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In molecular cloning, digoxigenin is used as a ligand that can be incorporated into DNA and RNA probes and detected after hybridization with an anti-digoxigenin-antibody enzyme conjugate. Methods to label nucleic acids with digoxigenin and to detect digoxigenin-labeled probes are introduced here., (© 2022 Cold Spring Harbor Laboratory Press.)

The polymerase chain reaction (PCR) can be used to produce both nonradiolabeled DNA probes and radiolabeled DNA probes with high specific activity. In this protocol, PCR is used to generate double-stranded probes. Related methods, including the generation of asymmetric probes by PCR, are also discussed., (© 2022 Cold Spring Harbor Laboratory Press.)

Northern hybridization is used to measure the amount and size of RNAs transcribed from eukaryotic genes and to estimate their abundance. No other method is capable of obtaining these pieces of information simultaneously from a large number of RNA preparations; northern analysis is therefore fundamental to studies of gene expression in eukaryotic cells. To prepare a northern blot for hybridization, RNA must be separated according to size through a denaturing agarose or polyacrylamide gel and transferred to a solid support in a way that preserves its topological distribution within the gel. These important steps in northern analysis are discussed here., (© 2022 Cold Spring Harbor Laboratory Press.)

In most cases, fractionation of RNA by agarose gel electrophoresis is but a prelude to hybridization of the fractionated population to specific labeled probes that detect particular target mRNAs. RNA is first transferred from an agarose gel to a 2D support, usually a nylon membrane. This protocol presents the steps involved in the transfer of RNA from an agarose gel to a membranous support, facilitated by the upward flow of buffer, followed by various methods for fixation of the RNA to the membrane in preparation for hybridization. An alternative method for transfer by downward capillary flow is also given., (© 2022 Cold Spring Harbor Laboratory Press.)

Dot blotting and slot blotting are techniques for immobilizing several preparations of nucleic acids on the same solid support, usually a charged nylon membrane. The concentrations of the target sequence of interest can be estimated by hybridizing the immobilized samples to an appropriate probe. The amounts of target sequence are estimated by comparing the intensity of signals emitted by dots containing the test samples with standards containing known concentrations of the target sequence. This protocol describes the blotting and subsequent hybridization of RNA that has been purified from cells or tissues., (© 2022 Cold Spring Harbor Laboratory Press.)

Samples of RNA may be denatured by treatment with formamide and separated by electrophoresis through agarose gels containing formaldehyde. In this method, RNA is fractionated by electrophoresis through an agarose gel containing 2.2 m formaldehyde., (© 2022 Cold Spring Harbor Laboratory Press.)

If labeled oligonucleotides are to be used only as probes in hybridization experiments, complete removal of unincorporated label is generally not necessary. However, to reduce background to a minimum, the bulk of the unincorporated label should be separated from the labeled oligonucleotide. Most of the residual unincorporated precursors can be removed from the preparation by differential precipitation with ethanol, as described in this protocol, if the oligonucleotide is >18 nt in length., (© 2022 Cold Spring Harbor Laboratory Press.)

Labeled nucleic acids and oligonucleotides are typically generated by enzymatic methods such as end-labeling, random priming, nick translation, in vitro transcription, and variations of the polymerase chain reaction (PCR). Some of these methods place the label in specific locations within the nucleic acid (e.g., at the 5' or 3' terminus); others generate molecules that are labeled internally at multiple sites. Some methods yield labeled single-stranded products, whereas others generate double-stranded nucleic acids. Finally, some generate probes of defined length, whereas others yield a heterogeneous population of labeled molecules. Options available for generating and detecting labeled nucleic acids, as well as advice on designing oligonucleotides for use as probes, is included here., (© 2022 Cold Spring Harbor Laboratory Press.)

In this protocol, hybridization is first performed in conventional aqueous solvents at a temperature well below the melting temperature, and the hybrids are then washed at higher stringency in buffers containing quaternary alkylammonium salts. TMACl is used with probes that are 14-50 nt in length, whereas TEACl is used with oligonucleotides that are 50-200 nt in length., (© 2022 Cold Spring Harbor Laboratory Press.)

