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Alzate O, editor. Neuroproteomics. Boca Raton (FL): CRC Press/Taylor & Francis; 2010.

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Chapter 5Mass Spectrometry for Proteomics

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"Proteomics" is a word coined in 1994 by Marc Wilkins as an alternative to “the protein complement of the genome” (1). Proteomics is still defined in various ways (2), from “the large-scale analysis of the proteome” to “the simultaneous study of all proteins in the cell.” In this chapter, we define it as the study of proteins and their interactions. Proteomics is a new field—only 10 years old—and the rapid evolution of this field is due in large part to many improvements in mass spectrometry (MS) that have occurred during the past several years.

Mass spectrometers do one thing—they measure mass. In proteomics, the mass gives information on the protein identity, its chemical modifications, and its structure. Every mass spectrometer has three main components: a source, an analyzer, and a detector. Mass spectrometers measure masses of charged species, so the source must be able to produce ions, the analyzer must be able to separate these ions based on their mass (or, more accurately, mass-to-charge ratio), and the detector must be able to detect charged particles and then amplify the response to give a measurable signal.

In 2002, the award for the Nobel Prize in chemistry was given to two scientists (John Fenn and Koichi Tanaka) responsible for the development of two ionization techniques that have revolutionized biomedical mass spectrometry in general, and proteomics in particular. These two techniques, electrospray (3) and MALDI (matrix-assisted laser desorption ionization) (4) mass spectrometry, were groundbreaking in that they allow the vaporization and ionization (and thus the analysis) of relatively large, non-volatile biomolecules such as proteins and peptides. In addition, simultaneous improvements to and development of mass analyzers and detectors have greatly increased the use of mass spectrometry for biological studies.


To really appreciate the dramatic changes in mass spectrometry that have resulted from these new ionization techniques, one has to understand the limitations of mass spectrometry for the analysis of biomolecules before MS was developed. As recently as the late 1970s, mass spectrometry was limited to the analysis of volatile and low-molecular-weight components. One of the most common ionization techniques was electron impact (EI), which uses an “open” low pressure source, in which an electron beam (from a filament) is used to “knock an electron off” a molecule to give a positive ion. This is a fairly high-energy process, often resulting in fragmentation and production of molecular ions (M+) at low relative abundances.

Another available ionization technique was positive ion chemical ionization (CI), in which a high-pressure (0.5–1.0 torr) reagent gas (usually methane) is ionized via a filament in a closed, higher-pressure source. The protonated reagent ions then transfer a proton to the target analytes, producing a positive ion and providing a more gentle ionization technique with less fragmentation than electron impact. In the late 1970s, negative chemical ionization was developed. This technique, similar to positive chemical ionization, uses methane (0.3–0.5 torr) to “slow” the electrons released from the filament to the point where they can be “captured” by electronegative analytes to form negative ions by electron capture (Mr), or by proton abstraction (M - H)-. Both EI and CI produce singly charged ions, M+ from electron impact and M-, (M-H)-, M+, and (M+H)+, from chemical ionization. These EI and CI sources were typically interfaced with mass analyzers that could detect only up to ~650 Da.

The ionization technique for which John Fenn received the Nobel Prize in 2002 was electrospray ionization (ESI) (3). In this technique, developed around 1985, the sample solution is passed through a needle which is kept at high voltage (~2–5 kV). This results in a spray of small charged droplets containing the analyte. As the solvent evaporates the droplet shrinks until completely desolvated, resulting in a highly-charged species. The extremely high charge density results in a “coulombic explosion” to produce multiply charged, yet stable analyte ions (Figure 5.1). Multiply charged analytes are a desirable feature especially for proteomics, since mass spectrometers separate ions based on their mass-to-charge (m/z) ratios. With multiple or higher charges, higher-molecular-weight compounds (that typically have masses outside the range of the mass spectrometer) could then be analyzed because they have an apparent low-molecular m/z. For example, if you have an instrument with a mass range of 1000 Da, with EI or CI, analysis was limited to analytes with molecular masses of 1000 Da or below. In contrast, if the sample was introduced by ESI, a compound of Mw 4000, with 10 charges, for example, would appear at [(4000 + 10 x 1.008 {the mass of a proton})/10] Da, or approximately 401.0 Da, which is within the detectable mass range.

FIGURE 5.1. Electrospray ionization.


Electrospray ionization.

