Lysing Yeast Cells with Glass Beads for Immunoprecipitation.

Yeast cells display cell walls that must first be broken before the addition of detergents for lysis. This method describes the use of glass beads in combination with a mechanical bead beater to disrupt cell walls of both Saccharomyces cerevisiae or Schizosaccharomyces pombe directly in a nonionic detergent Lysis buffer containing 0.1% Nonidet P-40. Alternatively, this protocol can be applied for the lysis of yeast cells in Lysis buffer without detergent; upon completion of the bead beating, Triton X-100 is added to complete lysis. Yeast cells are cultured and collected while in log phase before being washed once and mixed together with glass beads in a tube. The applied shaking process facilitates disruption of the cell walls, upon which separation of yeast and glass beads is accomplished by forcing lysed cells through a hole created in the bottom of the tube during the centrifugation process. An alternative bead-beating protocol details the use of Lysis Buffer 2, which does not contain detergents and calls for the addition of Triton X-100 after cell lysis in the presence of glass beads. Use of Lysis Buffer 2 without detergent may avoid bubble and foam formation during the bead-beating process that could potentially denature proteins.

Lysing Yeast Cells for Immunoprecipitation Using a Coffee Grinder.

This protocol describes the freezing of yeast in liquid nitrogen (LN2) to form small "beans" that can be ground using a simple propeller-blade coffee grinder. The method is ideally suited for lysate preparations from larger yeast cultures ranging from 50 mL to 5 L and displays the advantage that samples remain cold during the preparative steps. Cells are cultured and collected by centrifugation while in log phase, and the resultant cell pellets are mixed with deionized distilled water and dropped into LN2 to form small frozen beans. Before the freezing process, it is imperative to keep all cell pellets at 4°C on ice. The frozen yeast beans are ground by using a simple kitchen coffee grinder, and the yeast powder is collected for immediate lysis or storage at -80°C for subsequent use. Protective clothing and safety glasses should be worn at all times when working with liquid nitrogen. Plasticware may shatter upon repeated cooling in liquid nitrogen, and appropriate care should be taken.

Immunoprecipitation.

Immunoprecipitation, commonly referred to as IP, involves the binding of proteinaceous antigen in solution by an antigen-specific antibody followed by purification of the antigen-antibody complex via attachment to a solid-phase matrix such as Protein A or G agarose. This rather simplistic and rapid technique yields highly purified immune complexes from multifactorial solutions, including cell lysates or homogenized tissues, and is most commonly used to identify and determine the relative abundance of interacting proteins, referred to as coimmunoprecipitation or co-IP. Although methods encompassing immunoblotting or western blotting of cell lysate preparations can also be applied to determine the presence and quantity of a specific antigen, its relative molecular weight, rate of synthesis or degradation, and state of target-specific posttranslational modification, immunoprecipitation can significantly increase the sensitivity for these methodologies.

Chromatin Immunoprecipitation.

Chromatin immunoprecipitation, commonly referred to as ChIP, is a powerful technique for the evaluation of in vivo interactions of proteins with specific regions of genomic DNA. Formaldehyde is used in this technique to cross-link proteins to DNA in vivo, followed by the extraction of chromatin from cross-linked cells and tissues. Harvested chromatin is sheared and subsequently used in an immunoprecipitation incorporating antibodies specific to protein(s) of interest and thus coprecipitating and enriching the cross-linked, protein-associated DNA. The cross-linking process can be reversed, and protein-bound DNA fragments of optimal length ranging from 200 to 1000 base pairs (bp) can subsequently be purified and measured or sequenced by numerous analytical methods. In this protocol, two different fixation methods are described in detail. The first involves the standard fixation of cells and tissue by formaldehyde if the target antigen is highly abundant. The dual cross-linking procedure presented at the end includes an additional preformaldehyde cross-linking step and can be especially useful when the target protein is in low abundance or if it is indirectly associated with chromatin DNA through another protein.

Differential Detergent Lysis of Cellular Fractions for Immunoprecipitation.

Differential detergent fractionation of cells is a rapid method for extraction of cytoplasmic and nuclear proteins in preparation of an immunoprecipitation. This method can be applied for use of adherent or suspension cells and can significantly reduce nonspecific background in an immunoprecipitation by separation of cellular compartments into individual fractions. The lysis of cells by differential detergents permits the rapid extraction of proteins from the cytoplasm (digitonin), the cytoplasmic membranes, and organelles (Triton X-100), and nucleoplasm (Tween/DOC), facilitated through the use of distinct extraction buffers. Cytoplasmic and nuclear matrix proteins as well as DNA are left behind during the detergent-based extraction.

