bluuebunny/crossref_metadata_2025 huggingface dataset:
Phosphatases and Kinases
Authors: Tabor Stanley
Published: June 2004

The reaction conditions and applications of two phosphatases and one kinase are described in this unit. Bacterial alkaline phosphatase (BAP) from E. coli and calf intestine phosphatase (CIP) from veal are commonly used in nucleic acid research. Both enzymes catalyze the hydrolysis of 5′‐phosphate residues from DNA, RNA, and ribo‐ and deoxyribonucleoside triphosphates. The dephosphorylated products possess 5′‐hydroxyl termini which can subsequently be radioactively labeled using [γ‐32P]ATP and T4 polynucleotide kinase. T4 polynucleotide kinase has 3 activities: the forward reaction efficiently catalyzes the transfer of the terminal (γ) phosphate of ATP to the 5′‐hydroxyl termini of DNA and RNA. The exchange reaction catalyzes the exchange of 5'‐terminal phosphates. Lastly, T4 polynucleotide kinase is a 3′ phosphatase.
Exonucleases
Authors: Tabor Stanley
Published: June 2004

Reaction conditions for numerous exonucleases are detailed in this unit along with discussions of potential applications. Single‐stranded and double‐stranded 5′ => 3′ and 3′ => 5′ exonucleases are included.
Endonucleases
Authors: Tabor Stanley , Struhl Kevin
Published: June 2004

Reaction conditions for numerous ensonucleases are detailed in this unit along with discussions of potential applications. Specific enzymes include BAL 31 nuclease, S1 nuclease, mung bean nuclease, micrococcal nuclease and DNase I.
Ribonucleases
Authors: Struhl Kevin
Published: June 2004

Ribonucleases (RNases) with different sequence specificities are used for a variety of analytical purposes, including RNA sequencing, mapping, and quantitation. One very common application for RNase A is presented in this unit and involves hydrolyzing RNA that contaminates DNA preparations. Two other commonly used RNases, RNase H and RNase T1, are also described. In addition, many commercially available RNases are sequence‐specific endoribonucleases, a property has been used for enzymatic sequencing of RNA.
Labeling and Colorimetric Detection of Nonisotopic Probes
Authors: Boyle Ann , Perry‐O'Keefe Heather
Published: June 2004

Although a number of different nonisotopic labels have been described in the literature, biotin and digoxigenin are used most frequently and are commercially available. Either label can be easily incorporated into DNA probes and be detected colorimetrically; a number of fluorochromes, as well as alkaline phosphatase and horseradish peroxidase (which produce colored precipitates) are available directly conjugated to anti‐digoxigenin antibodies and to avidin. Chemiluminescent detection methods and indirect immunofluorescent techniques also provide sensitive alternatives for many molecular biology applications. This unit presents two protocols for incorporating biotinylated nucleotides into DNA probes by nick translation and random‐primed synthesis. A describes colorimetric detection of the probes, which also serves to check the extent of nucleotide incorporation. An describes adaptations of the basic protocols for incorporation of digoxigenin‐modified nucleotides.
Guanidine Methods for Total RNA Preparation
Authors: Kingston Robert E. , Chomczynski Piotr , Sacchi Nicoletta
Published: June 2004

Three different methods for RNA preparation using guanidine are presented in this unit ‐‐ a single‐step isolation method employing liquid‐phase separation to selectively extract total RNA from tissues and cultured cells and two methods that rely on a CsCl step gradient to isolate total RNA.
Preparation of Bacterial RNA
Authors: Reddy K.J. , Gilman Michael
Published: June 2004

Procedures for isolating RNA from bacteria involve disruption of the cells, followed by steps to separate the RNA from contaminating DNA and protein. Lysis strategies differ in the protocols presented in this unit, including chemical degradation of Gram‐negative cell walls using sucrose/detergent or lysozyme, and sonication to break open Gram‐positive cell walls. Combinations of enzymatic degradation, organic extraction, and alcohol or salt precipitation are employed in the procedures to isolate the RNA from other cellular components, and various inhibitors of ribonuclease activity (diethylpyrocarbonate, vanadyl‐ribonucleoside complex, and aurintricarboxylic acid) are described. If extremely high‐quality RNA is required (e.g., for gene expression studies), instructions are provided for CsCl step‐gradient centrifugation to remove all traces of contaminating DNA.
Preparation of Poly(A)+RNA
Authors: Kingston Robert E.
Published: June 2004

Most messenger RNAs contain a poly(A) tail, while structural RNAs do not. Poly(A) selection therefore enriches for messenger RNA. The technique has proved essential for construction of cDNA libraries. It is also useful when analyzing the structure of low‐abundance mRNAs. Removing the ribosomal and tRNAs from a preparation increases the amount of RNA that can be clearly analyzed by S1 analysis, for example, thus allowing detection of a low level message. This protocol separates poly(A)+ RNA from the remainder of total RNA, which is largely rRNA and tRNA. Total RNA is denatured to expose the poly(A) (polyadenylated) tails. Poly(A)‐containing RNA is then bound to oligo(dT) cellulose, with the remainder of the RNA washing through. The poly(A)+ RNA is eluted by removing salt from the solution, thus destabilizing the dT:rA hybrid. The column can then be repeated to remove contaminating poly(A)− RNA.
cDNA Libraries
Authors: Moore David D.
Published: June 2004