Figure 5.2A shows the ESI spectrum of a 38 kDa protein obtained by nanoelectrospray ionization. In this high-sensitivity technique, the sample is placed in a glass needle with a 10–20 μm orifice, and is pulled into the vacuum system without the use of any liquid chromatography pumps. The flow rates are very low (a few nanoliters per minute). Because each peak differs from the next by a single charge, a simple calculation can be used to “deconvolute” this spectrum into the average molecular weight of each form of the protein (Figure 5.2B).

FIGURE 5.2. ESI mass spectrum of a 38 kDa protein (collaborator: Hengming Ke).


ESI mass spectrum of a 38 kDa protein (collaborator: Hengming Ke). (a) Spectrum as acquired, showing charge states, (b) Deconvoluted spectrum showing average and apex masses.

The 2002 Nobel Prize was shared between Dr. Fenn and Koichi Tanaka, whose research led to the development of an ionization method called “matrix-assisted laser desorption/ionization” (MALDI) in which a laser and a UV-absorbing chemical compound are used to vaporize and ionize the analyte (Figure 5.3). In MALDI, a laser pulse, typically at a wavelength of 337 nm, is fired at a solid analyte that has been co-crystallized with a chemical matrix, usually a substituted benzoic acid, typically alpha-cyano-4-hydroxybenzoic acid or 2,5-dihydroxybenzoic acid (DHB). This creates a vapor plume of analyte, matrix, and their ions.

FIGURE 5.3. MALDI ionization.


MALDI ionization.

MALDI and ESI can be considered as complementary techniques. They deal with the analyte at two different physical states—solid and liquid. As with electrospray, MALDI produces multiply charged analytes, although typically with less charge than with electrospray. Figure 5.4 shows the MALDI spectrum of the same 38 kDa protein shown above (Figure 5.2A) with electrospray. When compared to Figure 5.2A you can see the difference between the ESI spectra and the MALDI spectra in the number of charges the protein incorporates, the measured m/z, and the improvement in mass resolution using the nanoelectrospray mode.

FIGURE 5.4. MALDI spectrum of the same 38 kDa protein as in Figure 5.


MALDI spectrum of the same 38 kDa protein as in Figure 5.2 (collaborator: Hengming Ke).

In terms of detecting intact proteins, electrospray is more limited, with an upper mass limit of 65–70 kDa in a standard commercial instrument, versus the 250 kDa practical mass limit with MALDI. These limits, however, may not be a consequence of the ionization process alone, but may also be due to the mass analyzers and detectors used. With design and software modifications to commercial ESI instruments, a few research groups have detected protein complexes as large as 800 kDa (5). Although MALDI is capable of volatilizing analytes as large as 250 kDa, the output is a “humpogram” rather than sharp peaks, both because of the lower high-mass sensitivity and the lower effective resolution (m/Δm) at very high mass. One disadvantage of electrospray is its sensitivity to salts and detergents, and the fact that it is not as “high throughput” as MALDI, where MS spectra of a series of samples can be obtained in only a few seconds per sample.

Because what is measured is the mass-to-charge ratio, it is essential to accurately determine the charge on the peptide. Higher resolution instruments make it possible to accurately determine the charge state of the ions. For example, the separation of peaks in the isotope cluster of a peptide with a +1 charge state are 1 Da apart, while those of a +2 ion are 0.5 Da apart (+3 charge state, 0.33 Da apart), and so on (Figure 5.5). This allows the correct determination of the peptide molecular weight and allows more confident identification of an analyte.

FIGURE 5.5. Different charge states of the same peptide ion (FKDLGEEHFK from BSA), showing different spacing of isotopes.


Different charge states of the same peptide ion (FKDLGEEHFK from BSA), showing different spacing of isotopes.

Not all peptides ionize equally (Figure 5.6A and C), and this ionization efficiency also depends on the ionization mode. On average, it has been found that approximately 25% of peptides ionize exclusively in one mode or the other (Figure 5.7). For projects requiring the highest possible sequence coverage (for example, modification site determination), both techniques are often used.

FIGURE 5.6. BSA digest spectrum in (a) ESI; (b) ESI, deconvoluted to +1 charge state; and (c) MALDI modes.


BSA digest spectrum in (a) ESI; (b) ESI, deconvoluted to +1 charge state; and (c) MALDI modes.

FIGURE 5.7. Selective ionization in (A) ESI (41.


Selective ionization in (A) ESI (41.2% coverage); (B) MALDI (25.2% coverage)—some peptides can only be detected in one mode or the other, while others can be detected in both ionization modes; (C) Venn diagram of peptide ionization in ESI and (more...)


