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Structural data on biological macromolecules provide invaluable insights into the interactions of proteins with nucleic acids. Data obtained at atomic resolution by X-ray diffraction or nuclear magnetic resonance (NMR) studies ultimately describes the exact folding of polypeptide chains and the contacts between proteins and DNA. However, many complexes are difficult to analyze at atomic resolution because either they are too large or too difficult to crystallize in three dimensions (3-D). Recent progress in electron microscopy, specimen preservation, and image processing provides the possibility to calculate detailed molecular envelopes that are complementary to X-ray crystallography with little theoretical limit on specimen size. In the case of membrane proteins organized into one-molecule-thick two-dimensional (2-D) arrays, atomic models could be elaborated from electron microscopy data (1,2). It is beyond the scope of this chapter to describe in detail the electron crystallographic methods and computer image analysis required to calculate a 3-D model of a crystallized protein complex; these aspects are described elsewhere (3). Here, we will focus on the formation of 2-D crystals of soluble proteins, an essential preliminary step in high-resolution structure determination by electron microscopy and provide the necessary information to set up, screen, and evaluate crystallization experiments. [ABSTRACT FROM AUTHOR]

The many detailed, though static, structures of protein-nucleic acid complexes have revealed the underlying structural determinants of the binding process: a high surface complementarity and a precise orientation of interacting roups. However, another important question not yet fully answered is why the molecules interact with each other. To answer such a question means that we have to rationalize a structure energetically. This requires a deconvolution of the relatively large negative value of the free energy of association (ΔG) into its enthalpic (ΔH) and entropic (ΔS) contributions. Only then can one appreciate the true nature of the forces that drive the interaction of protein and nucleic acid. Despite some promising theoretical developments and the steady accumulation of experimental data, the deconvolution of ΔG into its components remains a difficult task [ABSTRACT FROM AUTHOR]

The asymmetric carbon atoms present in the sugars of nucleotides and in all the amino acids (with the exception of glycine) results in nucleic acids and proteins displaying optical activity. Further contributions to the optical activity of the polymers result from their ability to form well-defined secondary structures, in particular helices, which themselves possess asymmetry. As a consequence, circular dichroism (CD) has found widespread use in secondary structure prediction of proteins (1). Similar studies, though less widespread, have sought to correlate structural parameters of DNA with their CD spectrum (2), with some success particularly in assigning quaternary structures to nucleic acids (e.g., in the case of DNA triplexes and G-quartet mediated structures) (3,4). It follows that the disruption of secondary structure by, for example, denaturation or ligand binding can be usefully followed by circular dichroism. [ABSTRACT FROM AUTHOR]

The scanning transmission electron microscope (STEM) at Brookhaven National Laboratory* (BNL) is nearly unique in its ability not only to image isolated unstained and unshadowed biological molecules but also to obtain quantitative information about them such as their oligomeric state (1) (see Note 1). This makes it ideal, in principle, for looking at protein-DNA complexes and obtaining information about the masses bound. However, there are very stringent requirements on the purity and stability of the samples. [ABSTRACT FROM AUTHOR]

DNA-protein interactions are instrumental in the replication, repair, recombination, and expression of genetic information. Their study under physiological conditions requires rapid methods that do not cause extensive cell damage. Among the available techniques, crosslinking by ultraviolet (UV) laser irradiation seems best placed to fulfill these requirements, but the crosslinking yield is still relatively low with the commonly used pulses in the nanosecond (ns) range (seeChapter 27). To attain higher crosslinking yields, laser parameters must be adjusted to the underlying two-photon process, which strongly depends on pulse intensity and pulse length (1). The highest crosslink yields are achieved using femtosecond (fs) laser pulses, but these also result in higher DNA damage (1). In order to reduce the UV-induced DNA damage, we have applied a combination of UV pulses for the first excitation step and blue pulses for the second step (2). This strategy has the advantage that the intensity of the UV pulse in the first step can be kept low, thus reducing DNA damage caused by the UV photons. High crosslinking efficiency can, however, still be attained by applying the second pulse with a very short time delay and at a visible wavelength too long to excite DNA bases from the ground state. This second pulse cannot alone damage DNA, but it can provide enough additional energy for the UV excited bases to pass their ionization threshold, leading to crosslinking. [ABSTRACT FROM AUTHOR]