This unit provides a basic description of issues that require careful consideration before undertaking construction of cDNA libraries. Topics include: generating a double‐stranded DNA copy of the mRNA, the quality of the mRNA, the relative abundance of the clone of interest, the choice of screening method, the size of the library that will be necessary to include the clone of interest, and choices of linkers or adaptors for insertion into the vector.
Size Fractionation Using Agarose Gels
Authors: Quertermous Thomas
Published: June 2004

In the , digested genomic DNA is size fractionated on a slab agarose gel, and the appropriate region of the gel is defined by Southern blot analysis (for subgenomic libraries) or by size (for genomic libraries) and the DNA is eluted. An describes electrophoresis using a Bull's‐eye apparatus. Digested genomic DNA is loaded onto a large preparative circular agarose gel in this apparatus, the DNA fragments are electrophoresed toward the center of the gel and then eluted. Fragments leaving the gel are pooled and constitute a fraction. Fractions containing the gene of interest are identified by Southern blotting of a small aliquot of alternate fractions. This procedure allows the purification of large amounts of size‐fractionated DNA that is particularly well suited for genomic library construction and normally allows creation of large numbers of recombinant clones.
Ligation of Linkers or Adapters to Double‐Stranded cDNA
Authors: Klickstein Lloyd B. , Neve Rachael L.
Published: June 2004

The describes the strategy of producing a genomic DNA library. A number of small‐scale ligations are performed using a set amount of vector and varying amounts of insert. Test ligations are transformed into bacteria (plasmid vectors) or packaged and plated on host bacteria (λ and cosmid vectors). The number of clones in the different ligations is compared, and the optimum ratio of vector to insert is indicated by the ligation with the most recombinant clones. A large‐scale ligation is then set up using this optimum ratio. This protocol employs a bacteriophage vector; however, cosmid or plasmid vectors can be used with minor modifications.
Production of a Genomic DNA Library
Authors: Quertermous Thomas
Published: June 2004

The describes the strategy of producing a genomic DNA library. A number of small‐scale ligations are performed using a set amount of vector and varying amounts of insert. Test ligations are transformed into bacteria (plasmid vectors) or packaged and plated on host bacteria (λ and cosmid vectors). The number of clones in the different ligations is compared, and the optimum ratio of vector to insert is indicated by the ligation with the most recombinant clones. A large‐scale ligation is then set up using this optimum ratio. This protocol employs a bacteriophage vector; however, cosmid or plasmid vectors can be used with minor modifications.
Amplification of a Bacteriophage Library
Authors: Klickstein Lloyd B.
Published: June 2004

This protocol may be used for genomic DNA or cDNA libraries. A freshly packaged and titered library is adsorbed to log phase plating bacteria. The mixture is then plated at high density and allowed to grow until the plaques are just subconfluent. The phage are eluted from the plate by overnight incubation with phage buffer and the library is titered and stored.
Plating and Transferring Bacteriophage Libraries
Authors: Quertermous Thomas
Published: June 2004

The usefulness of a recombinant phage library depends on the ability to screen a large number of phage and identify the clone that carries the DNA sequence of interest. This unit presents a protocol in which phage are allowed to multiply in host bacteria in a thin layer of agarose on regular bacterial plates. When nitrocellulose is applied to the agarose, phage particles and unpackaged DNA adsorb to the filter to produce a replica of the plate surface. If the agarose surface is not excessively wet, there will be little spreading of the phage on the filter. Subsequent treatment of the filter with sodium hydroxide destroys the phage particles and denatures the phage DNA which then binds to the nitrocellulose. Neutralization of the filters is required to maintain the integrity of the nitrocellulose. Hybridization of these filters to a DNA or RNA probe will identify the location of the phage plaque of interest, which can then be recovered from the plate.
Purification of Bacteriophage Clones
Authors: Quertermous Thomas
Published: June 2004

Careful purification of the clone of interest away from contaminating phage is required before growth and characterization of the clone can proceed. It is common for a “purified” clone to be contaminated by a second phage, leading to confusing results and wasted time. Several rounds of purification should be performed even if the phage appears pure as early as the secondary screening stage. In this unit, phage plates are correctly oriented to the autoradiograph film, and a region that should contain the clone of interest is sampled by toothpicking each phage plaque onto secondary plates containing a lawn of host cells. Alternatively, a plug of agarose can be taken from the primary plate, placed in suspension medium, and this solution used to plate a small secondary library. Plaques on the secondary plates are transferred to nitrocellulose filters, hybridized to a 32P‐labeled probe, and an isolated positive plaque is picked, diluted in suspension medium, and regrown. This process is repeated until the desired plaque is purified.
Use of Monoclonal Antibodies for Expression Cloning
Authors: Hollenbaugh Diane , Aruffo Alejandro , Jones Bryan , Linsley Peter
Published: June 2004

This unit details the use of transient expression in mammalian cells to screen cDNA libraries with monoclonal antibodies (MAb) to isolate cDNA clones encoding cell‐surface and intracellular proteins. The first protocol in this unit describes the cloning of cDNAs encoding cell‐surface antigens. Several steps in this protocol involve transfection procedures that are described in greater detail elsewhere in this volume. The second protocol is a modification that facilitates isolation of cDNAs encoding antigens that are expressed intracellularly. Both protocols are designed for use with the expression vector CDM8, which contains a polylinker for subcloning double‐stranded cDNA.
DNA Sequencing Strategies
Authors: Slatko Barton E. , Eckert Richard L. , Albright Lisa M. , Ausubel Frederick M.
Published: June 2004

