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Peptide Sequencing by MALDI 193‐nm Photodissociation TOF MS
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[6] Peptide Sequencing by MALDI 193‐nm Photodissociation TOF MS By JOSEPH W. MORGAN, JUSTIN M. HETTICK , and DAVID H. RUSSELL Abstract
Ultraviolet photodissociation time‐of‐flight (TOF) mass spectrometry (MS) is described as a method for determination of peptide ion primary structure. Monoisotopic selection and bond‐specific activation, combined with the rapidity of TOF MS analysis, render this technique invaluable to the rapidly expanding field of proteomics. Photofragment ion spectra of model peptides acquired using both post‐source decay (PSD) focusing and TOF‐TOF experimental methods are exhibited. Advantages of 193‐nm photodissociation for de novo sequencing of peptide ions are discussed.
Introduction
Advances in ionization methods and instrumentation that have occurred over the past decade enable MS to play a key role in the emerging area of proteomics. For example, matrix‐assisted laser desorption ionization (MALDI) (Karas et al., 1987; Tanaka et al., 1988) and electrospray ionization (ESI) (Whitehouse et al., 1985) are now routinely used to ionize peptides and proteins (Pandey and Mann, 1985), which can then be aimed at sequencing peptides and=or proteins (Chaurand et al., 1999; Cotter, 1997), determination of posttranslational modifications (Lennon and Walsh, 1999), or to study protein–protein complexes (Dikler, 2001; Young et al., 2000). Although various instrument platforms can be used for the mass analysis, over the past decade great strides have been made in the development of TOF instrumentation. Advances in instrument geometry and electronics used for TOF MS allow us to interrogate large gas‐phase ions with improved resolution and mass accuracy (Cotter, 1999; Mamyrin et al., 1973; Wiley and McLaren, 1955). Two extremely important TOF mass spectrometric techniques to the field of proteomics are peptide mass fingerprinting (PMF)=protein database searching (Henzel et al., 1993) and peptide sequencing, and both methods owe their widespread acceptance to the versatility and user friendliness of modern TOF instruments. PMF, combined with genomic database searching to identify proteins, is now widely used, and PMF based on accurate mass measurement is now a METHODS IN ENZYMOLOGY, VOL. 402 Copyright 2005, Elsevier Inc. All rights reserved.
0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)02006-9
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routine TOF analysis (Clauser et al., 1999; Edmondson and Russell, 1996; Jensen et al., 1996; Russell and Edmondson, 1997). Peptide identification based on PMF=protein database searching is especially powerful when combined with peptide sequencing by tandem MS. For example, determining a partial peptide sequence or identifying a specific posttranslationally modified amino acid dramatically increases the confidence level of identification. For proteins that are not listed in a database, peptide sequencing is essential for protein identification. Peptide sequencing using MS is based on fragmentation of the gas‐phase ion. For example, fragmentation along the peptide backbone yields a series of ions separated in mass by an amount equal to the mass of the eliminated residue (Biemann, 1990a). Consequently, the ions must be formed with sufficient internal energy to undergo unimolecular dissociation (metastable ion decay or PSD), or ions formed with very little initial internal energy (i.e., by ESI) must be activated to energies above the dissociation threshold. The most commonly used ion‐activation methods for peptide sequencing are collision‐induced dissociation (CID) (Russell and Edmondson, 1997) and surface‐induced dissociation (SID) (Cooks and Miller, 1999; Nikolaev et al., 2001; Stone et al., 2001); however, photodissociation has considerable potential for peptide sequencing, especially for peptide sequencing involving TOF instruments (Barbacci, 2000; Barbacci and Russell, 1999; Hettick, 2003; Hettick et al., 2001a; Hunt et al., 1987; Thompson et al., 2004). The physics of CID are not compatible with peptide sequencing via PSD focusing using the standard tandem TOF instrument. For instance, high‐energy CID (Chaurand et al., 1999; Yergey et al., 2002) tends to enhance the yield of low m=z fragment ions at the expense of higher mass (e.g., more structurally=sequence informative) fragment ions. Perhaps most troublesome for tandem TOF is the physics of converting kinetic energy into internal energy. CID yields ions having a range of velocities and a broad distribution of arrival times, which limits both m=z resolution and mass measurement accuracy of the TOF measurement (Hettick et al., 2001a). Another important consideration for CID of large ions is the effect of center‐of‐mass collision energy. In virtually all tandem TOF experiments, the mass of the ion undergoing CID is much larger than the target atom. Consequently, only a very small fraction of the laboratory energy can be partitioned as internal energy of the ion, and as the mass of the analyte ion increases, energy deposition becomes even less efficient (Shukla and Futrell, 1994; Uggerud et al., 1989). To circumvent some of the limitations of CID, several groups began investigating SID as an alternative method of ion activation (Mabud et al., 1985; McCormack et al., 1992). Under optimized conditions, SID yields
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very good fragment ion intensities; however, at higher collision energies, SID produces primarily low m=z ions, for example, immonium ions (Riederer et al., 2000; Stone et al., 2001). As the translational energy of the peptide ion is decreased, surface sticking, or ‘‘soft‐landing,’’ becomes competitive with fragmentation processes (Cooks et al., 2004; Grill et al., 2001). Also, because fragment ions in SID leave the surface with a range of velocities and angles, careful attention must be paid to instrument geometry. A comprehensive review of SID has been published by Grill et al. (2001). Several years ago we began evaluating 193‐nm photodissociation as a method to circumvent some of the limitations of CID. Radiation from an ArF excimer (193‐nm, 6.43 eV=photon) laser is particularly well suited to photodissociation of peptide ions, because the amide linkage is a natural chromophore (lmax 210 nm), and the photon imparts sufficient energy to cleave the bond. Relatively few tandem TOF photodissociation experiments have been reported (Gimon‐Kinsel et al., 1995; Seeterlin et al., 1993; Willey et al., 1994) This is in part due to logistical issues that must be addressed in a tandem TOF photodissociation experiment. First, the timing for the trigger pulse for the excimer laser with respect to the ionization=desorption event is critical. In order to intersect the ion beam with a pulsed laser beam, both the delay generator output and the intrinsic time delay in the laser trigger must be stable to 2 ns. (Gimon‐Kinsel et al., 1995), Second, the ions should be irradiated in a region where they are spatially separated and well defined, in terms of m=z. Third, they must be given sufficient time to undergo unimolecular decay in the field‐free region before entering the reflectron. Several years ago we began a series of studies to evaluate photodissociation of peptide and protein ions (Gimon‐Kinsel et al., 1995). The instrument consisted of two collinear TOF mass analyzers, with a photodissociation region positioned between the two analyzers. The instrument used for the preliminary studies was specifically designed to investigate photodissociation of MALDI‐generated peptide and protein ions; however, the instrument did not include a reflectron to mass analyze the resultant photofragments. These studies allowed us to differentiate between fragment ions and neutrals produced by photodissociation of the peptide and protein ions. We also showed that photodepletion of [M þ H]þ ions is a single‐photon process, whereas photodepletion of fragment neutrals, F0n , formed by photoionization of the [M þ H]þ is a two‐photon process. Finally, based on the observation that photodepletion occurred within 1 s of irradiation, it was determined that the rate constant for photodepletion was at least 106 s1.
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Early photodissociation studies were aimed at small organic molecules, and many of these studies were carried out using Fourier transform mass spectrometers (FT‐MSs). The FTMS photodissociation experiment is relatively simple; a defocused light source is passed through the cell and the photofragment ions are trapped and detected using conventional methods (Dunbar, 1989). The long trapping times (milliseconds to hundreds of seconds) and density of neutral molecules (baseline vacuum of 106 to 109 torr) is sufficient for the analyte ion and=or photofragment ions to undergo ion–molecule reactions, which can complicate the photofragment ion spectrum (Willey et al., 1994). Due to the ions being trapped in the FT‐MS ion cell and bathed in a continuous radiation field, it is difficult to control the photoexcitation process (e.g., single vs. multiphoton excitation). Multiphoton excitation deposits very large amounts of energy to the molecule, resulting in extensive fragmentation. In addition, photofragment ions can also absorb a photon and yield second‐generation photofragment ions. These complications do not apply to TOFMS photodissociation experiments. Instrumentation
The results from our preliminary experiments demonstrated efficient photodissociation of protonated peptide=protein ions and motivated us to design and construct an instrument for mass analyzing the photofragment ions. The instrument used for these studies is shown in Fig. 1, and it consists of a linear TOF‐1 and a reflectron that can be used as TOF‐2. The effective flight‐path length of TOF‐1 is 0.5 m and TOF‐2 is 3.4 m. Two turbo molecular pumps maintain baseline pressure of approximately 2 108 torr. It is important that such a high vacuum be maintained to eliminate the possibility of ion collisions with background gases, which contribute ‘‘background fragmentation’’ to the experiment. Ions are generated in the ion source by MALDI (337 nm) and extracted by pulsing the extraction grid potential to ground (Ingendoh et al., 1994). Ions of interest are selected by the timed‐ion selector and subsequently photodissociated by the excimer laser (Fig. 2). The output of the excimer laser is directed to the mass spectrometer via three reflective mirrors (front surface specific to 193 nm), and laser beam size is controlled by a mechanical iris positioned between the final mirror and the vacuum window. The laser output energy is controlled by the laser control panel, and fine control is achieved by passing the laser output through a sample cuvette filled with a methanol=water solution (Hettick et al., 2001b). Control of the laser photon density output is important, because at high laser beam energy, the photofragment ion spectrum is dominated by low m=z immonium ions,
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FIG. 1. Schematic of the MALDI Photofragment Tandem TOF Instrument (A) Source plate; (B) extraction grids; (C) reflectron; (D) stage manipulator; (E) timed ion selector; (F) quartz window; (G) reflected ion detector; (H) linear ion detector; (I) 170‐L=s turbo molecular pump; (J) 330‐L=s turbo molecular pump; (K) roughing pumps; (L) 1‐GHz digital oscilloscope.
FIG. 2. Expanded view of the laser–mass spectrometer interface.
and at lower laser beam energy, the abundance of structurally informative backbone and side‐chain cleavages are increased. Precautions are taken to ensure that the laser beam does not strike the surface of the vacuum chamber because this causes photoejection of electrons and neutral species that can enhance abundances of CID fragment ions.
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Photofragment ion spectra are calibrated using methods commonly employed for PSD spectra (Spengler, 1997). Briefly, photofragment spectra are taken at a number of PSD reflectron ratios and then are truncated to include only that portion of the spectrum within a few microseconds of the arrival time of the parent ion. The truncated spectra are spliced together to create a complete photofragment ion spectrum. A limitation of PSD experiments is the relatively low mass resolution of the timed‐ion selector (MS1). Currently, the timed‐ion selector is limited to a 300‐ns pulse. Thus, all ions that arrive at the timed‐ion selector within the 300 ns window are allowed to continue toward the reflectron, and any fragment ions formed by dissociation of these ions are observed in the photofragment ion spectrum. Although the mass resolution of the timed‐ ion selector is low enough that [M þ H]þ peptide ions of m=z 1000 (20‐keV translational energy) and [M þ Na]þ ions are not resolved, it is possible to selectively photoexcite only the [M þ H]þ or [M þ Na]þ ion (Fig. 3). For example, as the ions approach the laser‐ion beam intersection, the [M þ H]þ and [M þ Na]þ ions are separated by 6.0 mm, and during a single excimer laser pulse (17 ns), the ions travel approximately 1.0 mm. Thus, even if the
FIG. 3. Spatial separation of ions at the laser–mass spectrometer Interface Isotopes of a theoretical peptide [M þ H]þ of m=z 1000 (a, b, and c) are separated by approximately 0.3 mm, whereas the [M þ Na]þ ion (d) is approximately 6.0 mm behind. The excimer laser is fired at time tL when the [M þ H]þ isotopic distribution reaches the point dL. At time tL þ 17 ns, the excimer laser beam exits the instrument. All ions have advanced approximately 1.05 mm, and only the isotopic distribution of the theoretical peptide [M þ H]þ ion, having traversed through point dL, has absorbed 193‐nm photons.
