Edman sequencing. One of the earliest methods used for protein identification was microsequencing by Edman chemistry to obtain N-terminal amino acid sequences. Little has changed in Edman chemistry since its introduction, but improvements in sequencing technology have increased the sensitivity and ease of Edman sequencing. Although the use of Edman sequencing is waning in the field of proteomics, it is still a very useful tool for several reasons. First, because Edman sequencing existed before MS as a sequencing tool, a considerable number of investigators continue to use Edman sequencing. Second, Edman sequencing of relatively abundant proteins is a viable alternative to MS if a mass spectrometer is in high demand for the identification of low-copy proteins or is not available. Finally, Edman sequencing is used to obtain the N-terminal sequence of a protein (if possible) to determine its true start.

The N-terminal sequencing of proteins was introduced by Edman in 1949 (45). Today, Edman sequencing is most often used to identify proteins after they are transferred to membranes. The development of membranes compatible with sequencing chemicals allowed Edman sequencing to become a more applicable sequencing method for the identification of proteins separated by SDS-polyacrylamide gel electrophoresis (8, 159). One of the biggest problems that has limited the success of Edman sequencing in the past is N-terminal modification of proteins. Since it is difficult to tell if a protein is N-terminally blocked before it is sequenced, precious samples were often lost in failed sequencing attempts. To overcome this problem, we developed a novel approach called mixed-peptide sequencing (38). In mixed-peptide sequencing, a protein is converted into peptides by cleavage with cyanogen bromide (CNBr) or skatole and the peptides are sequenced in an Edman sequencer simultaneously (9, 38, 99).

Briefly, the process of mixed-peptide sequencing involves separation of a complex protein mixture by polyacrylamide gel electrophoresis (1-D or 2-D) and then transfer of the proteins to an inert membrane by electroblotting (Fig. 4). The proteins of interest are visualized on the membrane surface, excised, and fragmented chemically at methionine (by CNBr) or tryptophan (by skatole) into several large peptide fragments. On average, three to five peptide fragments are generated, consistent with the frequency of occurrence of methionine and tryptophan in most proteins. The membrane piece is placed directly into an automated Edman sequencer without further manipulation. Between 6 and 12 automated Edman cycles are carried out (4 to 8 h), and the mixed-sequence data are fed into the FASTF or TFASTF algorithms, which sort and match the data against protein (FASTF) and DNA (TFASTF) databases to unambiguously identify the protein. The FASTF and TFASTF programs were written in collaboration with William Pearson (Department of Biochemistry, University of Virginia). Because minimal sample handling is involved, mixed-peptide sequencing can be a sensitive approach for identifying proteins in polyacrylamide gels at the 0.1- to 1-pmol level. An example of mixed-peptide sequencing is shown in Fig. 5A. The mixed-sequence approach has the advantage of enabling subsequent searches to be carried out against unannotated or non-species-specific DNA databases as well as annotated protein databases. This is because the T/FASTF algorithms utilize actual amino acid sequence and are therefore able to tolerate errors in the database as well as polymorphisms or conservative substitutions. A recent variation of T/FASTF has been devised for MS (101) (Fig. 5B). The T/FASTF/S programs are available at http://fasta.bioch.virginia.edu/ (Table 1).

 
Mass spectrometry. MS enables protein structural information, such as peptide masses or amino acid sequences, to be obtained. This information can be used to identify the protein by searching nucleotide and protein databases (Fig. 4). It also can be used to determine the type and location of protein modifications. The harvesting of protein information by MS can be divided into three stages: (i) sample preparation, (ii) sample ionization, and (iii) mass analysis.

(i) Sample preparation. In most of proteomics, a protein is resolved from a mixture by using a 1- or 2-D polyacrylamide gel. The challenge is to extract the protein or its constituent peptides from the gel, purify the sample, and analyze it by MS. The extraction of whole proteins from gels is inefficient; however, if a protein is "in-gel" digested with a protease, many of the peptides can be extracted from the gel. A method for in-gel protein digestion was developed (149, 169) and is now commonly applied to both 1- and 2-D gels (136). In-gel digestion is more efficient at sample recovery than other common methods such as electroblotting (37). In addition, the conversion of a protein into its constituent peptides provides more information than can be obtained from the whole protein itself. For many applications, the peptides recovered following in-gel digestion need to be purified to remove gel contaminants. Common impurities from electrophoresis such as salts, buffers, and detergents can interfere with MS (172). In addition, peptide samples often require concentration before being analyzed by MS. One method of peptide purification commonly employed for this purpose is reverse-phase chromatography, which is available in a variety of formats. Peptides can be purified with ZipTips (Millipore) or Poros R2 perfusion material (PerSeptive Biosystems, Framingham, Mass.) (149, 169, 170) or by high-pressure liquid chromatography (HPLC).

