Research articles

Northern blotting and hybridization are used to study gene expression by detecting RNA species of interest, and to identify alternate RNA splicing patterns. It is analogous to Southern blotting, which is used to analyze DNA. This protocol describes the transfer of RNA from agarose gels to nylon membranes and the fixation of the RNA to the membrane^1,2,3.

In 1992, investigators working on mixing and fusing various domains of different DNA polymerases showed that, when combined with the robust reliability of Taq DNA polymerase, these enzymes produced longer amplicons. Long and accurate polymerase chain reaction (LA PCR) refers to the production of amplified product longer than ∼3 kilobases (kb) with high fidelity. Long PCR mixtures typically yield PCR products with some tenfold fewer mutations¹ than those observed in products resulting from conventional PCR. The mixture of DNA polymerases typically included either Taq or Klentaq1 (which have no 3′-exonuclease proofreading activity) as the major component and, as the minor component, an archaebacterial DNA polymerase (with proofreading activity) such as Deep Vent, Vent or Pfu¹. Among other factors that improve LA PCR are the enzyme deoxyuridine triphosphatase (dUTPase)², which prevents the incorporation of dUTP, the deaminated form of deoxycytosine triphosphate (dCTP) into DNA, and the chemical betaine. Although introduced for amplification of high G-C content targets³, betaine, when included at surprisingly high concentrations, usually helps to promote long PCR up to at least 20 kb. Since the introduction of mixtures of DNA polymerases, most PCRs of any length have improved in reliability and in yield of product. The following protocol is based on the use of ten various primer pairs for the amplification of a genomic template DNA of ∼5 kb in a reaction volume of 50 μl.

The immunoblotting method has evolved from early stages when antibodies were used to 'stain' polyacrylamide gels directly^1,2 to more versatile methods using replica techniques, in which the separated polypeptides are transferred to nitrocellulose membranes, chemically activated paper or nylon sheets. Although there are several variations on this basic theme, the most common and effective is electrophoretic transfer to nitrocellulose sheets^3,4. The separated proteins can then be probed with antibodies; this variation of the technique is known as immunoblotting (or western blotting). The membrane can also be probed with specific ligands, such as DNA, protein, small molecules (for example, heparin or GTP) or even whole cells. Electrophoretic transfer can be achieved either in a tank³ or in a semidry apparatus⁵, in which the buffer volume is reduced to filter paper pads. The following protocol presents the original method used for tank blotting.

Micrococcal nuclease (MNase) is unique among nucleases in its ability to induce double-strand breaks within nucleosome linker regions, but only single-strand nicks within the nucleosome itself. Because of this property, MNase can be used to determine whether a DNA fragment of interest is within a nucleosome^1,2. In addition, MNase can be used to determine the approximate positions of nucleosomes in a region of DNA, if the nucleosomes are consistently positioned. In brief, cell nuclei are isolated and limiting concentrations of MNase are added to the nuclei, resulting in cleavage at nucleosome linker regions. The genomic DNA is purified and the fragments are separated by agarose gel electrophoresis; the resulting ladder of stained bands corresponds in size to multiples of the nucleosome core plus the linker (∼200 base pairs (bp)). To determine whether a DNA fragment of interest is within a nucleosome, the genomic DNA is subjected to Southern blot analysis. If a probe derived from the DNA fragment hybridizes to the ladder of nucleosomal bands, the fragment may indeed be assembled into nucleosomes. To determine nucleosome positioning, the purified genomic DNA must be cleaved with a restriction enzyme before gel electrophoresis and Southern blot analysis.

This method is used to extend partial cDNA clones by amplifying the 5′ sequences of the corresponding mRNAs^1,2,3. The technique requires knowledge of only a small region of sequence within the partial cDNA clone. During PCR, the thermostable DNA polymerase is directed to the appropriate target RNA by a single primer derived from the region of known sequence; the second primer required for PCR is complementary to a general feature of the target—in the case of 5′ RACE, to a homopolymeric tail added (via terminal transferase) to the 3′ termini of cDNAs transcribed from a preparation of mRNA. This synthetic tail provides a primer-binding site upstream of the unknown 5′ sequence of the target mRNA. The products of the amplification reaction are cloned into a plasmid vector for sequencing and subsequent manipulation.

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.