Isolation of RNA from yeast is complicated by the need to first break the thick, rigid cell wall. The protocol provided here uses a cycle of heating and freezing of cells in the presence of phenol and the detergent sodium dodecyl sulfate (SDS). The extraction is performed in the presence of low salt so that, following separation of the aqueous and phenol phases by centrifugation, DNA can be collected from the interface while RNA remains in the aqueous phase. This protocol should yield ∼50-250 µg of RNA from 10 mL of culture. The RNA isolated using this approach is suitable for most follow-up applications such as northern blot hybridization, reverse transcriptase-polymerase chain reaction (RT-PCR), and cDNA construction., (© 2021 Cold Spring Harbor Laboratory Press.)

Plasmids occupy a place of honor in molecular cloning: They were used in the first recombinant DNA experiments and, 40 or more years later, they remain as the carriage horses of molecular cloning. After almost half a century of sequential improvement in design, today's plasmid vectors are available in huge variety, are often optimized for specific purposes, and bear only passing resemblance to their forebears. Here, various features of plasmid vectors and methods for transforming E. coli cells are introduced., (© 2021 Cold Spring Harbor Laboratory Press.)

Alkaline agarose gels are run at high pH, which causes each thymine and guanine residue to lose a proton and thus prevents the formation of hydrogen bonds with their adenine and cytosine partners. The denatured DNA is maintained in a single-stranded state and migrates through an alkaline agarose gel as a function of its size. Other denaturants such as formamide and urea do not work well because they cause the agarose to become rubbery., (© 2021 Cold Spring Harbor Laboratory Press.)

This protocol describes a convenient method for the preparation, use, and storage of competent Escherichia coli The reported transformation efficiency of this method is ∼5 × 10 7 transformants/µg of plasmid DNA., (© 2021 Cold Spring Harbor Laboratory Press.)

When labeled oligonucleotides are to be used in enzymatic reactions such as primer extension, virtually all of the unincorporated label must be removed from the oligonucleotide. For this purpose, chromatographic methods or gel electrophoresis are superior to differential precipitation of the oligonucleotide with ethanol or cetylpyridinium bromide (CPB). This protocol describes a method to separate labeled oligonucleotides from unincorporated label that takes advantage of differences in mobility between oligonucleotides and mononucleotides during size-exclusion chromatography. Although size-exclusion chromatography can, in principle, be used to purify either radiolabeled or nonradiolabeled oligonucleotides, this protocol is geared toward purifying radiolabeled oligonucleotides, whose elution from the column is monitored using a minimonitor and whose separation from unincorporated nucleotides is monitored by liquid scintillation counting., (© 2021 Cold Spring Harbor Laboratory Press.)

In this method, a short primer is hybridized to an oligonucleotide template whose sequence is the complement of the desired radiolabeled probe. The primer is then extended using the Klenow fragment to incorporate [α- 32 P]dNTPs in a template-directed manner. After the reaction, the template and product are separated by denaturation followed by electrophoresis through a polyacrylamide gel under denaturing conditions. With this method, it is possible to generate oligonucleotide probes that contain several radioactive atoms per molecule of oligonucleotide and to achieve specific activities as high as 2 × 10 10 cpm/µg of probe. Because the end product of the reaction is dsDNA, whose strands must be separated and the labeled product isolated, this method is generally not used to prepare nonradiolabeled oligonucleotides., (© 2021 Cold Spring Harbor Laboratory Press.)

Biotin is a water-soluble vitamin that can be attached to a variety of proteins and nucleic acids, often without altering their properties. Its use in molecular biology is introduced here., (© 2021 Cold Spring Harbor Laboratory Press.)

This introduction outlines various methods to clone amplified DNAs and to facilitate the construction of complex multicomponent genetic units. Because of the ease with which the termini of amplified DNAs can be tailored by polymerase chain reaction (PCR), many of the methods outlined here use PCR not only to synthesize DNAs but also to link them together into purpose-designed constructs. The most recent refinements however have been the development of modular genetic units that can be harnessed to target DNAs not by PCR but by site-specific recombination enzymes., (© 2021 Cold Spring Harbor Laboratory Press.)