This unit contains a general discussion of factors that should be considered before embarking on a DNA sequencing project. In general, any sequencing strategy should include plans for sequencing both strands of the DNA fragment. Complementary strand confirmation leads to higher accuracy, especially when sequencing regions where artifacts such as “compressions” are a problem. Sequencing the opposite strand is often required to obtain accurate data for such regions. The most commonly used methods for generating appropriately sized DNA fragments for dideoxy and chemical sequencing are discussed in this unit, and the biochemistry underlying these procedures, as well as how to choose between these and alternative sequencing methods, are discussed in the introduction to this chapter.
Constructing Nested Deletions for Use in DNA Sequencing
Authors: Slatko Barton , Heinrich Peter , Nixon B. Tracy , Voytas Daniel
Published: June 2004

Nested deletions useful for dideoxy DNA sequencing are a set of deletions originating at one end of a target DNA fragment and extending various lengths along the target DNA. Each successively longer deletion brings “new“ regions of the target DNA into sequencing range (about 300 bp for normal sequencing gels) of the primer site for a general discussion of nested deletions in DNA sequencing). Two protocols for generating nested subclones via enzymatic digestion are included in this unit. In the first, a set of nested deletions is generated by exonuclease III. The primary advantage of this method is that the deletion products generated from the original clone can be recircularized to generate functional plasmids and thus do not require subcloning into another vector. An alternate method utilizes Bal 31 nuclease to generate the deletions. This method requires subcloning of the deletion fragments into a separate vector for subsequent use. Both methods require the presence of unique restriction sites in the vector that are not present in the insert DNA. Bal 31 can also be used to generate nested deletions for chemical sequencing in conjunction with specialized chemical sequencing vectors.
Preparation of Templates for DNA Sequencing
Authors: Slatko Barton E. , Heinrich Peter , Nixon B. Tracy , Eckert Richard L.
Published: June 2004

This unit contains protocols for preparing DNA suitable for use as dideoxy sequencing templates and as material for end labeling and chemical sequencing. In all protocols, the starting material contains the recombinant molecule to be sequenced. DNA from M13mp‐derived phage is easily prepared and is currently the most reliable source of template for large‐scale dideoxy sequencing projects. Because it is occasionally necessary or convenient to use a λ‐derived phage as a source of DNA, a protocol for preparing λ phage DNA from plate lysates is provided. Two protocols for minipreps of plasmid DNA are provided, one intended for dideoxy sequencing, the other for end labeling and chemical sequencing; they differ primarily in the way in which cellular RNA is removed. Alkali denaturation of double‐stranded DNA (necessary prior to annealing) is described, and a final protocol describes the preparation of template for thermal cycle sequencing from a single phage plaque or bacterial colony.
DNA Sequencing by the Chemical Method
Authors: Eckert Richard L.
Published: June 2004

This procedure is written for a novice with no chemical sequencing experience. It is assumed, however, that the investigator has experience with DNA fragment isolation and DNA labeling with [32P]dNTPs. A DNA fragment labeled at only one end with 32P or 35S is divided into four aliquots and subjected to base‐specific modification reactions. Piperidine is then added to catalyze strand scission at the modified bases. Finally, the four reactions are subjected to electrophoresis on adjacent lanes of a high‐resolution denaturing polyacrylamide gel. The gel is then autoradiographed and the sequence is read from the film.
Computer Manipulation of DNA and Protein Sequences
Authors: Cherry J. Michael
Published: June 2004

This unit outlines a variety of methods by which DNA sequences can be manipulated by computers. Procedures for entering sequence data into the computer and assembling raw sequence data into a contiguous sequence are described first, followed by a description of methods of analyzing and manipulating sequences‐‐e.g., verifying sequences, constructing restriction maps, designing oligonucleotides, identifying protein‐coding regions, and predicting secondary structures. This unit also provides information on the large amount of software available for sequence analysis.The appendix to this unit lists some of the commercial software, shareware, and free software related to DNA sequence manipulation. The goal of this unit is to serve as a starting point for researchers interested in utilizing the tremendous sequencing resources available to the computer‐knowledgeable molecular biology laboratory.
Oligonucleotide‐Directed Mutagenesis without Phenotypic Selection
Authors: Kunkel Thomas A.
Published: June 2004

A DNA sequence can be specifically altered by synthesizing the desired sequence change within an oligonucleotide, and then converting this into a biologically active circular DNA strand by using the oligonucleotide to prime in vitro synthesis on a single‐stranded circular template. This protocol uses a DNA template containing a small number of uracil residues in place of thymine. Use of the uracil‐containing template allows rapid and efficient recovery of mutants; in principle this same template can be applied to most of the other mutagenesis protocols in use.
Gene Synthesis: Assembly of Target Sequences Using Mutually Priming Long Oligonucleotides
Authors: Moore David D.
Published: June 2004

