Van Eyk, Dunn - Proteomic and Genomic Analysis of Cardiovascular Disease - 2003 (522919), страница 52
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In parallel, equal concentrations ofboth control and diseased samples arepooled in one tube and labeled with Cy2.Both the mixture of Cy3 and Cy5 labeled proteins and the Cy2 labeled pooled sample areseparated on the same gel. The Cy2 labeledpooled sample is used for normalisation purposes. Each 2DE protein profile associatedwith each individual dye is visualized at a specific wavelength.18118211 Proteomics, a Step Beyond Genomicsmethods with similar detection sensitivities such as silver (>2 hours) and SyproRuby (> 3 hours). The main advantage to DIGE is the ability to compare two samples on the same gel. This reduces inter-gel variability, decreases the number ofgels required by 50% and allows for more accurate and rapid analysis of differences.
Improved accuracy of quantitation, for comparison of multiple samples,can be achieved using a pooled internal standard containing all experimental samples labeled with a third dye Cy2. This internal standard is included with experimental samples run on each gel (i.e. those labeled with Cy3 and Cy5 dyes)(Fig. 11.4) [35, 36].DIGE does not interfere with subsequent analysis by mass spectrometry, however there are some disadvantages associated with DIGE.
DIGE relies on the fluorescence of the dye for quantitation. Therefore carefully controlled labeling is critical for accurate quantitation. The number of lysine residues contained within aparticular protein will dictate the labeling efficiency of that protein. Proteins witha high percentage of lysine residues could be labeled more efficiently than proteins with little or no lysine.
Hence, DIGE cannot detect proteins without lysine.Therefore a protein spot can appear to be highly abundant when silver stainedbut appear significantly reduced when using DIGE.DIGE can affect protein spot patterns compared to patterns obtained with conventional systems. Proteins are physically labeled i.e. Cy3 and Cy5 molecules arecovalently linked to lysine residues. This can alter the location of a protein spotwithin a gel slightly.
The dyes used do not change the pI values of proteins because the dye molecules are positively charged which negates the loss of thecharge on the lysine group. However, the dye molecule has a molecular weight of0.5 kDa. An additional 0.5 kDa will not significantly affect the migration of largeproteins because only a small percentage of the protein becomes labeled. However this small increase in molecular weight becomes more evident as the molecular weight of proteins decreases.The labeling reaction is carried out at low efficiency (protein:dye, 5 : 1) such thatat most only one lysine residue per protein molecule is conjugated with dye. Thisminimizes any change in molecular weight.
Since only a small percentage (typically 2–5%) of a protein becomes labeled with Cy3 or Cy5, most of the proteinmolecules cannot be visualised in the Cy3 and Cy5 images. To locate unlabelledprotein the 2DE gel can be stained post-DIGE using Sypro Ruby [35, 37]. The protein of interest is identified by comparing the Sypro Ruby stained image and theCy3 and Cy5 images.Whilst the various staining methods described are common in most proteomicslaboratories, another technique used to detect proteins is radiolabelling. This toocan also achieve very high sensitivity, although not all proteins can be readilyradiolabelled. The use of phosphorimaging screens has increased the dynamicrange of radiolabelling and has overcome the problem of non-linearity associatedwith X-ray film images of radiolabelled 2DE separations.
The surface of phosphorimaging screens contain a thin layer of BaFbr: Eu2+ in a plastic support. A dried2-D gel is placed in contact with the screen. During this exposure step the b-particles emitted by the radiolabelled proteins pass through the layer converting Eu2+11.5 Protein Identification3+to Eu . After a suitable exposure time the screen is transferred to a phosphorimaging scanner where light from a high intensity HeNe laser (633 nm) is absorbed causing the excited state Eu3+ ions to decay back to the Eu2+ ground stateby the emission of blue (390 nm) luminescence proportional to the amount of radiation incident on the screen. This approach requires relatively short exposuretimes compared to conventional autoradiography, has a high dynamic range andgood linearity of response.
