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Formation of DNA damage is a crucial event in carcinogenesis. Irreparable DNA lesions have the potential to cause mispairing during DNA replication, thereby giving rise to mutations. Critically important mutations in cancer-related genes, i.e., oncogenes and tumor suppressor genes, are key contributors to carcinogenesis. Theoretically, co-localization(s) of persistent DNA lesions and mutational hotspots in cancer-relevant genes can be used for causality inference. The inferred causality can be validated if a suspected carcinogen can similarly produce corresponding patterns of DNA damage and mutagenesis in vitro and/or in vivo. DNA-lesion footprinting (mapping) in conjunction with mutagenicity analysis is used for investigating cancer etiology. Ligation-mediated polymerase chain reaction (LM-PCR) is a versatile DNA-lesion footprinting technique, which enables sensitive and specific detection of DNA damage, at the level of nucleotide resolution, in genomic DNA. Here, we describe an updated protocol for LM-PCR analysis of the mammalian genome. This protocol can routinely be used for DNA-lesion footprinting of a variety of chemical and/or physical carcinogens in mammalian cells.

We present a high-resolution mass spectrometric (MS) footprinting method enabling identification of contact amino acids in protein-protein complexes. The method is based on comparing surface topologies of a free protein versus its complex with the binding partner using differential accessibility of small chemical group selective modifying reagents. Subsequent MS analysis reveals the individual amino acids selectively shielded from modification in the protein-protein complex. The current report focuses on probing interactions between full-length HIV-1 integrase and its principal cellular partner lens epithelium-derived growth factor. This method has a generic application and is particularly attractive for studying large protein-protein interactions that are less amenable for crystallographic or NMR analysis.

Bacterial promoter identification and characterization is not as straightforward as one might presume. Promoters vary widely in their similarity to the consensus recognition element sequences, in their activities, and in their utilization of transcription factors, and multiple approaches often must be used to provide a framework for understanding promoter regulation. Characterization of RNA polymerase-promoter complex formation in the absence of additional regulatory factors (basal promoter function) can provide a basis for understanding the steps in transcription initiation that are ultimately targeted by nutritional or environmental factors. Promoters can be localized using genetic approaches in vivo, but the detailed properties of the RNAP-promoter complex are studied most productively in vitro. We first describe approaches for identification of bacterial promoters and transcription start sites in vivo, including promoter-reporter fusions and primer-extension. We then describe a number of methods for characterization of RNAP-promoter complexes in vitro, including in vitro transcription, gel mobility shift assays, footprinting, and filter binding. Utilization of these methods can result in determination of not only basal promoter strength but also the rates of transcription initiation complex formation and decay.

Footprinting is a simple method for assessing the sequence selectivity of DNA-binding ligands. The method is based on the ability of the ligand to protect DNA from cleavage at its binding site. This review describes the use of DNase I and hydroxyl radicals, the most commonly used footprinting probes, in footprinting experiments. The success of a footprinting experiment depends on using an appropriate DNA substrate and we describe how these can best be chosen or designed. Although footprinting was originally developed for assessing a ligand's sequence selectivity, it can also be employed to estimate the binding strength (quantitative footprinting) and to assess the association and dissociation rate constants for slow binding reactions.

We have optimized a recombinant chromatin assembly system that properly incorporates core histones and histone H1 into a chromatin template containing a natural promoter sequence. This article provides a step-by-step procedure for expression and purification of the proteins required for assembling well-defined chromatin templates. We describe how to measure the degree of chromatin assembly in the absence and presence of histone H1 using topological analysis and how to perform micrococcal nuclease digestion to confirm H1 incorporation and determine the quality of in vitro chromatin templates. Further, we describe the use of sucrose gradient ultracentrifugation to verify that no unincorporated H1 remains as a second means for deciding on the proper H1 to core histone ratio during assembly. Additionally, we discuss the use of both yeast and Drosophila NAP-1 (yNAP-1 and dNAP-1, respectively) in the assembly of H1-containing chromatin. Finally, we provide detailed description of functional assays for investigating the mechanism of transcriptional regulation in a chromatin context (transcription, histone acetyltransferase activity, and protein association with promoter-bound complexes using immobilized chromatin templates).

In vivo footprinting techniques are useful for the identification of regulatory elements mediating transcriptional control of a gene. However, regulation of a gene can differ between the two alleles, and further steps must be taken to distinguish between the regulatory elements occupied on one allele and those used on the second allele. Many hematologic malignancies result from chromosomal translocations, which, in some cases, relocate a gene to a transcriptionally active region leading to the deregulated expression of that gene. This situation provides an example of differential expression between two alleles. In studying the t(14; 18) and t(8; 14) translocations, which involve the bcl-2 and c-myc proto-oncogenes, respectively, we have been able to identify regulatory elements important in mediating the activation of the translocated alleles and the silencing of the normal alleles. Following in vivo methylation and isolation of genomic DNA, we were able to separate the translocated and normal alleles by electrophoresis. Using the ligation-mediated polymerase chain reaction (LMPCR) technique, we could then assess protein interactions on the two different alleles. A detailed description of this methodology with examples from our studies are provided with a discussion of how these techniques may be applied to the study of other genes., ((c) 2002 Elsevier Science (USA).)