This protocol uses pairs of oligonucleotides annealed at a short duplex segment at their 3' ends as both templates and primers (mutually primed synthesis) to generate desired sequences up to 400 bp in a single step. The procedure is divided into three sections: design of the oligonucleotides, extension by mutually primed synthesis, and cloning of the extension products. The strategy for design of oligonucleotides is discussed in the commentary.
Transfection by Electroporation
Authors: Potter Huntington
Published: June 2004

Electroporation‐‐the use of high‐voltage electric shocks to introduce DNA into cells‐‐can be used with most cell types, yields a high frequency of both stable transformation and transient gene expression, and, because it requires fewer steps, can be easier than alternate techniques. In this unit, the describes the electroporation of mammalian cells and an outlines modifications for preparation and transfection of plant protoplasts.
Overview of Genetic Reporter Systems
Authors: Kain Steven R. , Ganguly Subinay
Published: June 2004

One method to gauge changes in transcription is to link a presumed cis‐acting sequence(s) from a gene of interest to the coding sequence for an unrelated reporter gene. Following introduction of the chimeric reporter construct into an appropriate cell type or animal, measurement of reporter‐gene product provides an indirect estimate of the induction in gene expression directed by the regulatory sequences. Numerous in vitro and in vivo reporters genes are discussed in this overview with regard to important issues regarding the selection of a transcription reporter gene, and their applications and limitations.
Isotopic Assays for Reporter Gene Activity
Authors: Kingston Robert E. , Sheen Jen , Moore David
Published: June 2004

This unit describes two widely used reporter systems that are based on radioactive detection assays. The first assay uses chloramphenicol acetyltransferase (CAT) activity as a measure of the level of expression of a transfected gene. This bacterial enzyme catalyzes the transfer of an acyl group from acetyl CoA (or any of several other acyl CoA cofactors) to chloramphenicol. In the assays described here, transfected cells are harvested and lysed, and then acyl CoA and radioactively labeled chloramphenicol are added to cell lysate, and modified derivatives of the antibiotic are separated from the starting material using either thin‐layer chromatography or phase‐extraction. The second reporter system uses a kit to perform a simple wo‐site radioimmunoassay to quantitate the amount of human growth hormone (hGH) secreted into culture medium by transfected cells. Medium is incubated with 125I‐labeled antibody specific for hGH, and immune complexes are collected by an avidin‐coated bead. The quantity of hormone is determined based on comparison with a standard curve.
Detection of Helper Virus in Retrovirus Stocks
Authors: Cepko Constance , Pear Warren
Published: June 2004

Helper virus is a replication‐competent virus that is sometimes present in stocks of replication‐incompetent virus. There are several types of applications in which the presence of helper virus can be problematic. If animal infections are being done, helper virus can lead to leukemia, particularly if the infection is carried out pre‐ or neonatally. If retroviruses are being used for lineage analysis, helper virus may cause horizontal spread of the marker virus, creating false lineage relationships. This unit describes protocols for the detection of helper virus by a selectable marker assay, by rescue of an integrated provirus, or by measuring reverse transcriptase activity.
Two‐Dimensional Gel Electrophoresis Using the ISO‐DALT System
Authors: Adams Lonnie D.
Published: June 2004

Two high‐resolution electrophoretic procedures (isoelectric focusing and SDS‐polyacrylamide gel electrophoresis) are combined in this unit to provide much greater resolution than either of the individual procedures. Solubilized proteins are first separated according to their isoelectric point by isoelectric focusing in a tube gel. This first dimension gel is then applied to the top of an SDS‐polyacrylamide slab gel and electrophoresed. The proteins in the first‐dimension gel migrate into the second‐dimension gel where they are further separated on the basis of their molecular size. The ISO‐DALT system was specifically designed for running multiple high‐resolution two‐dimensional gels at one time.
Two‐Dimensional Gel Electrophoresis Using the O'Farrell System
Authors: Adams Lonnie D. , Gallagher (minigels) Sean R.
Published: June 2004

Two‐dimensional gel electrophoresis is the combination of two high‐resolution electrophoretic procedures (isoelectric focusing and SDS‐polyacrylamide gel electrophoresis) to provide much greater resolution than either procedure alone. In the first‐dimension gel, solubilized proteins are separated according to their isoelectric point (pI) by isoelectric focusing. This gel is then applied to the top of an SDS‐slab gel and electrophoresed. The proteins in the first‐dimension gel migrate into the second‐dimension gel where they are separated on the basis of their molecular weight. The basic protocols in this unit are based on the type of equipment originally described by O'Farrell in . For very basic or very acidic proteins, two alternate protocols are provided. A third alternate protocol describes how two‐dimensional electrophoresis can be performed using a minigel system. Protein sample preparation is presented in the support protocol.
Staining Proteins in Gels
Authors: Sasse Joachim , Gallagher Sean R.
Published: June 2004

This unit describes protocols for detecting protein in a gel by either Coomassie blue staining or silver staining. The former is easier and more rapid; however, silver staining methods are considerably more sensitive and thus can be used to detect smaller amounts of protein. Alternate rapid staining procedures are provided for each method and a support protocol describes how to photograph stained gels. Fluorescent staining is a popular alternative to traditional staining procedures, mainly because it is more sensitive than Coomassie staining, and often as sensitive as silver staining. Staining of proteins in SDS‐polyacrylamide gels is described, and an alternate protocol details variations in the procedure for proteins in nondenaturing gels. A final support protocol describes the photography of fluorescently stained proteins.
Gel‐Filtration Chromatography
Authors: Hagel Lars
Published: June 2004