Interactions of proteins with nucleic acids play important roles in biological phenomenon. Almost every stage in the regulation of gene expression involves the interaction of proteins with specific nucleic acids sequences. The identification of the nucleic acid recognition sequence of a given DNA-binding protein is therefore often the first step to be undertaken in the study of its biological function. Over the last 10 yr, the SELEX procedure (systematic evolution of ligands by exponential enrichment) has been used to identify high-affinity nucleic acids ligands for a large number of different proteins. The method was first described for the identification of the DNA and RNA target sequences of nucleic-acid-binding proteins (1,2) but has since been used for the selection of the nucleic acid sequence ligands for other kinds of molecules (3). SELEX uses the power of genetic selection while taking advantage of in vitro biochemistry. It is a rapid technique that is relatively easy to implement and can accelerate and simplify nucleic-acid/protein interaction studies. [ABSTRACT FROM AUTHOR]

Surface plasmon resonance (SPR) measures refractive index changes (Δn) at or near a surface and relates these to changes in mass at the surface Fig 1). This relationship is given by the Clausius Mossotti form (Eq. 2) of the Debye equation (Eq. 1): 1 where ɛ is the real part of the dielectric constant or permittivity constant related to the refractive index by ɛ = n2, N is the number density given by NAρ/Ma (NA is Avogadro's number, ρ is the density and Ma is the molecular mass). It is assumed that Δn/ΔC is a constant. [ABSTRACT FROM AUTHOR]

There are a number of proteins involved in DNA replication, recombination, or repair that bind stoichiometrically to single DNA strands irrespective of the nucleotide sequence, and some of these proteins also bind to single-stranded RNA. Some of the best known examples are the ssb protein of Escherichia coli, the gene 32 protein of phage T4, the gene 5 protein of the M13/fd/f1 filamentous bacterial viruses, and the more recently isolated human replication protein A (1-4). Complexes formed by these proteins contain protein bound to the nucleic acid at defined ratios of the number of nucleotides per molecule of bound protein; the ratios are determined by the interactive properties of the protein. These ratios, and the structures of the complexes that are formed, may vary with factors such as changes in solution conditions that alter the binding properties of the proteins. [ABSTRACT FROM AUTHOR]

Atomic force microscopy (AFM; also called scanning force microscopy [SFM]) is a rather novel technique that offers unique advantages in the potential for the very high resolution of DNA and small ligands in the absence of stains, shadows, and labels (1,2). Furthermore, the scanning can be performed in air or liquid. The latter is particularly important for resolving fully hydrated structures. The AFM is theoretically capable of resolving structural details at the level of atomic dimensions, provided that the specimen is not dynamic. [ABSTRACT FROM AUTHOR]

Most transcription activator proteins have three important features that can be probed at the molecular level: they bind to specific sequences near promoters, they can be interconverted between active and inactive forms by covalent or noncovalent modification, and, when bound at target promoters, they can stimulate the initiation of transcription by RNA polymerase (1). This chapter is concerned with in vitro methods for measuring the transcription activation function of this important class of proteins. In most cases, these methods are applied to activators that have been substantially purified, and for which the target sequences are known and the binding sites characterized (other chapters in this volume cover methods for locating and investigating the binding sites for such activators). Here we are concerned with the measurement of the products of activation. Because Escherichia coli transcription activators have been studied more than any others, we will take these as the paradigm, though, in principle, the techniques can be applied to any organism for which in vitro systems have been developed. [ABSTRACT FROM AUTHOR]

An increasing number of structural studies are aimed at identifying the principles that govern protein-DNA recognition in gene regulation (1). This work depends on the successful reconstitution of protein-DNA complexes from their purified components. X-ray crystallography and two-dimensional nuclear magnetic resonance (NMR) techniques require large amounts of pure protein and DNA. These can be supplied through expression in bacteria of the cDNA coding for intact proteins or their smaller DNA-binding domains and the automated chemical synthesis of DNA in the laboratory. [ABSTRACT FROM AUTHOR]

Understanding the underlying structural and physico-chemical basis for the recognition of specific DNA sequences by regulatory proteins is a central goal of modern biochemical genetics. A method for the rapid identification of mutant molecules altered in the affinity and/or specificity of such interactions could be a powerful tool in the hands of those studying this difficult problem. Conventional genetic approaches for obtaining and analyzing interesting mutant forms of specific DNA-binding proteins are often infeasible because of the genetic intractability of the species being studied or as a result of difficulties in identifying relevant and specific phenotypes associated with alterations in the interaction under investigation. High-resolution genetic analysis of DNA-protein interactions is particularly problematic in metazoans. We have devised an approach that makes use of the modularity in structure and function of eukaryotic transcription factors (1), the power of the polymerase chain reaction (PCR) to generate specific DNA fragments with defined levels of mutagenesis in vitro (2,3), and the recombinogenic potential of S. cerevisiae (2,4) to carry out a high-resolution genetic analysis of the sequence-specific DNA-binding properties of Xenopus transcription factor IIIA (TFIIIA) (5). It seems likely that this approach will be generally applicable to the study of many DNA-protein interactions. [ABSTRACT FROM AUTHOR]