Chemical footprinting methods have been used extensively to probe the structures of biologically important RNAs at nucleotide resolution. One of these methods, hydroxyl-radical footprinting, has recently been employed to study the kinetics of RNA folding. Hydroxyl radicals can be generated by a number of different methods, including Fe(II)-EDTA complexes, synchrotron radiation, and peroxynitrous acid disproportionation. The latter two methods have been used for kinetic studies of RNA folding. We have taken advantage of rapid hydroxyl-radical generation by Fe(II)-EDTA-hydrogen peroxide solutions to develop a benchtop method to study folding kinetics of RNA complexes. This technique can be performed using commercially available chemicals, and can be used to accurately define RNA folding rate constants slower than 6 min(-1). Here we report the method and an example of time-resolved footprinting on the hairpin ribozyme, a small endoribonuclease and RNA ligase.

We describe a method to reconstitute chromatin complexes from reversed-phase high-performance liquid chromatography (HPLC)-purified histones. The complexes reconstituted in this way exhibit the same structural characteristics as their equivalent native counterparts. Furthermore, this method works independently of the acid- or salt-extracted origin of the histones used for the HPLC fractionation. The potential of this method for the reconstitution of chromatin particles consisting of sequence-defined DNA templates and well-defined histone variants and/or their posttranslationally modified isoforms is discussed.

The study of chromatin, once thought to be a purely structural matrix serving to compact the DNA of the genome into the nucleus, is of increasing value for our understanding of how DNA functions in the cell. This article provides two basic procedures for the study of chromatin in vivo. The first is a DNase I-based method for the treatment of isolated nuclei to resolve the chromatin structure of a particular region; the second employs dimethyl sulfate footprinting of whole cells in vivo to determine the binding of factors to cis elements in the locus of interest. Specific examples illustrating the techniques described are given from our work on the regulation of the yeast PHO8 gene, but have also been successfully and reliably applied to the study of many other yeast loci. These procedures make it possible to correlate the binding of a transactivator with an altered or perturbed chromatin organization at a specific locus.

The unmatched power of Drosophila genetics revealed the complex regulatory network of gene activities that governs the development of higher eukaryotes. An understanding of gene control at the level of transcription requires insight into the protein/DNA interactions that regulate transcription in the developing embryo. Genomic footprinting allows the direct visualization of these protein/DNA interactions within intact nuclei or cells. In combination with other in vivo assays such as protein/DNA crosslinking and classical biochemistry, genomic footprinting can give valuable insight into the architecture of promoters in various states of activity. In this article we summarize our experience in analyzing Drosophila embryos by genomic footprinting and describe modifications of the ligation-mediated PCR procedure that have improved this analysis. Applications of genomic footprinting to embryos are currently limited by the fact that all target nuclei must be uniform with respect to the protein/DNA interactions at the chosen site. We discuss strategies that should allow the analysis of small numbers of cells derived from heterogeneous populations and tissues.

Regulatory factors important for the developmental control of genes have been identified by genetic studies or by examining the ontological expression profiles of proteins that were originally characterized in adult tissues; direct biochemical studies of transcription factors within small amounts of embryo tissues have been limited. We have found that the ligation-mediated PCR (LM-PCR) technique can detect specific dimethylsulfate modifications in genomic DNA from as few as several thousand cells, making it technically feasible to identify protein-DNA interactions in pools of nascent embryo tissues. Herein we show that LM-PCR can reveal methylation protections on the albumin gene enhancer in embryonic mouse hepatocytes, indicating occupancy of a C/EBP factor binding site. Comparison of the in vivo protection pattern with that obtained from the in vitro analysis of different C/EBP isoforms suggests that in embryonic hepatocytes, C/EBP beta is bound to the albumin gene enhancer. Detailed protocols are provided so that the approach can be used to study other genes in developing embryos.

Analysis of the interaction of proteins with either DNA or RNA sequences by in vivo footprinting involves two steps: (i) the in situ modification of nucleic acids by the footprinting reagent and (ii) the visualization of the footprints. Ligation-mediated PCR (LM-PCR) procedures provide a level of sensitivity and specificity that is suitable for visualization of footprints of single-copy genes or low-abundance mRNAs in higher eukaryotes. In this article, we discuss several of the technical aspects of these multistep procedures that contribute to the quality of the results, particularly the parameters that affect the specificity and fidelity of the reactions: (i) the design of the primers, which is important to achieve optimal specificity; (ii) the choice of polymerases so that the amplified material represents faithfully the initial nucleic acid population; and (iii) the impact of the plateau effect within the PCR on the interpretation of the data. We then discuss aspects of in vivo nucleic acid manipulation that may affect the quality of the footprinting image, in particular the choice of the footprinting reagent and its condition of use (e.g., on intact or permeabilized cells or prepared nuclei) and the extent of nucleic acid modification. Finally, we provide detailed experimental procedures corresponding to the techniques we have developed or modified: LM-PCR, reverse ligation-mediated PCR, and nuclease treatment of RNAs in vivo.