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[M þ Na]þ ions are allowed to pass the TIS to the photodissociation region, they are not irradiated, and their fragment ions should not appear in the photofragment ion spectrum. It is important that fragmentation from ions such as [M þ Na]þ be eliminated, because they produce product ions shifted in mass (þ23 Da), making spectrum analysis significantly more difficult. Using the PSD method to analyze photofragment ions has limitations as well. Metastable ions that are formed within the first field‐free drift region (TOF‐1) have the same velocity as their respective parent ions. The metastable ions arising from the ion of interest are activated by the photodissociation laser, giving rise to small‐neutral loss ions, which complicate the photofragment ion spectra. Our previous work also showed that metastable neutral species can be ionized by 193‐nm irradiation (Gimon‐Kinsel et al., 1995). The most significant drawback to the photodissociation experiment using PSD focusing is the time required for data acquisition and calibration. As with most PSD experiments, the reflectron voltage must be lowered in increments to acquire the entire fragment ion spectrum, so data acquisition must be repeated n times, where n represents the number of segments needed to stitch the spectrum. To circumvent this limitation, nonlinear reflectron fields have been proposed and used with some success (Cordero et al., 1996; Cornish and Cotter, 1993). Alternatively, ions could be activated ‘‘in‐source’’ so that all fragment ions are accelerated to the same final kinetic energy. In this case, acquisition of the photofragment ion spectrum would have the same duty cycle as a conventional TOF mass analysis. Our most recent photodissociation experiments used an instrument design that negates the limitations described earlier. A four‐grid deceleration=acceleration cell (10 cm in length) was added, centered about the photodissociation window. The MALDI source potential for these experiments was 15 kV, and 8 kV was applied to the central element of the photodissociation cell, and the outside cell grids are held at ground potential. Ions are separated from their metastable decay products as they traverse the decelerating electric field, and consequently, only the ions of interest are irradiated at the center of the cell. A fraction of photoexcited ions dissociate within 0.5 s as they traverse the remaining field‐free region of the photodissociation cell. As ions exit the photodissociation cell, they are then accelerated to near‐uniform kinetic energies and m=z analyzed by TOF‐2. Because differences in final kinetic energies between parent and product ions are small, a complete photofragment ion spectrum is acquired at a single reflectron voltage. Variation in final kinetic energies between photofragment ions is dependent on parent ion mass‐to‐charge ratio and is accounted for in a new calibration equation:
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peptide sequencing by MALDI 193‐nm photodissociation TOF MS
mf TOF ¼ a bmf þ c
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193
þ t0
Parameters a and c are constant for given mass spectrometer dimensions and source and cell potentials, and parameter b is inversely proportional to the m=z of the parent ion. The [M þ H]þ ions for a mixture of five known peptides (HLGLAR, des‐Arg9 bradykinin, angiotensin III, bradykinin, and substance P) were sequenced by both PSD focusing and TOF‐TOF photodissociation tandem MS methods. All mass spectra and photofragment ion spectra were acquired using sample preparation methods that reduce the abundance of PSD fragment ions (e.g., near‐threshold laser power for MALDI) and using fructose sample preparations (Beavis et al., 1988; Castoro and Wilkins, 1993; Hettick et al., 2001a; Ko¨ ster et al., 1992). CID spectra were acquired using an Applied Biosystems 4700 Proteomics Analyzer (Framingham, MA) (Medzihradsky et al., 2000). Instrument conditions used for CID included 1‐kV accelerating voltage and medium gas pressure.
Mechanism of Photodissociation
Absorption of ultraviolet radiation by a gas‐phase ion yields an electronically excited ion (Fig. 4), and the photoexcited ion may dissociate directly or decay by radiative and=or nonradiative channels. Although radiative decay of gas‐phase ions is an important route for rigid molecules (Maier, 1982), nonradiative decay is a much higher probability process for peptide ions. Because the gas‐phase ion is an isolated system (i.e., energy cannot be transferred to solvent or the local environment), the photoexcited ion may partition the electronic energy to vibrational modes, and the vibrationally excited ion can then undergo dissociation, provided sufficient energy and time are available. According to RRK theory (Steinfeld et al., 1999), the rate for unimolecular dissociation for a vibrationally activated ion having s degrees of freedom and internal energy E is calculated using the following equation: E E0 s1 kðEÞ ¼ V E Rates of dissociation are directly proportional to the amount of excess energy required for bond cleavage (E E0) and the frequency of the critical oscillator (). The process of electronic excitation of a specific oscillator via
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FIG. 4. Vibrational predissociation mechanism absorption of a photon of 193 nm is sufficient to induce a Franck‐Condon transition to the upper electronic state. The ion may then relax via internal conversion to the lower electronic state and fragment.
single‐photon absorption may increase the rate of dissociation for fragmentation pathways that are not accessible for lower energy activation methods (Tecklenburg et al., 1989). Photofragment ion spectra of peptides activated by 157‐nm radiation also provide evidence that photodissociation can occur promptly at bonds that are near the chromophore (Thompson et al., 2004). Peptide Sequencing
For peptide sequencing experiments, it is important to accurately determine the m=z values of fragment ions. Most tandem TOF sequencing experiments have average mass errors in the range of 0.1–1.0 Da (Kaufmann et al., 1996). Although the mass difference between most of the amino acids is more than 1 Da, there are several cases in which mass differences are much less. Leu and Ile have identical masses (113.084 Da) and cannot be distinguished based solely on fragmentation along the peptide backbone. Gln and Lys differ by only 0.036 Da (128.059 and 128.095 Da, respectively), so a mass measurement accuracy of 0.01 is required to distinguish these residues. Table I lists fragment ion mass measurements
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peptide sequencing by MALDI 193‐nm photodissociation TOF MS TABLE I FRAGMENT ION MASS ACCURACY DATA FOR THE HEXAPEPTIDE HLGLAR ACQUIRED ON THE MALDI PHOTOFRAGMENT TOF INSTRUMENT
TOF (ns) 49235.53 47248.32 49195.00 49110.40 49192.09 47897.94 47016.22 48858.32 47941.37 47616.95 48375.26 48517.89 47509.16 47372.89 47149.81 48868.05 46084.94 47695.02 48541.49 49028.12
PSD ratio 1.000 0.870 0.770 0.770 0.740 0.740 0.700 0.635 0.635 0.635 0.560 0.530 0.500 0.410 0.378 0.340 0.240 0.180 0.170 0.130
Ion identity þ
[M þ H] y5 y5‐NH3 b5 þ H2O b5 a5 b4 y4 y4‐NH3 a4 y3 y3‐NH3 b3 b2 y2‐NH3 a2 b1 R A1(H) L
Meas. m=z
Calc. m=z
Error
666.405 529.395 512.286 510.387 492.275 464.347 421.262 416.284 399.302 393.294 359.259 342.232 308.163 251.110 229.071 223.116 138.073 112.082 110.074 86.088
666.405 529.346 510.304 492.293 492.293 464.299 421.256 416.262 399.236 393.261 359.241 342.214 308.172 251.151 229.130 223.156 138.067 112.087 110.072 86.097
0.000 0.049 0.034 0.083 0.018 0.048 0.006 0.022 0.066 0.033 0.018 0.018 0.009 0.041 0.059 0.040 0.006 0.005 0.002 0.009
Average Std. Dev.
0.028 0.024
for the peptide HLGLAR acquired during a PSD focusing photodissociation experiment. The average mass measurement error of 0.028 Da is sufficient to elucidate the composition of this peptide (with the exception of differentiating between Leu and Ile) from either terminus. Both mass accuracy and resolution are higher for photodissociation TOF‐TOF experiments. The utility of photodissociation tandem TOF for peptide sequencing is illustrated by the photofragment ion spectra of bradykinin. Figure 5 contains the PSD focusing photofragment ion spectrum of bradykinin (RPPGFSPFR) taken from a five‐peptide mixture. As in the previous example, immonium ions corresponding to proline, phenylalanine, and arginine are observed, and a complete set of a‐type ions (except a7) are observed. In addition, several b‐type ions are observed as well, notably the b8 þ H2O ion at m=z 904. Ions corresponding to charge retention by the C‐terminus are prominent in this spectrum, as evidenced by several prominent y‐type ions. Because there are arginine residues at either terminus of
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FIG. 5. Post‐source decay (PSD) focusing photofragment ion spectrum of bradykinin (RPPGFSPFR) taken from the five peptide mixture.
this peptide, it is reasonable to expect significant populations of fragment ions exhibiting C‐terminal charge retention from this peptide. In addition to simple backbone cleavage products, numerous side‐chain cleavage products are observed in this spectrum, notably d6, d9, and w3. We also observe internal fragmentation in this spectrum, which is typical for proline‐containing peptides (Breci et al., 1963). Internal fragmentation is highly useful in reconstructing peptide sequence when a complete series of backbone cleavage products is not observed. For instance, using just the a‐type ions observed in this photofragment ion spectrum, the sequence would be assigned ‘‘RPPGFSXXR,’’ where XX can be either FP or PF, but knowledge of one of the internal cleavage ions such as FSP, GFSP, or PGFSP indicates the sequence must be RPPGFSPFR. Alternatively, this information could be extracted from C‐terminal sequence information given by the y‐type ion series. In this case, we are able to assign the sequence as XPXXXSPFR from the observed y‐ions. Thus, in the case of bradykinin, the information obtained from the a‐ and y‐type fragment ions is sufficient to unambiguously assign the complete sequence. Figure 6 contains the TOF‐TOF photofragment ion spectrum of bradykinin. Note that contributions from multiple fragmentations, such as internal cleavage ions and small neutral losses, are in much lower abundance in this spectrum. Products of prompt photodissociation, such as a‐type
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F IG . 6. Time‐of‐flight (TOF)‐TOF photofragment ion spectrum of bradykinin (RPPGFSPFR).
sequence informative ions and d‐type side‐chain cleavage ions, are now the major contributors to the photofragment ion spectrum. The CID spectrum of this species is shown in Fig. 7. Note the appearance of several ion types having charge retention on either side of the dissociating bond and internal fragment ions and other unidentified species. Although much information is contained within this spectrum, its utility for de novo sequencing is low because of difficulties in the assignment of peaks to fragment ions. Figure 8 contains the photofragment ion spectrum of des‐Arg9 bradykinin (RPPGFSPF) obtained from a five‐peptide mixture using the PSD focusing method. The immonium ions for proline, phenylalanine, and arginine verify the presence of these three amino acids in the peptide. A number of a‐ and b‐type fragment ions are also observed in the spectrum, which we attribute to charge retention at the N‐terminus of the peptide ion. We also observe sequence ions that arise by loss of NH3 from either the primary photofragment ions or the activated metastable ions, and these are labeled Fi‐NH3 ions. Also present are fragment ions that result from charge retention at the C‐terminus of the peptide ion or at a basic amino acid side chain near the C‐terminus. For example, y1, y2, and y3 are observed. In addition to simple backbone cleavage reactions (e.g., a‐, b‐, and y‐type ions), we also observe side‐chain cleavage ions. For instance, v4 and w6 arise by side‐chain cleavage products in which the charge is retained by the
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FIG. 7. Collision‐induced dissociation (CID) spectrum of bradykinin (RPPGFSPFR).