(ii) Sample ionization. For biological samples to be analyzed by MS, the molecules must be charged and dry. This is accomplished by converting them to desolvated ions. The two most common methods for this are electrospray ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI). In both methods, peptides are converted to ions by the addition or loss of one or more protons. ESI and MALDI are "soft" ionization methods that allow the formation of ions without significant loss of sample integrity. This is important because it enables accurate mass information to be obtained about proteins and peptides in their native states.

(a) Electrospray ionization. In ESI, a liquid sample flows from a microcapillary tube into the orifice of the mass spectrometer, where a potential difference between the capillary and the inlet to the mass spectrometer results in the generation of a fine mist of charged droplets (52, 72, 172). As the solvent evaporates, the sizes of the droplets decrease, resulting in the formation of desolvated ions (52). A significant improvement in ESI technology occurred with the development of nanospray ionization (169, 170). In nanospray ionization, the microcapillary tube has a spraying orifice of 1 to 2 µm and flow rates as low as 5 to 10 nl/min (170). The low flow rates possible with nanospray ionization reduce the amount of sample consumed and increase the time available for analysis (148, 149). For ESI, there are several ways to deliver the sample to the mass spectrometer. The simplest method is to load individual microcapillary tubes with sample. Because a new microcapillary tube is used for each sample, cross-contamination is avoided. In ESI, peptides require some form of purification after in-gel digestion, and this can be accomplished directly in the microcapillary tubes. The drawback to both the purification and manual loading of microcapillary tubes is that it is tedious and slow. As an alternative, electrospray sources have been connected in line with liquid chromatography (LC) systems that automatically purify and deliver the sample to the mass spectrometer. Examples of this method are LC (39, 55, 95, 106), reverse-phase LC (RP-LC) (64) and reverse-phase microcapillary LC (RP-µLC) (41).

(b) Matrix-assisted laser desorption/ionization. In MALDI, the sample is incorporated into matrix molecules and then subjected to irradiation by a laser. The laser promotes the formation of molecular ions (84). The matrix is typically a small energy-absorbing molecule such as 2,5-dihydroxybenzoic acid or -cyano-4-hydroxycinnamic acid. The analyte is spotted, along with the matrix, on a metal plate and allowed to evaporate, resulting in the formation of crystals. The plate, which can be 96-well format, is then placed in the mass spectrometer, and the laser is automatically targeted to specific places on the plate. Since sample application can be performed by a robot, the entire process including data collection and analysis can be automated. This is the single biggest advantage of MALDI. Another advantage of MALDI over ESI is that samples can often be used directly without any purification after in-gel digestion (131).

(iii) Mass analysis. Mass analysis follows the conversion of proteins or peptides to molecular ions. This is accomplished by the mass analyzers in a mass spectrometer, which resolve the molecular ions on the basis of their mass and charge in a vacuum.

(a) Quadrupole mass analyzers. One of the most common mass analyzers is the quadrupole mass analyzer. Here, ions are transmitted through an electric field created by an array of four parallel metal rods, the quadrupole (172). A quadrupole can act to transmit all ions or as a mass filter to allow the transmission of ions of a certain mass-to-charge (m/z) ratio. If multiple quadrupoles are combined, they can be used to obtain information about the amino acid sequence of a peptide. For a more detailed review of the operating principles of a quadrupole mass analyzer, the reader is directed to several excellent reviews (25, 109, 172).

(b) Time of flight. A time-of-flight (TOF) instrument is one of the simplest mass analyzers. It measures the m/z ratio of an ion by determining the time required for it to traverse the length of a flight tube. Some TOF mass analyzers include an ion mirror at the end of the flight tube, which reflects ions back through the flight tube to a detector. In this way, the ion mirror serves to increase the length of the flight tube. The ion mirror also corrects for small energy differences among ions (172). Both of these factors contribute to an increase in mass resolution.