This protocol describes the use of TOPO-activated TA vectors for cloning. Manufacturers of cloning kits provide excellent manuals that explain in detail what to do and why to do it. This makes TOPO cloning easy, but not foolproof. When setting up TOPO cloning for the first time, set up a trial experiment as described here., (© 2021 Cold Spring Harbor Laboratory Press.)

The removal of 5' phosphates from nucleic acids with phosphatases and their readdition in radiolabeled form by bacteriophage T4 polynucleotide kinase is a widely used technique for generating 32 P-labeled probes. When the reaction is performed efficiently, 40%-50% of the protruding 5' termini in the reaction becomes radiolabeled. However, the specific activity of the resulting probes is not as high as that obtained by other radiolabeling methods because only one radioactive atom is introduced per DNA molecule. Nevertheless, the availability of [γ- 32 P]ATP with specific activities in the 3000-7000 Ci/mmol range allows the synthesis of probes suitable for many purposes. This protocol includes procedures for labeling the 5' ends of dsDNA and oligonucleotides., (© 2021 Cold Spring Harbor Laboratory Press.)

This protocol describes a method for the separation of radiolabeled oligonucleotides from unincorporated radiolabel that takes advantage of the reversible affinity of oligonucleotides for silica gel. This protocol can be used only to purify oligonucleotides carrying a 5'-phosphate group, radiolabeled or unlabeled., (© 2021 Cold Spring Harbor Laboratory Press.)

Terminal deoxynucleotidyl transferase (TdT, also simply called terminal transferase) is a template-independent polymerase that catalyzes the addition of deoxynucleotides and dideoxynucleotides to the 3'-hydroxyl terminus of a DNA molecule. Cobalt (Co 2+ ) is a necessary cofactor for the activity of this enzyme. Incorporation at the 3' terminus can be limited to just 1 nt by using [α- 32 P]ddATP or biotin-, digoxigenin (DIG)-, or fluorescein-ddUTP. Because none of these molecules carries a 3'-hydroxyl group, no additional molecules can be incorporated. Alternatively, the enzyme is capable of adding several (2-100) nt to 3' ends in a so-called homopolymeric "tailing" reaction. A tailing reaction is performed in the presence of a mixture of labeled and unlabeled dNTPs. The rate of addition of dNTPs, and thus the length of the tail, is a function of the ratio of 3' DNA ends to dNTP concentration and, in addition, the specific dNTP that is used., (© 2021 Cold Spring Harbor Laboratory Press.)

In this protocol, restriction fragments that have been transferred to a membrane by Southern blotting are hybridized to a labeled probe. Methods for stripping the probe from the membrane are also included., (© 2021 Cold Spring Harbor Laboratory Press.)

In Southern blotting, DNA is digested with one or more restriction enzymes, and the resulting fragments are separated according to size by electrophoresis through a standard agarose gel. The DNA is then denatured in situ and transferred from the gel to a solid support (usually a nylon or nitrocellulose membrane). The relative positions of the DNA fragments are preserved during their transfer to the membrane. The DNA is then fixed to the membrane and prepared for hybridization. Alternatively, DNA can be simultaneously transferred from the top and bottom surfaces of a single agarose gel to two membranes. This procedure is useful when the need arises to analyze the same set of restriction fragments with two different probes. Transfer of DNA fragments is rapid, but the efficiency is low because the agarose gel quickly becomes dehydrated as fluid is withdrawn from both sides. The method therefore works best when the target sequences are present in high concentration (e.g., when analyzing cloned DNAs [plasmids, bacteriophages, cosmids, PACs, or BACs] or less complex genomes [those of Saccharomyces cerevisiae or Drosophila ]). Too little mammalian genomic DNA is transferred to allow signals from single-copy sequences to be detected in a reproducible or timely fashion., (© 2021 Cold Spring Harbor Laboratory Press.)