FIG. 8. Post‐source decay (PSD) focusing photofragment ion spectrum of des‐Arg9 bradykinin (RPPGFSPF) taken from the five peptide mixture. (Used, with permission, from Medzihradsky et al., 2000.)
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C‐terminus. In addition to the v‐ and w‐side‐chain cleavage ions, side‐chain cleavage product ions corresponding to N‐terminal charge retention, d2, d3, and d6 are also observed. The TOF‐TOF photofragment ion spectrum of des‐Arg9 bradykinin is shown in Fig. 9. Photofragment ions arising from multiple dissociations and=or fragmentation of metastable ions are in much lower abundance. C‐terminal charge retention products such as phenylalanine immonium ion and y‐type ions are observed in lower abundance also. Analysis of these promptly formed photofragment ions provides evidence that N‐terminal charge retention is dominant in the initial population of MALDI formed ions. Through comparison with results from the PSD focusing method, we postulate that b‐ and y‐type ions are more likely to be formed from lower energy, longer‐lived dissociating ions, as they appear in much lower relative abundance when the time allotted for observation of photofragment ions is decreased from 10 s to 0.5 s. Figure 10 contains the CID spectrum of fibrinopeptide A (ADSGEGDFLAEGGGVR). The spectrum consists of mostly y‐type sequence ions and w‐type side chain cleavage ions. Note the dominance of the y‐type ions corresponding to backbone cleavage at acidic residues. These same dominant ions are observed in the PSD focusing photofragment ion spectrum
FIG. 9. Time‐of‐flight (TOF)‐TOF Photofragment ion spectrum of des‐Arg9 bradykinin.
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FIG. 10. Collision‐induced dissociation (CID) spectrum of fibrinopeptide A (ADSGEGDFLAEGGGVR).
shown in Fig. 11. When photoactivation is used, side‐chain cleavage product w8a becomes more prominent in the tandem mass spectrum. The TOF‐TOF photofragment ion spectrum is shown in Fig. 12. Strong signals from y‐type ions are no longer observed, rather x‐type sequence ions are the most common. Side‐chain cleavage products in this spectrum are observed in abundance similar to that of the other sequence informative ions. Comparison of these spectra further confirms that y‐type ions are formed subsequent to internal energy reorganization, requiring a greater fragmentation time scale. Through comparison of the PSD focusing and TOF‐TOF methods, we observe that photoactivation using 193‐nm radiation causes both prompt fragmentations at the sites of the chromophores, as well as rotationally and vibrationally excited species, which dissociate over longer time scales. Although the isomass residues leucine and isoleucine cannot be distinguished on the basis of m=z, these residues can be distinguished by side‐ chain cleavages. For example, leucine exhibits a loss of 42 Da and isoleucine loses 28 (Biemann, 1990b) Side‐chain cleavage products are rarely observed by low‐energy activation or in PSD spectra of MALDI‐formed ions; however, side‐chain cleavage reactions are frequently observed by high‐energy CID and photodissociation (Barbacci and Russell, 1999; Johnson et al., 1988; Yergey et al., 2002) The TOF‐TOF photofragment ion spectrum of peptide HLGLAR is shown in Fig. 13. Because the most basic
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FIG. 11. Post‐source decay (PSD) focusing photofragment ion spectrum of fibrinopeptide A (ADSGEGDFLAEGGGVR).
FIG. 12. Time‐of‐flight (TOF)‐TOF photofragment ion spectrum of fibrinopeptide A (ADSGEGDFLAEGGGVR).
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FIG. 13. Time‐of‐flight (TOF)‐TOF photofragment ion spectrum of HLGLAR.
residue in this peptide is arginine, primarily C‐terminal charge‐retaining photofragment ions are observed. Side‐chain cleavage ions w3a and w5a appear in similar abundance to other sequence informative ions, which can be used to unambiguously assign leucine to the second and fourth amino acid residues. A complete x‐type ion series is observed, as well as other sequence informative ions. The TOF‐TOF photofragment ion spectra of HLGLAR, fibrinopeptide A, and des‐arg9‐bradykinin serve to further illustrate the analytical utility of de novo peptide sequencing using 193‐nm photoactivation. Peptides that are most basic at the N‐terminus produce predominantly a‐type ions. C‐terminal charge‐carrying peptides, such as peptides that result from protein digestion with trypsin, photodissociate into x‐type ions. Interpretation of tandem MS data is simplified when a single complete sequence ion series dominates the mass spectrum. Further sequence information is obtained in the form of side‐chain cleavage ions for both N‐ and C‐terminal charge carriers. To further evaluate the photodissociation tandem TOF for peptide sequencing, we analyzed a tryptic digest of the protein cytochrome C. A‐1 l aliquot of the tryptic digest (0.5 mg=ml) was mixed with matrix and spotted on the MALDI sample stage. It should be stressed that no sample pretreatment was performed; all buffers, salts, and all other impurities present in the digest solution are present in the MALDI sample
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FIG. 14. Matrix‐assisted laser desorption ionization (MALDI)–time‐of‐flight (TOF) mass spectrum of cytochrome C tryptic digest.
spot. Figure 14 contains the MALDI mass spectrum of this sample. Two peptides were selected, those at m=z 1633 and 1168. We previously assigned the sequence of the peptide at m=z 1633 to a heme‐containing peptide (CAQCHTVEK þ heme) based on accurate mass measurements of the parent ion (Edmondson et al., 1996), and this assignment is consistent with the photofragment ion spectrum (Fig. 15). Note that the dominant photofragment ion is the heme at m=z 616. Both a‐ and b‐ type cleavages are observed in the spectrum, but there are too few fragment ions to completely sequence this peptide. However, confirmation of the presence of the heme group supports our earlier assignment. Figure 16 contains the PSD focusing photofragment ion spectrum of the m/z 1168 peptide ion (TGPNLHGLFGR) of cytochrome C. This spectrum contains signals that correspond to the immonium ions of asparagine and phenylalanine. On the basis of a‐, b‐, and y‐type fragment ions, the sequence of this peptide is assigned as TGPNXHGXFGR, where X is either leucine or isoleucine. This spectrum also contains a number of internal and side‐chain cleavages. Of particular interest are the d8 and w7a ions, both of which correspond to leucine residues. Thus, we can assign positions 5 and 8 to leucine and the sequence can be unambiguously assigned to TGPNLHGLFGR. The TOF‐TOF photofragment ion spectrum of phosphorylated angiotensin II (DRVpYIHPF) is displayed in Fig. 17. Complete amino acid sequence information is obtained from a‐ and y‐type ion series. The isoleucine at the fifth position is unambiguously identified using the d5a ion. The identity of
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FIG. 15. Post‐source decay (PSD) focusing photofragment ion spectrum of m=z 1633 tryptic peptide (CAQCHTVEK þ heme).
FIG. 16. Post‐source decay (PSD) focusing photofragment ion spectrum of m=z 1168 tryptic peptide (TGPNLHGLFGR).
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FIG. 17. Time‐of‐flight (TOF)‐TOF photofragment ion spectrum of phosphorylated angiotensin II (DRVpYIHPF).
the modified amino acid is revealed by the phosphorylated tyrosine immonium ion, and its position is clearly denoted by the a‐type ion series. Photofragment ion spectra of phosphorylated peptides acquired using the PSD focusing method include ions identified as peptide fragments with loss of phosphoric acid. It was uncertain as to whether these fragment ions arise from photodissociation of the phosphorylated peptide ion or its phosphate‐ loss metastable ion. In the TOF‐TOF photodissociation experiments, sequence ions with phosphate loss are not observed since the phosphate‐loss metastable ions are not activated. Conclusions
Photodissociation of MALDI‐generated peptide [M þ H]þ ions with 193‐nm excimer radiation results in significant fragmentation. Current results demonstrate that this fragmentation may be used to reconstruct peptide sequences. In addition, photodissociation results in significant numbers of high‐energy side‐chain cleavages, which are particularly useful for differentiating between isomass residues. Our results demonstrate the ability to select peptides of interest from a digest solution and sequence them in the mass spectrometer with no prior sample treatment.
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Also, photodissociation is a reliable method for probing posttranslational modification. Acknowledgments The authors gratefully acknowledge the support of the U.S. Department of Energy, Division of Chemical Science, and the Center for Environmental and Rural Health, Texas A&M University College of Veterinary Medicine, an NIEHS (P30‐ESO9106) funded center, for supporting the photofragment project. Also, David L. McCurdy of Truman State University for assistance in acquiring some of the spectra contained herein, and Zee Yong Park for preparation of the cytochrome C tryptic digest.
References Barbacci, D. C. (2000). Photodissociation of peptide ions in a matrix‐assisted laser desorption ionization reflectron time‐of‐flight mass spectrometer. Ph.D. Dissertation, pp. 1–182. Texas A&M University. Barbacci, D., and Russell, D. (1999). Sequence and side‐chain specific photofragment (193 nm) ions from protonated substance P by matrix‐assisted laser desorption ionization time‐ of‐flight mass spectrometry. J. Am. Soc. Mass Spectrom. 10, 1038–1040. Beavis, R. C., Lindner, J., Grotemeyer, J., and Schlag, E. W. (1988). Sample‐matrix effects in infrared laser neutral desorption, multiphoton‐ionization mass spectrometry. Chem. Phys. Lett. 146, 310–314. Castoro, J. A., and Wilkins, C. L. (1993). Ultrahigh resolution matrix‐assisted laser desorption=ionization of small proteins by Fourier‐transform mass spectrometry. Anal. Chem. 65, 2621–2627. Chaurand, P., Luetzenkirchen, F., and Spengler, B. (1999). Peptide and protein identification by matrix‐assisted laser desorption ionization (MALDI) and MALDI‐post‐source decay time‐of‐flight mass spectrometry. J. Am. Soc. Mass Spectrom. 10, 91–103. Clauser, K. R., Baker, P., and Burlingame, A. L. (1999). Role of accurate mass measurement (10 ppm) in protein identification strategies employing MS or MS=MS and database searching. Anal. Chem. 71, 2871–2882. Cooks, R. G., Jo, S., and Green, J. (2004). Collisions of organic ions at surfaces. App. Surf. Sci. 231–232, 13–21. Cooks, R. G., and Miller, S. A. (1999). Fundamentals and applications of gas phase ion chemistry. NATO ASI Ser. 521, 55–114. Cordero, M. M., Cornish, T. J., and Cotter, R. J. (1996). Matrix‐assisted laser desorption=ionization tandem reflection time‐of‐flight mass spectrometry of fullerenes. J. Am. Soc. Mass Spectrom. 7, 590–597. Cornish, T. J., and Cotter, R. J. (1993). A curved‐field reflectron for improved energy focusing of product ions in time‐of‐flight mass spectrometry. Rap. Comm. Mass Spectrom. 7, 1037–1040. Cotter, R. J. (1999). The new time‐of‐flight mass spectrometry. Anal. Chem. 71, 445A–451A. Cotter, R. J. (1997). ‘‘Time‐of‐Flight Mass Spectrometry: Instrumentation and Applications in Biological Research.’’ pp. 253–274. American Chemical Society, Washington, DC. Dikler, S. (2001). Derivatization of peptides for structural studies by mass spectrometry. Ph.D. Dissertation, pp. 87–108. Texas A&M University.
