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Complexity of Protein-Protein Interactions

Curator: Larry H. Bernstein, MD, FCAP

Cracking the Complex

Using mass spec to study protein-protein interactions

By Jeffrey M. Perkel | November 1, 2015

http://www.the-scientist.com//?articles.view/articleNo/44317/title/Cracking-the-Complex/

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Mass spectrometry is a proteomics workhorse. By precisely measuring polypeptide masses, researchers can identify and sequence those molecules, and characterize whether and how they have been chemically modified. To twist a phrase, by their masses you shall know them.

But many proteins do not act in isolation. Critical biological processes such as DNA replication, transcription, translation, cell division, and energy generation rely on the action of massive protein assemblies, many of which comprise dozens of subunits. While these clusters are ripe for study, few traditional mass spectrometric methods can handle them.

Indeed, protein complexes are unwieldy for many types of analysis, says Philip Compton, director of instrumentation at the Proteomics Center of Excellence at Northwestern University in Evanston, Illinois. Most complexes are held together by noncovalent interactions, assemble only transiently, or are located in the cell membrane—all of which complicate sample preparation, he explains. Also, while some complexes are relatively abundant, others are rare, further thwarting detection and analysis.

For mass spectrometry specifically, however, the problem with analyzing protein complexes, which can weigh in at 500 kDa, is size. “In a mass spec, things of that size have traditionally been fairly difficult to handle,” Compton says. Even if you can deliver them into the spectrometer itself, you need a way to figure out which proteins are present, and in what stoichiometry. Plus, normal sample preparation procedures tend to denature proteins, ripping complexes apart.

Still, researchers are increasingly keen to train their mass specs on intact protein assemblies. The Scientistasked four protein-complex experts about the approaches they use in their own labs. This is what they said.

Determining subunit composition 

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GETTING TOGETHER: Lactate dehydrogenase from human skeletal muscle comprises four identical M subunits, shown here in different colors.  FVASCONCELLOS/WIKIMEDIA COMMONS

RESEARCHER: Philip Compton, Director of Instrumentation, Proteomics Center of Excellence, Northwestern University

PROJECT: High-throughput top-down proteomics

SOLUTION: If protein complexes are onions, Compton needs a way to iteratively peel off the layers to see what’s inside. Working with researchers at Thermo Fisher Scientific, Compton is developing an Orbitrap-based mass spectrometer that can do just that, or perform what is called an MS3 study.

Basically, an MS3 experiment involves weighing all the complexes in a sample fraction—there could be as many as 10 or 15 at a time—grabbing one, smashing it into inert-gas molecules to eject a subunit, weighing and sequencing the cast-off piece, and then repeating the process.

That’s the goal, but because that instrument is not yet built, Compton must temporarily content himself with what he calls a “pseudo-MS3” experiment. Basically, instead of one seamless workflow, the instrument shatters the complex, weighs the pieces that come off it, and then repeats the process, only this time capturing and fragmenting those ejected pieces for subsequent analysis (Anal Chem, 85:11163-73, 2013). “We’re kind of splitting it into these two different steps; that accomplishes essentially the same thing,” Compton says.

Compton and his team are still ironing out the kinks, but they have begun applying the approach to protein complexes involved in metabolism. One of these, lactate dehydrogenase (LDH), is a 145-kDa tetramer comprising M (muscle) and H (heart) subunits that can exist in any of five configurations (MMMM, MMMH, MMHH, MHHH, and HHHH). Using the MS3 workflow, Compton says he can differentiate these “multiproteoform assemblies,” as well as any posttranslational modifications those subunits may bear, and determine the abundance of each. Now he hopes to apply the approach to quantify LDH differences between cell and tissue types.

From Protein Complexes to Subunit Backbone Fragments: A Multi-stage Approach to Native Mass Spectrometry

EXTEND YOUR RANGE: Compton’s team uses a souped-up version of Thermo Fisher’s Orbitrap-based Q Exactive HF mass spectrometer, which among other things features a fourfold wider mass range. Other researchers can perform similar work using Thermo’s Exactive Plus EMR Orbitrap system, an off-the-shelf, “extended mass range” instrument. But, because the EMR lacks the “high-mass isolation capabilities” of Compton’s bespoke hardware, the application range is more limited, he says. “You can still do a similar experiment to us, provided that you have one clean [purified] complex.”

Mapping protein-protein interaction interfaces
RESEARCHER: Igor Kaltashov, Professor of Chemistry, University of Massachusetts Amherst

PROJECT: Probing the interactions of candidate protein therapeutics with their molecular targets

SOLUTION: Most attempts at studying protein complexes deliver them to the mass spec intact. Kaltashov takes a different approach, using a technique called hydrogen-deuterium exchange (HDX).

It works like this: proteins (like other molecules) pass hydrogen atoms back and forth with the solvent that surrounds them. Normally, one hydrogen is simply swapped for another, and nobody is the wiser. But in deuterated (“heavy”) water, as hydrogens are swapped at the protein surface, the protein gets slightly heavier as deuterium molecules replace some of the hydrogens. This allows researchers to probe how accessible different pieces of the protein are to the solvent, based on how much deuterium they pick up from the buffer, and how quickly they do so.

As Kaltashov explains, HDX can be used to study any event that might alter the accessibility of different protein regions to the solvent that surrounds them. Those events include protein folding and aggregation, but also protein-protein interactions. “Once two proteins bind to each other, solvent would be excluded from the interface, and that would be reflected in the hydrogen-deuterium exchange kinetics,” he says. That change is evident when compared to the proteins in isolation.

In a 2009 review, Kaltashov demonstrated the process with transferrin, an iron transport protein, and its receptor. After undergoing the exchange reaction, the proteins were fragmented to peptides and analyzed piecemeal. Some peptides exhibited no hydrogen-deuterium exchange, he says. That suggests they were never exposed to solvent because they were buried inside the protein core. Other peptides exchanged hydrogens with the solvent at the same rate regardless of receptor binding, indicating they are not part of the protein-receptor interface. A third set of peptides, though, exhibited clear differences in the presence and absence of receptor, marking those as elements of the protein-protein interaction domain (Anal Chem, 81:7892-99, 2009).