Southern transfer and hybridization are used to study how genes are organized within genomes by mapping restriction sites in and around segments of genomic DNA for which specific probes are available. Genomic DNA is first digested with one or more restriction enzymes, and the resulting fragments are separated according to size by electrophoresis through a standard agarose gel. The DNA is then denatured in situ and transferred from the gel to a solid support (usually a nylon or nitrocellulose membrane). The DNA attached to the membrane is hybridized to a labeled DNA, RNA, or oligonucleotide probe, and bands complementary to the probe are located by an appropriate detection system (e.g., by autoradiography). By estimating the size and number of the bands generated after digestion of the genomic DNA with different restriction enzymes, singly or in combination, it is possible to place the target DNA within a context of restriction sites., (© 2021 Cold Spring Harbor Laboratory Press.)

The nontemplate-dependent terminal transferase activity inherent in nonproofreading DNA polymerases such as Taq provides a highly efficient method to clone PCR products. The enzyme adds a single, unpaired residue-preferentially an adenosyl residue-to each 3' end of a double-stranded amplified product. The unpaired terminal (A) residues can pair with a linear T vector that carries an unpaired 3'-thymidyl residue at each end. The two chief advantages of TA cloning are speed and lack of reliance on restriction enzymes. The major disadvantage is an inability to clone directionally. For this reason, it is important to pick and analyze several transformed clones when a particular orientation of the amplified fragment is required., (© 2021 Cold Spring Harbor Laboratory Press.)

During polymerase chain reaction (PCR), DNA polymerases such as Taq add a single, unpaired residue-preferentially an adenosyl residue-to each 3' end of a double-stranded amplified product. The unpaired terminal (A) residues can pair with a linear T vector that carries an unpaired 3'-thymidyl residue at each end. T vectors can be prepared in the laboratory or they may be purchased ready-made from commercial suppliers. This protocol outlines two methods in common use to generate T vectors., (© 2021 Cold Spring Harbor Laboratory Press.)

For mapping the 5' termini of mRNA molecules, primer extension is the method of choice. A purified oligonucleotide is end-labeled using polynucleotide kinase. The probe and a population of mRNA are allowed to hybridize, and the primers and template are used to carry out reverse transcription using an enzyme cloned from the Moloney murine leukemia virus. The primer extension products are separated on a denaturing polyacrylamide gel and analyzed by radiography., (© 2021 Cold Spring Harbor Laboratory Press.)

In this protocol a randomly labeled single-stranded RNA probe is prepared and then hybridized to a population of mRNA molecules. The RNAs are digested with a mixture of RNase A and RNase T1. The hybrid molecules, which are resistant to the RNases, are separated and analyzed using gel electrophoresis and radiography., (© 2021 Cold Spring Harbor Laboratory Press.)

Details of the fine structure of a particular transcript may be assessed by S1 mapping or ribonuclease protection. The use of either of these techniques allows the detection of the 5' and 3' ends of a particular mRNA, as well as the splice junctions, precursors, and processing intermediates of mRNA. Primer extension provides a measure of the amount of a particular mRNA species and allows an exact determination of the 5' end of the mRNA. These three methods are introduced here., (© 2021 Cold Spring Harbor Laboratory Press.)

This protocol provides details for nuclease S1 mapping of mRNA using a uniformly labeled, single-stranded DNA probe. DNA-RNA hybrids are generated, which are subsequently digested with nuclease S1. The digestion products are separated using gel electrophoresis and analyzed by radiography., (© 2021 Cold Spring Harbor Laboratory Press.)

To generate polymerase chain reaction (PCR) products that can be directionally cloned into a vector, different restriction sites are built into the forward and reverse primers that are used in the PCR. After PCR, the amplified product is purified, cleaved with the appropriate restriction enzymes, ligated into a vector with compatible cohesive ends, and used to transform E. coli ., (© 2021 Cold Spring Harbor Laboratory Press.)

Restriction enzymes provided the foundation on which molecular cloning was built, and they remain as essential tools in current recombinant DNA technology. The three classes of restriction enzymes and their features are introduced here., (© 2021 Cold Spring Harbor Laboratory Press.)