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Dunbar, R. C. (1989). In ‘‘Gas Phase Inorganic Chemistry’’ (D. H. Russell, ed.), pp. 325–352. Plenum Press, New York and London. Edmondson, R. D., and Russell, D. H. (1996). Evaluation of matrix‐assisted laser desorption ionization‐time‐of‐flight mass measurement accuracy by using delayed extraction. J. Am. Soc. Mass Spectrom. 7, 995–1001. Edmondson, R. D., Barbacci, D. C., and Russell, D. H. (1996). ‘‘Accurate Mass Assignments of Protein Digest Fragments, Proceedings of the 44th ASMS Conference on Mass Spectrometry and Allied Topics, Portland, Oregon, May 12–16.’’ Elsevier, New York, NY. Gimon‐Kinsel, M. E., Kinsel, G. R., Edmondson, R. D., and Russell, D. H. (1995). Photodissociation of high molecular weight peptides and proteins in a two‐stage linear time‐of‐flight mass spectrometer. J. Am. Soc. Mass Spectrom. 6, 578–587. Grill, V., Shen, J., Evan, C., and Cooks, R. G. (2001). Collisions of ions with surfaces at chemically relevant energies: Instrumentation and phenomena. Rev. Sci. Instrum. 72, 3149–3179. Henzel, W. J., Billeci, T. M., Stults, J. T., Wong, S. C., Grimley, C., and Watanabe, C. (1993). Identifying proteins from two‐dimensional gels by molecular mass searching of peptide fragments in protein sequence databases. Proc. Natl. Acad. Sci. USA 90, 5011–5015. Hettick, J. M. (2003). Optimization and utilization of MALDI 193‐nm photofragment time‐of‐ flight mass spectrometry for peptide sequencing. Ph.D. Dissertation, pp. 1–271. Texas A&M University. Hunt, D. F., Shabanowitz, J., and Yates, J. R. (1987). Peptide sequence analysis by laser photodissociation Fourier transform mass spectrometry. J. Chem. Soc., Chem. Commun. 8, 548–550. Ingendoh, A., Karas, M., Hillenkamp, F., and Giessmann, U. (1994). Factors affecting the resolution in matrix‐assisted laser desorption‐ionization mass spectrometry. Int. J. Mass Spectrom. Ion Proc. 131, 345–354. Jensen, O. N., Podtelejnkiov, A., and Mann, M. (1996). Delayed extraction improves specificity in database searches by matrix‐assisted laser desorption/ionization peptide maps. Rapid Comm. Mass Spectrom. 10, 1371–1378. Johnson, R. S., Martin, S. A., and Biemann, K. (1988). Collision‐induced fragmentation of (M þ H)þ ions of peptides. Side chain specific sequence ions. Int. J. Mass Spectrom. Ion Processes. 86, 137–154. Karas, M., Bachmann, D., Bahr, U., and Hillenkamp, F. (1987). Matrix‐assisted ultraviolet laser desorption of non‐volatile compounds. Int. J. Mass Spectrom. Ion Processes. 78, 53–68. Kaufmann, R., Chaurand, P., Kirsch, D., and Spengler, B. (1996). Post‐source decay and delayed extraction in matrix‐assisted laser desorption=ionization‐reflectron time‐of‐flight mass spectrometry. Are there trade‐offs? Rapid Comm. Mass Spectrom. 10, 1199–1208. Ko¨ ster, C., Castoro, J. A., and Wilkins, C. L. (1992). High‐resolution matrix‐assisted laser desorption=ionization of biomolecules by Fourier transform mass spectrometry. J. Am. Chem. Soc. 114, 7572–7574. Lennon, J. J., and Walsh, K. A. (1999). Locating and identifying posttranslational modifications by in‐source decay during MALDI‐TOF mass spectrometry. Protein Sci. 8, 2487–2493. Mabud, M. A., Dekrey, M. J., and Cooks, R. G. (1985). Surface‐induced dissociation of molecular ions. Int. J. Mass Spectrom. Ion Processes. 67, 285–294. Maier, J. P. (1982). Open‐shell organic cations: Spectroscopic studies by means of their radiative decay in the gas phase. Acc. Chem. Res. 15, 18–23. Mamyrin, B. A., Karataev, V. I., Shmikk, D. V., and Zagulin, V. A. (1973). Mass reflectron. New nonmagnetic time‐of‐flight high‐resolution mass spectrometer. Zh. Eksp. Teor. Fiz. 64, 82–89.
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McCormack, A. L., Jones, J. L., and Wysocki, V. H. (1992). Surface‐induced dissociation of multiply protonated peptides. J. Am. Soc. Mass Spectrom. 3, 859–862. Medzihradsky, K. F., Campbell, J. M., Baldwin, M. A., Falick, A. M., Juhasz, P., Vestal, M. L., and Burlingame, A. L. (2000). The characteristics of peptide collision‐induced dissociation using a high‐performance MALDI‐TOF=TOF tandem mass spectrometer. Anal. Chem. 72, 552–558. Nikolaev, E. N., Somogyi, A., Smith, D. L., Gu, C., Wysocki, V. H., Martin, C. D., and Samuelson, G. L. (2001). Implementation of low‐energy surface‐induced dissociation (eV SID) and high‐energy collision‐induced dissociation (keV CID) in a linear sector‐TOF hybrid tandem mass spectrometer. Int. J. Mass Spectrom. 212, 535–551. Riederer, D. E., Haney, L. L., Hilgenbrink, A. R., and Beck, J. R. (2000). ‘‘Investigations into the Influence of Impact Velocity and Ion Mass on the Surface Induced Dissociation Behavior of Large Ions. Proceedings of the 48th ASMS Conference on Mass Spectrometry and Allied Topics, Long Beach, California, June 11–15.’’ Elsevier Inc., New York, NY. Russell, D. H., and Edmondson, R. D. (1997). High‐resolution mass spectrometry and accurate mass measurements with emphasis on the characterization of peptides and proteins by matrix‐assisted laser desorption=ionization time‐of‐flight mass spectrometry. J. Mass Spectrom. 32, 263–276. Seeterlin, M. A., Vlasak, P. R., Beussman, D. J., McLane, R. D., and Enke, C. G. (1993). High efficiency photo‐induced dissociation of precursor ions in a tandem time‐of‐flight mass spectrometer. J. Am. Soc. Mass Spectrom. 4, 751–754. Shukla, A. K., and Futrell, J. H. (1994). In ‘‘Experimental Mass Spectrometry’’ (D. H. Russell, ed.), pp. 71–112. Plenum Press, New York and London. Spengler, B. (1997). Post‐source decay analysis in matrix‐assisted laser desorption=ionization mass spectrometry of biomolecules. J. Mass Spectrom. 32, 1019–1036. Steinfeld, J., Francisco, J., and Hase, W. (1999). In ‘‘Chemical Kinetics and Dynamics,’’ pp. 340–343. Prentice Hall, NJ. Stone, E., Gillig, K. J., Ruotolo, B. T., Fuhrer, K., Gonin, M., Schultz, A., and Russell, D. H. (2001). Surface‐induced dissociation on a MALDI‐ion mobility‐orthogonal time‐of‐flight mass spectrometer: Sequencing peptides from an ‘‘in‐solution’’ protein digest. Anal. Chem. 73, 2233–2238. Tanaka, K., Waki, H., Ido, Y., Akita, S., Yoshida, Y., and Yohida, T. (1988). Protein and polymer analyses up to m=z 100,000 by laser ionization time‐of‐flight mass spectrometry. Rap. Comm. Mass Spectrom. 2, 151–153. Tecklenburg, R. E., Miller, M. N., and Russell, D. H. (1989). Laser ion beam photodissociation studies of model amino acids and peptides. J. Am. Chem. Soc. 111, 1161–1171. Thompson, M., Cui, W., and Reilly, J. (2004). Fragmentation of singly charged peptide ions by photodissociation at lambda ¼ 157 nm. Angew. Chem. Int. Ed. 43, 4791–4794. Uggerud, E., and Derrick, P. J. Z. (1989). Mechanism of energy transfer in collisional activation of kiloelectron‐volt macromolecular ions. Naturforsch A Phys. Sci. 44, 245–246. Whitehouse, C. M., Dreyer, R. N., Yamashita, M., and Fenn, J. B. (1985). Electrospray interface for liquid chromatographs and mass spectrometers. Anal. Chem. 57, 675–679. Wiley, W. C., and McLaren, I. H. (1955). Time‐of‐flight mass spectrometer with improved resolution. Rev. Sci. Instrum. 26, 1150–1157. Willey, K. F., Robbins, D. L., Yeh, C. S., and Duncan, M. A. (1994). In ‘‘Time‐of‐Flight Mass Spectrometry’’ (R. J. Cotter, ed.), pp. 61–72. American Chemical Society, Washington, DC. Yergey, A. L., Coorssen, J. R., Backlund, P. S., Blank, P. S., Humphrey, G. A., Zimmerberg, J., Campbell, J. M., and Vestal, M. L. (2002). De novo sequencing of peptides using MALDI=TOF‐TOF. J. Am. Soc. Mass Spectrom. 13, 784–791.
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Young, M. M., Tang, N., Hempel, J. C., Oshiro, C. M., Taylor, E. W., Kuntz, I. D., Gibson, B. W., and Dollinger, G. (2000). High throughput protein fold identification by using experimental constraints derived from intramolecular cross‐links and mass spectrometry. Proc. Natl. Acad. Sci. USA 97, 5802–5806.
Further Reading Biemann, K. (1990a). Peptides and proteins: Overview and strategy. In ‘‘Methods in Enzymology,’’ (J. A. McClosky, ed.), pp. 351–360. Biemann, K. (1990b). Appendix 5. In ‘‘Methods in Enzymology.’’ pp. 886–887. Elsevier Inc., New York, NY. Breci, L. A., Tabb, D. L., Yates, J. R., and Wysocki, V. H. (2003). Cleavage N‐terminal to proline: Analysis of a database of peptide tandem mass spectra. Anal. Chem. 75, 1963–1971. Hettick, J. M., McCurdy, D. L., Barbacci, D. C., and Russell, D. H. (2001a). Optimization of sample preparation for peptide sequencing by MALDI‐TOF photofragment mass spectrometry. Anal. Chem. 73, 5378–5386. Hettick, J. M., McCurdy, D. L., and Russell, D. H. (2001b). ‘‘Book of Abstracts, 1901P, The Pittsburgh Conference on Analytical Chemistry and Applied Spectroscopy.’’ The Pittsburg Conference, Pittsburg, PA. Pandey, A., and Mann, M. (2000). Proteomics to study genes and genomes. Nature. 405, 837–846. Stone, E. G., Gillig, K. J., Ruotolo, B. T., and Russell, D. H. (2001). Optimization of a matrix‐ assisted laser desorption ionization‐ion mobility‐surface‐induced dissociation‐orthogonal‐ time‐of‐flight mass spectrometer: Simultaneous acquisition of multiple correlated MS1 and MS2 spectra. Int. J. Mass Spectrom. 212, 519–533.
[7] Peptide Sequence Analysis By KATALIN F. MEDZIHRADSZKY Abstract
In mass spectrometry (MS)–based protein studies, peptide fragmentation analysis (i.e., MS=MS experiments such as matrix‐assisted laser desorption ionization [MALDI]–post‐source decay [PSD] analysis, collision‐induced dissociation [CID] of electrospray‐ and MALDI‐generated ions, and electron‐capture and electron‐transfer dissociation analysis of multiply charged ions) provide sequence information and, thus, can be used for (i) de novo sequencing, (ii) protein identification, and (iii) posttranslational or other covalent modification site assignments. This chapter offers a qualitative overview on which kind of peptide fragments are formed under different MS=MS conditions. High‐quality PSD and CID spectra provide illustrations of de novo sequencing and protein identification. The MS=MS METHODS IN ENZYMOLOGY, VOL. 402 Copyright 2005, Elsevier Inc. All rights reserved.
