UNIT 22.1 Overview of Proteome Analysis (David W. Speicher, The Wistar Institute, Philadelphia, Pennsylvania). This unit reviews the new discipline of proteomics, which includes any large-scale protein-based systematic analysis of the proteome or defined sub-proteome from a cell, tissue, or entire organism. Proteomics originated in the mid-1990s due to two key enabling advances, availability of complete genome sequences, and mass spectrometry advances that allowed high-sensitivity identifications of proteins. Proteome analyses can be broadly categorized into three types of studies: quantitative protein profile comparisons, analysis of protein-protein interactions, and compositional analysis of simple proteomes or subproteomes such as organelles or large protein complexes. The complexity of different types of proteomes, the merits of targeted versus global proteome studies, and the advantages of alternative separation and analysis technologies are discussed.

UNIT 22.2 Protein Profiling Using Two-Dimensional Difference Gel Electrophoresis (2D DIGE) (Kathryn S. Lilley, University of Cambridge, Cambridge, United Kingdom). 2D-DIGE relies on pre-electrophoretic labeling of samples with one of three spectrally distinct fluorescent dyes, followed by electrophoresis of all samples in one gel. The dye-labeled samples are then viewed individually by scanning the gel at different wavelengths, which circumvents problems with spot matching between gels. Image analysis programs can then be used to generate volume ratios for each spot, which essentially describe the intensity of a particular spot in each test sample, and thus enable expression differences to be identified and quantified. This unit describes the DIGE procedure in terms of sample preparation from various types of cells, labeling of proteins, and points to consider in the downstream processing of fluorescently labeled samples.

UNIT 22.3 Laser Capture Microdissection for Proteome Analysis (Rachel A. Craven and Rosamonde E. Banks, St. James's University Hospital, Leeds, United Kingdom). Laser capture microdissection is being employed increasingly to isolate specific cell types from tissues, thus overcoming problems of experimental interpretation due to tissue heterogeneity of samples. This unit describes protocols which have been optimized to allow laser capture microdissection of tissues with minimal effect on protein integrity, and their subsequent analysis by techniques including 2D-PAGE, immunoblotting, and SELDI, and discusses the relative merits of this approach.

UNIT 5.17 Use of the Gateway System for Protein Expression in Multiple Hosts (James L. Hartley, SAIC/NCI, Frederick, Maryland). The Gateway cloning method allows a gene to be cloned and subsequently transferred into any vector by in vitro site specific recombination. It does not necessarily follow, however, that a gene can be cloned once and expressed in all the available host-vector combinations with uniformly satisfactory results. This is because different organisms have different mechanisms of translating mRNA into protein, and also because choices always have to be made when designing an expression construct— for example, the presence or absence of a stop codon. This unit reviews Gateway cloning, summarizes aspects of protein expression that limit the universality of the use of one clone in many vectors and hosts, and discusses how conflicts between the structure of a Gateway clone of a gene and the rules of protein expression can be minimized or resolved.

UNIT 21.13 Expression, Purification and Characterization of Caspases (Jean-Bernard Denault and Guy S. Salvesen, The Burnham Institute, La Jolla, California). This unit describes a protocol to obtain milligram amounts of enzymatically active pure recombinant caspases. Specific details for the expression, purification of caspase-3, -6, -7, -8, -9 and -10 are discussed along with strategies to obtain particular forms (e.g., the zymogen) of some of them.