Gel filtration (GF) chromatography separates proteins solely on the basis of molecular size. Separation is achieved using a porous matrix to which the molecules, for steric reasons, have different degrees of access‐‐i.e., smaller molecules have greater access and larger molecules are excluded from the matrix. Hence, proteins are eluted from the GF column in decreasing order of size. This unit describes the experimental theory behind gel filtration and contains many useful tables listing the properties and characteristics of currently available matrices.
Ion‐Exchange Chromatography
Authors: Williams Alan , Frasca Verna
Published: June 2004

Ion‐exchange chromatography separates biomolecules on the basis of charge characteristics. Charged groups on the surface of a protein interact with oppositely charged groups immobilized on the ion‐exchange medium. This unit provides thorough explanations of the theory behind the technology, troubleshooting tips for making the procedure work best for you, and detailed listings of column matrices and separation properties.
Immunoaffinity Chromatography
Authors: Springer Timothy A.
Published: June 2004

Immunoaffinity purification is a powerful technique for isolating proteins. Purifications of 10,000‐fold or more can often be achieved in one step. This unit describes the isolation of soluble or membrane‐bound protein antigens from cells or homogenized tissue by immunoaffinity chromatography. The technique involves the elution of a single protein from an immunoaffinity column after prior elution of nonspecifically adsorbed proteins. An alternate protocol describes the use of batch purification of antigens which can shorten the column loading time.
Metal‐Chelate Affinity Chromatography
Authors: Petty Kevin J.
Published: June 2004

Recombinant proteins engineered to have six consecutive histidine residues on either the amino or carboxyl terminus can be purified using a resin containing nickel ions (Ni2+) that have been immobilized by covalently attached nitrilotriacetic acid (NTA). This technique, known as metal‐chelate affinity chromatography (MCAC), can readily be performed with either native or denatured protein. Techniques are discussed for creating a fusion protein consisting of the protein of interest with a histidine tail attached (for purification by MCAC). The basic protocol describes expression of histidine‐tail fusion proteins and their purification in native form by MCAC. Two alternate protocols describe purification of histidine‐tail fusion proteins by MCAC under denaturing conditions and their renaturation by either dialysis or solid‐phase renaturation. Support protocols are provided for analysis of the purified product and regeneration of the NTA resin. All of these protocols are easily adaptable to any protein expression system.
HPLC of Peptides and Proteins: Standard Operating Conditions
Authors: Boysen Reinhard I. , Hearn Milton T.W.
Published: June 2004

The standard operating conditions for the eight basic modes of HPLC are presented in this unit. They include: size‐exclusion chromatography (HP‐SEC), ion‐exchange chromatography (HP‐IEX), normal phase chromatography (HP‐NPC), hydrophobic interaction chromatography (HP‐HIC), reversed‐phase chromatography (RP‐HPLC), hydrophilic interaction chromatography (HP‐HILIC), immobilized metal ion affinity chromatography (HP‐IMAC), and biospecific/biomimetic affinity chromatography (HP‐BAC). In addition, some subsets of these chromatographic modes, e.g., mixed mode chromatography (HP‐MMC), charge transfer chromatography (HP‐CTC), or ligand‐exchange chromatography (HP‐LEC) are described. Procedures for multimodal column switching are also included, as are guidelines for a systematic approach to method development. Example separations help illustrate the procedures.The standard operating conditions for the eight basic modes of HPLC are presented in this unit.
Reversed‐Phase Isolation of Peptides
Authors: Henzel William J. , Stults John T.
Published: June 2004

In reversed‐phase HPLC, peptides are separated on a hydrophobic stationary phase and eluted with a gradient of increasing organic solvent concentration. Protocols describing the separation of peptides in 5‐ to 500‐pmol quantities via narrow‐bore (2‐mm‐i.d.) or microbore (1‐mm‐i.d.) columns, as well as for the separation of peptides in quantities <5 pmol are provided in this unit. Capillary HPLC columns require a gradient flow rate of 3 to 5 omponents present in a small sample prior to automated sequencing is possible via the procedures for matrix‐assisted laser desorption/ionization (MALDI) mass spectrometry and capillary electrophoresis
Purification of Recombinant Proteins and Study of Protein Interaction by Epitope Tagging
Authors: Zhang Ning , Chen Jin‐Long
Published: June 2004

A protein molecule can be engineered to include a short stretch of residues corresponding to an epitope to facilitate its subsequent biochemical and immunological analysis; a technique often referred to as “epitope tagging.” This unit presents a protocol for small‐scale immunoprecipitation of epitope‐tagged recombinant proteins expressed in transiently transfected mammalian cells. The immunoprecipitant can then be analyzed by SDS‐PAGE. An immunoprecipitation protocol is also provided that has been optimized for use with a baculovirus overexpression system. An describes how multisubunit complexes can be assembled by starting with a core protein affixed to beads via an epitope tag, and adding the other members of the complex in a stepwise manner.
Immunoprecipitation
Authors: Bonifacino Juan S. , Dell'Angelica Esteban C. , Springer Timothy A.
Published: June 2004