At the same time as these ionization techniques were being developed, there was growing interest in coupling liquid chromatography (LC) with mass spectrometry as an alternative to gas chromatography since many biomolecules decompose when analyzed by gas chromatography. While the advantages of this technique were clear (less decomposition, and the reduction of “suppression effects” by having few types of ions in the source at a time), the challenge of this was enormous because it involves coupling a liquid-based separation technique with a gas phase technique (6). The first method to accomplish this coupling was the “moving belt” interface (7). This was an on-line technique, and the 1-mL/min flow rate, which was common at the time, meant that significant amounts of solvent had to be removed. This was accomplished by depositing the LC effluent on a belt that moved through various vacuum chambers, and the solvent was evaporated before entering the MS source. Inside the source, the analyte was volatilized by flash evaporation and ionized by electron impact or chemical ionization. One problem with this technique was carryover of one sample to the next due to incomplete cleaning of the belt. Interestingly, one early application of this technique was for the analysis of peptides, but only after derivatization (8).

The other early on-line approaches (ca. 1980) were based on the use of a chemical ionization source. The first technique, called direct liquid injection (DLI), relied on splitting the LC effluent so that only a few microliters per minute entered the source (the LC solvent becomes the CI reagent gas) (9). In this technique, the effluent is sprayed through a tapered capillary or 10-μm pinhole in a diaphragm, producing a fine spray of droplets that can be desolvated in the source. This technique led to the interest in the development of microbore liquid chromatography-mass spectrometry (LC-MS) (10), and eventually capillary LC-MS systems, where the entire LC effluent (200 nL/min to 1 μL/min) can enter the CI source without the need for splitting.

Alternately, heat could be used to desolvate a higher flow rate (1 mL/min) of reversed-phase solvent in a technique called “thermospray,” which was perhaps the first easily used LC interface (11). This technique, developed in 1982, used ammonium acetate to provide CI-type ionization, with or without the use of a filament. Although the moving belt interface, the DLI interface, and thermospray are no longer used, these techniques laid the groundwork for the LC-MS methods currently in use today, and provided the impetus for the development of low-flow-rate LC systems which are commonly used in modern LC-MS techniques.

The solution-based technique of ESI was the easiest to couple to mass spectrometry, for the analysis of both proteins and for peptides. MALDI is still most often used for the analysis of single analytes or digests of a single protein. However, LC-MALDI of mixtures would be a very useful technique for mixtures, and although on-line LC-MALDI has been attempted, the more promising approach appears to be off-line LC-MALDI (the basic concept for on-line versus off-line LC methods is discussed in Chapter 3). Various LC-MALDI spotting devices have been developed, using elec-trodeposition or simply depositing droplets of the eluent on a target. Matrix can be added later, or the matrix solution can be added to the eluent before the droplet is formed. This technique can be thought of as a direct descendant of the “moving belt” interface, but since such low flow rates (usually 200 nL/min to 1 μL/min) are used, and spots are collected every 10–15 seconds, no additional vacuum or heating device is needed. Also, the off-line “decoupling” of the MS from the deposition step greatly simplifies the procedure.


5.4.1. Quadrupole Mass Filters

The revolution in biological mass spectrometry was not only due to improvements in ionization techniques, in particular MALDI and ESI, but also from improvements to existing mass analyzers and the design of new and hybrid instruments. Early mass analyzers, such as the quadrupole mass Alter, had a mass range of only ~650 Da, while modern quadrupole instruments can scan up to 4000 Da. Also, while earlier quadrupole mass spectrometers were capable of only unit resolution (which means that they could distinguish ions 1 Da apart), new advances and the coupling of quadrupole and time-of-flight (qQTOF) analyzers made it possible to achieve higher resolution, up to 0.01 Da in these hybrid instruments.

In quadrupole mass analyzers (otherwise known as quadrupole mass Alters), a combination of RF and DC voltages on the four parallel rods allows only ions with specific m/z to pass through to the detector (Figure 5.8). The quadrupole can scan a specified m/z range by ramping the voltages to produce MS spectra, or they can be set to allow the passage of only preselected masses (for peptide sequencing [MS/ MS], or selected ion monitoring).

FIGURE 5.8. Diagram of single quadrupole mass analyzer.


Diagram of single quadrupole mass analyzer.