UNIT 13.10 Use of Protein Phosphatase Inhibitors (Shirish Shenolikar, Duke University Medical Center, Durham, North Carolina). Reversal of protein phosphorylations by cellular phosphatases hinders the investigation of many physiological events. Phosphatase inhibitors open new avenues for slowing or preventing the turnover of protein-bound phosphate and facilitate the analysis of cellular phosphoproteins. This unit describes the treatment of cells with cell-permeable compounds that preserve phosphorylation on serine/threonine and tyrosine residues. Protocols describe inhibitors of protein serine/threonine phosphatases, PP1/PP2A and PP2B, respectively. These compounds have provided insights into signaling events where the physiological stimuli are unknown and enhance the actions of known stimuli to facilitate the mapping of signaling pathways. Other inhibitors enter cells poorly and are better used to preserve phosphorylations following cell lysis, fractionation or protein isolation. Additional protocols describe compounds that preserve serine/threonine phosphates, allowing detailed analysis of biochemical/biophysical consequences of protein modifications. Also, an inhibitor of protein tyrosine phosphatases is discussed that is widely used to study events activated by growth factors and cytokines.

UNIT 7.11 Rapid Screening of E. coli Extracts by Heteronuclear NMR (Angela M. Gronenborn, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland). A simple and efficient method for screening of overexpressed proteins in crude E.coli extracts is provided. Structural integrity and/or intermolecular interactions can be assessed by 1H-15N HSQC NMR spectroscopy.

UNIT 20.7 Analytical Ultracentrifugation: Sedimentation Velocity (Walter Stafford,Boston Biomedical Research Institute, Watertown, Massachusetts and Massachusetts General Hospital, Boston, Massachusetts). The analytical ultracentrifuge is a high speed centrifuge with an optical system allowing observation of the concentration of macromolecules as a function of radius and time. In sedimentation velocity experiments, relatively high-speeds are used so that a boundary is formed between the solution of sedimenting macromolecule and the buffer in which it is dissolved. Analysis of the rate boundary movement and evolution of its shape can yield information about the molar masses of species present as well as stoichiometries and equilibrium constants for their interactions. This overview discusses crucial issues pertaining to sedimentation velocity experiments, including noninteracting and interacting systems, ideality and nonideality, and reversible versus kinetically limited equilibrium scenarios.

UNIT 11.8 C-terminal Sequence Analysis (Tomas Bergman, Ella Cederlund, Hans Jörnvall, Karolinska Institutet, Stockholm, Sweden; and Elizabeth Fowler, Millennium Pharmaceuticals, Cambridge, Massachusetts). This unit has been updated to describe performance improvements to standard automated C-terminal sequencer protocols. Polypeptides ranging from 10 to 600 residues have been tested in amounts as low as 10 pmol. Extended runs (up to 15 residues using 100-500 pmol), detection of Pro and phenylisocyanate PTC-Lys, and efficient combination of N- and C-terminal degradation are possible. The technique provides quantitative characterization of C-terminal sequences and truncation patterns, and is efficient for analysis of N-terminally blocked polypeptides. Two methods of manual C-terminal sequence determination are also presented that employ one or more carboxypeptidases. One method involves mass spectrometry to analyze the set of peptides generated by sequential removal of C-terminal amino acids, where upon the sequence is deduced from the mass differences between adjacent peaks in the mass profile. A second manual method entails peptide digestion, separation of the released amino acids from the protein, and finally amino acid analysis.

UNIT 6.1 Overview of the Purification of Recombinant Proteins Produced in Escherichia coli (Paul T. Wingfield, National Institute of Arthritis and Musculoskeletal & Skin Diseases, Bethesda, Maryland). The updated version of this unit presents an overview of recombinant protein purification with special emphasis on proteins expressed in E. coli. The first section deals with information pertinent to protein purification that can be derived from translation of the cDNA sequence. This is followed by a brief discussion of some of the common problems associated with bacterial protein expression. A flow chart is included to summarize approaches for establishing solubility and localization of bacterially produced proteins. Purification strategies for both soluble and insoluble proteins are also reviewed and summarized in flow charts. A section on glycoproteins produced in bacteria in the nonglycosylated state is included to emphasize that, although they may not be useful for in vivo studies, such proteins are well suited for structural studies. The final sections deal with protein handling, scale and aims of purification, and specialized equipment needed for recombinant protein purification and characterization. The methodologies and approaches described here are essentially suitable for laboratory-scale operations.