Southwestern blotting was first described by Bowen et al. (1) and was used to identify DNA-binding proteins that specifically interact with a chosen DNA fragment in a sequence-specific manner. In this technique, mixtures of proteins such as crude nuclear extracts or partially purified preparations are first fractionated on a sodium dodecyl sulfate (SDS) denaturing gel; the gel is then equilibrated in a SDS-free buffer to remove detergent and the proteins transferred by electroblotting to an immobilizing membrane. During the transfer the proteins renature and hence DNA-binding proteins may subsequently be detected on the membrane by their ability to bind radiolabeled DNA. Fractionation of crude nuclear extracts on an SDS gel followed by electroblotting and analysis for sequence-specific DNA binding directly on the blot combines the advantages of a high-resolution fractionation step with the ability to rapidly analyze for a large number of different DNA-binding specificities. [ABSTRACT FROM AUTHOR]

Changes in the fluorescence emission spectrum of a protein upon binding to DNA can often be used to determine the stoichiometry of binding and equilibrium binding constants; in some cases the data can also give an indication of the location of particular residues within the protein. The experiments are generally quick and easy to perform, requiring only small quantities of material (1). Spectroscopic techniques allow one to measure binding at equilibrium (unlike, for example, gel retardation assays and other separation techniques that are strictly nonequilibrium methods). Fluorescence is one of the most sensitive of spectroscopic techniques, allowing the low concentrations (typically in the nanomolar to micromolar range) required for estimation of binding constants for many protein-DNA interactions. Considerable care, however, needs to be exercised in the experiment itself and in the interpretation of results. The fundamental principles of fluorescence are discussed briefly in the remainder of the Introduction. [ABSTRACT FROM AUTHOR]

Type II restriction endonucleases are familiar to most investigators in the biological sciences as indispensable reagents for a variety of molecular biology techniques. Usually, these dimeric enzymes cut double-stranded DNA at defined, palindromic, sequences 4, 6, or 8 base pairs in length (1). Restriction endonucleases have extremely high specificities, a key property that underlies their applications in genetic engineering and which has attracted the attention of scientists interested in sequence-specific DNA discrimination. Considerable effort has been expended in trying to elucidate the mechanisms that underlie specificity, often using the "structural perturbation" approach (2,3). Here, an alteration is made to the protein by site-directed mutagen-esis, or to the nucleic acid by substituting with natural base pairs, or with synthetic base, sugar and phosphate analogs and the effects on binding and cleavage observed. A useful measure of the perturbation can be obtained by evaluation of the parameter (kst/KD)modified/(kst/KD)unmodified where "unmodified" refers to the native endonuclease and the natural DNA target sequence and "modified" indicates that either the protein or the nucleic acid has been changed in order to delete a selected interaction. The parameter kst is the first-order rate constant obtained under single-turnover conditions. Provided that the association of DNA and the essential cofactor Mg2+with the protein are rapid (see Note 1), kst measures the slowest step following the assembly of the endonuclease-DNA-Mg2+complex up to and including the hydrolysis step. KD (the equilibrium dissociation constant). [ABSTRACT FROM AUTHOR]

Interference studies are just the inverse approach of "footprinting" experiments. In one type of experiment, the effect of a chemical modification of a single base on the subsequent binding of a sequence-specific protein is determined, whereas in the other, it is the accessibility of the protein-bound DNA to modification that is determined. Thus, the experiments necessarily differ in the order in which the protein binding and DNA modification steps occur. The "interference" approach is characterized first by chemical modification of the DNA and by subsequent protein binding. Such studies provide information on the change of the binding strength following single-base modification. This change can either be positive or negative and can be quantified by the gel shift assay (1) (seeChapters 2 and 5). [ABSTRACT FROM AUTHOR]

In the phage display method, peptides (1) or protein domains (2,3) cloned as fusions to the coat proteins of filamentous bacteriophage are displayed on the capsid, which encloses the viral genome. Proteins of interest and their associated phage can be selected from a large pool of variants (a library) by affinity purification using an appropriate ligand bound to a solid support. Thus, while weakly interacting phage are removed by washing, strongly bound phage are retained and can be subsequently amplified by passage through a bacterial host. Sequential rounds of selection and amplification lead to enrichment of those clones with the highest affinity for the target ligand. The identities of these clones can then be deduced by sequencing part of the phage genome. [ABSTRACT FROM AUTHOR]