0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)02007-0
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[6] Peptide Sequencing by MALDI 193‐nm Photodissociation TOF MS By JOSEPH W. MORGAN, JUSTIN M. HETTICK , and DAVID H. RUSSELL Abstract
Ultraviolet photodissociation time‐of‐flight (TOF) mass spectrometry (MS) is described as a method for determination of peptide ion primary structure. Monoisotopic selection and bond‐specific activation, combined with the rapidity of TOF MS analysis, render this technique invaluable to the rapidly expanding field of proteomics. Photofragment ion spectra of model peptides acquired using both post‐source decay (PSD) focusing and TOF‐TOF experimental methods are exhibited. Advantages of 193‐nm photodissociation for de novo sequencing of peptide ions are discussed.
Introduction
Advances in ionization methods and instrumentation that have occurred over the past decade enable MS to play a key role in the emerging area of proteomics. For example, matrix‐assisted laser desorption ionization (MALDI) (Karas et al., 1987; Tanaka et al., 1988) and electrospray ionization (ESI) (Whitehouse et al., 1985) are now routinely used to ionize peptides and proteins (Pandey and Mann, 1985), which can then be aimed at sequencing peptides and=or proteins (Chaurand et al., 1999; Cotter, 1997), determination of posttranslational modifications (Lennon and Walsh, 1999), or to study protein–protein complexes (Dikler, 2001; Young et al., 2000). Although various instrument platforms can be used for the mass analysis, over the past decade great strides have been made in the development of TOF instrumentation. Advances in instrument geometry and electronics used for TOF MS allow us to interrogate large gas‐phase ions with improved resolution and mass accuracy (Cotter, 1999; Mamyrin et al., 1973; Wiley and McLaren, 1955). Two extremely important TOF mass spectrometric techniques to the field of proteomics are peptide mass fingerprinting (PMF)=protein database searching (Henzel et al., 1993) and peptide sequencing, and both methods owe their widespread acceptance to the versatility and user friendliness of modern TOF instruments. PMF, combined with genomic database searching to identify proteins, is now widely used, and PMF based on accurate mass measurement is now a METHODS IN ENZYMOLOGY, VOL. 402 Copyright 2005, Elsevier Inc. All rights reserved.
0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)02006-9
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routine TOF analysis (Clauser et al., 1999; Edmondson and Russell, 1996; Jensen et al., 1996; Russell and Edmondson, 1997). Peptide identification based on PMF=protein database searching is especially powerful when combined with peptide sequencing by tandem MS. For example, determining a partial peptide sequence or identifying a specific posttranslationally modified amino acid dramatically increases the confidence level of identification. For proteins that are not listed in a database, peptide sequencing is essential for protein identification. Peptide sequencing using MS is based on fragmentation of the gas‐phase ion. For example, fragmentation along the peptide backbone yields a series of ions separated in mass by an amount equal to the mass of the eliminated residue (Biemann, 1990a). Consequently, the ions must be formed with sufficient internal energy to undergo unimolecular dissociation (metastable ion decay or PSD), or ions formed with very little initial internal energy (i.e., by ESI) must be activated to energies above the dissociation threshold. The most commonly used ion‐activation methods for peptide sequencing are collision‐induced dissociation (CID) (Russell and Edmondson, 1997) and surface‐induced dissociation (SID) (Cooks and Miller, 1999; Nikolaev et al., 2001; Stone et al., 2001); however, photodissociation has considerable potential for peptide sequencing, especially for peptide sequencing involving TOF instruments (Barbacci, 2000; Barbacci and Russell, 1999; Hettick, 2003; Hettick et al., 2001a; Hunt et al., 1987; Thompson et al., 2004). The physics of CID are not compatible with peptide sequencing via PSD focusing using the standard tandem TOF instrument. For instance, high‐energy CID (Chaurand et al., 1999; Yergey et al., 2002) tends to enhance the yield of low m=z fragment ions at the expense of higher mass (e.g., more structurally=sequence informative) fragment ions. Perhaps most troublesome for tandem TOF is the physics of converting kinetic energy into internal energy. CID yields ions having a range of velocities and a broad distribution of arrival times, which limits both m=z resolution and mass measurement accuracy of the TOF measurement (Hettick et al., 2001a). Another important consideration for CID of large ions is the effect of center‐of‐mass collision energy. In virtually all tandem TOF experiments, the mass of the ion undergoing CID is much larger than the target atom. Consequently, only a very small fraction of the laboratory energy can be partitioned as internal energy of the ion, and as the mass of the analyte ion increases, energy deposition becomes even less efficient (Shukla and Futrell, 1994; Uggerud et al., 1989). To circumvent some of the limitations of CID, several groups began investigating SID as an alternative method of ion activation (Mabud et al., 1985; McCormack et al., 1992). Under optimized conditions, SID yields
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very good fragment ion intensities; however, at higher collision energies, SID produces primarily low m=z ions, for example, immonium ions (Riederer et al., 2000; Stone et al., 2001). As the translational energy of the peptide ion is decreased, surface sticking, or ‘‘soft‐landing,’’ becomes competitive with fragmentation processes (Cooks et al., 2004; Grill et al., 2001). Also, because fragment ions in SID leave the surface with a range of velocities and angles, careful attention must be paid to instrument geometry. A comprehensive review of SID has been published by Grill et al. (2001). Several years ago we began evaluating 193‐nm photodissociation as a method to circumvent some of the limitations of CID. Radiation from an ArF excimer (193‐nm, 6.43 eV=photon) laser is particularly well suited to photodissociation of peptide ions, because the amide linkage is a natural chromophore (lmax 210 nm), and the photon imparts sufficient energy to cleave the bond. Relatively few tandem TOF photodissociation experiments have been reported (Gimon‐Kinsel et al., 1995; Seeterlin et al., 1993; Willey et al., 1994) This is in part due to logistical issues that must be addressed in a tandem TOF photodissociation experiment. First, the timing for the trigger pulse for the excimer laser with respect to the ionization=desorption event is critical. In order to intersect the ion beam with a pulsed laser beam, both the delay generator output and the intrinsic time delay in the laser trigger must be stable to 2 ns. (Gimon‐Kinsel et al., 1995), Second, the ions should be irradiated in a region where they are spatially separated and well defined, in terms of m=z. Third, they must be given sufficient time to undergo unimolecular decay in the field‐free region before entering the reflectron. Several years ago we began a series of studies to evaluate photodissociation of peptide and protein ions (Gimon‐Kinsel et al., 1995). The instrument consisted of two collinear TOF mass analyzers, with a photodissociation region positioned between the two analyzers. The instrument used for the preliminary studies was specifically designed to investigate photodissociation of MALDI‐generated peptide and protein ions; however, the instrument did not include a reflectron to mass analyze the resultant photofragments. These studies allowed us to differentiate between fragment ions and neutrals produced by photodissociation of the peptide and protein ions. We also showed that photodepletion of [M þ H]þ ions is a single‐photon process, whereas photodepletion of fragment neutrals, F0n , formed by photoionization of the [M þ H]þ is a two‐photon process. Finally, based on the observation that photodepletion occurred within 1 s of irradiation, it was determined that the rate constant for photodepletion was at least 106 s1.
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Early photodissociation studies were aimed at small organic molecules, and many of these studies were carried out using Fourier transform mass spectrometers (FT‐MSs). The FTMS photodissociation experiment is relatively simple; a defocused light source is passed through the cell and the photofragment ions are trapped and detected using conventional methods (Dunbar, 1989). The long trapping times (milliseconds to hundreds of seconds) and density of neutral molecules (baseline vacuum of 106 to 109 torr) is sufficient for the analyte ion and=or photofragment ions to undergo ion–molecule reactions, which can complicate the photofragment ion spectrum (Willey et al., 1994). Due to the ions being trapped in the FT‐MS ion cell and bathed in a continuous radiation field, it is difficult to control the photoexcitation process (e.g., single vs. multiphoton excitation). Multiphoton excitation deposits very large amounts of energy to the molecule, resulting in extensive fragmentation. In addition, photofragment ions can also absorb a photon and yield second‐generation photofragment ions. These complications do not apply to TOFMS photodissociation experiments. Instrumentation
The results from our preliminary experiments demonstrated efficient photodissociation of protonated peptide=protein ions and motivated us to design and construct an instrument for mass analyzing the photofragment ions. The instrument used for these studies is shown in Fig. 1, and it consists of a linear TOF‐1 and a reflectron that can be used as TOF‐2. The effective flight‐path length of TOF‐1 is 0.5 m and TOF‐2 is 3.4 m. Two turbo molecular pumps maintain baseline pressure of approximately 2 108 torr. It is important that such a high vacuum be maintained to eliminate the possibility of ion collisions with background gases, which contribute ‘‘background fragmentation’’ to the experiment. Ions are generated in the ion source by MALDI (337 nm) and extracted by pulsing the extraction grid potential to ground (Ingendoh et al., 1994). Ions of interest are selected by the timed‐ion selector and subsequently photodissociated by the excimer laser (Fig. 2). The output of the excimer laser is directed to the mass spectrometer via three reflective mirrors (front surface specific to 193 nm), and laser beam size is controlled by a mechanical iris positioned between the final mirror and the vacuum window. The laser output energy is controlled by the laser control panel, and fine control is achieved by passing the laser output through a sample cuvette filled with a methanol=water solution (Hettick et al., 2001b). Control of the laser photon density output is important, because at high laser beam energy, the photofragment ion spectrum is dominated by low m=z immonium ions,
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FIG. 1. Schematic of the MALDI Photofragment Tandem TOF Instrument (A) Source plate; (B) extraction grids; (C) reflectron; (D) stage manipulator; (E) timed ion selector; (F) quartz window; (G) reflected ion detector; (H) linear ion detector; (I) 170‐L=s turbo molecular pump; (J) 330‐L=s turbo molecular pump; (K) roughing pumps; (L) 1‐GHz digital oscilloscope.
FIG. 2. Expanded view of the laser–mass spectrometer interface.
and at lower laser beam energy, the abundance of structurally informative backbone and side‐chain cleavages are increased. Precautions are taken to ensure that the laser beam does not strike the surface of the vacuum chamber because this causes photoejection of electrons and neutral species that can enhance abundances of CID fragment ions.
[6]
peptide sequencing by MALDI 193‐nm photodissociation TOF MS
191
Photofragment ion spectra are calibrated using methods commonly employed for PSD spectra (Spengler, 1997). Briefly, photofragment spectra are taken at a number of PSD reflectron ratios and then are truncated to include only that portion of the spectrum within a few microseconds of the arrival time of the parent ion. The truncated spectra are spliced together to create a complete photofragment ion spectrum. A limitation of PSD experiments is the relatively low mass resolution of the timed‐ion selector (MS1). Currently, the timed‐ion selector is limited to a 300‐ns pulse. Thus, all ions that arrive at the timed‐ion selector within the 300 ns window are allowed to continue toward the reflectron, and any fragment ions formed by dissociation of these ions are observed in the photofragment ion spectrum. Although the mass resolution of the timed‐ ion selector is low enough that [M þ H]þ peptide ions of m=z 1000 (20‐keV translational energy) and [M þ Na]þ ions are not resolved, it is possible to selectively photoexcite only the [M þ H]þ or [M þ Na]þ ion (Fig. 3). For example, as the ions approach the laser‐ion beam intersection, the [M þ H]þ and [M þ Na]þ ions are separated by 6.0 mm, and during a single excimer laser pulse (17 ns), the ions travel approximately 1.0 mm. Thus, even if the
FIG. 3. Spatial separation of ions at the laser–mass spectrometer Interface Isotopes of a theoretical peptide [M þ H]þ of m=z 1000 (a, b, and c) are separated by approximately 0.3 mm, whereas the [M þ Na]þ ion (d) is approximately 6.0 mm behind. The excimer laser is fired at time tL when the [M þ H]þ isotopic distribution reaches the point dL. At time tL þ 17 ns, the excimer laser beam exits the instrument. All ions have advanced approximately 1.05 mm, and only the isotopic distribution of the theoretical peptide [M þ H]þ ion, having traversed through point dL, has absorbed 193‐nm photons.