5.4.2. Time-of-Flight Analyzers

With time-of-flight (TOF) instruments, the mass-to-charge ratio is indirectly calculated from the length of time it takes for the ion to reach the detector in a field-free vacuum. “Packets” of ions are released into the TOF analyzer at time = 0, and then ions drift toward the detector and generate a signal at t = final. The mass of the analyte is proportional to the flight time (measured in nanoseconds) and heavier ions travel more slowly, taking more time to reach the detector.

The first TOF design was the linear TOF in which the ions traveled a linear path from the opening or orifice on the mass analyzer to the detector (Figure 5.9, top). However, the nature of MALDI ionization produces a plume of ions that may experience slightly different acceleration voltages depending on where they were formed in the “plume” prior to reaching the field-free vacuum. These ions may therefore reach the detector at slightly different times. This translates into broader peaks and reduces resolution. TOF analyzers were greatly improved by two modifications: “delayed extraction” of ions leaving the source, and the addition of a set of reflectron lenses to the standard linear design. The increasing voltages applied to these successive ring electrodes “reflect” ions to a second detector. Since ions with higher kinetic energy penetrate “deeper” into the reflectron, and thus have to travel farther, this helps correct for the variability in ion kinetic energies, resulting in sharper, better resolved peaks (Figure 5.9, bottom).

FIGURE 5.9. Diagram of TOF analyzers in (A) linear and (B) reflection configurations.


Diagram of TOF analyzers in (A) linear and (B) reflection configurations.

5.4.3. Ion Traps

During the 1950s, Paul and Steinwedel patented a novel alternation to the quadrupole mass analyzer (12,13). The principles of mass filtering were similar in that RF and DC voltages were used to filter out masses. However the physical design was quite different (Figure 5.10). Instead of using four rods arranged in parallel the new design had only three electrodes. The hyperbolically shaped entrance and exit electrodes allowed selective flow of ions to and from the mass analyzer, and a third electrode between them—the ring electrode—was used to trap ions along this path. The design effectively acts as a three-dimensional quadrupole. Ion traps normally have unit mass resolution.

FIGURE 5.10. Diagram of an ion trap.


Diagram of an ion trap.

5.4.4. Fourier Transform MS

A type of mass spectrometer less commonly used for proteomics, but which has the highest mass resolution, is the Fourier transform (FT) MS. These are the most expensive instruments and are available in two types: FT-ICR and the Orbitrap.

FT-ICR (Fourier transform ion cyclotron resonance) MS was first described in the late 1940s by Hipple, Thomas, and Sommer (14,15), but only became practical in the 1970s when multiple ions could be detected (16). The principal components of ICR mass spectrometers are an ICR cell, which typically consists of pairs of plates arranged as a cube or capped cylinder, and a surrounding strong magnetic field (Figure 5.11). ICR-MS requires ion trapping, excitation, detection, and data transformation.

FIGURE 5.11. Diagram of an FT-ICR.


Diagram of an FT-ICR.

Ions generated in an external source are directed into the ICR cell made of two trapping plates, two excitation plates, and two detector or receiver plates. The ions are trapped in the cell by an electric field applied to the trapping electrodes and by the magnetic field (7–18 tesla). Within the cell the trapped ions travel in a circular orbit (cyclotron motion) with orbital frequencies inversely proportional to the ions' mass-to-charge ratio. An RF sweep is then applied across the excitation plates. Ions with cyclotron frequencies in resonance with the applied RF are accelerated and increase their orbit radius bringing them closer to detector plates, which then record the image current. The complex image current is deconvoluted to the frequency domain by fast Fourier transformation from which the mass-to-charge ratio is then determined.

By far FT-ICR instruments are the most expensive primarily due to the cost of the superconducting magnet commercially available in field strengths of 7 tesla ($800,000) and 15 tesla for $2.4 million. However, the cost may be justified for certain projects that require the increased sensitivity due to the trapping nature of the technology, the high mass accuracy (up to 0.5 ppm), and the high resolution (100,000 full-width half-mass).

In 2005 Alexander Makarov developed the newest type of mass analyzer to use FT technology, having properties similar to ion traps but with results approaching those of FT-ICR instruments (17). In an Orbitrap, ions enter the trap in a tangent to an inner axial electrode. The voltage of the axial electrode is increased as the ion enters, causing it to spiral. The spiral moves toward the center of the axis then the voltage increase stops and the spiral becomes a ring of ions (Figure 5.12). The rings of ions cycle at different frequencies and the frequencies are converted to ion masses by Fourier transformation of the ion current.