Bending of DNA by proteins plays an important role in transcription initiation, DNA replication, and recombination. The degree of protein-induced DNA-bending can be simply and conveniently determined by combining gel electrophoresis of DNA-protein complexes with the use of special plasmid vectors carrying the bendable DNA sequence (1,2). The vectors contain duplicate sets of restriction sites in a direct repeat order and cloning sites for insertion of protein-binding sequences between the two sets. Restriction enzyme digestion readily generates DNA fragments, which are identical in size but differ in the location of the protein-binding site Fig. 1). [ABSTRACT FROM AUTHOR]

Photochemical crosslinking is a powerful method for studying all types of protein-nucleic acids interactions. In particular UV-induced crosslinking has been successfully applied to the study of protein-DNA interactions (e.g., ref. 1, seeChapters 23-26 and 43). Ultraviolet (UV) light is a zero-length crosslinking agent. It is therefore not subject to the steric problems that can be associated with chemical crosslinking agents and provides strong evidence for close protein-DNA interactions. However, crosslinking with conventional UV-light sources requires exposure times ranging from minutes to several hours (e.g., seerefs. 1-3), permitting protein redistribution and the crosslinking of UV-damaged molecules. Because UV-laser irradiation is intense enough to induce crosslinking after very short exposure times, artifactual crosslinking can be avoided. The typically nanosecond or picosecond exposures times also allow UV-laser-induced crosslinking to be applied to study the intermediate states in rapid protein-DNA binding reactions (4,5). [ABSTRACT FROM AUTHOR]

Site-specific protein-DNA photocrosslinking has proved to be the method of choice for analysis of the formation of nucleoprotein complexes such as those involved in transcription by mammalian RNA polymerase II (RNA Pol II). The method has two principal advantages. First, it yields structural information on large, multisubunit complexes that in general cannot be analyzed using standard high-resolution techniques such as X-ray crystallography or nuclear magnetic resonance (NMR). For example, site-specific protein-DNA photocrosslinking, in conjunction with complementary methods such as protein-affinity chromatography and electron microscopy, has produced information on both the molecular organization and the composition of the RNA Pol II pre-initiation complex on promoter DNA (1-4). This complex contains RNA Pol II and the general transcription factors TBP, TFIIA, TFIIB, TFIIE, TFIIF (RAP74 and RAP30), and TFIIH, and is composed of more than 25 polypep-tides ranging in Mr from 10 to 220 kDa (5). Neither X-ray crystallography nor NMR, which can only resolve the structure of complexes containing short protein fragments bound to small pieces of promoter DNA, could provide any detailed structural information on this complex. Second, the method has sufficient technical flexibility so as to allow the rapid analysis of complexes assembled under various conditions. Over the past few years, we have analyzed a large collection of complexes assembled in the presence of various combinations of the general transcription factors (wild-type or different deletion mutants) and RNA Pol II (1-4). These experiments have enabled us to draw conclusions on the dynamics of RNA Pol II pre-initiation complex assembly and has led to the notion that isomerization of the RNA Pol II pre-initiation complex proceeds through wrapping of the promoter DNA around the enzyme (4). [ABSTRACT FROM AUTHOR]

Site-specific DNA photoaffinity labeling is a useful technique for mapping interactions of proteins with DNA in complex systems such as the yeast RNA polymerase III (Pol III) transcription complex, which consists of at least 25 different proteins (1,2). This technique allows probing of protein-DNA interactions across large stretches of DNA and can be done in relatively crude extracts. The regions or domains of the protein contacting DNA can be identified by peptide mapping of the photoaffinity labeled protein. Our discussion of DNA photoaffinity labeling will focus on (1) the synthesis of photoreactive nucleotide analogs, (2) the manner in which the photoreactive nucleotide is incorporated into DNA, and (3) experimental details of DNA photoaffinity labeling. [ABSTRACT FROM AUTHOR]