The following is an elegant and simple protocol for generating and cloning blunt-ended DNA. Incubation of a ligation reaction in the presence of an excess amount of restriction enzyme can dramatically increase the yield of recombinant plasmids. The role of the restriction enzyme is to cleave circular and linear concatemers at restriction sites that are regenerated when plasmid molecules ligate to themselves. The method requires that ligation of the plasmid to a target DNA molecule destroys the restriction site, so preventing the restriction enzyme from digesting recombinants generated during the ligation reaction. The net effect of constant reclamation of unit-length linear vector molecules is to drive the equilibrium of the ligation reaction strongly in favor of recombinants between vector and insert. The method is efficient because regeneration of vector DNA, ligation, and polishing the termini of PCR-generated fragments of DNA all occur simultaneously in the same reaction mixture., (© 2021 Cold Spring Harbor Laboratory Press.)

In this procedure, synthesis of cDNA is performed in the presence of saturating concentrations of all four dNTPs and trace amounts of a single radiolabeled dNTP. After subtraction hybridization, the enriched single-stranded cDNA is radiolabeled to high specific activity in a second synthetic reaction by extension of random oligonucleotide primers using the Klenow fragment of Escherichia coli DNA Pol I. Because the concentrations of dNTP in the first reaction are nonlimiting, both the amounts and size of cDNA generated are greater than those achieved in standard labeling protocols. The subtractive hybridization step can therefore be performed with higher efficiency. Because the resulting population of cDNA is not vulnerable to radiolytic cleavage, it can be stored indefinitely and radiolabeled to higher specific activity when needed. The protocol works best when the cDNA synthesized in the initial synthetic reaction is full length or close to it. For this reason, synthesis of cDNAs is primed by oligo(dT) rather than random hexanucleotide primers. In contrast, the subsequent radiolabeling reaction is primed by random oligonucleotides, yielding shorter DNA products whose size is ideal for hybridization., (© 2021 Cold Spring Harbor Laboratory Press.)

Polyacrylamide gels, unlike agarose gels, have a highly cross-linked matrix that does not permit efficient, quantitative, or reproducible transfer by passive diffusion. Polyacrylamide gels should therefore be transferred by electroblotting (also known as "electrophoretic transfer"). Because this approach requires the use of a transfer buffer of low ionic strength, nylon membranes are generally used. The membrane is placed directly on the gel, and the gel together with its membrane is then sandwiched between layers of filter paper and placed in a special cassette that, in turn, is placed in a tank of buffer; the cassette is oriented such that the membrane side of the gel faces the positively charged anode. A voltage gradient is applied perpendicular to the direction of the initial electrophoresis of the gel, forcing the sample to migrate out of the gel and onto the membrane., (© 2021 Cold Spring Harbor Laboratory Press.)

Thin (0.4-1.5 mm) polyacrylamide-urea gels provide high resolution of RNAs up to 1000 nt in size and are capable of resolving single-stranded fragments of RNA that differ in length by as little as 1 nt. The polyacrylamide gel is cast between two glass plates that are separated by two thin Teflon or nylon spacers. A so-called shark's tooth comb or, less frequently, a standard slotted comb forms the sample wells into which the RNA samples are loaded before electrophoresis. In contrast to electrophoresis using agarose gels, which occurs while the gel is horizontal, polyacrylamide gels are run while in the vertical position. Gels are also typically run at 45°C-55°C, which is the melting temperature of RNA, and in the presence of 6-8 m urea. The gel recipe and protocol presented here for 8 m urea/TBE polyacrylamide gels can be used for a variety of applications including mapping RNA with nuclease S1, ribonuclease protection assay, or analysis of RNA by primer extension., (© 2021 Cold Spring Harbor Laboratory Press.)