Denaturing Lysis of Cells for Immunoprecipitation.

The only way to solubilize many antigens for immunoprecipitation is by denaturation. This cell lysis protocol is ideally suited for this purpose to release proteins from complex structures or reveal antibody epitopes hidden within native proteins. Short linear epitopes may not be accessible to antibodies within the native tertiary and quaternary protein structures, but they become exposed upon the unraveling of proteins, exposing their secondary structure. Antibodies otherwise not suitable for the immunoprecipitation of proteins prepared under nondenaturing conditions are now able to bind these antigens of interest in cell lysates prepared under denaturing conditions. These antibodies may also work well for immunoblotting purposes when the protein target is completely denatured. Harvested cells in this protocol are washed in tris-buffered saline (TBS) before lysis in 2% sodium dodecyl sulfate (SDS)-containing Lysis buffer for 10 min at 100°C. The resulting sample is diluted 20-fold in TBS to reduce the SDS concentration to ≤0.1% before the addition of an antibody for immunoprecipitation. Addition of 2% bovine serum albumin (BSA) or 0.1% Nonidet P-40 to the TBS before an immunoprecipitation, respectively, ensures either removal of SDS from the target protein or retaining denatured proteins in solution.

Using Dounce Homogenization to Lyse Cells for Immunoprecipitation.

Dounce homogenization in combination with hypotonic buffers facilitates the lysis of adherent and suspension cells. Addition of hypotonic buffers results in the swelling of the cell's cytoplasm, allowing for the gentle rupture of cell membranes by mechanical force. Dounce homogenization releases cytoplasmic proteins that can be processed separately from the remaining intact nuclei, which can undergo high-salt extraction for detergent-free extraction of nuclear proteins. In this protocol, cells are initially swollen by incubation in hypotonic buffer, making them susceptible to dounce lysis. Douncing is continued until most cells are lysed, leaving free intact nuclei behind from which nuclear proteins can be extracted. This method does not facilitate the extraction of histones; however, it is effective in extracting transcription factors and other chromatin-bound proteins. It is important to keep buffer volumes to a minimum to maintain high protein concentrations. Upon completion of hypotonic and high-salt extraction, respective cytoplasmic and nuclear fractions undergo dialysis to achieve physiological salt conditions before further use.

Cross-Linking Antibodies to Beads Using Dimethyl Pimelimidate (DMP).

This protocol describes the cross-linking of antibodies to either Protein A or G agarose beads using dimethyl pimelimidate (DMP). DMP contains an imidoester at each end of a 7-carbon spacer arm and forms an amidine bond with amino groups at alkaline pH; however, cross-linking is more efficient when performed at pH >8. DMP will react with primary amines; thus, it is important that the cross-linking procedure is conducted using nonamine-containing buffers. Following the antibody-bead incubation, beads are washed in Borate buffer to remove residual amines from the Tris buffer. After completion of the cross-linking process in the presence of DMP, unreacted DMP is quenched with ethanolamine, and beads are washed extensively to remove residual noncross-linked antibody before immediate use or storage at 4°C.

Cross-Linking Antibodies to Beads with Disuccinimidyl Suberate (DSS).

This protocol describes the cross-linking of antibodies to either Protein A or G agarose beads using disuccinimidyl suberate (DSS), a bifunctional cross-linker capable of directly reacting with two different amines to form stable amide bonds. Proteins, including antibodies, generally display several primary amines in the side chains of lysine (K) residues and the amino terminus of each polypeptide that represent available potential targets for N-hydroxysuccinimide (NHS)-ester cross-linking reagents. The antibody-bead cross-linking process generates a reusable resource of antibody and beads, commonly referred to as an antibody-specific resin, and can be repeatedly used for the immunoprecipitation of specific proteins if treated and stored correctly.

Tandem Immunoaffinity Purification Using Anti-FLAG and Anti-HA Antibodies.