Immunoprecipitation is a technique in which an antigen is isolated by binding to a specific antibody attached to a sedimentable matrix. It is also used to analyze protein fractions separated by other biochemical techniques such as gel filtration or density gradient sedimentation. The source of antigen for immunoprecipitation can be unlabeled cells or tissues, metabolically or intrinsically labeled cells, or in vitro‐translated proteins. This unit describes a wide range of immunoprecipitation techniques, using either suspension or adherent cells lysed by various means (e.g., with and without detergent, using glass beads, etc.). Flow charts and figures give the user a clear‐cut explanation of the options for employing the technology.
Synthesizing Proteins In Vitro by Transcription and Translation of Cloned Genes
Authors: Struhl Kevin
Published: June 2004

The availability of a cloned gene makes it possible to synthesize the encoded protein by in vitro transcription and translation. The basis of the method presented in this unit is to clone the protein‐coding sequences into a vector containing a promoter for SP6 or T7 RNA polymerase, to produce messenger RNA by transcribing the DNA template with this enzyme, and to synthesize the desired protein (often as an 35S‐labeled species) by translation of this mRNA in vitro. A major advantage of this method is that any desired mutant protein can be generated simply by altering the DNA template.
Capillary Electrophoresis of Proteins and Peptides
Authors: Burgi Dean , Smith Alan J.
Published: June 2004

Capillary electrophoresis (CE) is a high‐resolution technique for the separation of a wide variety of molecules of biological interest such as metabolites, drugs, amino acids, nucleic acids, and carbohydrates. This unit focuses on the use of CE to separate proteins and peptides. As detailed here, CE can also be used to determine the isoelectric point of a protein, either in purified form or in a mixture, by focusing the sample in a pH gradient that is generated within the capillary during electrophoresis. In addition, CE can be used as a micropreparative technique, with either multiple separations that are pooled or a single, larger‐scale separation, for the isolation of peptides from a protease digestion (in much the same way that RP‐HPLC is currently used). In most of these examples the same capillary column can be used for all the separations. Only changes in buffer composition, ionic strength, and the presence or absence of additives are required for each specific application.
Enzyme‐Linked Immunosorbent Assays (ELISA)
Authors: Hornbeck Peter , Winston Scott E. , Fuller Steven A.
Published: June 2004

This unit describes six different ELISA systems for the detection of specific antibodies, soluble antigens, or cell‐surface antigens. In all six systems, soluble reactants are removed from solution after specifically binding to solid‐phase reactants. In the first four protocols, solid‐phase reactants are prepared by adsorbing an antigen or antibody onto plastic microtiter plates; in the next two protocols, the solid‐phase reactants are cell‐associated molecules. In all protocols, the solid‐phase reagents are incubated with secondary or tertiary reactants covalently coupled to an enzyme. Unbound conjugates are washed out and a chromogenic or fluorogenic substrate is added. As the substrate is hydrolyzed by the bound enzyme conjugate, a colored or fluorescent product is generated. Finally, the product is detected visually or with a microtiter plate reader. The amount of product generated is proportional to the amount of analysate in the test mixture. Support protocols are provided for optimizing the different ELISAs and preparing lysates for use as test antigen from bacterial cultures containing expressed protein.
Immunization of Mice
Authors: Fuller Steven A. , Takahashi Miyoko , Hurrell John G.R.
Published: June 2004

In the protocols in this unit, antigen is prepared for injection either by emulsifying an antigen solution with Freunds adjuvant or by homogenizing a polyacrylamide gel slice containing the protein antigen. Mice are immunized at 2‐ to 3‐week intervals. Test bleeds are collected 7 days after each booster immunization to monitor serum antibody levels. Mice are chosen for hybridoma fusions when a sufficient antibody titer is reached.
Cloning of Hybridoma Cell Lines by Limiting Dilution
Authors: Fuller Steven A. , Takahashi Miyoko , Hurrell John G.R.
Published: June 2004

Cloning by limiting dilution is a method based on the Poisson distribution. Dilution of cells to an appropriate number per well can maximize the proportion of wells that contain one single clone. Two or more cloning procedures are carried out until >90%
of the wells containing single clones are positive for antibody production.
Production of Monoclonal Antibody Supernatant and Ascites Fluid
Authors: Yokoyama Wayne M.
Published: June 2004

Procedures detailing the production of monoclonal antibody supernatants, including the production of larger amounts (liters) are presented in this unit. An alternate protocol for large‐scale production of hybridomas or cells (e.g., for isolation of cellular proteins) involves a similar procedure. A method for producing and obtaining ascites fluid containing the monoclonal antibody is also presented.
In Vitro Antibody Production
Authors: Mond James J. , Brunswick Mark
Published: June 2004

This unit describes the antigenic stimulation of in vitro antibody production by B cells and the subsequent measurement of secreted antibodies. A generalized system for inducing in vitro antibody production is presented along with a procedure for quantifying the number of antibody‐producing cells by plaque‐forming cell (PFC) assays: the Cunningham‐Szenberg technique and the Jerne‐Nordin technique. The assay can be modified as described to measure all classes of antibodies or to enumerate total immunoglobulin‐secreting B cells. A protocol for preparing the resting B cells by Percoll gradient centrifugation is also described.
Identification of Polyol‐Responsive Monoclonal Antibodies for Use in Immunoaffinity Chromatography
Authors: Thompson Nancy E. , Burgess Richard R.
Published: June 2004