“You can actually localize these sites and obtain information both on the strength of the binding [interactions] and the structural characteristics of the interface region,” Kaltashov says.

H/D exchange and mass spectrometry in the studies of protein conformation and dynamics: Is there a need for a top-down approach?

Introduction: HDX MS in the context of structural proteomics

Since the majority of HDX MS studies target protein dynamics under near-native conditions, the experiments are typically carried out at physiological pH, where the progress of the exchange is followed by monitoring the protein mass change. The direct infusion scheme offers the simplest way to carry out such measurements, either in real time⁷ or by using on-line rapid mixing.¹⁰ However, in many cases these straightforward approaches cannot be used, as they limit the choice of exchange buffer systems to those compatible with electrospray ionization (ESI). To avoid this, HDX can be carried out in any suitable buffer followed by rapid quenching (lowering pH to 2.5-3 and temperature to near 0°C). Dramatic deceleration of the intrinsic exchange rate for backbone amides under these conditions allows the protein solution to be de-salted prior to MS analysis. Additionally, the slow exchange conditions denature most proteins, resulting in facile removal of various binding partners, ranging from small ligands to receptors (their binding to the protein of interest inevitably complicates the HDX MS data interpretation by making accurate mass measurements in the gas phase less straightforward).

An example of such experiments is shown in Figure 2, where HDX is used to probe the higher order structure and conformational dynamics of metal transporter transferrin (Fe₂Tf) alone and in the receptor-bound form. Both Tf-metal and Tf-receptor complexes dissociate under the slow exchange conditions prior to MS analysis; therefore, the protein mass evolution in each case reflects solely deuterium uptake in the course of exchange in solution. The extra protection afforded by the receptor binding to Tf persists over an extended period of time, and it may be tempting to assign it to shielding of labile hydrogen atoms at the protein-receptor interface. However, this view is overly simplistic, as the conformational effects of protein binding are frequently felt well beyond the interface region. The difference in the backbone protection levels of receptor-free and receptor-bound forms of Fe₂Tf appears to grow during the initial hour of the exchange (Figure 2), reflecting significant stabilization of Fe₂Tf higher order structure by the receptor binding. Indeed, while the fast phase of HDX is typically ascribed to frequent local fluctuations (transient perturbations of higher order structure) affecting relatively small protein segments, the slower phases of HDX usually reflect relatively rare, large-scale conformational transitions (transient partial or complete unfolding). This is why global HDX MS measurements similar to those presented in Figure 2 are can be used to obtain quantitative thermodynamic characteristics for protein interaction with a variety of ligands, ranging from metal ions¹¹ and small organic molecules ¹² to other proteins¹³ and oligonucleotides.¹⁴

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HDX MS of Fe₂Tf in the presence (blue) and the absence (red) of the cognate receptor. The exchange was carried out by diluting the protein stock solution 1:10 in exchange solution (100 mM NH₄HCO₃ in D₂O, pH adjusted to 7.4) and incubating for a certain period of time as indicated on each diagram followed by rapid quenching (lowering pH to 2.5 and temperature to near 0°C). The black trace shows unlabeled protein.

While global HDX MS measurements under near-native conditions provide valuable thermodynamic information on proteins and their interaction with binding partners, structural studies (e.g., localizing the changes in Tf that occur as a result of receptor binding) must rely on the knowledge of exchange kinetics at the local level. This is typically accomplished by carrying out proteolysis under the slow exchange conditions following the quench of HDX.⁶ Here we will refer to this approach as “bottom-up” HDX MS, by drawing analogy to a bottom-up approach to obtain sequence information.¹⁵ An example is shown in Figure 3, where Fe₂Tf undergoes exchange in solution in the absence and in the presence of the receptor, followed by rapid quenching of HDX reactions, protein reduction and digestion with pepsin and LC/MS analysis of the deuterium content of individual proteolytic peptides.

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Localizing the influence of the receptor binding on backbone protection of Fe₂Tf using bottom-up HDX MS on the physiologically relevant time scale. The panels show isotopic distributions of representative peptic fragments derived from the protein subjected to HDX in the presence (blue) and the absence (red) of the receptor and followed by rapid quenching. Dotted lines indicate deuterium content of unlabeled and fully exchanged peptides. Colored segments within the Fe₂Tf/receptor complex show location of the peptic fragments.

Evolution of deuterium content of various peptic fragments in Figure 3 reveals a wide spectrum of protection, which is clearly distributed very unevenly across the protein sequence. While some peptides exhibit nearly complete protection of backbone amides (e.g., segment [396-408] sequestered in the core of the protein C-lobe), exchange in some other segments is fast (e.g., peptide [612-621] in the solvent-exposed loop of the C-lobe). The influence of the receptor binding on the backbone protection is also highly localized. While many segments appear to be unaffected by the receptor binding, there are a few regions where exchange kinetics noticeably decelerates (e.g., segment [71-81] of the N-lobe, which contains several amino acid residues that form Tf/receptor interface according to the available model of the complex based on low-resolution cryo-EM data¹⁶).

Although the increased protection of backbone amides proximal to the protein/receptor binding interface is hardly surprising, HDX MS data also reveal a less trivial trend, acceleration of exchange kinetics in some segments of the protein as a result of receptor binding (such behavior is illustrated in Figure 3 with segment [113-134], a part of the N-lobe that is distal to the receptor). Therefore, in addition to mapping binding interface regions, HDX MS also provides a means to localize the protein segments that are affected by the binding indirectly via allosteric mechanisms. However, this example also highlights one of the limitations of HDX MS, namely inadequate spatial resolution. This peptic fragment spans several distinct regions of the protein (an α-helical segment, a β-strand, and two loops). The moderate level of protection observed in this segment in the absence of the receptor binding (fast exchange of three protons followed by slow exchange of the rest) is likely to be a result of averaging out very uneven protection patterns across this peptide. Even smaller peptides may comprise two or more distinct structural elements, such as segment [71-81] spanning three distinct regions of the protein (an α-helical segment, a β-strand, and a loop connecting them).