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[M þ Na]þ ions are allowed to pass the TIS to the photodissociation region, they are not irradiated, and their fragment ions should not appear in the photofragment ion spectrum. It is important that fragmentation from ions such as [M þ Na]þ be eliminated, because they produce product ions shifted in mass (þ23 Da), making spectrum analysis significantly more difficult. Using the PSD method to analyze photofragment ions has limitations as well. Metastable ions that are formed within the first field‐free drift region (TOF‐1) have the same velocity as their respective parent ions. The metastable ions arising from the ion of interest are activated by the photodissociation laser, giving rise to small‐neutral loss ions, which complicate the photofragment ion spectra. Our previous work also showed that metastable neutral species can be ionized by 193‐nm irradiation (Gimon‐Kinsel et al., 1995). The most significant drawback to the photodissociation experiment using PSD focusing is the time required for data acquisition and calibration. As with most PSD experiments, the reflectron voltage must be lowered in increments to acquire the entire fragment ion spectrum, so data acquisition must be repeated n times, where n represents the number of segments needed to stitch the spectrum. To circumvent this limitation, nonlinear reflectron fields have been proposed and used with some success (Cordero et al., 1996; Cornish and Cotter, 1993). Alternatively, ions could be activated ‘‘in‐source’’ so that all fragment ions are accelerated to the same final kinetic energy. In this case, acquisition of the photofragment ion spectrum would have the same duty cycle as a conventional TOF mass analysis. Our most recent photodissociation experiments used an instrument design that negates the limitations described earlier. A four‐grid deceleration=acceleration cell (10 cm in length) was added, centered about the photodissociation window. The MALDI source potential for these experiments was 15 kV, and 8 kV was applied to the central element of the photodissociation cell, and the outside cell grids are held at ground potential. Ions are separated from their metastable decay products as they traverse the decelerating electric field, and consequently, only the ions of interest are irradiated at the center of the cell. A fraction of photoexcited ions dissociate within 0.5 s as they traverse the remaining field‐free region of the photodissociation cell. As ions exit the photodissociation cell, they are then accelerated to near‐uniform kinetic energies and m=z analyzed by TOF‐2. Because differences in final kinetic energies between parent and product ions are small, a complete photofragment ion spectrum is acquired at a single reflectron voltage. Variation in final kinetic energies between photofragment ions is dependent on parent ion mass‐to‐charge ratio and is accounted for in a new calibration equation:
[6]
peptide sequencing by MALDI 193‐nm photodissociation TOF MS
mf TOF ¼ a bmf þ c
12
193
þ t0
Parameters a and c are constant for given mass spectrometer dimensions and source and cell potentials, and parameter b is inversely proportional to the m=z of the parent ion. The [M þ H]þ ions for a mixture of five known peptides (HLGLAR, des‐Arg9 bradykinin, angiotensin III, bradykinin, and substance P) were sequenced by both PSD focusing and TOF‐TOF photodissociation tandem MS methods. All mass spectra and photofragment ion spectra were acquired using sample preparation methods that reduce the abundance of PSD fragment ions (e.g., near‐threshold laser power for MALDI) and using fructose sample preparations (Beavis et al., 1988; Castoro and Wilkins, 1993; Hettick et al., 2001a; Ko¨ ster et al., 1992). CID spectra were acquired using an Applied Biosystems 4700 Proteomics Analyzer (Framingham, MA) (Medzihradsky et al., 2000). Instrument conditions used for CID included 1‐kV accelerating voltage and medium gas pressure.
Mechanism of Photodissociation
Absorption of ultraviolet radiation by a gas‐phase ion yields an electronically excited ion (Fig. 4), and the photoexcited ion may dissociate directly or decay by radiative and=or nonradiative channels. Although radiative decay of gas‐phase ions is an important route for rigid molecules (Maier, 1982), nonradiative decay is a much higher probability process for peptide ions. Because the gas‐phase ion is an isolated system (i.e., energy cannot be transferred to solvent or the local environment), the photoexcited ion may partition the electronic energy to vibrational modes, and the vibrationally excited ion can then undergo dissociation, provided sufficient energy and time are available. According to RRK theory (Steinfeld et al., 1999), the rate for unimolecular dissociation for a vibrationally activated ion having s degrees of freedom and internal energy E is calculated using the following equation: E E0 s1 kðEÞ ¼ V E Rates of dissociation are directly proportional to the amount of excess energy required for bond cleavage (E E0) and the frequency of the critical oscillator (). The process of electronic excitation of a specific oscillator via
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FIG. 4. Vibrational predissociation mechanism absorption of a photon of 193 nm is sufficient to induce a Franck‐Condon transition to the upper electronic state. The ion may then relax via internal conversion to the lower electronic state and fragment.
single‐photon absorption may increase the rate of dissociation for fragmentation pathways that are not accessible for lower energy activation methods (Tecklenburg et al., 1989). Photofragment ion spectra of peptides activated by 157‐nm radiation also provide evidence that photodissociation can occur promptly at bonds that are near the chromophore (Thompson et al., 2004). Peptide Sequencing
For peptide sequencing experiments, it is important to accurately determine the m=z values of fragment ions. Most tandem TOF sequencing experiments have average mass errors in the range of 0.1–1.0 Da (Kaufmann et al., 1996). Although the mass difference between most of the amino acids is more than 1 Da, there are several cases in which mass differences are much less. Leu and Ile have identical masses (113.084 Da) and cannot be distinguished based solely on fragmentation along the peptide backbone. Gln and Lys differ by only 0.036 Da (128.059 and 128.095 Da, respectively), so a mass measurement accuracy of 0.01 is required to distinguish these residues. Table I lists fragment ion mass measurements
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peptide sequencing by MALDI 193‐nm photodissociation TOF MS TABLE I FRAGMENT ION MASS ACCURACY DATA FOR THE HEXAPEPTIDE HLGLAR ACQUIRED ON THE MALDI PHOTOFRAGMENT TOF INSTRUMENT
TOF (ns) 49235.53 47248.32 49195.00 49110.40 49192.09 47897.94 47016.22 48858.32 47941.37 47616.95 48375.26 48517.89 47509.16 47372.89 47149.81 48868.05 46084.94 47695.02 48541.49 49028.12
PSD ratio 1.000 0.870 0.770 0.770 0.740 0.740 0.700 0.635 0.635 0.635 0.560 0.530 0.500 0.410 0.378 0.340 0.240 0.180 0.170 0.130
Ion identity þ
[M þ H] y5 y5‐NH3 b5 þ H2O b5 a5 b4 y4 y4‐NH3 a4 y3 y3‐NH3 b3 b2 y2‐NH3 a2 b1 R A1(H) L
Meas. m=z
Calc. m=z
Error
666.405 529.395 512.286 510.387 492.275 464.347 421.262 416.284 399.302 393.294 359.259 342.232 308.163 251.110 229.071 223.116 138.073 112.082 110.074 86.088
666.405 529.346 510.304 492.293 492.293 464.299 421.256 416.262 399.236 393.261 359.241 342.214 308.172 251.151 229.130 223.156 138.067 112.087 110.072 86.097
0.000 0.049 0.034 0.083 0.018 0.048 0.006 0.022 0.066 0.033 0.018 0.018 0.009 0.041 0.059 0.040 0.006 0.005 0.002 0.009
Average Std. Dev.
0.028 0.024
for the peptide HLGLAR acquired during a PSD focusing photodissociation experiment. The average mass measurement error of 0.028 Da is sufficient to elucidate the composition of this peptide (with the exception of differentiating between Leu and Ile) from either terminus. Both mass accuracy and resolution are higher for photodissociation TOF‐TOF experiments. The utility of photodissociation tandem TOF for peptide sequencing is illustrated by the photofragment ion spectra of bradykinin. Figure 5 contains the PSD focusing photofragment ion spectrum of bradykinin (RPPGFSPFR) taken from a five‐peptide mixture. As in the previous example, immonium ions corresponding to proline, phenylalanine, and arginine are observed, and a complete set of a‐type ions (except a7) are observed. In addition, several b‐type ions are observed as well, notably the b8 þ H2O ion at m=z 904. Ions corresponding to charge retention by the C‐terminus are prominent in this spectrum, as evidenced by several prominent y‐type ions. Because there are arginine residues at either terminus of
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FIG. 5. Post‐source decay (PSD) focusing photofragment ion spectrum of bradykinin (RPPGFSPFR) taken from the five peptide mixture.
this peptide, it is reasonable to expect significant populations of fragment ions exhibiting C‐terminal charge retention from this peptide. In addition to simple backbone cleavage products, numerous side‐chain cleavage products are observed in this spectrum, notably d6, d9, and w3. We also observe internal fragmentation in this spectrum, which is typical for proline‐containing peptides (Breci et al., 1963). Internal fragmentation is highly useful in reconstructing peptide sequence when a complete series of backbone cleavage products is not observed. For instance, using just the a‐type ions observed in this photofragment ion spectrum, the sequence would be assigned ‘‘RPPGFSXXR,’’ where XX can be either FP or PF, but knowledge of one of the internal cleavage ions such as FSP, GFSP, or PGFSP indicates the sequence must be RPPGFSPFR. Alternatively, this information could be extracted from C‐terminal sequence information given by the y‐type ion series. In this case, we are able to assign the sequence as XPXXXSPFR from the observed y‐ions. Thus, in the case of bradykinin, the information obtained from the a‐ and y‐type fragment ions is sufficient to unambiguously assign the complete sequence. Figure 6 contains the TOF‐TOF photofragment ion spectrum of bradykinin. Note that contributions from multiple fragmentations, such as internal cleavage ions and small neutral losses, are in much lower abundance in this spectrum. Products of prompt photodissociation, such as a‐type
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F IG . 6. Time‐of‐flight (TOF)‐TOF photofragment ion spectrum of bradykinin (RPPGFSPFR).
sequence informative ions and d‐type side‐chain cleavage ions, are now the major contributors to the photofragment ion spectrum. The CID spectrum of this species is shown in Fig. 7. Note the appearance of several ion types having charge retention on either side of the dissociating bond and internal fragment ions and other unidentified species. Although much information is contained within this spectrum, its utility for de novo sequencing is low because of difficulties in the assignment of peaks to fragment ions. Figure 8 contains the photofragment ion spectrum of des‐Arg9 bradykinin (RPPGFSPF) obtained from a five‐peptide mixture using the PSD focusing method. The immonium ions for proline, phenylalanine, and arginine verify the presence of these three amino acids in the peptide. A number of a‐ and b‐type fragment ions are also observed in the spectrum, which we attribute to charge retention at the N‐terminus of the peptide ion. We also observe sequence ions that arise by loss of NH3 from either the primary photofragment ions or the activated metastable ions, and these are labeled Fi‐NH3 ions. Also present are fragment ions that result from charge retention at the C‐terminus of the peptide ion or at a basic amino acid side chain near the C‐terminus. For example, y1, y2, and y3 are observed. In addition to simple backbone cleavage reactions (e.g., a‐, b‐, and y‐type ions), we also observe side‐chain cleavage ions. For instance, v4 and w6 arise by side‐chain cleavage products in which the charge is retained by the
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FIG. 7. Collision‐induced dissociation (CID) spectrum of bradykinin (RPPGFSPFR).