One of the limitations of immunoaffinity chromatography as been that high‐affinity antigen‐antibody complexes are difficult to dissociate, often leading to inactivation of the protein product during elution from the immobilized antibody. As described in this unit, some antigen‐antibody complexes can be dissociated in the presence of a combination of a low‐molecular‐weight polyhydroxylated compound (polyol) and a nonchaotropic salt. These conditions seem to be generally nondenaturing and, in some cases, even protein‐stabilizing. This type of antibody is designated “polyol‐responsive.“ These antibodies can be easily identified and isolated as monoclonal antibodies (MAbs) from a typical fusion, using standard hybridoma procedures. They have proven to be very valuable reagents for the immunoaffinity purification of active, labile, multi‐subunit protein complexes.
DNase I Footprint Analysis of Protein‐DNA Binding
Authors: Brenowitz Michael , Senear Donald F. , Kingston Robert E.
Published: June 2004

Deoxyribonuclease I (DNase I) protection mapping, or footprinting, is a valuable technique for locating the specific binding sites of proteins on DNA. The basis of this assay is that bound protein protects the phosphodiester backbone of DNA from DNase I‐catalyzed hydrolysis. Binding sites are visualized by autoradiography of the DNA fragments that result from hydrolysis, following separation by electrophoresis on denaturing DNA sequencing gels. Footprinting has been developed further as a quantitative technique to determine separate binding curves for each individual protein‐binding site on the DNA. For each binding site, the total energy of binding is determined directly from that site's binding curve. For sites that interact cooperatively, simultaneous numerical analysis of all the binding curves can be used to resolve both the intrinsic binding and cooperative components of these energies.

DNase I footprint titration is described in this unit and involves (1) preparation of a singly end‐labeled DNA restriction fragment, (2) equilibration of the protein with DNA, (3) exposure of the equilibrium mixture to DNase I, and (4) electrophoretic separation on gels of the denatured hydrolysis products, followed by autoradiography. A describes (1) densitometric analysis of the autoradiograms to obtain binding data and (2) numerical analysis of the binding data to yield binding curves and equilibrium constants for the interactions at each of the separate sites. An describes the qualitative use of footprinting to identify DNA‐binding proteins in crude extracts.
Purification of DNA‐Binding Proteins Using Biotin/Streptavidin Affinity Systems
Authors: Chodosh Lewis A.
Published: June 2004

Short fragments of DNA‐either natural or formed from oligonucleotides‐containing a high‐affinity site for a DNA‐binding protein provide a powerful tool for purification. The biotin/streptavidin purification system is based on the tight and essentially irreversible complex that biotin forms with streptavidin. In this procedure, a DNA fragment is prepared that contains a high‐affinity binding site for the protein of interest, and a molecule of biotinylated nucleotide is then incorporated into one of the ends of the DNA fragment. The protein of interest binds to the DNA, and then this complex binds (via the biotin moiety) to the tetrameric protein streptavidin. Next, the protein/biotinylated fragment/streptavidin ternary complex is efficiently isolated by adsorption onto a biotin‐containing resin. Since streptavidin is multivalent, it is able to serve as a bridge between the biotinylated DNA fragment and the biotin‐containing resin. Proteins remaining in the supernatant are removed by washing, and the resin‐bound protein is then eluted with a high‐salt buffer. An alternate protocol describes a microcolumn method that is useful for larger volumes of biotin‐cellulose resin. This method is also used to elute the protein in as small a volume (i.e., as high a concentration) as possible. Another variation on the basic procedure is provided in which streptavidin‐agarose is employed.
Detection, Purification, and Characterization of cDNA Clones Encoding DNA‐Binding Proteins
Authors: Singh Harinder
Published: June 2004

In this unit, an appropriate recombinant clone is detected in an expression library with a DNA recognition‐site probe, purified, and shown to encode a DNA‐binding domain of defined sequence specificity. The strategy described below obviates purification of a sequence‐specific DNA‐binding protein for the purpose of isolating its gene; it simply requires a suitable cDNA library constructed in the expression vector lgt11 and a DNA recognition‐site probe. The basic protocol enables the detection and purification of clones encoding sequence‐specific DNA‐binding proteins. The alternate protocol describes a denaturation/renaturation procedure that can increase detection of certain clones. The support protocol provides a rapid means of characterizing the DNA‐binding activities of the proteins encoded by the cloned cDNAs.
Rapid Separation of Protein‐Bound DNA from Free DNA Using Nitrocellulose Filters
Authors: Papoulas Ophelia
Published: June 2004

Nitrocellulose binds proteins but not double‐stranded DNA. Use of radioactively labeled double‐stranded DNA fragments allows quantitation of DNA bound to the protein, permitting kinetic and equilibrium studies of DNA‐binding interactions. In the basic procedure, purified protein is mixed with double‐stranded DNA and then the mixture is filtered through nitrocellulose, allowing unbound DNA to pass through the filter while the protein (and any DNA interacting with it) is retained. When the binding site of a protein is unknown, the pure protein can be added to a mixture of fragments to select those fragments of DNA for which it has the greatest affinity. Specificity of binding can be influenced by the buffer conditions and filtering regimen. An is provided that creates conditions that disrupt weaker, presumably nonspecific binding interactions, while retaining the stronger binding interactions. The goal is to recover enough of a single input fragment to visualize by subsequent autoradiography. In some cases the quantitation (by scintillation counting) of DNA retained is not sufficient information. A describes how the DNA can be recovered from the filters for further analysis by gel electrophoresis or amplification and cloning.
Purification of Sequence‐Specific DNA‐Binding Proteins by Affinity Chromatography
Authors: Kerrigan Leslie A. , Kadonaga James T.
Published: June 2004