UNIT 1.5 Overview of the Physical State of Proteins Within Cells (Howard R. Petty, Wayne State University, Detroit, Michigan). Proteins, the end product of gene expression, play a pivotal role in cellular structure and function. To understand how proteins work it is necessary to understand their physical state within the cell. This unit reviews the classification of proteins, how that is related to the hydropathicity of the protein, other factors that affect the heterogeneity of proteins, protein assemblies, methods for altering the solubility of proteins, and limitations of in vitro manipulations of proteins.

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UNIT 21.16 Monitoring Peptidase Activity Using Synthetic Fluorogenic Substrates (Linda Troeberg and Hideaki Nagase, Imperial College London, United Kingdom). Fluorogenic synthetic substrates are commonly used to monitor the activity of peptidases in vitro. This unit presents a representative protocol that employs (7-methoxycoumarin-4-yl)acetyl-Pro-Leu-Gly~Leu-(3-[2,4-dinitrophenyl]-L-2,3-diaminopropionyl)-Ala-Arg-NH2 (Mca-Pro-Leu-Gly~Leu-Dpa-Ala-Arg-NH2) as a substrate to assay matrix metallopeptidases (MMPs). This substrate was first described for the assay of MMP-1, -2 and -3 and it is now widely used as a general MMP substrate. Protocols are given for both stopped-time assays (suitable for assaying MMP activity in a large number of samples) and continuous assays (commonly used when establishing an assay protocol or investigating kinetic aspects of enzyme behavior). Other fluorogenic peptides and protein substrates, together with nonfluorogenic alternatives, are also discussed.

UNIT 22.4 Preparing Protein Extracts For Quantitative 2-D Gel Comparison (Mireille Chevallet, Christophe Tastet, Sylvie Luche and Thierry Rabilloud, DRDC/BECP, Commission de L'Energie Atomique, Grenoble, France). This unit describes the basic protocols needed for efficient and reproducible protein solubilization from a variety of biological samples, including cultured animal cells, animal tissues, plasma and serum, cell nuclei, other subcellular organelles, bacteria, plant tissues and cells, and other biological fluids. The optimized extraction process is strongly sample-dependent and cannot be described for every type of sample. However, typical protocols are provided as general guidelines and illustrate good starting points for sample preparation optimization. These solubilization procedures take into account the constraints introduced by two-dimensional electrophoresis and are thus well suited for proteomics approaches using this technique.

UNIT 21.14 Aspartic Peptidases (Bret B. Beyer, Nathan E. Goldfarb, and Ben M. Dunn, University of Florida, Gainesville, Florida). The unit describes a basic protocols utilized to obtain milligram amounts of enzymatically active, pure recombinant Plasmodium plasmepsins and "short" human pseudocathepsin D. Specific details for the expression and purification of Plasmodium falciparum plasmepsin 2 and "short" human pseudocathepsin D in zymogen form are described in this chapter. The plasmepsin 2 protocols are also applicable to Plasmodium vivax, P. ovale, and P. malariae plasmepsins, as well as P. falciparum plasmepsin 4.

UNIT 21.15 Zymography of Peptidases (Linda Troeberg and Hideaki Nagase, Imperial College London, United Kingdom). Zymography is an electrophoretic technique enabling visualization of the number and approximate size of peptidases in a sample on the basis of their hydrolysis of a protein substrate within the gel. The technique is particularly useful for analyzing the peptidase composition of complex biological samples because visualization depends directly on proteolytic activity. This unit presents a representative zymography protocol for the study of matrix metallopeptidases (MMPs).

