Research articles

The electrophoretic mobility shift assay (EMSA), one of the most sensitive methods for studying the DNA-binding properties of a protein, can be used to deduce the binding parameters and relative affinities of a protein for one or more DNA sites or for comparing the affinities of different proteins for the same sites¹. It is also useful for studying higher-order complexes containing several proteins, observed as a 'supershift assay'. EMSA also can be used to study protein- or sequence-dependent DNA bending². In an EMSA, or simple 'gel shift', a ³²P-labeled DNA fragment containing a specific DNA site is incubated with a candidate DNA-binding protein. The protein-DNA complexes are separated from free (unbound) DNA by electrophoresis through a nondenaturing polyacrylamide gel. The protein retards the mobility of the DNA fragments to which it binds; thus, the free DNA migrates faster through the gel than does the DNA-protein complex. An image of the gel reveals the positions of the free and bound ³²P-labeled DNA.

Many protein-protein associations that exist within the cell remain intact when a cell is lysed under nondenaturing conditions. Thus, if protein X is immunoprecipitated, then protein Y, which stably associated with X, may also precipitate. Coimmunoprecipitation is most commonly used to test whether two proteins of interest are associated in vivo, but it can also be used to identify interacting partners of a target protein. In both cases, the cells, labeled with [³⁵S]methionine, are collected and lysed under conditions that preserve protein-protein interactions. The target protein is specifically immunoprecipitated from the cell extracts, and the immunoprecipitates are fractionated by SDS-PAGE. Coimmunoprecipitated proteins are detected by autoradiography and/or by western blotting with an antibody directed against that protein. The identity of interacting proteins may be established or confirmed by Edman degradation of tryptic peptides. Some early examples of this method include the use of antibodies to viral antigens to determine the host cellular proteins that interact with these viral transforming oncoproteins. Two interacting proteins of particular note are the tumor suppressor proteins p53 and pRB^1,2,3. This protocol was used to identify pVHL-associated proteins; conditions should be optimized for the protein of interest.

Preparations of RNA containing an mRNA of interest are hybridized to a complementary single-stranded DNA probe. At the end of the reaction period, nuclease S1 is used to degrade unhybridized regions of the probe, and the surviving DNA-RNA hybrids are then separated by gel electrophoresis and visualized by either autoradiography or Southern hybridization. The method can be used to quantify RNAs, to map the positions of introns and to identify the locations of 5′ and 3′ ends of mRNAs on cloned DNA templates^1,2,3.

The uptake of DNA is markedly enhanced when the nucleic acid is presented as a coprecipitate of calcium phosphate and DNA¹. The insoluble precipitate attaches to the cell surface and is brought into the cells by endocytosis. Since the publication of the original method¹, increases in efficiency have been achieved by including additional steps such as glycerol shock² and/or chloroquine treatment³. This protocol is a modified version of a published method⁴, in which calcium phosphate–based transfection methods for Chinese hamster ovary cells and the 293 line of human embryonic kidney cells were rigorously optimized. The protocol is easily adapted for use with other types of cells, both adherent and nonadherent.

In situ hybridization (ISH) is used to visualize defined nucleic acid sequences in cellular preparations by hybridization of complementary probe sequences. Probe sequences can be labeled with isotopes, but nonisotopic ISH is used increasingly as it is considerably faster, usually has greater signal resolution, and provides many options to simultaneously visualize different targets by combining various detection methods. The most popular protocols use fluorescence detection, as described here. These protocols have many applications, from basic gene mapping and diagnosis of chromosomal aberrations^1,2,3 to detailed studies of cellular structure and function, such as the painting of chromosomes in three-dimensionally preserved nuclei^4,5. This protocol describes fluorescence in situ hybridization (FISH) of biotin- or digoxigenin-labeled probes to denatured metaphase chromosomes and interphase nuclei. The hybridized probes are detected and visualized using fluorochrome-conjugated reagents.

The basic strategy for enzymatic conversion of RNA into DNA has changed little since the 1970s; however, there have been great improvements to the efficiency of the overall process. In this method^1,2,3, the product of a first-strand synthesis (the cDNA-mRNA hybrid) is used as template for a nick translation reaction. Ribonuclease (RNase) H produces nicks and gaps, creating a series of RNA primers used by Escherichia coli DNA polymerase I during the synthesis of the second-strand DNA. Residual nicks are then repaired by E. coli DNA ligase, and the frayed termini of the double-stranded cDNA are polished by a DNA polymerase. Finally, bacteriophage T4 polynucleotide kinase catalyzes the phosphorylation of the ends of the cDNAs to facilitate cloning.

Two-dimensional polyacrylamide gel electrophoresis (PAGE) is used for separation of complex protein mixtures by the independent parameters of isoelectric point and molecular weight. Isoelectric focusing (IEF) separates proteins in a pH gradient. Each protein is 'focused' because it moves under the influence of the electric field until it reaches its isoelectric point, the pH at which it has no net charge. After IEF in the presence of urea and a nonionic detergent, the IEF gel is equilibrated in sodium dodecyl sulfate (SDS) to prepare the proteins for SDS-PAGE. The method described here¹ uses carrier ampholytes to form a pH gradient in a long, thin (1.2-mm) focusing gel composed of a low percentage (2.7%) of acrylamide and containing 9.5 M urea and 2% Nonidet P-40 to maintain protein solubility. After IEF, the gel is briefly equilibrated in SDS and placed directly onto the top edge of a second-dimension slab gel. Because the time between the end of IEF and the start of SDS-PAGE is only a few minutes (thereby minimizing diffusion), the spots are highly resolved and nearly round in shape.

Glutathione-S-transferase (GST) fusion proteins have had a range of applications since their introduction as tools for synthesis of recombinant proteins in bacteria¹. Typically, GST pull-down experiments are used to identify interactions between a probe protein and unknown targets and to confirm suspected interactions between a probe protein and a known protein^2,3. The probe protein is a GST fusion, whose coding sequence is cloned into an isopropyl-β-D-thiogalactoside (IPTG)-inducible expression vector. This fusion protein is expressed in bacteria and purified by affinity chromatography on glutathione-agarose beads. Target proteins are usually lysates of cells, either labeled with [³⁵S]methionine or unlabeled, depending on the method used to assay the interaction between the target and the probe. The cell lysate and the GST fusion protein are incubated together with glutathione-agarose beads. Complexes recovered from the beads are resolved by SDS-PAGE and analyzed by western blotting, autoradiography or staining.

The 'megaprimer' method of site-directed mutagenesis uses three oligonucleotide primers and two rounds of polymerase chain reaction (PCR)¹. One oligonucleotides is mutagenic; the others are forward and reverse primers that lie upstream and downstream from the binding site for the mutagenic oligonucleotide. The mutagenic primer and the nearer of the external primers are used in the first PCR to generate and amplify a mutated fragment of DNA. This amplified fragment—the megaprimer—is used in the second PCR in conjunction with the remaining external primer to amplify a longer region of the template DNA. This protocol is based on a method that uses forward and reverse external primers with significantly different melting temperatures (T_m)².

Southern transfer and hybridization¹ is used to study how genes are organized within genomes by mapping restriction sites in and around segments of genomic DNA. This protocol describes the first stages of Southern blotting: digestion of genomic DNA with restriction enzymes, separation of the resulting fragments by gel electrophoresis, and capillary transfer of the denatured fragments to a membrane².