Over the last two decades, mass spectrometry has made tremendous strides in analyzing protein samples derived from a variety of different sample types. Coupled with electrospray ionization and various separation techniques, thousands of proteins may be identified and quantitated in a single sample. The most common approach used in the laboratory today involves some form of protein extraction followed by proteolytic digestion of protein sample of interest. The use of proteolytic enzymes like trypsin produces peptides that can easily be analyzed by a variety of different instrument configurations. This approach termed “bottom-up” proteomics, can be used to study the state of living cells as a function of their environment. One of the major advantages of the “bottom-up” approach is that the peptides produced have very similar physiochemical properties which makes for a straight forward separation of thousands of peptides in complex samples. Any separation approach coupled with tandem mass spectrometry can then be used to produce amino acid sequence information that is utilized to identify the proteins in a given sample. Although this technique is routine in many laboratories, there are limitations as to the amount of information that can be obtained when reducing intact proteins to their constituent peptides.
In contrast to “bottom-up” proteomics, “top-down” proteomics refers to methods of analysis in which protein samples are introduced intact into a mass spectrometer, without enzymatic, chemical or other means of digestion. Top-down analysis enables the study of the intact protein, allowing identification, primary structure determination and localization of post-translational modifications (PTMs) directly at the protein level. Top-down proteomic analysis typically consists of introducing an intact protein into the ionization source of a mass spectrometer, fragmenting the protein ions and measuring the mass-to-charge ratios and abundances of the various fragments so-generated. The resulting fragmentation is many times more complex than a peptide fragmentation, which may, in the absence of the methods taught herein, necessitate the use of a mass spectrometer with very high mass accuracy and resolution capability in order to interpret the fragmentation pattern with acceptable certainty. The interpretation generally includes comparing the observed fragmentation pattern to either a protein sequence database that includes compiled experimental fragmentation results generated from known samples or, alternatively, to theoretically predicted fragmentation patterns. For example, Liu et al. (“Top-Down Protein Identification/Characterization of a Priori Unknown Proteins via Ion Trap Collision-Induced Dissociation and Ion/Ion Reactions in a Quadrupole/Time-of-Flight Tandem Mass Spectrometer”, Anal. Chem. 2009, 81, 1433-1441) have described top-down protein identification and characterization of both modified and unmodified unknown proteins with masses up to ≈28 kDa
An advantage of a top-down analysis over a bottom-up analysis is that a protein may be identified directly, rather than inferred as is the case with peptides in a bottom-up analysis. Another advantage is that alternative forms of a protein, e.g. post-translational modifications and splice variants, may be identified. However, top-down analysis has a disadvantage when compared to a bottom-up analysis in that many proteins can be difficult to isolate and purify. Thus, each protein in an incompletely separated mixture can yield, upon mass spectrometric analysis, multiple ion species, each species corresponding to a different respective degree of protonation and a different respective charge state, and each such ion species can give rise to multiple isotopic variants.
The process of analyzing intact proteins in cell lysates by mass spectrometry (MS) is associated with a number of difficulties. Firstly, electrospray ionization (ESI) of protein mixtures from cell lysates can generate extremely complex mass spectra due to the presence of multiple proteins, each comprising its own charge state envelope, where each charge state envelope is the collection of mass spectral lines corresponding to plural charge states, and where each charge state correlates directly with the number of positively charged protons that are adducted to an otherwise charge-free molecule. Consequently, multiple charge state envelopes may be overlapping within any given mass-to-charge (m/z) range. In this example, multiple proteins overlap at the same m/z value that have different molecular weights and charges. Commonly used techniques in MS are often insufficient for simplifying these spectra because of the inherent peak overlapping as well as the inherent wide range of magnitudes of MS lines of ionized constituents, where such constituents may range from uninteresting small molecules to interfering biomolecules to the proteins of interest, themselves. Isolation of a specified charge state of a protein within such complex spectra does not typically alleviate the burden of multiple protein peaks overlapping, since the isolation of ions of a particular protein charge state will generally result in co-isolation of one or more additional ions. This co-isolation makes it a challenge not only to dissociate the protein in an attempt to identify it based on the fragments produced, but also to accurately determine the intact mass and sequence coverage of that protein.