UNIT 22.5 Isolation of Organelles and Prefractionation of Protein Extracts Using Free-Flow Electrophoresis (Peter J. A. Weber, Gerhard Weber, Christoph Eckerskorn, Tecan Munich GmbH, Germany). One of the major obstacles in the analysis of proteomes is the extreme complexity of any given cell or biological fluid. Free-flow electrophoresis (FFE) is a powerful tool for the reduction of this complexity, which is a prerequisite for systematic and comprehensive protein analyses. This unit describes methods for partial purification of complex mixtures at two different stages: on the protein level by isoelectric focusing FFE-fractionation of crude protein mixtures, e.g. whole-cell lysates, and on a subcellular level by zone electrophoretical FFE-purification of organelles.

UNIT 23.2 Proteomics Using 2-D Liquid Separations of Intact Proteins From Whole-Cell Lysates (Kan Zhu, Fang Yan, Kimberly A. O’Neil, Rick L. Hamler, University of Michigan, Ann Arbor, Michigan; Linda Lin and Timothy J. Barder, Eprogen, Darien, Illinois; and David M. Lubman, University of Michigan, Ann Arbor, Michigan). Protocols for protein isoelectric point (pI) fractionation in the first dimension include the use of liquid isoelectric focusing (IEF) and chromatofocusing. Liquid IEF provides a pI-based fractionation using a batch-phase electrophoretic method while chromatofocusing uses a column-based chromatographic method to generate the pH gradient. Using either method, a second-dimension fractionation is performed in the liquid phase using nonporous silica reversed-phase HPLC (NPS-RP-HPLC) to generate a 2-D liquid map of the protein content of the cell. Procedures for HPLC separation of the pI fractions of intact proteins are also described. The eluent of the 2-D liquid fractionation is directly coupled to a mass spectrometer for on-line detection of the intact molecular weights of proteins. As a result, a multidimensional map of protein expression is obtained that characterizes cellular proteins by pI, hydrophobicity, and intact molecular weight. Such expression maps are useful for differential proteomic comparison between cell samples.

UNIT 23.3 Quantitative Protein Analysis Using Proteolytic H2[18O] Labeling (Kristy J. Reynolds and Catherine Fenselau, University of Maryland, College Park, Maryland). This unit describes the procedure for proteolytic H2[18O] labeling of peptides in order to quantitate relative protein levels for a comparative proteomic experiment.

UNIT 23.4 Quantitative Protein Profile Comparisons Using ICAT and Related Methods (Eugene C. Yi and David R. Goodlett, Institute for Systems Biology, Seattle, Washington). Current methods for measuring pair-wise changes in protein expression involve differential stable isotopic labeling of proteins or peptides either in vivo or in vitro followed by identification and quantification using a mass spectrometer. In comparison to the methods for measuring changes in mRNA expression that observe two colors, the mass spectrometer observes two different masses. Changes in protein expression are observed when the identical peptide from each of two biological conditions is identified and subsequent area under the curve measurements comparing the peptide labeled with "heavy" isotopes to the one with a "normal" isotopic distribution of chemical elements show a difference. These approaches provide a means to measure the changes in expression of many proteins simultaneously between two different biological states; e.g. yeast grown on galactose versus glucose or normal versus cancer. This unit describes one of these popular methods for quantitative protein profiling, the Isotope Coded Affinity Tag (ICAT).

UNIT 4.3 Subcellular Fractionation of Tissue Culture Cells (Fernando Aniento, University of Valencia, Valencia, Spain; and Jean Gruenberg, University of Geneva Sciences II, Geneva, Switzerland). Cell fractionation techniques include some of the most important and widely used analytical tools in cell and molecular biology, and are essential for the development of cell-free assays that reconstitute complicated cellular processes. In addition to simple gradient systems, this unit discusses the immunopurification of organelles, in particular endosomes. As antigens, purification can be achieved using endogenous or ectopically expressed proteins, provided that appropriate antibodies are available. Alternatively, tagged proteins can be used, when combined with anti-tag antibodies. Now that sequencing of the genomes of several organisms has been completed, biochemical strategies, and in particular fractionation and in vitro transport assays, are more necessary than ever for studying the numerous protein and protein complexes that are being discovered.