Retinoic acid (RA) treatment of embryonal carcinoma (EC) cells initiates a cascade of alterations in gene regulation, leading to their differentiation into various cell types. In P19 EC cells RA treatment stimulates induction of the RAR beta gene, while it represses Oct3/4 gene expression. Here we present dimethylsulfate-based genomic footprinting analyses of these two genes. We found that the RAR beta promoter is not occupied prior to RA treatment, but following RA treatment all regulatory elements in this promoter become occupied. On the other hand, the Oct3/4 promoter is occupied at all three known elements before RA treatment, but this occupancy is coordinately lost following the treatment. Thus, factor occupancy coincides with expression of the genes. It is likely that the presence of factor binding or its absence revealed here represents a mechanism of the regulated expression of these genes in vivo. Our results demonstrate the power of genomic footprinting for studying regulatory events for transcription in vivo. In contrast, with in vitro protein-DNA binding assay, factors for both promoters are present in these cells regardless of RA treatment. It has been shown that RA receptor (RAR) and retinoid X receptor (RXR), by heterodimerization, mediate the RA action in EC cells. To elucidate the role of RAR/RXR heterodimers in the RAR beta promoter occupancy in vivo, genomic footprinting has been performed in P19 cells stably expressing dominant negative mutants of RXR. Two such mutants, lacking either the DNA binding domain or the C-terminal activation domain, inhibit RA induction of the RAR beta gene in these cells. RA-induced factor occupancy is also markedly inhibited at all elements in the RAR beta promoter in these cells. Our results show that binding of liganded RAR/RXR heterodimers to RARE is required for other factors to gain access to their respective elements in the promoter.

Protein-DNA interactions in mammalian cells can be analyzed at the nucleotide level of resolution by genomic sequencing techniques. The most sensitive genomic sequencing method uses ligation-mediated polymerase chain reaction (LMPCR) for signal amplification. Several footprinting methods are compatible with LMPCR. Here we describe in detail the use of UV irradiation for in vivo footprinting. The distribution of the two major types of UV-induced DNA photoproducts [cyclobutane pyrimidine dimers and (6-4) photoproducts] can be analyzed by LMPCR and a wide variety of protein-DNA contacts can be detected by analyzing both photoproducts. A comparison of UV photofootprinting data with data from experiments using other probing techniques shows that UV light has the potential to reveal all protein-DNA interactions provided that there is a dipyrimidine sequence on either DNA strand within a factor binding site. The simplicity and nondisruptiveness of this probing method together with its ability to detect a large number of different transcription factors should make UV light a generally useful tool for in vivo footprinting. We provide protocols for UV footprinting and LMPCR.

Chromatin structure is becoming increasingly recognized as important for a full understanding of gene function and cell memory. With regard to cell memory, which involves the transfer of chromatin-encoded epigenetic information from one cell generation to another, the detailed structure of metaphase chromatin is of crucial importance. In this paper we describe methods for the use of dimethyl sulfate, DNase I, and potassium permanganate for in vivo footprinting and chromatin analysis, with special emphasis on studies of metaphase cells. We review the use of ligation-mediated PCR for the analysis of chromatin, including the human phosphoglycerate kinase promoter, and also report initial studies of a matrix attachment region near the human beta-interferon gene.

The identification of relevant DNA regulatory sequences involved in transcriptional control is critical to establishing which proteins mediate cell-specific gene expression. As an approach to investigating the mechanisms of gene regulation, in vivo footprinting studies reveal protein-DNA interactions as they actually occur in situ. We have used in vivo footprinting to complement in vitro studies of the human globin locus control regions (LCRs) in erythroid cells. To further enhance the detection of protein contacts with this technique, we have modified the dimethyl sulfate-based ligation-mediated PCR (LMPCR) in vivo footprinting procedure to permit the assessment of protein binding at guanine and adenine residues, rather than exclusively at guanines. This modification, termed GA-LMPCR in vivo footprinting, was essential for the analysis of GATA-1 motifs in the alpha-LCR and HS-3 of the beta-LCR. Moreover, GA-LMPCR in vivo footprinting provided high-resolution analysis of AP-1/NF-E2 elements and revealed protein contacts at sequences that are not coincident with previously described regulatory motifs. A comprehensive discussion of the GA-LMPCR in vivo footprinting methodology and representative analyses from our studies, including GATA-1 and AP-1/NF-E2 motifs, are presented to illustrate the modified technique.