structural analyses. HX MS has been used to probe
the location(s) of binding sites of specific ligands on target
proteins for which X-ray crystallographic and NMR methods are
not applicable. Interpretation of these data, however, must be
done with caution as binding is known to cause changes in protein
dynamics and conformation at sites distant from the actual
binding site or interface. In the discussion below there are a few
examples of this observable fact [see also ref. (Mayne et al.,
1992)]. An label and chase method that tries to compensate for
this problem has been described (Garcia, Pantazatos, &
Villarreal, 2004).
There are many examples in recent years of HX MS being
used in conjunction with high resolution structures to explore the
organization and dynamics of complex molecular assemblies.
The 42 kDa eukaryotic protein actin, for example, has been
investigated with HX MS (Chik & Schriemer, 2003). G-actin, Factin (formed by polymerization of monomeric G-actin), F-actin
bound to phalloidin, and a DNaseI:G-actin complex were studied
to provide further information about the structure of F-actin and
the structural effects of phalloidin and DNaseI ligand interactions. Note that the size and nature of polymerized actin (F-actin)
excludes use of X-ray diffraction and NMR. HX MS results
suggested a conformational transition from a ‘‘closed’’ to an
‘‘open’’ state of actin. The changes were localized to the
phosphate binding loops upon polymerization of G-actin.
Additionally, phalloidin binding to F-actin induced the monomer
conformation to adopt a more G-actin-like state with the
phosphate groups excluded from solution, a conformational
change that inhibited phosphate release thereby reducing the rate
of monomer dissociation. HX MS also provided evidence for
conformational changes that occurred away from the DNaseI
binding site of G-actin. These distal changes indicated a possible
alteration of conformational flexibility consistent with previously published data in which the C-terminal residues were
found more accessible to trypsin digestion.
A study aimed at identifying regions in MAP kinases
specific for binding two peptide docking motifs (DEJL and DEF)
included the presentation of the distal effects of these ligand
interactions (Lee et al., 2004). The data revealed that DEJL motif
interactions with p38a MAPK induced enhanced backbone
flexibility in the activation lip, an effect that was shown to be
conserved between different MAP kinases. There were backbone
conformational changes away from the region of the DEF motif
interaction with ERK2 kinase. However, it remained unknown
whether the interactions were attributable solely to allosteric
effects or occurred as a result of alternate binding sites or nonspecific binding of the synthetic peptide to the kinase.
Protein binding to small molecules is of great interest
especially for development of small-molecule therapeutics. A
recent HX MS study illustrating this type of experiment involved
the binding of ATP to a-crystallin (Hasan, Smith, & Smith,
2002). ATP decreased the accessibility of amide hydrogens in
multiple regions of both aA and aB subunits of a-crystallin. Four
regions of a-crystallin, two in aA and two in aB, showed a
significant decrease in the uptake of deuterium. Location
information from HX MS data allowed for the comparison of
&
the regions affected by ATP binding with proposed substrate
binding sites. The authors concluded that ATP binding releases
substrate by means of both direct displacement and a global
conformational change.
Recent HX MS binding studies between papain (target
enzyme) and cystatin (thiol protease inhibitor) demonstrated that
enzyme–inhibitor interactions can be characterized by HX MS
coupled to CID in a hexapole ion-guide using ESI-FTICR MS
(Akashi & Takio, 2000). Binding of cystatin to papain reduced
the flexibility throughout the papain molecule, data that are
consistent with previous structural studies.
HX MS protein:protein binding studies with the complex of
UBC9 and SUMO-1 illustrated several key issues about protein
binding experiments by HX MS (Engen, 2003). First, HXMS
titrations can be used to estimate the Kd for complexes. Further,
backbone amide hydrogen exchange may not be altered in all
proteins during complex formation, especially if the protein
complex is formed primarily via electrostatic side-chain interactions. Finally, by combining site-directed mutagenesis with
HX MS, much more information about the nature of complexes
can be obtained.
XI. LOOKING AHEAD
While HX MS has come a long way in the past 12–15 years,
many more improvements can be made. It has only recently been
widely realized that this technique can provide valuable
information about proteins and protein dynamics that cannot
be easily obtained with other techniques. Improvements in
the method, as suggested by the circles in Figure 1, will only
increase its power. We hope that many new researchers will
realize this and join in the analysis of all proteins with HX MS
technologies.
ACKNOWLEDGMENTS
We gratefully acknowledge support for this work from the
National Institutes of Health (R01-GM070590, R01-GM068901,
R24-CA088339, and P20-RR016480).
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Dr. Thomas E. Wales received his B.A. degree in Chemistry and Spanish (1998) from
Assumption College and his Ph.D. in Analytical Chemistry (2003) from Duke University.
He is currently a postdoctoral fellow with John R. Engen at the University of New Mexico.
His research has focused on the analysis of the protein backbone from both a chemical and
structural perspective. Currently he is investigating the minute time-scale unfolding of
protein backbone motions using hydrogen-deuterium exchange mass spectrometry.
Dr. John R. Engen received B.S. degrees in Molecular Biology (1994) and Biochemistry
(1995) from Union College and his Ph.D. in Analytical Chemistry (1999) from the
University of Nebraska-Lincoln. As an EMBO Fellow, he did postdoctoral work (2000) in
molecular biology and cellular signaling at the European Molecular Biology Laboratory in
Heidelberg, Germany followed by a second postdoctoral appointment (2001) at Los Alamos
National Laboratory. He was appointed an Assistant Professor of Chemistry at the
University of New Mexico in January 2002. His research centers around the analysis of
protein function with mass spectrometry, specifically hydrogen exchange analysis of
structural activation in oncogenic kinases and protein folding during cellular processes.
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