RNA samples that have been transferred and fixed to a membrane may be hybridized with a specific probe to locate the RNA species of interest. Any one of a large number of methods can be used to label and detect probes, at the discretion of the investigator. After treating the membrane with blocking agents that suppress nonspecific absorption of the probe, the membrane is incubated under conditions that favor hybridization of the labeled probe to the immobilized target RNA. The membrane is then washed extensively to remove adventitiously bound probe and finally manipulated to yield an image of the distribution of the tightly bound probe on the membrane. After analysis of the results, the probe may be stripped from the membrane, and the membrane can then be used again in another hybridization experiment., (© 2021 Cold Spring Harbor Laboratory Press.)

Cross-linked chains of polyacrylamide can be used as electrically neutral gels to separate double-stranded DNA fragments according to size and single-stranded DNAs according to size and conformation. Polyacrylamide gels have the following three major advantages over agarose gels: (1) Their resolving power is so great that they can separate molecules of DNA whose lengths differ by as little as 0.1% (i.e., 1 bp in 1000 bp). (2) They can accommodate much larger quantities of DNA than agarose gels. Up to 10 µg of DNA can be applied to a single slot (1 cm × 1 mm) of a typical polyacrylamide gel without significant loss of resolution. (3) DNA recovered from polyacrylamide gels is extremely pure and can be used for the most demanding purposes (e.g., microinjection of mouse embryos). However, polyacrylamide gels have the disadvantage of being more difficult to prepare and handle than agarose gels. Methods are presented here for preparing and running nondenaturing polyacrylamide gels and for detection of DNA in these gels by staining., (© 2020 Cold Spring Harbor Laboratory Press.)

Many of the commonly used techniques in molecular cloning depend on methods to map accurately the distribution of radioactive atoms on two-dimensional (2D) surfaces. Without this ability, methods such as Southern blotting, northern hybridizations, radiolabeled DNA sequencing, and library screening would not have been possible. In the 1970s and 1980s-the pioneering days of molecular cloning-imaging of 2D surfaces was obtained using autoradiography. In this technique, β-particles emitted by radioactive specimens were recorded on X-ray film, producing a latent image that can be converted to a true image by developing and fixing the film. Autoradiography was a lot of fun, but it was also messy. In the impatient excitement of wanting to see how an experiment had turned out, people used to hold the newly developed X-ray films in their metal frames up to the darkroom light. Drips of the final wash would run down their arms, clothes would be stained, and shoes ruined. It is hardly surprising that autoradiography was quickly abandoned when sensitive phosphorimagers came onto the market at the end of the 1990s., (© 2020 Cold Spring Harbor Laboratory Press.)

Bands of radioactive DNA separated by polyacrylamide gel electrophoresis may be detected by autoradiography or phosphorimaging. Analytical polyacrylamide gels containing radioactive DNA are usually fixed and dried before autoradiography. However, if bands of radioactive DNA are to be recovered from the gel, the gel should generally not be fixed or dried., (© 2020 Cold Spring Harbor Laboratory Press.)

This protocol describes procedures for cloning blunt-ended DNA fragments into linearized plasmid vectors. To obtain the maximum number of "correct" ligation products when cloning blunt-ended target fragments, the two components of DNA in the ligation reaction must be present at an appropriate ratio. If the molar ratio of plasmid vector to target DNA is too high, then the ligation reaction may generate an undesirable number of circular empty plasmids, both monomeric and polymeric; if too low, the ligation reaction may generate an excess of linear and circular homopolymers and heteropolymers of varying sizes, orientations, and compositions. For this reason, the orientation of the foreign DNA and the number of inserts in each recombinant clone must always be validated by restriction endonuclease mapping or some other means., (© 2020 Cold Spring Harbor Laboratory Press.)

This protocol presents a method to remove 5'-phosphate residues from protruding or blunt termini of linearized plasmids. This suppresses the recircularization of plasmid DNA during subsequent ligation procedures., (© 2020 Cold Spring Harbor Laboratory Press.)

This protocol describes the standard, old-fashioned but reliable procedure for cloning linear DNA fragments whose ends are incompatible with each other but are compatible with those of the linearized vector., (© 2020 Cold Spring Harbor Laboratory Press.)