FIG. 8. Post‐source decay (PSD) focusing photofragment ion spectrum of des‐Arg9 bradykinin (RPPGFSPF) taken from the five peptide mixture. (Used, with permission, from Medzihradsky et al., 2000.)
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C‐terminus. In addition to the v‐ and w‐side‐chain cleavage ions, side‐chain cleavage product ions corresponding to N‐terminal charge retention, d2, d3, and d6 are also observed. The TOF‐TOF photofragment ion spectrum of des‐Arg9 bradykinin is shown in Fig. 9. Photofragment ions arising from multiple dissociations and=or fragmentation of metastable ions are in much lower abundance. C‐terminal charge retention products such as phenylalanine immonium ion and y‐type ions are observed in lower abundance also. Analysis of these promptly formed photofragment ions provides evidence that N‐terminal charge retention is dominant in the initial population of MALDI formed ions. Through comparison with results from the PSD focusing method, we postulate that b‐ and y‐type ions are more likely to be formed from lower energy, longer‐lived dissociating ions, as they appear in much lower relative abundance when the time allotted for observation of photofragment ions is decreased from 10 s to 0.5 s. Figure 10 contains the CID spectrum of fibrinopeptide A (ADSGEGDFLAEGGGVR). The spectrum consists of mostly y‐type sequence ions and w‐type side chain cleavage ions. Note the dominance of the y‐type ions corresponding to backbone cleavage at acidic residues. These same dominant ions are observed in the PSD focusing photofragment ion spectrum
FIG. 9. Time‐of‐flight (TOF)‐TOF Photofragment ion spectrum of des‐Arg9 bradykinin.
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FIG. 10. Collision‐induced dissociation (CID) spectrum of fibrinopeptide A (ADSGEGDFLAEGGGVR).
shown in Fig. 11. When photoactivation is used, side‐chain cleavage product w8a becomes more prominent in the tandem mass spectrum. The TOF‐TOF photofragment ion spectrum is shown in Fig. 12. Strong signals from y‐type ions are no longer observed, rather x‐type sequence ions are the most common. Side‐chain cleavage products in this spectrum are observed in abundance similar to that of the other sequence informative ions. Comparison of these spectra further confirms that y‐type ions are formed subsequent to internal energy reorganization, requiring a greater fragmentation time scale. Through comparison of the PSD focusing and TOF‐TOF methods, we observe that photoactivation using 193‐nm radiation causes both prompt fragmentations at the sites of the chromophores, as well as rotationally and vibrationally excited species, which dissociate over longer time scales. Although the isomass residues leucine and isoleucine cannot be distinguished on the basis of m=z, these residues can be distinguished by side‐ chain cleavages. For example, leucine exhibits a loss of 42 Da and isoleucine loses 28 (Biemann, 1990b) Side‐chain cleavage products are rarely observed by low‐energy activation or in PSD spectra of MALDI‐formed ions; however, side‐chain cleavage reactions are frequently observed by high‐energy CID and photodissociation (Barbacci and Russell, 1999; Johnson et al., 1988; Yergey et al., 2002) The TOF‐TOF photofragment ion spectrum of peptide HLGLAR is shown in Fig. 13. Because the most basic
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FIG. 11. Post‐source decay (PSD) focusing photofragment ion spectrum of fibrinopeptide A (ADSGEGDFLAEGGGVR).
FIG. 12. Time‐of‐flight (TOF)‐TOF photofragment ion spectrum of fibrinopeptide A (ADSGEGDFLAEGGGVR).
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FIG. 13. Time‐of‐flight (TOF)‐TOF photofragment ion spectrum of HLGLAR.
residue in this peptide is arginine, primarily C‐terminal charge‐retaining photofragment ions are observed. Side‐chain cleavage ions w3a and w5a appear in similar abundance to other sequence informative ions, which can be used to unambiguously assign leucine to the second and fourth amino acid residues. A complete x‐type ion series is observed, as well as other sequence informative ions. The TOF‐TOF photofragment ion spectra of HLGLAR, fibrinopeptide A, and des‐arg9‐bradykinin serve to further illustrate the analytical utility of de novo peptide sequencing using 193‐nm photoactivation. Peptides that are most basic at the N‐terminus produce predominantly a‐type ions. C‐terminal charge‐carrying peptides, such as peptides that result from protein digestion with trypsin, photodissociate into x‐type ions. Interpretation of tandem MS data is simplified when a single complete sequence ion series dominates the mass spectrum. Further sequence information is obtained in the form of side‐chain cleavage ions for both N‐ and C‐terminal charge carriers. To further evaluate the photodissociation tandem TOF for peptide sequencing, we analyzed a tryptic digest of the protein cytochrome C. A‐1 l aliquot of the tryptic digest (0.5 mg=ml) was mixed with matrix and spotted on the MALDI sample stage. It should be stressed that no sample pretreatment was performed; all buffers, salts, and all other impurities present in the digest solution are present in the MALDI sample
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FIG. 14. Matrix‐assisted laser desorption ionization (MALDI)–time‐of‐flight (TOF) mass spectrum of cytochrome C tryptic digest.
spot. Figure 14 contains the MALDI mass spectrum of this sample. Two peptides were selected, those at m=z 1633 and 1168. We previously assigned the sequence of the peptide at m=z 1633 to a heme‐containing peptide (CAQCHTVEK þ heme) based on accurate mass measurements of the parent ion (Edmondson et al., 1996), and this assignment is consistent with the photofragment ion spectrum (Fig. 15). Note that the dominant photofragment ion is the heme at m=z 616. Both a‐ and b‐ type cleavages are observed in the spectrum, but there are too few fragment ions to completely sequence this peptide. However, confirmation of the presence of the heme group supports our earlier assignment. Figure 16 contains the PSD focusing photofragment ion spectrum of the m/z 1168 peptide ion (TGPNLHGLFGR) of cytochrome C. This spectrum contains signals that correspond to the immonium ions of asparagine and phenylalanine. On the basis of a‐, b‐, and y‐type fragment ions, the sequence of this peptide is assigned as TGPNXHGXFGR, where X is either leucine or isoleucine. This spectrum also contains a number of internal and side‐chain cleavages. Of particular interest are the d8 and w7a ions, both of which correspond to leucine residues. Thus, we can assign positions 5 and 8 to leucine and the sequence can be unambiguously assigned to TGPNLHGLFGR. The TOF‐TOF photofragment ion spectrum of phosphorylated angiotensin II (DRVpYIHPF) is displayed in Fig. 17. Complete amino acid sequence information is obtained from a‐ and y‐type ion series. The isoleucine at the fifth position is unambiguously identified using the d5a ion. The identity of
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FIG. 15. Post‐source decay (PSD) focusing photofragment ion spectrum of m=z 1633 tryptic peptide (CAQCHTVEK þ heme).
FIG. 16. Post‐source decay (PSD) focusing photofragment ion spectrum of m=z 1168 tryptic peptide (TGPNLHGLFGR).
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FIG. 17. Time‐of‐flight (TOF)‐TOF photofragment ion spectrum of phosphorylated angiotensin II (DRVpYIHPF).
the modified amino acid is revealed by the phosphorylated tyrosine immonium ion, and its position is clearly denoted by the a‐type ion series. Photofragment ion spectra of phosphorylated peptides acquired using the PSD focusing method include ions identified as peptide fragments with loss of phosphoric acid. It was uncertain as to whether these fragment ions arise from photodissociation of the phosphorylated peptide ion or its phosphate‐ loss metastable ion. In the TOF‐TOF photodissociation experiments, sequence ions with phosphate loss are not observed since the phosphate‐loss metastable ions are not activated. Conclusions
Photodissociation of MALDI‐generated peptide [M þ H]þ ions with 193‐nm excimer radiation results in significant fragmentation. Current results demonstrate that this fragmentation may be used to reconstruct peptide sequences. In addition, photodissociation results in significant numbers of high‐energy side‐chain cleavages, which are particularly useful for differentiating between isomass residues. Our results demonstrate the ability to select peptides of interest from a digest solution and sequence them in the mass spectrometer with no prior sample treatment.
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Also, photodissociation is a reliable method for probing posttranslational modification. Acknowledgments The authors gratefully acknowledge the support of the U.S. Department of Energy, Division of Chemical Science, and the Center for Environmental and Rural Health, Texas A&M University College of Veterinary Medicine, an NIEHS (P30‐ESO9106) funded center, for supporting the photofragment project. Also, David L. McCurdy of Truman State University for assistance in acquiring some of the spectra contained herein, and Zee Yong Park for preparation of the cytochrome C tryptic digest.
References Barbacci, D. C. (2000). Photodissociation of peptide ions in a matrix‐assisted laser desorption ionization reflectron time‐of‐flight mass spectrometer. Ph.D. Dissertation, pp. 1–182. Texas A&M University. Barbacci, D., and Russell, D. (1999). Sequence and side‐chain specific photofragment (193 nm) ions from protonated substance P by matrix‐assisted laser desorption ionization time‐ of‐flight mass spectrometry. J. Am. Soc. Mass Spectrom. 10, 1038–1040. Beavis, R. C., Lindner, J., Grotemeyer, J., and Schlag, E. W. (1988). Sample‐matrix effects in infrared laser neutral desorption, multiphoton‐ionization mass spectrometry. Chem. Phys. Lett. 146, 310–314. Castoro, J. A., and Wilkins, C. L. (1993). Ultrahigh resolution matrix‐assisted laser desorption=ionization of small proteins by Fourier‐transform mass spectrometry. Anal. Chem. 65, 2621–2627. Chaurand, P., Luetzenkirchen, F., and Spengler, B. (1999). Peptide and protein identification by matrix‐assisted laser desorption ionization (MALDI) and MALDI‐post‐source decay time‐of‐flight mass spectrometry. J. Am. Soc. Mass Spectrom. 10, 91–103. Clauser, K. R., Baker, P., and Burlingame, A. L. (1999). Role of accurate mass measurement (10 ppm) in protein identification strategies employing MS or MS=MS and database searching. Anal. Chem. 71, 2871–2882. Cooks, R. G., Jo, S., and Green, J. (2004). Collisions of organic ions at surfaces. App. Surf. Sci. 231–232, 13–21. Cooks, R. G., and Miller, S. A. (1999). Fundamentals and applications of gas phase ion chemistry. NATO ASI Ser. 521, 55–114. Cordero, M. M., Cornish, T. J., and Cotter, R. J. (1996). Matrix‐assisted laser desorption=ionization tandem reflection time‐of‐flight mass spectrometry of fullerenes. J. Am. Soc. Mass Spectrom. 7, 590–597. Cornish, T. J., and Cotter, R. J. (1993). A curved‐field reflectron for improved energy focusing of product ions in time‐of‐flight mass spectrometry. Rap. Comm. Mass Spectrom. 7, 1037–1040. Cotter, R. J. (1999). The new time‐of‐flight mass spectrometry. Anal. Chem. 71, 445A–451A. Cotter, R. J. (1997). ‘‘Time‐of‐Flight Mass Spectrometry: Instrumentation and Applications in Biological Research.’’ pp. 253–274. American Chemical Society, Washington, DC. Dikler, S. (2001). Derivatization of peptides for structural studies by mass spectrometry. Ph.D. Dissertation, pp. 87–108. Texas A&M University.