In some favorable cases spatial resolution in HDX MS of small proteins (<15 kDa) may be enhanced up to a single residue level by analyzing deuterium content of a set of overlapping proteolytic fragments.¹⁷However, single-residue resolution has never been demonstrated in HDX MS studies of proteins falling out of the mass range routinely accessible by NMR, although overlapping peptic fragments frequently provide moderate improvement of spatial resolution.

In addition to limited spatial resolution, the “classic” HDX MS scheme frequently suffers from incomplete sequence coverage, especially when applied to larger and extensively glycosylated proteins. Proteins with multiple disulfide bonds constitute another class of targets for which adequate sequence coverage is difficult to achieve, although certain changes in experimental protocol can alleviate this problem, at least for smaller proteins.¹⁸ Typically, an 80% level of sequence coverage is considered good, although significantly lower levels may also be adequate, depending on the context of the study.

Protein processing in HDX MS experiments is carried out under the conditions that minimize the exchange rates for backbone amides. Since these slow exchange conditions are highly denaturing for most proteins, both intact protein and its proteolytic fragments lack any protection and inevitably begin to lose their labile isotopic labels, despite low (but finite) intrinsic exchange rates.¹⁹ This phenomenon, known as “back-exchange,” may be accelerated during various stages of protein processing, e.g. during the chromatographic step.²⁰ Although back-exchange was frequently evaluated in early HDX MS studies using unstructured model peptides, the utility of this procedure is questionable, since the intrinsic exchange rates are highly sequence-dependent. In many instances, back-exchange may be estimated using algorithms based on context-specific kinetics data (e.g., http://hx2.med.upenn.edu/download.html); it may also be determined experimentally for each proteolytic fragment by processing a fully labeled protein using a series of steps that precisely reproduce those used in HDX MS measurements.⁹ Typical back-exchange levels reported in recent literature range from 10% to 50%, although significantly higher numbers have also been reported. Even if back-exchange can be accounted for, it nonetheless has very detrimental influence on the quality of HDX MS measurements by reducing the available dynamic range.

Finally, the classic HDX MS scheme is poorly suited for measurements that are carried out under conditions favoring correlated exchange, when HDX kinetics follows the so-called EX1 regime, leading to appearance of bimodal and convoluted multi-modal isotopic distributions of protein ions.²¹ Carrying out HDX MS measurements under these conditions provides a unique opportunity to visualize and characterize distinct conformational states, which can be populated either transiently¹⁰ or at equilibrium.²² The distinction among such states can be made based on the differences in their deuterium contents. However, proteolysis in solution almost always leads to a loss of correlation between the deuterium content of fragment peptides and specific conformers with distinct levels of backbone protection. Therefore, the classic HDX MS scheme does not allow protein higher order structure and dynamics to be characterized in a conformer-specific fashion.

“Top-down” HDX MS: tandem MS allows protein structure to be probed in the conformer-specific fashion but raises the specter of hydrogen scrambling

The problem of characterizing protein conformation and dynamics in a conformer-specific fashion can be addressed using methods of tandem mass spectrometry (the so-called “top-down” HDX MS). Indeed, replacement of proteolysis in solution with protein ion fragmentation in the gas phase following mass selection of precursor ions provides a means to obtain fragment ions originating from a particular conformer with a specific level of deuterium incorporation. Deuterium content of fragment ions would then provide a measure of local protection patterns, assuming there is no internal re-arrangement of labile hydrogen and deuterium atoms during ion activation (vide infra). Although the idea to use polypeptide ion dissociation in the gas phase as an alternative to proteolysis was originally proposed in early 1990s,²³ its implementation for proteins only became possible²⁴ following dramatic improvements in FTMS and hybrid TOF analyzers in the late 1990s.

An example of conformer-specific characterization of protein higher order structure using a top-down HDX MS approach is illustrated in Figure 4. The isotopic profile of a fully deuterated 18 kDa protein wt*-CRABPI is recorded following its brief exposure to the ¹H-based exchange buffer. The bimodal appearance of the isotopic distribution of the molecular ion (top trace in Figure 4A) clearly indicates the presence of at least two conformers with different levels of backbone protection. Collisional activation of the entire protein ion population generates a set of fragment ions with convoluted isotopic distributions (top trace in Figure 4B). However, mass selection of precursor ions with a specific level of deuterium content allows the top-down HDX MS measurements to be carried out in a conformation-specific fashion, taking full advantage of the HDX MS ability to detect distinct conformers. For example, selective fragmentation of protein ions representing a highly protected conformation is achieved by mass-selecting a narrow population of intact protein ions with high level of retained deuterium (the blue trace in Figure 4A). Mass-selection and subsequent fragmentation of a narrow population of protein ions with significantly lower deuterium content (the red trace in Figure 4A) generates a set of fragment ions whose isotopic distributions provide information on backbone protection within non-native protein states. For example, the data presented in Figure 4 clearly indicate that the C-terminal segment of the protein represented by the y₁₇²⁺ ions retains significant structure even within the partially unfolded conformers: the amount of retained deuterium atoms reduces by only 30% as a result of switching from the precursor ion from highly protected (blue) to less protected (red). At the same time, selection of the precursor ion has a much more dramatic effect on the protection levels exhibited by the N-terminal segment (represented by the b₄₂⁵⁺ ion), where more than a two-fold decrease in the amount of retained deuterium atoms is observed. Extending this analysis to other protein fragments may allow detailed backbone protection maps to be created for each protein conformer, provided there is no hydrogen scrambling prior to protein ion fragmentation (vide infra).