So-called “front-end” separation techniques, such as liquid chromatography (LC) or ion mobility spectrometry (IMS), performed prior to introduction of samples into a mass spectrometer, may be implemented to reduce the overall complexity and provide an additional major benefit, which is the reduction of ionization competition at an ionization source. Unlike mixtures of proteolytic peptides typically analyzed in bottom-up experiments, intact proteins mixtures contain a wide range of molecular weights, isoelectric points, hydrophobicities, and other physiochemical properties that make it challenging to analyze these mixtures via any single separation technique in a comprehensive manner. Both of the above separation methods are associated with their own benefits and pitfalls. Liquid chromatography tends to require significant amounts of time per sample to separate individual proteins, although it is still common to have two or more proteins co-elute. Enhanced separation can reach the point of becoming more of “an art” than a standardized method, and the enhanced separation may be dependent on the user skill in the state-of-the-art. The latter technique, IMS, can rapidly separate certain proteins and/or charge states from others but IMS spectra are at least partially correlative with (i.e., not “orthogonal to”) mass spectra. The IMS method also suffers from ionization competition, requires extensive optimization and typically involves dynamic conditions to observe a full mass spectrum containing all charge states.
Proton transfer reactions, a type of ion-ion reaction that has been used extensively in biological applications for rapid separations of complex mixtures, addresses many of these aforementioned concerns. Experimentally, proton transfer is accomplished by causing multiply-positively-charged protein ions from a sample to react with introduced singly-charged reagent anions so as to reduce the charge of the multiply-charged protein ions. These reactions proceed with pseudo-first order reaction kinetics when the anions are present in large excess over the protein ion population. The rate of reaction is directly proportional to the square of charge of the protein ion (or other multiply-charged cation) multiplied by the charge on the anion. The same relationship holds for reactions of the opposite polarity as well. This produces a series of pseudo-first order consecutive reaction curves as defined by the starting multiply-charged protein ion population. Although the reactions are highly exothermic (in excess of 100 kcal/mol), proton transfer is an even-electron process performed in the presence of 1 mtorr of background gas (i.e. helium) and thus does not fragment the starting multiply-charged protein ion population. The collision gas serves to remove the excess energy on the microsecond time scale (108 collisions per second), thus preventing fragmentation of the resulting product ion population.
Proton transfer reactions (PTR) have been used successfully to identify individual proteins in mixtures of proteins. This mixture simplification process has been employed to determine charge state and molecular weights of high mass proteins. PTR has also been utilized for simplifying product ion spectra derived from the collisional-activation of multiply-charged precursor protein ions. Although PTR reduces the overall signal derived from multiply-charged protein ions, this is more than offset by the significant gain in signal-to-noise ratio of the resulting PTR product ions. The PTR process is 100% efficient leading to only single series of reaction products, and no side reaction products that require special interpretation and data analysis.
Various aspects of the application of PTR to the analysis of peptides, polypeptides and proteins have been described in the following documents: U.S. Pat. No. 7,749,769 B2 in the names of inventors Hunt et al., U.S. Patent Pre-Grant Publication No. 2012/0156707 A1 in the names of inventors Hartmer et al., U.S. Pre-Grant Publication No. 2012/0205531 A1 in the name of inventor Zabrouskov; McLuckey et al., Anal. Chem. 1998, 70:1198-1202; Stephenson et al., J. Am. Soc. Mass Spectrom. 1998, 8:637-644; Stephenson et al., J. Am. Chem. Soc. 1996, 118:7390-7397; McLuckey et al., Anal. Chem. 1995, 67:2493-2497; Stephenson et al., Anal. Chem. 1996, 68:4026-4032; Stephenson et al., J. Am. Soc. Mass Spectrom. 1998, 9:585-596; Stephenson et al., J. Mass Spectrom. 1998, 33:664-672; Stephenson et al., Anal. Chem., 1998, 70:3533-3544 and Scalf et al., Anal. Chem. 2000, 72:52-60. Various aspects of general ion/ion chemistry have been described in McLuckey and Stephenson, Mass Spec Reviews 1998, 17:369-407 and U.S. Pat. No. 7,550,718 B2 in the names of inventors McLuckey et al. Apparatus for performing PTR and for reducing ion charge states in mass spectrometers have been described in U.S. Pre-Grant Publication No. 2011/0114835 A1 in the names of inventors Chen et al., U.S. Pre-Grant Publication No. 2011/0189788 A1 in the names of inventors Brown et al., U.S. Pat. No. 8,283,626 B2 in the names of inventors Brown et al. and U.S. Pat. No. 7,518,108 B2 in the names of inventors Frey et al. Adaptation of PTR charge reduction techniques to detection and identification of organisms has been described by McLuckey et al. (“Electrospray/Ion Trap Mass Spectrometry for the Detection and Identification of Organisms”, Proc. First Joint Services Workshop on Biological Mass Spectrometry, Baltimore, Md., 28-30 Jul. 1997, 127-132).