In work carried out in collaboration with the laboratory of D. Reinberg (University of Medicine and Dentistry of New Jersey), we have developed a site-specific protein-DNA photocrosslinking procedure to define positions of proteins relative to DNA in protein-DNA and multiprotein-DNA complexes (1-3). The procedure has four parts Fig. 1): 1.Chemical (4-6) and enzymatic (7) reactions are used to prepare a DNA fragment containing a photoactivatible crosslinking agent and an adjacent radiolabel incorporated at a single, defined DNA phosphate (with a 9.7 Å linker between the photoreactive atom of the crosslinking agent and the phosphorus atom of the phosphate, and with an approximately 11 Å maximum "reach" between potential crosslinking targets and the phosphorus atom of the phosphate).2.The multiprotein-DNA complex of interest is formed using the site-specifically derivatized DNA fragment, and the multiprotein-DNA complex is ultraviolet (UV)-irradiated, initiating covalent crosslinking with proteins in direct physical proximity to the photoactivatible crosslinking agent.3.Extensive nuclease digestion is performed, eliminating uncrosslinked DNA and converting crosslinked DNA to a crosslinked, radiolabeled 3-5 nucleotide "tag."4.The "tagged" proteins are identified. [ABSTRACT FROM AUTHOR]

In biological systems, photoreactive derivatives have been widely applied to study specific interactions of receptor molecules with their ligands by photoaffinity labeling (1-3). While the receptors are generally proteins (e.g., enzymes, immunoglobulins, or hormone receptors), the ligands differ widely in their molecular structure (e.g., sugars, amino acids, nucleotides, or oligomers of these compounds). [ABSTRACT FROM AUTHOR]

Limited Proteolysis is a useful structural probe for investigating the globular nature of proteins by preferentially digesting the more accessible regions often found between domains. Generally, proteases require a small region of polypeptide chain possessing conformational flexibility for accommodation in the active site (1). The regions of a protein possessing conformational flexibility are often found between tightly folded domains and are, therefore, preferential sites for proteolysis. In practice, limited proteolysis is achieved by dilution of the enzyme sufficiently so that it will only digest the most accessible regions leaving the domains intact. Digestion of protein-nucleic acid is often advantageous in that the DNA may provide steric protection of the DNAbinding domain not afforded by the free protein. The generation of domains by limited proteolysis relies directly on the tertiary structure of the protein under investigation and provides much firmer evidence for their existence than that provided by sequence homology. [ABSTRACT FROM AUTHOR]

The basic side chains of lysine residues often play essential roles in DNA-protein recognition. They are able to contribute to the overall affinity of an interaction through nonspecific charge-charge interactions with the phosphate backbone and contribute substantially to the specificity of the interaction by forming direct hydrogen bonds with functional groups on the edges of the bases. This dual role and their almost ubiquitous presence in the interface of DNA-protein complexes make them very attractive targets for chemical modification experiments. [ABSTRACT FROM AUTHOR]

Chemical modification is a powerful tool for investigating the accessibility and function of specific amino acids within folded proteins. It has provided significant information regarding the role of different amino acids at the binding sites of numerous enzymes and DNA-binding proteins. The identification of such residues by chemical modification has then often be used to plan subsequent site-directed mutagenesis experiments. These data complement those from crystallographic and nuclear magnetic resonance (NMR) studies in determining the residues located at the active site; thus, one needs to consider all these techniques when elucidating protein structure and function. For example, chemical modification of leukotriene A4 hydrolase, 3-hydroxyisobutyrate dehydrogenase, and lactate dehydrogenase (1-3) have contributed significantly to the understanding of active-site mechanisms in these proteins and in elucidating the mechanisms of DNA binding in the Fd and Pf 1 gene 5 proteins (4-5). [ABSTRACT FROM AUTHOR]

The ordered and regular packaging of eukaryotic DNA within the chromatin complex allows the efficient utilization of this substrate for nuclear processes such as DNA replication, transcription, recombination, and repair (1,2). Thus, an understanding of the organization of protein-DNA interactions and associations within the chromatin complex is a prerequisite for a complete molecular description of these processes. For example, the linker histone protein plays crucial roles in the stability and organization of the chromatin fiber (3,4). This multidomain protein undoubtedly makes complicated and diverse but poorly understood interactions within the chromatin fiber (1). Currently, there is disagreement regarding the site of association of the globular domain of this protein within the nucleosome proper (5,6). Moreover, the molecular interactions and chemical activities of its N- and C-terminal tails are most likely modulated by the multiple posttranslational phosphorylation events known to occur within these domains (11,2). Thus, the linker histone tails represent critical points for signal transduction within the chromatin complex likely to be manifested as structural alterations within chromatin. [ABSTRACT FROM AUTHOR]