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Characterization of local dynamics in wt*-CRABP I in a conformer-specific fashion using top-down HDX MS (fully deuterated protein was exposed to ¹H₂O/CH₃CO₂N¹H₄ at pH 3.1 for 10 min; the gray trace at the bottom corresponds to HDX end-point). A: mass selection of precursor ions for subsequent CAD (from top to bottom): broad-band selection of the entire ionic population (not conformer-specific); highly protected conformers; narrow population of less protected conformers; HDX end-point. B: isotopic distributions of two representative fragment ions generated by CAD of precursor ions shown in panel A. Selection of different ion populations as precursor ions for subsequent fragmentation was achieved by varying the width of a mass selection window of a quadrupole filter (Q) in a hybrid quadrupole/time-of-flight mass spectrometer (Qq-TOF MS).

The example shown above illustrates a great promise of top-down HDX MS as a technique uniquely capable of probing structure and dynamics of populations of protein conformers coexisting in solution with high selectivity. Furthermore, this approach often allows one to avoid protein handling under the slow exchange conditions prior to MS analysis, thereby eliminating back-exchange as a factor adversely influencing the quality of measurements. Nonetheless, applications of top-down HDX MS have been limited due to concerns over the possibility of hydrogen scrambling accompanying collision-activated dissociation (CAD) of protein ions. Indeed, several reports pointed out that proton mobility in the gas phase may under certain conditions influence the outcome of top-down HDX MS measurements when CAD is employed to fragment protein ions.^{25, 26}

The occurrence (or the absence) of hydrogen scrambling in the gas phase can be reliably detected by using built-in scrambling indicators. One particularly convenient indicator is a Histag, a 6-30 residues long, histidine-rich segment appended to wild-type sequences to facilitate protein purification on metal affinity columns. Such segments are fully unstructured in solution and, therefore, should lack any backbone protection.²⁷ Alternatively, intrinsic scrambling indicators (e.g., internal flexible loops²⁶), as well as other approaches²⁵ can be used to detect occurrence of scrambling. The available experimental evidence suggests that slow protein ion activation (e.g., SORI CAD) always leads to hydrogen scrambling, while fast activation allows it to be minimized or eliminated in top-down HDX MS experiments.²⁶

Another shortcoming of top-down HDX MS schemes utilizing CAD is the limited extent of protein ion fragmentation, which may lead to sizeable gaps in sequence coverage, particularly for larger proteins,²⁸ and insufficient level of spatial resolution (even for smaller proteins²⁹). Our earlier attempts to solve this problem by employing multi-stage CAD (MSⁿ) were unsuccessful due to massive hydrogen scrambling exhibited by the second generation of fragments.

Electron-induced ion fragmentation in top-down schemes: keeping hydrogen scrambling at bay while enhancing sequence coverage and spatial resolution

Some time ago we suggested that the specter of hydrogen scrambling in top-down HDX MS measurements may be alleviated by using non-ergodic fragmentation processes, where dissociation is induced by ion-electron interaction, rather than collisional activation.³⁰ Indeed, the results of earlier work combining hydrogen exchange of polypeptide ions in the gas phase and electron capture dissociation (ECD) were consistent with the notion of intramolecular rearrangement of hydrogen atoms occurring on a slower time scale compared to ion dissociation.³¹ A recent study demonstrated that the extent of scrambling was indeed negligible when ECD was used as a means to obtain fragment ions in top-down HDX MS characterization of a small protein ubiquitin.³²

Our own recent work suggests that hydrogen scrambling can be avoided when top-down HDX MS employs ECD in characterizing higher order structure of larger proteins (approaching 20 kDa), although experimental conditions must be carefully controlled to minimize proton mobility induced by ion-molecule collisions in the ESI interface. The point in question is illustrated in Figure 5, which shows the results of top-down HDX MS analysis of higher order structure of wt*-CRABP I. The protein retains a significant proportion of labile deuterium label following its complete deuteration and then brief exposure to the ¹H-based exchange buffer, as indicated by the isotopic distribution of the surviving molecular ions (red and blue traces in Figure 5A). However, the deuterium content of fragment ions derived from the 21-residue long His-tag region of the protein (e.g., c₂₂ in Figure 5B) is indistinguishable from that of the exchange reaction endpoint, as long as moderate ion desolvation conditions are kept in the ESI interface. This clearly signals that hydrogen scrambling does not affect the outcome of local HDX MS measurements. However, once collision-assisted desolvation of protein ions is attempted in the ESI interface, the appearance of isotopic distributions of larger fragment ions derived from the His-tag region (e.g., c₂₂, red trace in Figure 5B) shifts, indicating apparent deuterium retention and signaling the occurrence of limited hydrogen scrambling. We also demonstrated that deuterium distribution across the protein backbone is preserved when another recently introduced fragmentation technique based on cation-electron interactions, electron transfer dissociation (ETD), is used in top-down HDX MS schemes.³³

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2805115/bin/nihms-140835-f0005.jpg

Top-down HDX MS of wt*-CRABP I using ECD of the entire protein ion population (fully deuterated protein was exposed to¹H₂O/CH₃CO₂N¹H₄ at pH 3.5 for varying time periods); the black trace at the bottom of corresponds to HDX end-point). A: isotopic distributions of surviving intact protein ions. B: two representative c-ions. Minimal collision-and temperature-induced desolvation was used for acquisition of all mass spectra, except the one top (red trace).

In addition to allowing scrambling to be easily eliminated in top-down HDX MS experiments, both ECD and ETD appear to be superior to CAD in terms of sequence coverage, at least for the proteins in the 20 kDa range. Unlike CAD, protein backbone cleavage in ECD and ETD is less specific,³⁴ leading to a higher number of fragment ions. This translates not only to improved sequence coverage, but also enhanced spatial resolution. Indeed, in some cases it becomes possible to generate patterns of deuterium distribution across the protein backbone down to the single residue level.