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Dunbar, R. C. (1989). In ‘‘Gas Phase Inorganic Chemistry’’ (D. H. Russell, ed.), pp. 325–352. Plenum Press, New York and London. Edmondson, R. D., and Russell, D. H. (1996). Evaluation of matrix‐assisted laser desorption ionization‐time‐of‐flight mass measurement accuracy by using delayed extraction. J. Am. Soc. Mass Spectrom. 7, 995–1001. Edmondson, R. D., Barbacci, D. C., and Russell, D. H. (1996). ‘‘Accurate Mass Assignments of Protein Digest Fragments, Proceedings of the 44th ASMS Conference on Mass Spectrometry and Allied Topics, Portland, Oregon, May 12–16.’’ Elsevier, New York, NY. Gimon‐Kinsel, M. E., Kinsel, G. R., Edmondson, R. D., and Russell, D. H. (1995). Photodissociation of high molecular weight peptides and proteins in a two‐stage linear time‐of‐flight mass spectrometer. J. Am. Soc. Mass Spectrom. 6, 578–587. Grill, V., Shen, J., Evan, C., and Cooks, R. G. (2001). Collisions of ions with surfaces at chemically relevant energies: Instrumentation and phenomena. Rev. Sci. Instrum. 72, 3149–3179. Henzel, W. J., Billeci, T. M., Stults, J. T., Wong, S. C., Grimley, C., and Watanabe, C. (1993). Identifying proteins from two‐dimensional gels by molecular mass searching of peptide fragments in protein sequence databases. Proc. Natl. Acad. Sci. USA 90, 5011–5015. Hettick, J. M. (2003). Optimization and utilization of MALDI 193‐nm photofragment time‐of‐ flight mass spectrometry for peptide sequencing. Ph.D. Dissertation, pp. 1–271. Texas A&M University. Hunt, D. F., Shabanowitz, J., and Yates, J. R. (1987). Peptide sequence analysis by laser photodissociation Fourier transform mass spectrometry. J. Chem. Soc., Chem. Commun. 8, 548–550. Ingendoh, A., Karas, M., Hillenkamp, F., and Giessmann, U. (1994). Factors affecting the resolution in matrix‐assisted laser desorption‐ionization mass spectrometry. Int. J. Mass Spectrom. Ion Proc. 131, 345–354. Jensen, O. N., Podtelejnkiov, A., and Mann, M. (1996). Delayed extraction improves specificity in database searches by matrix‐assisted laser desorption/ionization peptide maps. Rapid Comm. Mass Spectrom. 10, 1371–1378. Johnson, R. S., Martin, S. A., and Biemann, K. (1988). Collision‐induced fragmentation of (M þ H)þ ions of peptides. Side chain specific sequence ions. Int. J. Mass Spectrom. Ion Processes. 86, 137–154. Karas, M., Bachmann, D., Bahr, U., and Hillenkamp, F. (1987). Matrix‐assisted ultraviolet laser desorption of non‐volatile compounds. Int. J. Mass Spectrom. Ion Processes. 78, 53–68. Kaufmann, R., Chaurand, P., Kirsch, D., and Spengler, B. (1996). Post‐source decay and delayed extraction in matrix‐assisted laser desorption=ionization‐reflectron time‐of‐flight mass spectrometry. Are there trade‐offs? Rapid Comm. Mass Spectrom. 10, 1199–1208. Ko¨ ster, C., Castoro, J. A., and Wilkins, C. L. (1992). High‐resolution matrix‐assisted laser desorption=ionization of biomolecules by Fourier transform mass spectrometry. J. Am. Chem. Soc. 114, 7572–7574. Lennon, J. J., and Walsh, K. A. (1999). Locating and identifying posttranslational modifications by in‐source decay during MALDI‐TOF mass spectrometry. Protein Sci. 8, 2487–2493. Mabud, M. A., Dekrey, M. J., and Cooks, R. G. (1985). Surface‐induced dissociation of molecular ions. Int. J. Mass Spectrom. Ion Processes. 67, 285–294. Maier, J. P. (1982). Open‐shell organic cations: Spectroscopic studies by means of their radiative decay in the gas phase. Acc. Chem. Res. 15, 18–23. Mamyrin, B. A., Karataev, V. I., Shmikk, D. V., and Zagulin, V. A. (1973). Mass reflectron. New nonmagnetic time‐of‐flight high‐resolution mass spectrometer. Zh. Eksp. Teor. Fiz. 64, 82–89.
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McCormack, A. L., Jones, J. L., and Wysocki, V. H. (1992). Surface‐induced dissociation of multiply protonated peptides. J. Am. Soc. Mass Spectrom. 3, 859–862. Medzihradsky, K. F., Campbell, J. M., Baldwin, M. A., Falick, A. M., Juhasz, P., Vestal, M. L., and Burlingame, A. L. (2000). The characteristics of peptide collision‐induced dissociation using a high‐performance MALDI‐TOF=TOF tandem mass spectrometer. Anal. Chem. 72, 552–558. Nikolaev, E. N., Somogyi, A., Smith, D. L., Gu, C., Wysocki, V. H., Martin, C. D., and Samuelson, G. L. (2001). Implementation of low‐energy surface‐induced dissociation (eV SID) and high‐energy collision‐induced dissociation (keV CID) in a linear sector‐TOF hybrid tandem mass spectrometer. Int. J. Mass Spectrom. 212, 535–551. Riederer, D. E., Haney, L. L., Hilgenbrink, A. R., and Beck, J. R. (2000). ‘‘Investigations into the Influence of Impact Velocity and Ion Mass on the Surface Induced Dissociation Behavior of Large Ions. Proceedings of the 48th ASMS Conference on Mass Spectrometry and Allied Topics, Long Beach, California, June 11–15.’’ Elsevier Inc., New York, NY. Russell, D. H., and Edmondson, R. D. (1997). High‐resolution mass spectrometry and accurate mass measurements with emphasis on the characterization of peptides and proteins by matrix‐assisted laser desorption=ionization time‐of‐flight mass spectrometry. J. Mass Spectrom. 32, 263–276. Seeterlin, M. A., Vlasak, P. R., Beussman, D. J., McLane, R. D., and Enke, C. G. (1993). High efficiency photo‐induced dissociation of precursor ions in a tandem time‐of‐flight mass spectrometer. J. Am. Soc. Mass Spectrom. 4, 751–754. Shukla, A. K., and Futrell, J. H. (1994). In ‘‘Experimental Mass Spectrometry’’ (D. H. Russell, ed.), pp. 71–112. Plenum Press, New York and London. Spengler, B. (1997). Post‐source decay analysis in matrix‐assisted laser desorption=ionization mass spectrometry of biomolecules. J. Mass Spectrom. 32, 1019–1036. Steinfeld, J., Francisco, J., and Hase, W. (1999). In ‘‘Chemical Kinetics and Dynamics,’’ pp. 340–343. Prentice Hall, NJ. Stone, E., Gillig, K. J., Ruotolo, B. T., Fuhrer, K., Gonin, M., Schultz, A., and Russell, D. H. (2001). Surface‐induced dissociation on a MALDI‐ion mobility‐orthogonal time‐of‐flight mass spectrometer: Sequencing peptides from an ‘‘in‐solution’’ protein digest. Anal. Chem. 73, 2233–2238. Tanaka, K., Waki, H., Ido, Y., Akita, S., Yoshida, Y., and Yohida, T. (1988). Protein and polymer analyses up to m=z 100,000 by laser ionization time‐of‐flight mass spectrometry. Rap. Comm. Mass Spectrom. 2, 151–153. Tecklenburg, R. E., Miller, M. N., and Russell, D. H. (1989). Laser ion beam photodissociation studies of model amino acids and peptides. J. Am. Chem. Soc. 111, 1161–1171. Thompson, M., Cui, W., and Reilly, J. (2004). Fragmentation of singly charged peptide ions by photodissociation at lambda ¼ 157 nm. Angew. Chem. Int. Ed. 43, 4791–4794. Uggerud, E., and Derrick, P. J. Z. (1989). Mechanism of energy transfer in collisional activation of kiloelectron‐volt macromolecular ions. Naturforsch A Phys. Sci. 44, 245–246. Whitehouse, C. M., Dreyer, R. N., Yamashita, M., and Fenn, J. B. (1985). Electrospray interface for liquid chromatographs and mass spectrometers. Anal. Chem. 57, 675–679. Wiley, W. C., and McLaren, I. H. (1955). Time‐of‐flight mass spectrometer with improved resolution. Rev. Sci. Instrum. 26, 1150–1157. Willey, K. F., Robbins, D. L., Yeh, C. S., and Duncan, M. A. (1994). In ‘‘Time‐of‐Flight Mass Spectrometry’’ (R. J. Cotter, ed.), pp. 61–72. American Chemical Society, Washington, DC. Yergey, A. L., Coorssen, J. R., Backlund, P. S., Blank, P. S., Humphrey, G. A., Zimmerberg, J., Campbell, J. M., and Vestal, M. L. (2002). De novo sequencing of peptides using MALDI=TOF‐TOF. J. Am. Soc. Mass Spectrom. 13, 784–791.
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Young, M. M., Tang, N., Hempel, J. C., Oshiro, C. M., Taylor, E. W., Kuntz, I. D., Gibson, B. W., and Dollinger, G. (2000). High throughput protein fold identification by using experimental constraints derived from intramolecular cross‐links and mass spectrometry. Proc. Natl. Acad. Sci. USA 97, 5802–5806.
Further Reading Biemann, K. (1990a). Peptides and proteins: Overview and strategy. In ‘‘Methods in Enzymology,’’ (J. A. McClosky, ed.), pp. 351–360. Biemann, K. (1990b). Appendix 5. In ‘‘Methods in Enzymology.’’ pp. 886–887. Elsevier Inc., New York, NY. Breci, L. A., Tabb, D. L., Yates, J. R., and Wysocki, V. H. (2003). Cleavage N‐terminal to proline: Analysis of a database of peptide tandem mass spectra. Anal. Chem. 75, 1963–1971. Hettick, J. M., McCurdy, D. L., Barbacci, D. C., and Russell, D. H. (2001a). Optimization of sample preparation for peptide sequencing by MALDI‐TOF photofragment mass spectrometry. Anal. Chem. 73, 5378–5386. Hettick, J. M., McCurdy, D. L., and Russell, D. H. (2001b). ‘‘Book of Abstracts, 1901P, The Pittsburgh Conference on Analytical Chemistry and Applied Spectroscopy.’’ The Pittsburg Conference, Pittsburg, PA. Pandey, A., and Mann, M. (2000). Proteomics to study genes and genomes. Nature. 405, 837–846. Stone, E. G., Gillig, K. J., Ruotolo, B. T., and Russell, D. H. (2001). Optimization of a matrix‐ assisted laser desorption ionization‐ion mobility‐surface‐induced dissociation‐orthogonal‐ time‐of‐flight mass spectrometer: Simultaneous acquisition of multiple correlated MS1 and MS2 spectra. Int. J. Mass Spectrom. 212, 519–533.
[7] Peptide Sequence Analysis By KATALIN F. MEDZIHRADSZKY Abstract
In mass spectrometry (MS)–based protein studies, peptide fragmentation analysis (i.e., MS=MS experiments such as matrix‐assisted laser desorption ionization [MALDI]–post‐source decay [PSD] analysis, collision‐induced dissociation [CID] of electrospray‐ and MALDI‐generated ions, and electron‐capture and electron‐transfer dissociation analysis of multiply charged ions) provide sequence information and, thus, can be used for (i) de novo sequencing, (ii) protein identification, and (iii) posttranslational or other covalent modification site assignments. This chapter offers a qualitative overview on which kind of peptide fragments are formed under different MS=MS conditions. High‐quality PSD and CID spectra provide illustrations of de novo sequencing and protein identification. The MS=MS METHODS IN ENZYMOLOGY, VOL. 402 Copyright 2005, Elsevier Inc. All rights reserved.
0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)02007-0