Fluorescence spectroscopy is a useful technique for investigating the interaction of DNA-binding proteins with DNA. Generally, use is made of the intrinsic fluorescence of the protein arising from the aromatic amino acids, which is frequently perturbed in a DNA-protein complex (seeChapter 33). In some cases, however, changes in the intrinsic fluorescence emission of a protein arising from its interaction with nucleic acid may not be detectable. For example, if tryptophan and/or tyrosine residues are not located in the proximity of the DNA-binding site, the emission spectrum may not be perturbed by the interaction. Furthermore, in the presence of a large number of tryptophan and tyrosine residues, a relatively small perturbation in the overall emission spectrum brought about by DNA binding may be masked. [ABSTRACT FROM AUTHOR]

Structural studies of DNA-protein complexes have now made it clear that specific sequence recognition in these systems is accomplished in two ways, either directly by the formation of hydrogen bonds to base-pair edges from amino acid side chains located on a DNA-binding motif, such as a helix-turnhelix, or indirectly as a result of sequence-dependent distortions of the DNA conformation (1). These contacts occur in the context of oriented complexes between macromolecules that juxtapose the specific recognition elements. As part of these processes, proteins make a large number of contacts to the phosphodiester backbone of DNA, as was predicted from biochemical assays of the ionic strength dependence of DNA binding. [ABSTRACT FROM AUTHOR]

The in vivo analysis of DNA-protein interactions and chromatin structure can provide several kinds of critical information regarding regulation of gene expression and gene function. For example, DNA sequences spanned by nuclease-hypersensitive sites or bound by transcription factors often correspond to genetic regulatory elements. Using the ligation-mediated polymerase chain reaction (LMPCR) technology it is possible to map such DNA sequences and to demonstrate the existence of unusual DNA structures directly in living cells. LMPCR analyses can thus be used as a primary investigative tool to identify the regulatory sequences involved in gene expression. Once specific promoter sequence sites are shown to be bound by transcription factors in living cells, it is often possible to establish the identity of these factors simply by comparison with the consensus binding sites of known factors such as Sp1, AP-1, NF-1, and so forth. The identity of each factor can then be confirmed using in vitro gel shift (electrophoretic mobility shift assay [EMSA]) or footprinting assays. [ABSTRACT FROM AUTHOR]

Dimethyl sulfate (DMS) is an effective and widely used probe for sequencespecific protein-DNA interactions. It is the only probe routinely used both for in vitro (methylation protection, methylation interference) and in vivo (DMS genomic footprinting) applications because it rapidly reacts with DNA at room temperature and readily penetrates intact cells (1). DMS methylates predominantly the 7-nitrogen of guanine and the 3-nitrogen of adenine. Thus reactivity with G residues occurs in the major groove and with A residues in the minor groove. In standard Maxam and Gilbert protocols (2), the methylated bases are subsequently converted to strand breaks and displayed on sequencing gels. [ABSTRACT FROM AUTHOR]

A large number of processes within the cell, in particular the regulation of gene expression, rely on the binding of proteins to specific sites on the DNA. A primary ingredient to understanding these processes is the characterization of the protein-DNA interaction (1). Such a characterization consists in determining the position of the binding site on the DNA and measuring the affinity of the protein for this recognition site. A wide variety of footprinting techniques can accomplish this task (2,3). [ABSTRACT FROM AUTHOR]

The DNA double helix is highly malleable and, when constrained, either as a small circle or by the action of a protein, can be readily distorted from its energetically favored conformation. Such distortions may be relatively moderate, as exemplified by smooth bending (which maintains base stacking), or more extreme when this stacking can be disrupted. Deformations of this latter type include kinks, where the direction of the double helical axis is changed abruptly at a single-base step, and localized strand separation, which may be a direct consequence of protein-induced unwinding or of high negative superhelicity in free DNA. Both kinks and localized unwinding can arise transiently during the enzymatic manipulation of DNA by recombinases and by protein complexes involved in the establishment of unwound regions during the initiation of transcription or of DNA replication. [ABSTRACT FROM AUTHOR]

The interaction of cell-type-specific or inducible transcription factors with regulatory DNA sequences in gene promoters or enhancers is a pivotal step in genetic reprograming during cell proliferation and differentiation and in response to extracellular stimuli. The study of these interactions and the characterization of the factors involved are, therefore, a critical aspect of gene control. Transcription factor-DNA interactions in eukaryotes have been demonstrated by a wide variety of biochemical approaches, including deoxyribonuclease I (DNase I) and chemical nuclease footprinting (1-3) (Chapter 3), methylation protection (4) (Chapter 14), electrophoretic mobility-shift (5,6) (Chapter 2), and Southwestern (SW) assays (7) (Chapter 17). Despite their broad applicability, these techniques provide only partial information about the DNA-protein system under investigation. The first three techniques identify either the site(s) of transcription factor binding within the DNA (size and location of nucleotide stretches or atoms on individual bases) or the complexity of the binding pattern (stoichiometry), but do not yield information about the protein(s) involved. On the other hand, the SW assay reveals the relative molecular mass of renaturable (on a membrane support) active species in heterogeneous protein mixtures facilitating their identification, but fails to localize the exact target element within the probing DNA sequence. [ABSTRACT FROM AUTHOR]