One example of such work is shown in Figure 6, where ETD was used as a protein ion fragmentation tool in top-down HDX MS characterization of a 16 kDa variant of CRABP I. The bar graph shows the levels of deuterium retention in a series of c-ions derived from the N-terminal segment of the protein. The bar height at position n in this diagram shows mass difference between two c_n-1 fragments, one derived from the fully deuterated protein that was exposed to the protiated exchange buffer at pH 7 for 5 min and then placed under the slow exchange conditions for the duration of the data acquisition cycle, and another one representing the HDX endpoint (raw data for bars at n=14 and 35 are shown in Figure 7). Unchanged height between two adjacent bars at residues n and n+1 indicates no difference in deuterium content of c_n-1 and c_n fragments, signaling no backbone amide deuterium retention at residue n+1, while bar height increase by one unit indicates complete retention of deuterium at the n^th amide.

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2805115/bin/nihms-140835-f0006.jpg

Backbone protection pattern of CRABPI mutant (without N-terminal His-tag) obtained from top-down HDX MS measurements using ETD of the entire protein ion population. HDX was initiated by exposing the fully deuterated protein to ¹H₂O/CH₃CO₂N¹H₄ at pH 3.5 for 5 min followed by rapid quenching.

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2805115/bin/nihms-140835-f0007.jpg

An example of raw HDX MS data used to generate the protection plot shown in Figure 6. Isotopic distributions of c₁₃ and c₃₄ fragments derived from protein subjected to 5 min HDX exchange in solution (red trace) and protein at the HDX end-point (blue trace) were used to calculate the bar heights at n=12 and 35.

The resulting backbone protection pattern in Figure 6 shows clear correlation with the known higher order structure of the protein (the amino acid sequence and the secondary structure assignment are shown at the top of the graph). Furthermore, the diagram clearly shows uneven distribution of backbone protection even within single structural elements (e.g., lower protection at the fringes vs. the middle of helix α1), as well as unequal protection of similar structural elements participating in the same structural motif (e.g., lower protection of helix α2 vs. helix α1, consistent with the available NMR data). A comparable level of spatial resolution can be achieved with ECD, as shown recently in top-down HDX MS analysis of higher order structure of myoglobin.³⁵

The ability to characterize protein conformation and dynamics at the single residue level is certainly very exciting; however, it comes at a price. Since the protein fragmentation is carried out entirely in the gas phase, no fragment separation can be done prior to mass analysis. A large number of fragment ions with different masses and charges are usually confined to a relatively narrow m/z region, leading to inevitable overlaps of fragment ion isotopic distributions (Figure 7). This places rather stringent requirements on the resolving power of the mass analyzer, effectively narrowing the selection of mass spectrometers suitable for this work to FTMS.

Meeting in the middle: integration of top-down strategies into bottom-up HDX MS schemes

The top-down approach to HDX MS measurements clearly shows a promise to solve many problems that mar the commonly employed bottom-up methodology. The fragmentation efficiency afforded by ECD and ETD provides better spatial resolution, at least for proteins in the 20 kDa range, and this number is likely to grow as there are numerous examples of successful use of these fragmentation techniques to obtain sequence information on significantly larger proteins.³⁶ Unlike the classic bottom-up approach, top-down HDX MS provides an elegant solution to the problem of characterizing higher order structure and dynamics in a conformer-specific fashion (see Figure 4 and discussion in the text). Finally, back-exchange can be eliminated, as outsourcing protein fragmentation to the gas phase often eliminates the need to manipulate the protein in solution under the slow exchange conditions prior to MS analysis.

The top-down/bottom-up dichotomy in HDX MS should not be viewed through the “eitheror” prism. In fact, gas phase fragmentation can enhance the quality of HDX MS data derived from experiments that are built around the bottom-up approach. The suggestion to supplement proteolysis in solution with peptide ion fragmentation in the gas phase to achieve better spatial resolution was made over 10 years ago.³⁷ However, earlier attempts to implement this idea using CAD on a variety of platforms yielded mixed results due to apparent scrambling in some (but not all) fragment ions.^{37, 38} Later reports showed even more extensive scrambling in small peptide ions subjected to collisional activation,³⁹ an obvious anathema to the proposed marriage of CAD and bottom-up HDX MS. Nonetheless, continued search for a scrambling-free solution to this problem has yielded very encouraging results, with both ECD and ETD showing minimal scrambling when applied to short peptides under carefully controlled conditions^{40, 41} and feasibility of supplementing proteolytic fragmentation in solution with ETD in the gas phase was recently demonstrated using a small model protein.⁴² Although these initial steps are relatively modest, they certainly warrant further work in this field.

The two complementary approaches to HDX MS measurements share a set of common challenges that inevitably arise as these techniques gain popularity and the scope of their applications expands. One such challenge is presented by membrane proteins, a notoriously difficult class of biological objects. HDX MS has been shown to have a great potential in this field.⁴³ Interestingly, some initial work in this field was done nearly ten years ago using then-infant top-down HDX MS technique,⁴⁴ while more recent work in this field utilizes both bottomup¹⁸ and top-down⁴⁵ approaches. Another challenge faced by HDX MS is presented by highly heterogeneous proteins, such as proteins conjugated to other biopolymers and/or synthetic polymers, which constitute a significant fraction of the next generation of biopharmaceuticals. Presently, there are no biophysical techniques capable of characterizing conformation and dynamics of these systems, and there is an urgent need to fill this gap. Finally, nearly all HDX MS work reported to date was carried out in vitro under conditions that some regard as “reductionist.” Although initial HDX work with living objects was carried out over 75 years ago,⁴⁶ as the years passed only one report on in vivo HDX MS studies was published.⁴⁷ As mass spectrometry at large is being increasingly used in both in vivo and ex vivo studies, there is a growing pressure on HDX MS to follow the trend, although it remains to be seen how this will be done.