(c) Ion trap. Ion trap mass analyzers function to trap molecular ions in a 3-D electric field. In contrast to a quadrupole mass analyzer, in which ions are discarded before the analysis begins, the main advantage of an ion trap mass analyzer is the ability to allow ions to be "stored" and then selectively ejected from the ion trap, increasing sensitivity (172). For a review of the operating principles of an ion trap mass spectrometer, see reference 34.

(iv) Types of mass spectrometers. Most mass spectrometers consist of four basic elements: (i) an ionization source, (ii) one or more mass analyzers, (iii) an ion mirror, and (iv) a detector. The names of the various instruments are derived from the name of their ionization source and the mass analyzer. Some of the most common mass spectrometers are discussed; for a more comprehensive review of mass spectrometers, the reader is directed to references (76 and 172). The analysis of proteins or peptides by MS can be divided into two general categories: (i) peptide mass analysis and (ii) amino acid sequencing. In peptide mass analysis or peptide mass fingerprinting, the masses of individual peptides in a mixture are measured and used to create a mass spectrum (70). In amino acid sequencing, a procedure known as tandem mass spectrometry, or MS/MS, is used to fragment a specific peptide into smaller peptides, which can then be used to deduce the amino acid sequence.

(a) Triple quadrupole. Triple-quadrupole mass spectrometers are most commonly used to obtain amino acid sequences. In the first stage of analysis, the machine is operated in MS scan mode and all ions above a certain m/z ratio are transmitted to the third quadrupole for mass analysis (Fig. 6) (82, 173). In the second stage, the mass spectrometer is operated in MS/MS mode and a particular peptide ion is selectively passed into the collision chamber. Inside the collision chamber, peptide ions are fragmented by interactions with an inert gas by a process known as collision-induced dissociation or collisionally activated dissociation. The peptide ion fragments are then resolved on the basis of their m/z ratio by the third quadrupole (Fig. 6). Since two different mass spectra are obtained in this analysis, it is referred to as tandem mass spectrometry (MS/MS). MS/MS is used to obtain the amino acid sequence of peptides by generating a series of peptides that differ in mass by a single amino acid (71, 73).

View larger version (28K): [in this window] [in a new window]   FIG. 6. MS/MS. Conventional and MS/MS modes of analysis in a triple-quadrupole mass spectrometer are shown. (A) In the normal scanning mode, all ions of a certain m/z range are transmitted through the first two quadrupoles for mass analysis in the third quadrupole. From this MS spectrum, a parent ion is selected for fragmentation in the collision cell. (B) In MS/MS mode, the parent ion is selectively transmitted into the collision chamber and fragmented, and the resulting daughter ions are resolved in the third quadrupole.

 
(b) Quadrupole-TOF. In recent years, several "hybrid" mass spectrometers have emerged from the combination of different ionization sources with mass analyzers. One example is the quadrupole-TOF mass spectrometer (111, 112, 162). In this machine, the first quadrupole (Q₁) and the quadrupole collision cell (q) of a triple-quadrupole machine have been combined with a time-of-flight analyzer (TOF) (145). The main applications of a QqTOF mass spectrometer are protein identification by amino acid sequencing and characterization of protein modifications. However, because it is coupled to electrospray, it is not typically utilized for large-scale proteomics.

(c) MALDI-TOF. The principal application of a MALDI-TOF mass spectrometer is peptide mass fingerprinting because it can be completely automated, making it the method of choice for large-scale proteomics work (48). Because of its speed, MALDI-TOF is frequently used as a first-pass instrument for protein identification. If proteins cannot be identified by fingerprinting, they can then be analyzed by electrospray and MS/MS. A MALDI-TOF machine can also be used to obtain the amino acid sequence of peptides by a method known as post-source decay (152). However, peptide sequencing by post-source decay is not as reliable as sequencing with competing electrospray methods because the peptide fragmentation patterns are much less predictable (85, 111).

(d) MALDI-QqTOF. The MALDI-QqTOF mass spectrometer was developed to permit both peptide mass fingerprinting and amino acid sequencing (97, 147). It was formed by the combination of a MALDI ion source with a QqTOF mass analyzer (63, 91, 97, 147, 162). Thus, if a sample is not identified by peptide mass fingerprinting in the first step, the amino acid sequence can then be obtained without having to use a different mass spectrometer. However, in our experience, the amino acid sequence information obtained using this instrument was more difficult to interpret than that obtained from a nanospray-QqTOF mass spectrometer.