The immunoaffinity purification of target proteins followed by the identification and characterization of associated proteins by mass spectrometry is a widely used technique. An immunoaffinity purification bears resemblance to a standard immunoprecipitation; however, the end product for mass spectrometric analysis in the femtomole (10-15) to attomole (10-18) range is required to be of exceptional purity. This high degree of sensitivity in detection renders it of extreme importance to eliminate most if not all of the nonspecific background proteins and can be achieved by performing a tandem affinity purification (TAP). In TAP, the cDNA of the target protein is engineered to contain at least two different epitope tags, and the target protein is extracted under nondenaturing conditions upon expression using an appropriate protein expression platform (CHO cells, HEK 293 cells, or yeast). The expressed protein is initially immunoprecipitated using an antibody against one epitope tag and is eluted in the presence of excess peptide by competition for antibody-binding sites, before being reimmunoprecipitated using an antibody that specifically recognizes the second epitope. These sequential immunoprecipitations significantly reduce the presence of associated nonspecific proteins. Numerous combinations of epitope tags have been applied for tandem affinity purification, and this protocol illustrates the use of tandem hemagglutinin (HA) and FLAG epitope tags. The first immunoprecipitation uses an anti-FLAG antibody followed by the elution in the presence of a competing FLAG peptide before the reimmunoprecipitation of the protein using an anti-HA antibody. Numerous high-quality antiepitope tag antibodies are commercially available from different antibody manufacturers.

Pulse-Chase Labeling of Protein Antigens with [35S]Methionine.

Pulse-chase labeling of antigens with [35S]methionine is used to determine the relative half-life of a protein. In this protocol, intracellular unlabeled methionine levels are reduced by starvation of cells for 0.5-1 h, and then the cells are briefly labeled with [35S]methionine to create the pulse of newly synthesized proteins. Upon completion of cell labeling, the addition of Chasing medium containing an excess of unlabeled methionine is used to create the chase, reducing the likelihood that any remaining [35S]methionine will be incorporated into newly synthesized proteins. Labeling and chasing reactions of adherent cells can be directly performed in cell culture dishes in an incubator, whereas suspension cells are labeled and chased in a polypropylene tube kept in a water bath set at 37°C. At intervals after the pulse, aliquots of chased labeled cells are collected and pelleted with the option of immediately preparing cell lysates or freezing and storing the cell pellets at -80°C. Upon cell lysis and antigen purification by immunoprecipitation, SDS-PAGE-resolved proteins can be fixed on the gel and enhanced with fluorography or can be transferred to a nitrocellulose or polyvinylidene fluoride (PVDF) membrane followed by autoradiography or exposure in a phosphorimager. Membrane blotting has the advantage of allowing for detection of the target of interest by probing with an antigen-specific antibody to confirm that equal amounts of steady-state levels of the target protein were immunoprecipitated at each interval.

Metabolic Labeling of Protein Antigens with [32P]Orthophosphate.

In this protocol, cells are cultured in the presence of [32P]orthophosphate used by the cell to phosphorylate proteins. Cells must be maintained at 37°C at all times during this procedure. A drop in temperature will significantly reduce cellular metabolic activity and subsequently decrease the uptake of [32P]orthophosphate. Once the cells are labeled, cell lysates are prepared for immunoprecipitation, followed by SDS-PAGE analysis. The gel can be dried directly or transferred onto a nitrocellulose or polyvinylidene fluoride (PVDF) membrane for analysis by western blotting and imaging by autoradiography or phosphorimager.

Detergent Lysis of Tissue Culture Cells for Immunoprecipitation.

This protocol describes the lysis of tissue culture cells for the solubilization of proteins of interest for immunoprecipitation. Upon collection of cells by centrifugation and depending on the use of either Tris- or phosphate-based cell lysis buffers, cells are rinsed, respectively, in either TBS or PBS before lysis. If possible, the pH of the Wash buffer should match that of the Lysis buffer. Adherent cells can be directly lysed on the plate. This is particularly useful upon lysis encompassing mild nonionic detergents, leaving the cytoskeleton intact. Alternatively, adherent cells can be scraped off the plate and directly resuspended in Lysis buffer or Wash buffer for transfer to a tube followed by the addition of Lysis buffer.

Detergent Lysis of Animal Tissues for Immunoprecipitation.