It probably is not an exaggeration to say that we are witnessing a renaissance of HDX MS, with the emergence of the top-down approach not only expanding our experimental arsenal by offering new capabilities, but also serving as a catalyst in enhancing the classic bottom-up methodology. The two techniques are highly complementary, and their synergism will certainly bring about new exciting discoveries and accelerate our progress in solving a variety of problems ranging from very fundamental questions in biophysics to applied problems in drug design.

see more at  http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2805115/

WATCH OUT FOR DISULFIDES: If you’re going to try bottom-up HDX experiments, be careful of disulfide bonds, Kaltashov says. Pepsin is one of the very few proteinases that can efficiently digest a protein into its composite peptides under HDX experimental conditions, but it struggles when multiple disulfide bonds are present. In 2014, Kaltashov’s lab published two solutions to that problem. The first employs a fragmentation technique called electron capture dissociation (ECD) to break the disulfide linkage in the mass spec (Anal Chem, 86:5225-31, 2014); the second skips the pepsin digestion altogether—a strategy called top-down analysis (Anal Chem, 86:7293-98, 2014).

Enhancing the Quality of H/D Exchange Measurements with Mass Spectrometry Detection in Disulfide-Rich Proteins Using Electron Capture Dissociation

see more at   http://www.ncbi.nlm.nih.gov/pmc/articles/PMC4051250/

Approach to Characterization of the Higher Order Structure of Disulfide-Containing Proteins Using Hydrogen/Deuterium Exchange and Top-Down Mass Spectrometry

Guanbo Wang† and Igor A. Kaltashov*
http://www.chem.umass.edu/people/kaltashovlab/papers/Approach.pdf

Top-down hydrogen/deuterium exchange (HDX) with mass spectrometric (MS) detection has recently matured to become a potent biophysical tool capable of providing valuable information on higher order structure and conformational dynamics of proteins at an unprecedented level of structural detail. However, the scope of the proteins amenable to the analysis by top-down HDX MS still remains limited, with the protein size and the presence of disulfide bonds being the two most important limiting factors. While the limitations imposed by the physical size of the proteins gradually become more relaxed as the sensitivity, resolution and dynamic range of modern MS instrumentation continue to improve at an ever accelerating pace, the presence of the disulfide linkages remains a much less forgiving limitation even for the proteins of relatively modest size. To circumvent this problem, we introduce an online chemical reduction step following completion and quenching of the HDX reactions and prior to the top-down MS measurements of deuterium occupancy of individual backbone amides. Application of the new methodology to the top-down HDX MS characterization of a small (99 residue long) disulfide-containing protein β2- microglobulin allowed the backbone amide protection to be probed with nearly a single-residue resolution across the entire sequence. The high-resolution backbone protection pattern deduced from the top-down HDX MS measurements carried out under native conditions is in excellent agreement with the crystal structure of the protein and high-resolution NMR data, suggesting that introduction of the chemical reduction step to the top-down routine does not trigger hydrogen scrambling either during the electrospray ionization process or in the gas phase prior to the protein ion dissociation.

Since its initial introduction in the late 1990s,1−3 top-down hydrogen/deuterium exchange (HDX) with mass spectrometric (MS) detection evolved to become a potent biophysical tool capable of providing valuable information on higher order structure and conformational dynamics of proteins at an unprecedented level of structural detail. Among the many advantages offered by top-down HDX MS compared to conventional (bottom-up) measurements are significant reduction or indeed complete elimination of the back exchange,4 high spatial resolution,5,6 and the ability to study conformational dynamics in the conformer-specific fashion.7,8 However, despite the spectacular recent advances and the broader acceptance of this technique, the scope of the proteins amenable to the analysis by top-down HDX MS remains limited, with the protein size and the presence of disulfide bonds being the two most important limiting factors. The limitations imposed by the physical size of the proteins gradually become more relaxed as the sensitivity, resolution, and dynamic range of modern MS instrumentation continue to improve at an ever accelerating pace. However, the presence of disulfides remains a much less forgiving limitation even for the proteins of relatively modest size.

In this work we demonstrated feasibility of applying top-down HDX MS measurements to characterize higher order structure and conformational dynamics of disulfide-containing proteins, which have been out of the reach of this technique so far. Use of a moderate amount of a reducing agent TCEP is compatible with the ESI process, while allowing a fraction of the protein molecules to be reduced in solution thereby enabling nearcomplete sequence coverage at high resolution. The agreement between the top-down HDX MS and NMR data sets demonstrate that the new experimental approach is capable of capturing the dynamic picture of protein conformation at high spatial resolution without compromising the quality of the data by triggering hydrogen scrambling in the gas phase. Despite its modest size, β2m is known to be able to populate a non-native state,35 which might be a key player in a variety of processes, including amyloidosis. However, the structure of this non-native state of β2m remains elusive since this conformer exists in dynamic equilibrium with the native state of the protein.36,37 Recently we demonstrated that top-down HDX MS provides an elegant way to selectively probe structure of protein states coexisting in solution at equilibrium;8 however, β2m remained out of reach of this technique until recently due to the presence of a disulfide bond. The ability to expand the scope of top-down HDX MS to disulfide-containing proteins opens up a host of exciting possibilities to explore the structure of β2m, interferon, lysozyme, and a variety of other disulfidecontaining proteins in a conformer-specific fashion, where physiologically important non-native states may play important roles in processes as diverse as folding, recognition, signaling, and amyloidosis. ■ ASSOCIATED CONTENT *S Supporting Information Representative examples of isotopic distributions of fragment ions that have (Supplementary Figure 1) and have not (Supplementary Figure 2) been used to calculate the deuterium occupancy at individual backbone amides of β2m in top-down HDX MS measurements. This material is available free of charge via the Internet at http://pubs.acs.org.