Affinity chromatography is a very effective and straightforward means of purifying a protein based on its sequence‐specific DNA‐binding properties. The affinity chromatography procedure described in this unit uses DNA containing specific recognition sites for the desired protein that has been covalently linked to a solid support. The first basic protocol describes preparation of a DNA affinity resin, including cyanogen bromide (CNBr) activation of the agarose support. An provides a method to couple DNA to commercially available CNBr‐activated Sepharose, and a support protocol describes how to purify crude synthetic oligonucleotides by gel electrophoresis prior to preparation of the affinity resin. The second basic protocol outlines the affinity chromatography procedure. A second support protocol describes determination of the appropriate type and quantity of nonspecific competitor DNA that should be used in the procedure and its preparation. Parameters essential to the success of an affinity chromatography experiment are discussed in detail in the Commentary.
Yeast One‐Hybrid Screening for DNA‐Protein Interactions
Authors: Ouwerkerk Pieter B.F. , Meijer Annemarie H.
Published: June 2004

One‐hybrid screening in yeast is a powerful method to rapidly identify heterologous transcription factors that can interact with a specific regulatory DNA sequence of interest (the bait sequence). In this technique, the interaction between two proteins (bait and prey) is detected via in vivo reconstitution of a transcriptional activator that turns on expression of a reporter gene. Detection is based on the interaction of a transcription factor (prey) with a bait DNA sequence upstream of a reporter gene. To ensure that DNA binding results in reporter‐gene activation, cDNA expression libraries are used to produce hybrids between the prey and a strong trans‐activating domain. The advantage of cloning transcription factors or other DNA‐binding proteins via one‐hybrid screenings, compared to biochemical techniques, is that the procedure does not require specific optimization of in vitro conditions.
Yeast Vectors and Assays for Expression of Cloned Genes
Authors: Reynolds Ann , Lundblad Victoria , Dorris David , Keaveney Marie
Published: June 2004

This unit describes some of the most commonly used yeast vectors, as well as the cloned yeast genes that form the basis for these plasmids. Yeast vectors can be grouped into five general classes, based on their mode of replication in yeast: YIp, YRp, YCp, YEp, and YLp plasmids. With the exception of the YLp plasmids (yeast linear plasmids), all of these plasmids can be maintained in E. coli as well as in S. cerevisiae and thus are referred to as shuttle vectors. The nomenclature of different classes of yeast vectors, as well as details about their mode of replication in yeast are discussed.
Cloning Yeast Genes by Complementation
Authors: Lundblad Victoria
Published: June 2004

This unit presents a generalized protocol and describes the principles involved in cloning yeast genes by complementation in yeast. The protocol is presented using a hypothetical mutation of yeast, the cdc101‐1 mutation. This mutation was isolated as a cell cycle mutant and is both recessive and temperature‐sensitive for growth: it can grow relatively normally at 30oC but is unable to make a colony at 37oC. A genomic DNA clone that complements this mutation will be isolated by transforming the cdc101‐1 strain with a yeast genomic library and subsequently screening for temperature‐resistant colonies. Once isolated, two steps are necessary to prove that the insert present on the plasmid contains the wild‐type CDC101 gene. First, segregation of the complementing plasmid must result in co‐loss of both the plasmid‐borne selectable marker and the complementing phenotype, demonstrating that the observed complementation is plasmid‐specific and is not due to reversion of the
cdc101‐1 mutation. Second, it must be ruled out whether the cloned gene encodes a phenotypic suppressor of the mutation, rather than the wild‐type gene. This is done via a complementation test, which demonstrates whether or not a disruption of the cloned gene that is integrated into the genome can complement the original mutation.
Manipulation of Cloned Yeast DNA
Authors: Lundblad Victoria , Hartzog Grant , Moqtaderi Zarmik
Published: June 2004

A major advantage of working with yeast is the ability to replace the wild‐type chromosomal copy of a gene with a mutant derivative that is constructed in vitro using a cloned copy of the gene. This techniqueunavailable in most other eukaryotesallows the phenotype of the mutation to be studied under accurate in vivo conditions, with the mutation present in single copy at its normal chromosomal location. In the first protocol, a plasmid harboring both a selectable marker and a cloned gene of interest is integrated at the chromosomal location of the cloned gene via homologous recombination (integrative trandformation). Four methods are described for constructing a mutation in vitro in a cloned gene and reintroducing this mutation at the correct chromosomal site. This allows assessment of the genetic consequences of a mutation, and is often used to determine whether or not a gene is essential (by determining if a complete gene deletion is viable). Two of these techniquesintegrative disruption and one‐step gene disruptiongenerate either insertion or deletion mutations. The third techniquetransplacementis more generally applicable: it can be used to introduce insertion or deletion mutations containing a selectable marker, but it can also be used to introduce nonselectable mutations, such as conditional lethal mutations in an essential gene. Protocols are also provided to allow creation of modified genes by one‐step integrative replacement, and also conditional alleles by a copper‐inducible double‐shutoff procedure.