MS basics

Despite the significant achievements in the MS-based identification of intact proteins (top-down proteomics) ²²⁶, the vast majority of proteomic data has been generated by bottom-up proteomics. The general workflow consists of several steps (Figure 12):

! Sample preparation

Proteins must be extracted from the tissue samples by lysis and solubilization. This is not a requirement for biofluids such as plasma or urine because proteins are already soluble, but purification and concentration steps might be necessary in those cases. Proteins are then denatured by heat or by using denaturation reagents such as urea, the disulfide bonds are reduced and the free cysteins are alkylated in order to break the tridimensional structure of proteins and thus allow for a more efficient proteolysis by providing the proteolytic enzymes maximum access to cleavage sites within the proteins. Finally, proteins are cleaved into peptides by a protease or a mixture of proteases with high specificity ^227,228. Trypsin is the most widely used protease and it specifically cleaves peptide chains at the carboxyl side of

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the amino acids lysine (K) and arginine (R), generating peptides of an average size of 14 amino acids with a predictable MS fragmentation pattern ²²⁹. Tryptic peptides are also convenient to separate by reverse phase liquid chromatography, a separation technique fully compatible with MS analysis. Apart from trypsin, other proteases such as Lys-C or a combination of multiple enzymes can be used in order to increase protein sequence coverage ²²⁹.

Figure 12. Bottom-up MS-based proteomics workflow. Proteins are digested with trypsin and resulting peptides are analyzed by reverse phase liquid chromatography coupled with mass spectrometry detection.

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MS-based approaches achieve a good sensitivity, but the depletion of the most abundant proteins or, conversely, an enrichment of the proteins of interest can be used to improve the detection limit in clinical samples. A number of methodologies have been proposed, such as depletion of the most abundant proteins (e.g., albumin and immunoglobulin G (IgG) in plasma), glycoprotein enrichment, capture of peptides with infrequent amino-acids, or immunoaffinity purification (to be able to detect proteins at or below the ng/ml concentration level) ²³⁰.

! Separation prior to MS analysis

Due to the complexity of the proteome, the identification and quantification of peptides by MS is less efficient if the sample is not previously fractionated or separated ²³¹. The concomitance of too many compounds can generate a competition for the ionization in the source and can lead to signal suppression (i.e., matrix effect) ²³². In addition, prior sample separation reduces the risk of interferences caused by isobaric compounds. Therefore, separation techniques are employed to improve the analytical performance including the sensitivity and the coverage of the proteome. In the early stages of proteomics, two dimensional gel electrophoresis was the method of choice for the separation of proteins from complex samples prior to digestion and identification by MS (see section 5.1.3.1., page 50).

Nowadays, this approach has been mostly replaced by the separation of peptides by reverse phase high pressure liquid chromatography (HPLC) that separates peptides on the basis of their hydrophobic interactions with the analytical column ²²⁷. In this configuration, the outlet of the HPLC system is connected to a mass spectrometer (LC-MS) or several analyzers (LC-MS/MS) where the eluted peptides are directly analyzed. Off-gel fractionation is an alternative technique that fractionates proteins and/or peptides on the basis of their isoelectric point without the need of a gel-based matrix ^233,234. The fractions are further treated as regular proteomic samples and analyzed by LC-MS for an improved proteome coverage.

! MS analysis

Mass spectrometers measure the ratios between the masses and the charges of ions (m/z values). Therefore, peptides must be first ionized and vaporized by the ion source. The resultant ions are then separated according to their m/z by the mass analyzer and finally detected to generate a "mass spectrum", which is a plot of ions abundance against m/z values ²³⁵ (Figure 12). In the field of proteomics, the ionization process is mostly performed in positive mode, by addition of protons. The electrospray ionization (ESI) is well adapted for the analysis of peptides and proteins as it can be directly coupled with a liquid chromatography device and it generates multicharged peptides which promote efficient

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fragmentation at low energy collision induced dissociation (CID). After being ionized and converted to gas phase, the peptide ions are analyzed by one or several mass analyzers, depending on the mass spectrometer configuration. These include low resolution analyzers like the quadrupole and the ion trap, and high resolution/accurate mass analyzers like the time-of-flight (TOF), the orbitrap and the Fourier transform ion cyclotron resonance (FTICR) analyzers ^231,236. Nowadays, proteomic studies employ tandem mass spectrometers which are built by a combination of several analyzers and hence, both LC-MS and LC-MS/MS terminologies usually refer to a combination of mass analyzers. Common tandem mass spectrometers configurations for proteomics studies are the quadrupole time of flight (QqTOF), the quadrupole orbitrap (Q-OT), the triple quadrupole (QqQ) and the ion trap / orbitrap, with different performances in terms of mass accuracy, resolving power, sensitivity and dynamic range ²³⁶. In these configurations, peptides are selected in the first mass analyzer, fragmented by CID ²³⁷, and the resulting fragment ions are analyzed in the second mass analyzer. Fragmentation of peptides generates b- and y- fragment ions that allow to determine the peptide sequence ²³⁸ which, combined with the accurate mass of the precursor ion, dramatically improve the confidence of the identification (Figure 12).