It has long been known that the uranyl(VI) ion (UO22+) forms strong complexes with various inorganic and organic anions, including phosphates, and that the photochemically excited state of this ion is a very strong oxidant (1). For instance, uranyl-mediated photooxidation of alcohols has been studied in detail (2,3). It is also widely recognized that uranyl chemistry and photophysics/photochemistry are very complex. Thus monomeric UO22+is only present at low pH (pH approx 2), whereas polynuclear species and various "hydroxides," which often precipitate, form at a higher pH (4). [ABSTRACT FROM AUTHOR]

Osmium tetroxide is an foul-smelling chemical used as a fixative in electron microscopy that can also be used to modify thymidine residues within DNA (1,2). The ability of osmium tetroxide to modify DNA is very sensitive to DNA conformation. In particular, osmium tetroxide will attack thymidines that are unstacked, either because they are in a single-stranded region or for some other reason (e.g., because the DNA is bent or because it is overwound [3]). [ABSTRACT FROM AUTHOR]

The existence of cell-type specific promoter and enhancer elements has been known for several years. However, the mechanisms responsible for the remarkable specificity of such elements, in comparison to the ubiquitously active promoters and enhancers of "housekeeping" genes and DNA tumor viruses, have remained elusive until recently. Although transfection and mutagenesis experiments have taught us a great deal about the structure of cell-type-specific cis-acting elements, the breakthrough in understanding the molecular basis for the differential activity of these elements has come from the analysis of their recognition by specific DNA-binding proteins. [ABSTRACT FROM AUTHOR]

In the search for methods to explore the interaction between proteins and DNA, a plethora of footprinting techniques have been developed, many of which are discussed elsewhere in the present work. Most footprinting techniques are based on the simple premise of specific DNA regions being protected from the reagent by the bound protein or molecule of interest. However, a number of studies over the past decade have revealed remarkable distortion of the DNA molecule, including bending and strand separation, in response to the bound protein. These distortions are often within the classical footprint but are rarely detected by these classical techniques. Thus, their detection requires alternative approaches. Unlike enzymatic methods, the chemical probes diethyl pyrocarbonate (DEPC) and potassium permanganate can access and react with the entire sequence of the DNA, distinguishing DNA distortions " under the foot" of a typical footprinting experiment. Combining the information obtained from these chemical probing techniques with the spatial information afforded by traditional footprinting gives an in-depth account of the various ways proteins and other molecules interact with DNA. [ABSTRACT FROM AUTHOR]

Within the last few years footprinting techniques have become increasingly important in the study of protein-nucleic acid interactions. This is partly the result of a fast-growing number of known nucleic acid-binding proteins but also because of an increase in the available probes that can be chosen in order to tackle a specific problem. There are two major groups of probes—the chemical probes and the enzymatic probes. The enzymatic probes, such as DNase I or exonuclease III, have the advantage of acting specifically on the DNA. Chemical probes are often less specific and may also react with the protein, possibly disturbing the correct interaction of protein with DNA. For the study of very fragile protein-DNA complexes, enzymatic probes are therefore often preferable. [ABSTRACT FROM AUTHOR]

The basic principle of the DNA footprinting technique is the measurement of accessibility of the DNA using a probe. The probe can be any enzyme or a chemical reagent that is able to cut the DNA backbone. When the target DNA is a fragment containing a signal sequence for a sequence-specific binding protein, sites on the DNA that interact with the protein are inaccessible to the probe. After electrophoretic separation based on molecular weight, these inaccessible sites appear as blanks in an otherwise regular DNA cleavage pattern, thus revealing the characteristic interaction footprint for the binding protein. [ABSTRACT FROM AUTHOR]