FIGURE 5.12. Diagram of the Orbitrap.


Diagram of the Orbitrap.

5.4.5. Ion-Mobility MS

Ion-mobility mass spectrometry is a technology which separates ions based on their collisional cross sections (18). A new commercial instrument based on this technology has recently been introduced by Waters Corporation. A “wave guide” was developed to replace the original RF-only quadrupole-based collision cell, which had been originally developed by ABI. This new wave guide was observed to have the property that it could be used to separate ions based on their shapes instead of to refocus them. This technology is so new that it is difficult to predict its impact on proteomics.


The earliest mass spectrometers used photoplate detection, hence the old term “mass spectroscopy,” which is no longer appropriate. In the 1920s, multi-stage analyzers were developed, which relied on an electron “cascade” or “avalanche” to provide amplification of the ion signals. Early designs were a kind of “Venetian blind” device with each plate at an increasing voltage compared to the previous level. In the 1980s, this design was replaced with a “continuous dynode,” which replaced the plates with a cone-shaped device. Typically, analyte ions strike the detector surface, which releases electrons from the surface. The released electrons in turn strike the surface, emitting more electrons. Each of the newly emitted electrons repeats the process so that a final stream of electrons strikes the cathode (Figure 5.13). For negative ion detection, a “conversion dynode” is placed above the entrance to the multiplier cone. The conversion dynode “converts” the ion polarity of the negative ion. The incoming negative ion strikes the surface of the conversion dynode and produces a positive ion, which then initiates the electron cascade within the electron multiplier. The voltage of the conversion dynode is the same (negative) for both positive and negative ion detection modes. Most TOF instruments now use multichannel plate (MCP) detectors, which are similar in concept to the original electron multipliers but consist of several multipliers in an array (Figure 5.14). These devices are so efficient that they are capable of detecting single ions ("ion counting").

FIGURE 5.13. Continuous dynode detector.


Continuous dynode detector.

FIGURE 5.14. Diagram of multichannel plate detector.


Diagram of multichannel plate detector.

With the FT-based instruments, the ICR and the Orbitrap, the ion current generated from the orbiting ions is detected. In FT-ICR, it is measured by the trapping electrodes. With Orbitrap instruments, split outer electrodes flanking the center of the axial electrode measure the current of the ions oscillating along the axial electrode. With both ICR and Orbitrap instruments, the detected signals are then typically amplified prior to processing by fast Fourier transformation.


Mass spectrometers can be built with a single mass analyzer (single-stage) or can be built with multiple analyzers (multi-stage). The single-stage instruments simply measure the mass of the ion generated by the ion source. Multi-stage instruments can consist of the same mass analyzers such as a QQQ (triple-quadrupole, Figure 5.15) or dissimilar analyzers such as the ion-trap TOF (IT-TOF) or quadrupole-quadrupole/collision cell-TOF (QqTOF). These instruments are very powerful and can manipulate the ions to increase sensitivity, fragment selected ions for sequence and modification information, and provide better mass resolution. For example, single-stage ion traps normally have unit mass resolution. Hybrid QqTOF instruments normally have mass resolution (m/Δm) of 100–200 ppm.

FIGURE 5.15. Diagram of triple quadrupole mass spectrometer.


Diagram of triple quadrupole mass spectrometer.


If enough internal energy (20–30 eV) is added to a peptide, it will fragment. We are indeed fortunate that normally this fragmentation occurs along the peptide backbone, although occasionally this fragmentation occurs in the ion source (insource fragmentation, “skimmer-induced” fragmentation, or “prompt” fragmentation) (19–23). Fragmentation typically occurs in a cell and is induced or activated by colliding the analyte with inert gas molecules such as nitrogen or air. Thus, gasphase sequencing of peptides is usually done by performing MS/MS—two stages of mass spectrometry surrounding a collision cell. This technique, also called tandem mass spectrometry, involves selection of an ion with the first MS (usually a quadrupole mass filter), fragmenting the peptide ions in a collision cell containing an RF-only quadrupole, and finally detecting the fragments of the selected peptide.

Fragmentation resulting from collision of the analyte with neutral gas is termed collision-induced dissociation (CID). It is a low-energy process, typically resulting in cleavage of the amide bond to form “b” or “y”-type fragment ions, which are most useful for determining the peptide sequence. The fragment ions produced, which contain the N-terminus, are called b ions; those containing the C-terminus are called y-ions (Roepstorff nomenclature) (24) (Figure 5.16).