Determining surface topology of protein complexes

http://www.the-scientist.com/November2015/MS-surface-topology.jpg

SUSSING OUT THE SURFACE: Protein topology can be probed by firing low-energy electrons (white circles) at intact protein complexes within a high-resolution mass spectrometer. That reaction, called electron capture dissociation, causes the protein complex to fracture on its surface, revealing the exposed amino acid residues.     COURTESY OF PIRIYA WONGKONGKATHEP AND HUILIN LI, UCLA

RESEARCHER: Joseph Loo, Professor of Biological Chemistry, David Geffen School of Medicine, University of California, Los Angeles)

PROJECT: Studying protein-ligand and protein-protein interactions

SOLUTION: Loo is less interested in complex identification than in how the protein subunits assemble. Specifically, he wants to know which amino acid residues lie on the complex’s surface and which are buried inside or interacting with ligands.

It’s a question of structural biology, he explains: “How is this thing folded in a way that these residues are on the outside?”

To work that out, Loo combines high-resolution Fourier transform ion cyclotron resonance mass spectrometry (FTICR) with electron-capture dissociation (ECD), a mass spec fragmentation method in which an ion in the mass spectrometer interacts with free electrons, causing the protein to fracture along its peptide backbone. By measuring the mass of those fragments with high precision, researchers can determine the protein’s amino acid sequence.

In Loo’s case, though, that fragmentation is not uniform along the length of the protein. Proteins usually are denatured for mass spectrometry analysis, but the protein complexes in his studies are intact—a process called native mass spectrometry. Fragmentation thus occurs preferentially on the surface of the complex, like the cracks in the shell of a hard-boiled egg. “You get limited sequence information, but that sequence information comes from regions that are specific to its 3-D structure,” he says (Anal Chem, 86:317-20, 2014).

Native Top-Down ESI-MS of 158 kDa Protein Complex by High Resolution Fourier Transform Ion Cyclotron Resonance Mass Spectrometry

……

Native top-down CAD and ISD were performed for the aldolase tetramer; dissociation of the tetramer to yield monomer was observed in both approaches and no sequence information was obtained. The cleavage sites from ECD (colored in red) and CAD (colored in green) of the denatured aldolase monomer (26+) are overlaid with the native ECD results for aldolase tetramer (Figure 2B). As shown in Figure 2B, in contrast to the limited number of c-ion fragments observed in the ECD of aldolase tetramer, ECD of denatured aldolase monomer induces extensive c-ion fragments in the N-terminal region and enables the assignment of first 156 N-terminal AA residues. Surprisingly, the number of z^•-ions observed from ECD of the denatured aldolase monomer is much less compared to the ECD of the native aldolase tetramer. Although it may be possible that the z^•-ions may undergo secondary fragmentation due to excess available energy, electrons, or long ion-electron reaction times during the ECD experiment, ECD experiments with reduced reaction time and bias voltages were performed and the results argue against this assumption. Overall, 58% of the total number of backbone bonds are cleaved from combining top-down MS of native aldolase complex and denatured aldolase monomer (20% for native ECD of aldolase tetramer, 37% for ECD of denatured aldolase, and 5% for CAD of denatured aldolase).

The three dimensional structure of the aldolase tetramer is shown in Figure 3. To compare the flexibility of the structure to the data from ECD of the aldolase tetramer, one of the subunits (B-chain) is presented as B-factor putty and the D-chain is shown with its native ECD backbone cleavage regions colored in red. The remaining A- and C-chains are shown in grey. Although the C-terminal region (AA 340–363) of each subunit is highly flexible based on the crystallography B-factor (see B-chain in Figure 3A), only 4 out of 75 backbone cleavage sites are from the AA 340–363 region. Instead, the native ECD fragments largely originate from surface regions of the protein structure (see D-chain in Figure 3A). The N-terminal regions are not directly involved in the interfaces between subunits, but they are located in regions that are partially buried, which is consistent with the limited c-ions observed. To better show the native ECD backbone cleavage regions, the D-chain is rotated 90 degrees clockwise (Figure 3B). It is clear that, although protein structure flexibility might play a role in the native top-down ECD fragmentation pattern, for aldolase the ECD cleavage sites are not limited to the flexible region. In addition, backbone cleavage regions from CAD (yellow) and ECD (cyan) of denatured aldolase are complementary with the native ECD results.

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3908771/bin/nihms548404f3.jpg

A) Structure of tetrameric aldolase (1ZAH)²⁹. A- and C-chains are shown as grey ribbons, the B-chain is shown in B-factor putty, and the D-chain is in cartoon with native ECD cleavage sites colored in red, CAD cleavage sites of denatured aldolase in yellow, and ECD cleavage sites of the N-terminal region from ECD of denatured aldolase in cyan. B) The D-chain is rotated 90 degrees clockwise to show the outer surface region of the subunit structure.

Also evident in such data sets are protein–small molecule interactions. As the proteins break apart, Loo explains, ligands often remain attached to the polypeptide shards that are produced. In one recent publication, for instance, his team mapped zinc binding sites in eukaryotic alcohol dehydrogenase, a 147-kDa tetrameric complex (J Am Soc Mass Spectrom, 25:2060-8, 2014).

Revealing Ligand Binding Sites and Quantifying Subunit Variants of Non-Covalent Protein Complexes in a Single Native Top-Down FTICR MS Experiment

see more at   http://www.ncbi.nlm.nih.gov/pmc/articles/PMC4444062/

Still, Loo admits, the technique “is not really ready for prime time.” His team is collecting ECD data on a bank of proteins of known structure to ensure the data they collect really do reflect protein topology. In the meantime, they are working to extend the size of the complexes they can analyze. The technique’s current limit is 800 kDa.

GO NATIONAL: FTICR mass spectrometers offer top-of-the-line accuracy and resolution, with price tags to match. Few researchers have direct access to them, Loo says, but they can always try the national laboratories. Both the National High Magnetic Field Laboratory at Florida State University and the Environmental Molecular Sciences Laboratory at the Pacific Northwest National Laboratory have user facilities open to worthy projects.