A typical mammalian cell contains ∼10 -5 µg of RNA, 80%-85% of which is ribosomal RNA (rRNA; chiefly the 28S, 18S, 5.8S, and 5S species). Most of the remaining 15%-20% consists of a variety of low-molecular-weight species (e.g., transfer RNAs [tRNAs] and small nuclear RNAs). These abundant RNAs are of defined size and sequence. In contrast, messenger RNA (mRNA), which makes up between 1% and 5% of the total cellular RNA, is heterogeneous in both size-from a few hundred bases to many kilobases in length-and sequence. In this introduction, the development and use of monophasic lysis reagents for the isolation of total RNA from eukaryotic cells is discussed., (© 2020 Cold Spring Harbor Laboratory Press.)

Of the methods available for extracting total RNA from Drosophila , monophasic lysis reagents (such as TRIzol) yield consistent and reliable results and are considerably cheaper than other kit-based techniques for RNA isolation. In this protocol, up to 50 mg of Drosophila tissue is homogenized in a microcentrifuge tube using a plastic pestle in the presence of the monophasic lysis reagent. For larger amounts of starting material, homogenization can be performed using a mortar and pestle or a Dounce homogenizer, and the rest of the protocol can be readily scaled up., (© 2020 Cold Spring Harbor Laboratory Press.)

This protocol is for isolating total RNA from either larvae (L1-L4) or adult worms using a monophasic lysis reagent. It is preferable when multiple samples have to be processed., (© 2020 Cold Spring Harbor Laboratory Press.)

Several types of alkaline phosphatases (or alkaline phosphomonoesterase) are commonly used in molecular cloning, including bacterial alkaline phosphatase (BAP) and calf intestinal alkaline phosphatase (CIP, CIAP, or CAP). Similar enzymes isolated from more esoteric cold-blooded organisms (e.g., SAP from shrimp) have become available in recent years and have the advantage of being easier to inactivate than BAP or CIP at the end of dephosphorylation reactions. The uses and properties of these enzymes are introduced here., (© 2020 Cold Spring Harbor Laboratory Press.)

The removal of 5' phosphates from nucleic acids is used to enhance subsequent labeling with [γ- 32 P]-ATP, reduce the circularization of plasmid vectors in ligation reactions, and render DNA susceptible or resistant to other enzymes that act on nucleic acids (e.g., λ exonuclease). Essentially, any nucleotide phosphatase (e.g., bacterial alkaline phosphatase, calf intestinal alkaline phosphatase [CIP], placental alkaline phosphatase, shrimp alkaline phosphatase [SAP], or several acid phosphatases such as sweet potato and prostate acid phosphatase) will catalyze the removal of 5' phosphates from nucleic acid templates. In fact, these enzymes prefer small substrates such as p -nitrophenyl phosphate (PNPP) and the exposed 5' phosphates of nucleic acids to bulky globular protein substrates., (© 2020 Cold Spring Harbor Laboratory Press.)

In this method, E. coli DNA Pol I binds to a nick or short gap in duplex DNA. The 5' → 3' exonuclease activity of Pol I then removes nucleotides from one strand of the DNA, creating a template for the synthesis of DNA by the 5' → 3' polymerase activity of Pol I. The simultaneous elimination of nucleotides from the 5' side and the addition of nucleotides to the 3' side result in movement of the nick (nick translation) along the DNA, which becomes labeled to high specific activity., (© 2020 Cold Spring Harbor Laboratory Press.)

Preparing electrocompetent bacteria is considerably easier than preparing cells for transformation by chemical methods. Bacteria are simply grown to mid-log phase, chilled, centrifuged, washed extensively with ice-cold buffer or H 2 O to reduce the ionic strength of the cell suspension, and then suspended in an ice-cold buffer containing 10% glycerol. DNA may be introduced immediately into the bacteria by exposing them to a short high-voltage electrical discharge. Alternatively, the cell suspension may be snap-frozen and stored at -70°C for up to 6 mo before electroporation, without loss of transforming efficiency., (© 2020 Cold Spring Harbor Laboratory Press.)