DNase I footprinting was developed by Galas and Schmitz in 1978 as a method to study the sequence-specific binding of proteins to DNA (1). In the technique, a suitable uniquely end-labeled DNA fragment is allowed to interact with a given DNA-binding protein and then the complex partially digested with DNase I. The bound protein protects the region of the DNA with which it interacts from attack by the DNase. Subsequent molecular-weight analysis of the degraded DNA by electrophoresis and autoradiography identifies the region of protection as a gap in the otherwise continuous background of digestion products; for examples see Fig. 1. The technique can be used to determine the site of interaction of most sequence-specific DNA-binding proteins but has been most extensively applied to the study of transcription factors. Because the DNase I molecule is relatively large as compared to other footprinting agents (seeChapters 5 and 6 on the use of hydroxy radicals and diethylpyrocarbonate), its attack on the DNA is relatively easily sterically hindered. Thus, DNase I footprinting is the most likely of all the footprinting techniques to detect a specific DNA-protein interaction. This is clearly demonstrated by our studies on the transcription factor xUBF (see Fig. 1B). The xUBF interaction with the Xenopus ribosomal DNA enhancer can be easily detected by DNase I footprinting but has still not been detected by other footprinting techniques. [ABSTRACT FROM AUTHOR]

Several nuclear mechanisms involve specific DNA-protein interactions. The electrophoretic mobility shift assay (EMSA, also known as the gel mobility shift or gel retardation assay), first described almost two decades ago (1,2), provides a simple, efficient and widely used method to study such interactions. Its ease of use, its versatility, and especially its high sensitivity (10−18 mol of DNA [2]) make it a powerful method that has been successfully used in a variety of situations not only in gene regulation analyzes but also in studies of DNA replication, repair, and recombination. Although very useful for qualitative purposes, EMSA has the added advantage of being suitable for quantitative and kinetic analyzes (3). Furthermore, because of its very high sensitivity, EMSA makes it possible to resolve complexes of different protein or DNA stoichiometry (4) and even to detect conformational changes. [ABSTRACT FROM AUTHOR]

Membrane filtration has a long history in the analysis of protein-nucleic acid complex formation, having first been used to examine RNA-protein interactions (1), before being introduced to DNA-protein interaction studies by Jones and Berg in 1966 (2). The principle of the technique is straightforward. Under a wide range of buffer conditions, nucleic acids pass freely through membrane filters, whereas proteins and their bound ligands are retained. Thus, if a particular protein binds to a specific DNA sequence, passage through the filter will result in retention of a fraction of the protein-DNA complex by virtue of the protein component of the complex. The amount of DNA retained can be determined by using radioactively labeled DNA to form the complex and then determining the amount of radioactivity retained on the filter by scintillation counting. The technique can be used to analyze both binding equilibria and kinetic behavior, and if the DNA samples retained on the filter and in the filtrate are recovered for further processing, the details of the specific binding site can be probed by interference techniques. [ABSTRACT FROM AUTHOR]

Photochemical crosslinking is a powerful method for studying all types of protein-nucleic acids interactions. In particular, UV-induced crosslinking has been successfully applied to the study of protein-DNA interactions (1). Ultraviolet (UV) light is a zero-length crosslinking agent. It is therefore not subject to the steric problems that can be associated with chemical crosslinking agents and provides strong evidence for close protein-DNA interactions. However, to achieve an acceptable degree of crosslinking with conventional UV light sources, exposure times ranging from minutes to several hours have had to be used (1-3). Such prolonged irradiation allows for redistribution of proteins and the artifactual crosslinking of UV-damaged molecules, and it also precludes kinetic studies. The use of UV lasers overcomes these difficulties, since crosslinking is achieved after only nano- or picosecond exposures (4-5). [ABSTRACT FROM AUTHOR]

DNase I footprinting was developed by Galas and Schmitz in 1978 as a method to study the sequence-specific binding of proteins to DNA (1). In this technique a suitable uniquely end-labeled DNA fragment is allowed to interact with a given DNA-binding protein and then the complex is partially digested with DNase 1. The bound protein protects the region of the DNA with which it interacts from attack by the DNase. Subsequent molecular weight analysis of the degraded DNA by electrophoresis and autoradiography identifies the region of protection as a gap in the otherwise continuous background of digestion products (for examples, see Fig. 1). The technique can be used to determine the site of interaction of most sequence-specific DNAbinding proteins but has been most extensively applied to the study of transcription factors. Since the DNase I molecule is relatively large compared to other footprinting agents (seeChapters 3 and Chapters 7 in this volume), its attack on the DNA is more readily prevented by steric hindrance. Thus DNase I footprinting is the most likely of all the footprinting techniques to detect a specific DNA-protein interaction. This is clearly demonstrated by our studies on the transcription factor xUBF (see Fig. 1B). The xUBF interaction with the xenopus ribosomal DNA enhancer can be easily detected by DNase I footprinting but has not yet been detected by other footprinting techniques. [ABSTRACT FROM AUTHOR]