The product ions produced by the PTR process can be accumulated into one or into several charge states by the use of a technique known as “ion parking”. Ion parking uses supplementary AC voltages to consolidate the PTR product ions formed from the original variously protonated ions of any given protein molecule into a particular charge state or states at particular m/z values during the reaction period. This technique can be used to concentrate the product ion signal into a single or limited number of charge states (and, consequently, into a single or a few respective mass-to-charge [m/z] values) for higher sensitivity detection or further manipulation using collisional-activation, ETD, or other ion manipulation techniques. Various aspects of ion parking have been described in U.S. Pat. No. 8,440,962 B2 in the name of inventor Le Blanc and in the following documents: McLuckey et al., Anal. Chem. 2002, 74:336-346; Reid et al., J. Am. Chem. Soc. 2002, 124:7353-7362; He et al., Anal. Chem. 2002, 74:4653-4661; Xia et al., J. Am. Soc. Mass. Spectrom. 2005, 16:71-81; Chrisman et al., Anal. Chem. 2005, 77:3411-3414 and Chrisman et al., Anal. Chem. 2006, 78:310-316.
Another difficulty associated with the mass spectrometric analysis of proteins in cell lysates by (MS) is that the fragmentation behavior for each charge state of a protein is generally unknown prior to the dissociation event. In particular, ions comprising some charge states can dissociate well while ions comprising other charge states may dissociate poorly. Isolation and dissociation of ions of a particular charge state therefore does not guarantee efficient dissociation or dissociation into a set of diagnostic fragments.
A third challenge associated with intact protein analysis is the wide distribution of charge states produced for high molecular weight proteins typically in excess of 50 kDa. Here the starting signal can be divided into over 30 plus charge states, making tandem mass spectrometry of any given charge state produce a spectrum with low signal-to-noise ratio. The ability to produce ample sequence coverage for protein identification can therefore be difficult with a single tandem mass spectrum.
A variety of ion activation (fragmentation) techniques can be used to produce structural information on intact proteins. The most commonly used approach termed collision-induced dissociation (CID) involves collisions of an isolated population of multiply-charged precursor ions with a neutral background gas. Most commonly, the multiply-charged precursor ions are accelerated using the fundamental frequency of motion of the defined ion population in order to collide with the neutral background gas so as to produce unimolecular dissociation events. This process leads to fragmentation along the amide backbone of the protein thus yielding amino acid sequence information. More extensive fragmentation of proteins can be obtained with higher collision energy processes termed HCD or high energy collision induced dissociation. Many times this involves multiple fragmentation events inside the collision cell thus producing more extensive sequence coverage. Another approach used to produce protein sequence coverage via ion activation is that of photodissociation (PD), where photons of a defined wavelength are used to excite the ion of interest. Two common types employed include ultra-violet (UV-PD) and infrared multiphoton dissociation (IRMPD). The latter is a high energy process where the rate of energy deposition in the ion far exceeds that of the dissociation process. Here fragmentation can be produced along any point in the protein backbone, or may yield amino acid side chain fragmentation as well. For IRMPD, this is a much lower energy process that is characterized by the presence of cleavages at amide bonds and losses of ammonia and water from the intact protein and fragment ions generated during irradiation. The time frame of the IRMPD experiment can be expanded to produce more extensive fragmentation as well. Ion-ion reactions using electron transfer reagent ions can also be employed as a fragmentation approach for intact proteins. Here an electron transfer event from the multiple-charged protein to the singly-charged anion produces backbone fragmentation of the protein with any posttranslational modifications still intact.
Taken together, these ion activation approaches for tandem mass spectrometry produce many different complementary forms of fragmentation that can provide protein sequence information. Ideally, these approaches can be applied in a broad band fashion in order to increase sequence coverage of proteins and provide additional information on modifications, splice variants, and expression of single amino acid mutations. The application of these approaches in a broadband format (i.e. covering multiple charge states of the same intact protein) would provide a more comprehensive view of protein characterization and identification.