Determining the architecture of protein complexes

RESEARCHER: Vicki Wysocki, Ohio Eminent Scholar and Professor of Chemistry and Biochemistry, Ohio State University

PROJECT: Instrumentation development for whole-complex analysis

SOLUTION: An analytical chemist by training, Wysocki focuses on instrumentation development for protein-complex analysis. Among the discoveries in her lab is a method called surface-induced dissociation (SID).

http://www.the-scientist.com/November2015/LT1_last.jpg

HIT THE WALL, JACK: When it comes to molecular collision in a mass spectrometer, size matters. Collide a complex with small gas molecules, and proteins in the complex will simply unravel (top). By smacking them into a “wall”—a process called surface-induced dissociation—the complex dissociates to reveal its underlying architecture.  COURTESY OF VICKI WYSOCKI

Like many other fragmentation approaches, SID works by forcing an ion in the mass spectrometer to collide with another object. Usually that object is a small gas molecule, with the energy of collision sufficient to crack the peptide backbone. But for large protein complexes, bigger is better, and the collision partner in SID is as big as it can get: the method slams protein ions of interest into a nonreactive surface inside the instrument—essentially, a wall—causing complexes to fracture into subcomplexes that reveal the assembly’s inner architecture.

Wysocki combined this approach with ion-mobility separation—a kind of gas-phase electrophoresis that resolves molecules by their size and shape—to dissect an enzyme involved in antibiotic production. The enzyme, they found, has two copies each of three subunits, alpha, beta, and gamma, arranged as a pair of triads sitting on top of one another, with the alpha and beta subunits of one triad linked more tightly to each other than either is to gamma (Anal Chem, 83:2862-65, 2011).

Such information can be valuable to protein engineers, Wysocki says, especially as this particular complex otherwise falls into a structural biology knowledge gap: “It doesn’t crystallize, and it’s too small for the cryoEM and a little bit large for NMR,” she says. “And so, mass spec turned out to be a great tool.”

Revealing the Quaternary Structure of a Heterogeneous Noncovalent Protein Complex through Surface-Induced Dissociation

Anne E. Blackwell, Eric D. Dodds,† Vahe Bandarian, and Vicki H. Wysocki*
https://research.cbc.osu.edu/wysocki.11/wp-content/uploads/2012/09/Blackwell-2011-Revealing-the-Quater.pdf

As scientists begin to appreciate the extent to which quaternary structure facilitates protein function, determination of the subunit arrangement within noncovalent protein complexes is increasingly important. While native mass spectrometry shows promise for the study of noncovalent complexes, few developments have been made toward the determination of subunit architecture, and no mass spectrometry activation method yields complete topology information. Here, we illustrate the surface-induced dissociation of a heterohexamer, toyocamycin nitrile hydratase, directly into its constituent trimers. We propose that the single-step nature of this activation in combination with high energy deposition allows for dissociation prior to significant unfolding or other large-scale rearrangement. This method can potentially allow for dissociation of a protein complex into subcomplexes, facilitating the mapping of subunit contacts and thus determination of quaternary structure of protein complexes.

http://pubs.acs.org/appl/literatum/publisher/achs/journals/content/ancham/2011/ancham.2011.83.issue-8/ac200452b/production/pdfimages_v02/normal.img-000.jpg

The majority of proteins exist and perform their functions as multimers of varing stoichiometries and architecture.1 However, very few methods are available that can provide insights into subunit interactions. Native mass spectrometry (MS) is increasingly being used to study noncovalent protein complexes, as many structural features found in solution may be maintained in the gas phase.2,3 While subunit stoichiometries are readily obtainable by mass measurement alone, the determination of subunit arrangement within protein complexes remains a significant challenge. This is particularly true for heterogeneous complexes with multiple types of subunits. Considerable progress has been made using solution-phase disruption to divide the original protein complex into smaller subcomplexes, which may be readily measured by MS.4,5 The composition of the stable subcomplexes provides insight on the topology of the protein complex. However, MS activation methods used to date have fallen short of providing subunit topology. Here, we present the first evidence for subunit arrangement obtained directly from gas-phase experiments on a heterogeneous complex via surfaceinduced dissociation (SID). We have demonstrated previously the ability of SID to yield unique dissociation pathways for protein complexes, resulting in complementary information to collision-induced dissociation (CID).68 While the SID process is not yet well understood for macromolecules, there is a large body of work concerning SID of small molecules; influential factors such as collision energy, surface composition, and translational-to-vibrational energy conversion have been well-studied.911 The higher effective mass of a surface relative to that of neutral gas atoms used in CID (typically argon) results in significantly higher energy deposited through a single surface collision.9 As SID is a single-collision activation process, rather than activation via thousands of less energetic collisions as in CID, dissociation pathways other than those of the lowest energies become accessible

……

This is the only study to date demonstrating an ion activation method capable of yielding extensive dissociation, as well as the release of intact subcomplexes, thus providing relevant substructure information on a noncovalent, hetero-oligomeric protein complex. The capacity to produce intact, charge-symmetric subcomplexes suggests that dissociation occurs faster than subunit unfolding and that a significant degree of secondary and tertiary structure is maintained up to the point of dissociation and for some period of time afterward. Identification of trimeric substructure in TNH provides insight into a protein with little previous structural characterization and indicates a promising advancement of MS as a tool for structural biology.

Such information can be valuable to protein engineers, Wysocki says, especially as this particular complex otherwise falls into a structural biology knowledge gap: “It doesn’t crystallize, and it’s too small for the cryoEM and a little bit large for NMR,” she says. “And so, mass spec turned out to be a great tool.”

CHOOSE MASS: Mass spec may not be the only method for quickly working out protein structure, but it surely is the fastest, Wysocki says. She recalls one instance when a colleague sent over a complex that his group couldn’t crack. “In one afternoon, my student gave them a prediction of the structure: this one’s a heptamer, with a large subunit sitting atop a hexameric ring.” Even if the experiment doesn’t work, she adds, that fast turnaround time can be a boon, as collaborators can get rapid feedback for tweaking their experimental conditions. “Mass is a great thing.”

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