FIGURE 5.16. MS/MS fragmentation of peptides; nomenclature and schematic of “b” an “y” fragmentation.


MS/MS fragmentation of peptides; nomenclature and schematic of “b” an “y” fragmentation.

Fragmentation of a peptide is size dependent since the energy of the collision is absorbed by all the bonds of the peptide. Thus, in practice, if a peptide is larger than 2500 Da, it is difficult to get enough energy into a single bond to cause it to break. For this reason, although MALDI-TOF mass spectrometers are capable of a high mass range, some MALDI TOF/TOF instruments (for example, the 4700 Proteomics Analyzer), which are designed specifically for fragmenting peptides, have effective mass ranges of only 2500 Da.


Three fragmentation techniques, electron capture dissociation (ECD), infrared multiphoton dissociation (IRMPD), and electron-transfer dissociation (ETD), were recently introduced as means of fragmenting larger peptides and even proteins with mass spectrometry. In ECD, low-energy electrons are captured by peptide cations to form odd-electron peptides which then dissociate to fragments primarily of the c-and z-type (25) (Figure 5.17). Unlike CID processes, ECD cleavage does not increase the analyte’s internal vibrational energy. Therefore the weakest bonds, such as labile phosphate bonds, are maintained. Similar fragments are formed with ETD, but due to the interaction of the protonated peptide with radical anions rather than with a beam of electrons (26), fragments of b- and y-type are produced by IRMPD when the peptide absorbs energy from IR photons.

FIGURE 5.17. Top-down fragmentation: MS/MS spectrum of 115 residue protein obtained by ECD on 12T FT-ICR.


Top-down fragmentation: MS/MS spectrum of 115 residue protein obtained by ECD on 12T FT-ICR. Source: Reprinted from Borchers, C. H., Thapar, R., Petrotchenko, E. V. et al. (2006) Combined top-down and bottom-up proteomics identifies a phosphorylation (more...)

ECD and IRMPD are used only on FT instruments. Both of these are used primarily for “top-down” sequencing of proteins, which means that these proteins are fragmented and the molecular weights of the pieces are used to determine the sequence of the protein. This is in contrast to the “bottom-up” approach, which first uses enzymes to cleave the proteins into peptides followed by gasphase sequencing. ETD is more flexible in that it was first demonstrated in an ion trap but can be used in FT instruments as well.


For the identification of proteins in gel spots or bands, the gel plugs or slices are usually “dehydrated” by soaking them in acetonitrile, and then they are “rehydrated” with a solution containing trypsin. The proteins are digested into peptides, and the peptides are extracted, lyophilized, and analyzed by MALDI-TOF/TOF. The peptide masses and their partial sequences are compared with the theoretical masses in a database, and the protein is identified. MALDI is the method of choice for these analyses because of the high throughput. Each gel spot ends up as a single MALDI spot on the MALDI target. MALDI is a high-throughput technique, since no separation step is involved, and there is also no possibility of “carryover” of one sample to another.

In our laboratory, ESI is the method of choice for high-sensitivity peptide analysis and high mass-accuracy molecular weight determination. LC-ESI-MS/MS and LC-MALDI-MS/MS are used for modification site determination where high sequence coverage must be obtained. Modification site determination will be discussed in Chapter 6.


Although the advances in instrumentation described above are clearly critical for the development of MS-based proteomics, proteomics is also dependent on advances in other fields. Genomes had to be sequenced before they could be used to predict protein and peptide sequences. Software packages had to be written to compare the observed peptide molecular weights and sequences with those in the databases. Computers and storage devices need to be developed that can handle the acquisition rates and huge data files (often several gigabytes per analysis) resulting from these analyses. Bioinformatics techniques capable of interpreting the enormous amounts of data generated are also being developed.


Proteomics is a new field, and all of its components are constantly being improved. New mass spectrometers are still being developed, and existing designs continue to be improved by making them more sensitive, and by increasing their mass resolution, mass accuracy, stability, and scan speed. New genomes are being sequenced, and new software packages are being developed to provide protein identifications with lower false-positive and false-negative rates. Computer systems and data storage systems capable of handling terabytes of data—which would have been prohibitively expensive only a few years ago—are becoming common. Thus, this chapter can at best only be an overview of the history and evolution of a rapidly developing field.


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Copyright © 2010 by Taylor and Francis Group, LLC.
Bookshelf ID: NBK56011PMID: 21882443


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