This protocol details protein extraction from mouse tissues for immunoprecipitation purposes and has been applied for the performance of large-scale immunoprecipitations of target proteins from various tissues for the identification of associated proteins by mass spectroscopy. The key factors in performing a successful immunoprecipitation directly relate to the abundance of target protein in a particular tissue type and whether or not the embryonic, newborn, or adult mouse-derived tissues contain fibrous and other insoluble material. Several tissue types, including lung and liver as well as carcinomas, contain significant amounts of fibrous tissue that can interfere with an immunoprecipitation.

Immunoprecipitation.

This immunoprecipitation protocol details individual steps for the enrichment and purification process of specific proteins from a complex cell lysate using an antibody bound to a solid matrix. Purified antigen(s) can be eluted by various methods, and the resultant protein target can be analyzed and/or identified by numerous assays, including the enzyme-linked immunosorbent assay (ELISA), western blotting, or mass spectrometry.

Metabolic Labeling of Protein Antigens with [35S]Methionine.

Metabolic labeling of antigens can be achieved by supplementation of the cell culture medium with radioactive amino acid precursors such as [35S]methionine during the incubation period of target cells. In this protocol, intracellular unlabeled methionine levels are reduced by starvation of cells for 0.5-1 h before the addition of labeled [35S]methionine and incubation for 0.5-4 h. Upon completion of the metabolic labeling process, cells can be prepared for immunoprecipitation by lysis or alternatively pelleted and frozen for cell lysate preparations at a later time.

Indirect Competitive Enzyme-Linked Immunosorbent Assay (ELISA).

The indirect competitive ELISA (indirect cELISA) pits plate-immobilized antigen against antigens in solution for binding to antigen-specific antibody. The antigens in solution are in the test sample and are first incubated with antigen-specific antibody. These antibody-antigen complexes are then added to microtiter plates whose wells have been coated with purified antigen. The wells are washed to remove unbound antigen-antibody complexes and free antigen. A reporter-labeled secondary antibody is then added followed by the addition of substrate. Substrate hydrolysis yields a signal that is inversely proportional to antigen concentration within the sample. This is because when antigen concentration is high in the test sample, most of the antibody is bound before adding the solution to the plate. Most of the antibody remains in solution (as complexes) and is thus washed away before the addition of the reporter-labeled secondary antibody and substrate. Thus, the higher the antigen concentration in the test sample, the weaker the resultant signal in the detection step. The indirect cELISA is often used for competitive detection and quantification of antibodies against viral diseases in biological samples.

Direct Competitive Enzyme-Linked Immunosorbent Assay (ELISA).

The competitive enzyme-linked immunosorbent assay (ELISA) (cELISA; also called an inhibition ELISA) is designed so that purified antigen competes with antigen in the test sample for binding to an antibody that has been immobilized in microtiter plate wells. The same concept works if the immobilized molecule is antigen and the competing molecules are purified labeled antibody versus antibody in a test sample. Direct cELISAs incorporate labeled antigen or antibody, whereas indirect assay configurations use reporter-labeled secondary antibodies. The cELISA is very useful for determining the concentration of small-molecule antigens in complex sample mixtures. In the direct cELISA, antigen-specific capture antibody is adsorbed onto the microtiter plate before incubation with either known standards or unknown test samples. Enzyme-linked antigen (i.e., labeled antigen) is also added, which can bind to the capture antibody only when the antibody's binding site is not occupied by either the antigen standard or antigen in the test samples. Unbound labeled and unlabeled antigens are washed away and substrate is added. The amount of antigen in the standard or the test sample determines the amount of reporter-labeled antigen bound to antibody, yielding a signal that is inversely proportional to antigen concentration within the sample. Thus, the higher the antigen concentration in the test sample, the less labeled antigen is bound to the capture antibody, and hence the weaker is the resultant signal.

Immunoassays.

The enzyme immunoassay (EIA) is one of the most powerful of all immunochemical techniques. First described in the early 1970s, these assays are now used routinely in laboratory analyses and diagnostics. In biology and biotechnology, the EIA is a valuable and versatile tool used to detect and quantitate antigens and antibodies. Application of the appropriate EIA permits rapid quantification of different antigens and antibodies (referred to here as analytes) present at very low concentrations within a mixture. These assays are extremely sensitive and provide valuable information that would be difficult to determine by other techniques. Here we detail the development and optimization of the enzyme-linked immunosorbent assay (ELISA), a term generally used for any plate-based immunoassay that incorporates enzyme-, chemiluminescence-, or fluorescence-based reporters. It is amenable to standardization, automation, and large-scale sampling.