Proteomic Analysis of Zymogen Granules

María Gómez-Lázaro; Cornelia Rinn; Miguel Aroso; Francisco Amado; Michael Schrader


Expert Rev Proteomics. 2010;7(5):735-747. 

In This Article

Proteomics of ZG Proteins: Separation, Identification & Quantitation

Introduced by George Scheele, 2D separation of proteins in slab gels using isoelectric focusing (IEF) in the first dimension and SDS gel electrophoresis in the second dimension[50] was used for the analysis of the complex mixture of soluble proteins secreted by the exocrine pancreas. It allowed, for the first time, the simultaneous analysis of the major exportable proteins in subcellular fractions obtained after homogenization and in extracellular fluids (e.g., pancreatic juice [PJ] or incubation medium) and contributed to the determination of the intracellular transport of pulse-labeled proteins from one intracellular compartment to another along the secretory pathway.[1] In combination with the development of pancreatic lobules[51,52] as an in vitro model of biological response and, thus, the efficient study of protein synthesis, 2D-SDS-PAGE has furthermore provided valuable insights into the differential regulation of gene expression in the rat pancreas in response to hormones[53,54] and different nutrients.[55,56] In addition, 2D-SDS-PAGE has been used to identify many of the abundant ZG proteins.[8,57,58] In the 1980s, guinea pig and human exocrine pancreatic proteins were the most thoroughly described systems analyzed by 2D-SDS-PAGE. Owing to limited resolution and sensitivity, however, a relatively simple protein composition for the ZGM was proposed. Immunoblotting and immunocytochemistry studies led to the identification of some low-abundance ZGM proteins (e.g., Rab3D or SNARE proteins).[6,21,59] The recent improvements in organellar proteomics revealed a more-complex picture of the components of the ZGM.[34,35,60]

Alternative procedures besides the traditional 2D-SDS-PAGE, such as double SDS-PAGE, have demonstrated their capacity to resolve membrane proteins,[61] but the main developments were due to the application of nongel-based techniques, such as high-performance LC (HPLC) with improved capacity to resolve membrane and membrane-associated proteins, mostly applied after in-solution digestion.[34,35,60] For 1D-LC separation, the most widely used technique is a reversed-phase (RP)-LC based on hydrophobic interactions with the column (e.g., C4, C8 and C18). The need to improve the separation step led to the combination of more than one chromatographic method (2D-LC), with the combination of a strong cation-exchange (SCX) chromatography (separation by positive charge) followed by RP-LC representing the most-common method.[62,63] The combination of 2D-SDS-PAGE and 2D-LC allowed, for the first time, the separation and subsequent identification of a total of 101 proteins from purified ZGM by MS.[34] Direct comparison of the gel-based and LC-based approaches demonstrates the higher capacity of the LC-method to resolve membrane proteins, but also indicates that both methods are complementary. In another proteomics approach, the separation of ZG proteins was achieved by combining 1D-SDS-PAGE with RP-LC, allowing the identification of a total of 371 proteins by MS.[35]

The technical innovations in MS, protein-identification methods and database annotation over the last 20 years have allowed scientists to rapidly and systematically detect thousands of proteins in complex biological samples.[64,65] In this respect, MS has become the method of choice for the analysis of complex protein samples,[64,66] where the development of 'soft' ionization methods is the main method responsible for its success in the proteomics field. Presently, two ionization techniques, namely electrospray ionization (ESI)[67] and MALDI,[68,69] are playing a significant role in MS-based peptide analysis.[65,70] ESI ionizes molecules directly from solution and is therefore easily interfaced with liquid-based separation methods. MALDI ionizes samples that are co-crystallized with an organic matrix via laser pulses and is more often used to analyze relatively simple (but also complex) peptide mixtures.[64,71] After the ionization of the sample (proteins/peptides), the next key step is the determination of their m:z ratios by a mass analyzer. In proteomics research, there are now five basic types of mass analyzer in use: ion trap, time-of-flight (TOF), quadrupole (Q), Fourier transform ion cyclotron resonance (FTICR) and Orbitrap. The most relevant parameters of the analyzers are the sensitivity, resolution, mass accuracy and the possibility to fragment peptide ions (MS/MS).[64,65,71,72]

Until recently, the major ZG proteins have been identified by the purification of proteins, N-terminal peptide sequencing techniques and the specific immunolocalization of granule components. MS was only recently applied to the proteomic study of ZGs.[73] The introduction of MALDI-TOF/TOF, MALDI-TOF or ESI-Q/TOF in this field resulted in a better characterization of the ZG proteome, and contributed mainly to the discovery of additional ZGM proteins.[31,34,35] Valuable progress has been made in the identification of proteins involved in regulated exocytosis and ZG trafficking (e.g., Rab proteins and effectors, and SNARE proteins) as well as membrane channels and transporters. In addition, new enzymes have also been identified (Table 1).

To gain new insights regarding fundamental biological questions, accurate protein quantification is also required[74] (see also the 'Proteomics of ZG proteins in pancreatic juice' section). Scheele and co-workers searched early on for methods to quantify the discharge of 'individual' secretory proteins separated by 2D-SDS-PAGE and developed a 2D scanning approach of Coomassie Blue-stained gels and computer analysis of the scanning data.[58] MS can also be applied for protein quantification, but the quantity of analyte is not directly related to the ion-current intensity of the MS signal. To enable differential quantitation of proteins with MS, an additional processing of the sample is necessary, that most of the time involves the labeling of peptides with stable isotopes by biosynthetic or chemical methods.[71] To date, three different ways of stable isotope labeling of proteins/peptides are used: chemical,[75–78] metabolic[79,80] or enzymatic labeling.[74,81] The most popular metabolic method is stable isotope labeling by amino acids in cell culture (SILAC). For chemical labeling, the isotope-coded affinity tag (ICAT), the iTRAQ and the isotope-coded protein label (ICPL) are used, whereas enzymatic labeling is achieved by trypsin-catalyzed 18O-labeling. Alternatively, differential quantitation of proteins can be achieved by MS-based label-free quantification. With label-free methods, protein quantification is generally based on two categories of measurement: ion-intensity changes, such as peptide peak areas or peak heights in chromatography,[82,83] or spectral counting of identified proteins after MS/MS analysis.[63,84,85] To our knowledge, so far, just iTRAQ has been applied to the quantification of ZGM, first as a quantitative measurement of the enrichment of ZGM proteins through purification (see 'Isolation and subfractionation of ZGs') and later by combining a global protease protection assay with iTRAQ-based quantification and a statistical model for the assignment of membrane protein topology.[60,86] The tryptic peptides of ZGM proteins were separated into two clusters according to the iTRAQ ratios. The first cluster (with a lower ratio) included cytoplasm-orientated membrane and membrane-associated proteins, whereas the second one (with an unchanged ratio) included predominantly luminal proteins. This approach allowed the mapping of protein domains with cytoplasmic or luminal orientation from the same transmembrane domain.