The major disadvantage is the high cost of the phosphorimaging screens and the dedicated phosphorimaging device required. Thehigh cost aspect also applies to using fluorescently labeled proteins since a dedicated fluorescent imager is required for their imaging.11.5Protein IdentificationIn this section we aim to provide the reader with a brief overview of methods andtechniques used to identify and characterize proteins from 2-D gels.
For a morecomprehensive review we refer the reader to Chapters 12 and 13.Although 2DE can effectively separate all the component proteins of a proteome, providing quantitative data, protein identification and function will remainunknown. Conventional methods of identifying proteins from 2-D gels have reliedon Western blotting, microsequencing by automated Edman degradation [38] andamino acid compositional analysis [39]. The disadvantages of these techniques isthat they are very time consuming with limited sensitivity.Over the past few years a variety of sensitive methods, centered around massspectrometry (MS), have become available for the identification and characterization of proteins and peptides (Fig.
11.5). The development of techniques such asmatrix-assisted laser desorption/ionization (MALDI) and electrospray (ESI) ionization methods have brought with them very high sensitivity requiring smallamounts of sample (a sensitivity of femtomole to attomole) and the capacity forhigh sample throughput [40, 41]. Both MALDI and ESI are capable of ionizingvery large molecules with little or no fragmentation. The type of analyzer usedwith each can vary.
MALDI sources are usually coupled to time-of-flight (TOF)analyzers, whilst ESI sources can be coupled to analyzers such as quadropole, iontrap and hybrid quadropole time-of-flight (Q-TOF).Peptide mass fingerprinting (PMF) is the primary tool for MS identification ofproteins in proteomic studies.
Prior to analysis, an excised protein spot is digestedwith a protease, typically trypsin, which cleaves proteins at basic amino acids (arginine/lysine residues), if present, breaking proteins down into a mixture of peptides. The masses of the peptides present can then be measured by mass spectrometry to produce a characteristic mass profile or ‘fingerprint’ of that protein(Fig.
11.6). The mass profile is then compared with peptide masses predictedfrom theoretical digestion of known protein sequences contained within currentdatabases or predicted from nucleotide sequence databases. This approach provesvery effective when trying to identify proteins from species whose genomes are18318411 Proteomics, a Step Beyond GenomicsN-terminal sequencing tag(three residues)Fig. 11.5 Methods used for the identificationand characterization of proteins separted by2DE.
The following abbreviations have beenused: 2DE, two-dimensional electrophoresis;ESI, electrospray ionization; HPLC, high performance liquid chromatography; IR, infrared;Mr, relative molecular mass; MALDI, matrixassisted laser desorption ionization; MS,mass spectrometry; MS/MS tandem massspectrometry; pI, isoelectric point; PSD, postsource decay.completely sequenced, but is not so reliable for organisms whose genomes havenot been completed.
This has been a problem in the past for proteomic studies ofsome animal models of heart disease, for example those involving rats, dogs, pigsand cows. This problem has been shown to be overcome effectively by improvingPMF by adopting an orthogonal approach combined with amino acid compositional analysis [42].However, in some instances it is impossible to assign an unequivocal identity toa protein based on PMF. Amino acid sequence information is then required toconfirm an identity. This can be generated by conventional automated chemicalEdman microsequencing but is now most readily accomplished using tandemmass spectrometry (MS/MS). MS/MS is a two-stage process, either by means ofMALDI-MS with post-source decay (PSD) or ESI-MS/MS triple-quadropole, iontrap or Q-TOF machines, to induce fragmentation of peptide bonds.
Oneapproach, termed peptide sequence tagging, is based on the interpretation of aportion of the ESI-MS/MS or PSD-MALDI-MS fragmentation data to generate ashort partial sequence or ‘tag’. Using the ‘tag’ in combination with the mass ofthe intact parent peptide ion provides significant additional information for the11.6 BioinformaticsFig. 11.6 Peptide mass profiling of a silverstained spot from a 2DE separation of humanheart (ventricle) proteins. A MALDI-TOF massspectrum of tryptic peptides is shown, analyzed using a Micromass Tofspec 2E spectrometer (Manchester, UK) operated in thepositive ion reflectron mode at 20 kV acceler-ating voltage with “time-lag focusing” enabled.
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