(e) FT-ICR. A Fourier transform ion cyclotron resonance (FT-ICR) mass spectrometer is an ion-trapping instrument that can achieve higher mass resolution and mass accuracy than any other type of mass spectrometer (10). Recently, FT-ICR has been employed in the analysis of biomolecules ionized by both ESI and MALDI. The unique abilities of FT-ICR provide certain advantages compared to other mass spectrometers. For example, because of its high resolution, FT-ICR can be used for the analysis of complex mixtures. FT-ICR, coupled to ESI, is also being employed in the study of protein interactions and protein conformations. A high-throughput, large-scale proteomics approach involving FT-ICR has recently been developed by Smith et al. (150). For a review of the operating principles of FT-ICR and its applications, the reader is directed to reference 104.

(v) Peptide fragmentation. As peptide ions are introduced into the collision chamber, they interact with the collision gas (usually nitrogen or argon) and undergo fragmentation primarily along the peptide backbone (71, 73, 172). Since peptides can undergo multiple types of fragmentation, nomenclature has been created to indicate what type of ions have been generated (Fig. 7). If, after peptide bond cleavage, the charge is maintained on the N-terminus of the ion, it is designated a b-ion, whereas if the charge is maintained on the C terminus, it is a y-ion (Fig. 7) (18, 135, 173). The difference in mass between adjacent y- or b-ions corresponds to that of an amino acid. This can be used to identify the amino acid and hence the peptide sequence, with the exception of isoleucine and leucine, which are identical in mass and therefore indistinguishable (103). Both y- and b-type ions can also eliminate NH₃ (-17 Da), H₂O (-18 Da) and CO (-28 Da), resulting in pairs of signals observed in the mass spectrum (Fig. 7). In addition to fragmentation along the peptide backbone, cleavage can occur along amino acid side chains, and this information can be used to distinguish isoleucine and leucine (172).

View larger version (13K): [in this window] [in a new window]   FIG. 7. Peptide ion fragmentation nomenclature. Low-energy collisions promote fragmentation of a peptide primarily along the peptide backbone (73). Peptide fragmentation which maintains the charge on the C terminus is designated a y-ion, whereas fragmentation which maintains the charge on the N terminus is designated a b-ion. Additional types of fragmentation are also indicated.

 
(vi) Our approach to mass spectrometry. The sensitivity of a mass spectrometer is probably the single most important feature of the instrument. What is the sensitivity of a modern mass spectrometer? How much protein is needed to make an unambiguous identification? Many factors can affect sensitivity, such as sample preparation, sample ionization, the type of mass spectrometer used, the sample itself, and the type of database search employed. In our laboratory, we rely on 1- or 2-DE electrophoresis for the isolation and visualization of protein targets. We typically stain our gels with either Coomassie blue or silver stain. For most proteins, staining with Coomassie blue will give a dark band for 1µg of protein and a discernible one for 200 ng. With silver staining, we can detect a dark band at 50 ng and faint yet discernible bands at 5 to 10 ng. However, a significant number of proteins do not stain well by these methods and larger proteins tend to bind more stain (mole/mole) than small proteins. In addition, MS is not a quantitative technique because peptide ionization is not quantitative. Therefore, some proteins that are barely visible on gels can give stronger signals by MS than do some darkly staining proteins. For example, one of the most frequently sequenced proteins in MS is human keratin, a component of dust. It is a contaminant that will often appear on polyacrylamide gels as faint silver-stained bands with a variety of molecular weights. It can be introduced simply from the glass plates or gel combs used for protein gels; therefore, it is a good idea to wash these items in concentrated acid before use.

We have found in our laboratory that most proteins applied to the gel at 5 to 10 ng (100 to 200 fmol for a 50-kDa protein) can be identified by MS. However, the ability to identify a protein depends on the protein itself and its presence in the database. Below 5 to 10 ng, the success rate decreases because fewer peptides are obtained for sequencing. Several prominent MS laboratories routinely report record-breaking sequencing sensitivity to the attomolar level. However, this sensitivity is usually toward a purified peptide sample that is directly introduced into the mass spectrometer. Since most proteins are isolated from gels for identification, this is not an accurate measure of sensitivity. In another case, it was reported that an amino acid sequence was obtained after the in-gel digestion of 25 fmol (1.7 ng) of pure bovine serum albumin (90). Again, since the protein was known before the analysis began, this is not a fair assessment of sensitivity. For unknown proteins, more protein is required because several peptides have to be sequenced before a confident assignment can be made.

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