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INTRODUCTIONLarge multiprotein complexes involved in transcription generally cannot be resolved by the methods conventionally used to study cooperative DNA binding by smaller proteins. The immobilized template recruitment assay can be used to circumvent this limitation. In this procedure, a biotinylated DNA fragment is immobilized on streptavidin-coated magnetic beads. DNA-binding proteins are incubated with the immobilized template, and proteins not specifically assembled in a complex are removed by separating the immobilized template with a magnetic particle concentrator and subsequent washing of the magnetic bead/template pellet. Bound proteins are detected by immunoblotting. To detect cooperative binding, subsaturating amounts of two or more proteins or protein complexes are added to the immobilized template. Template binding of a protein in concert with its cooperative binding partner is then compared to the binding of each individual protein or protein complex alone. In addition to studying the binding of protein complexes, the immobilized promoter template can also be used in functional assays such as in vitro transcription, allowing a correlation of complex assembly with function.

INTRODUCTIONThe Mediator (Med) complex plays a key role in promoter-specific activation of transcription by RNA polymerase II (Pol II). Med is a major target of activators and can be used in many types of affinity binding and immobilized template studies to evaluate interactions with individual activators. Additionally, all of the Med subunits have been cloned and can be subjected to individual structure-function analyses to learn how a specific activator interacts with a particular subunit. This protocol presents a simple affinity-based method to purify Med complex from HeLa cells stably expressing the Flag-tagged Intersex protein.

INTRODUCTIONWhen a transient or stable transfection assay is developed for a promoter, a primary objective is to quantify promoter strength. Because transfection efficiency in such assays can be low, promoters are commonly fused to heterologous reporter genes that encode enzymes that can be quantified using highly sensitive assays. The reporter protein’s activity or fluorescence within a transfected cell population is approximately proportional to the steady-state mRNA level. Although the Escherichia coli lacZ gene, encoding β-galactosidase (β-gal), can be used as a standard reporter for monitoring the strength of a promoter or enhancer in a transient or stable transfection assay, it is predominantly used as an internal control during transient transfection experiments. When used in this manner, cells are usually transfected with the control plasmid (containing a ubiquitously active viral promoter fused to the E. coli lacZ gene) and an experimental plasmid containing another reporter gene (e.g., luciferase or chloramphenicol acetyltransferase [CAT]) under the control of the promoter or enhancer of interest. The basic colorimetric assay described here is the simplest and least expensive assay for quantifying β-gal activity. The cells are lysed and, after determining the total protein concentration in the extracts, an aliquot of the extract is mixed with the reaction substrate, O-nitrophenyl-β-D-galactopyranoside (ONPG), in a buffer containing sodium phosphate and magnesium chloride. When the yellow product becomes visible, the optical densities of the samples are determined spectrophotometrically.

INTRODUCTIONWhen a transient or stable transfection assay is developed for a promoter, a primary objective is to quantify promoter strength. Because transfection efficiency in such assays can be low, promoters are commonly fused to heterologous reporter genes that encode enzymes that can be quantified using highly sensitive assays. The reporter protein’s activity or fluorescence within a transfected cell population is approximately proportional to the steady-state mRNA level. In this protocol, cells transfected with an Escherichia coli transposon chloramphenicol acetyltransferase (CAT) reporter plasmid are lysed by repeated cycles of freezing and thawing and cellular debris is removed by centrifugation. The lysate is incubated with [14C]chloramphenicol and acetyl-coenzyme A; CAT catalyzes the acetylation of chloramphenicol. The acetylated products and the unmodified reactants are separated from the aqueous solution by organic extraction with ethyl acetate. Acetylation is monitored by autoradiography following thin-layer chromatography (TLC) to separate the acetylated from the unacetylated forms. The percent conversion of [14C]chloramphenicol to acetyl-[14C]chloramphenicol can be measured by PhosphorImager analysis of the TLC plate, by excising the radioactive spots from the TLC plate and counting in a scintillation counter, or by densitometry analysis of an autoradiograph. The acetylated 14C-labeled product can also be quantified without TLC by organic extraction and scintillation counting using reagent-grade chemicals.

INTRODUCTIONIn electroporation-based transfection methods, the DNA and cells are mixed in an empirically defined buffer, which establishes the resistance during the electric shock. The solution is placed in a cuvette that is compatible with the electroporation device being used. The mixture is then subjected to a brief high-voltage shock, which induces or stabilizes pores in the plasma membrane through which the DNA enters the cell. The cells are allowed to recover by a brief incubation at room temperature and then transferred to a tissue culture plate or T-flask along with growth medium and serum. Electroporation is an attractive method for transfection because it is more rapid and involves fewer manipulations than other methods and can be used for many types of adherent and nonadherent cells. This protocol describes transfection by electroporation of RAW 264.7 macrophages.

INTRODUCTIONIn this protocol for Dignam and Roeder nuclear extract preparation, HeLa cells are harvested by centrifugation, washed, and resuspended in a hypotonic buffer, which causes the cells to swell. Subsequently, the cells are lysed with a handheld Dounce homogenizer. The nuclei are pelleted by centrifugation, the cytoplasmic supernatant is decanted, and the nuclear pellet is resuspended by Dounce homogenization in a moderate salt buffer. After stirring the suspension for 30 min to allow extraction of transcription factors, the nuclei are centrifuged again, and the resulting supernatant or extract is dialyzed for use in transcription experiments. Generally, 1 L of culture (i.e., 1-2 g of cells, depending on cell density at harvest) yields 1-2 mL of extract at 6-10 mg/mL. This should provide enough extract for ~50-100 in vitro transcription reactions, depending on the scale.

INTRODUCTIONPrimer extension is a popular method for measuring transcription. Among the methods for start-site mapping in vivo, it is the one most easily adapted for in vitro transcription. Typically, DNA templates bearing activator binding sites are co-incubated with nuclear extracts (which provide the general transcription factors) and the corresponding activators to synthesize mRNA. In some cases, the activator is present in the extract (e.g., Sp1) and templates lacking or containing sites are compared. In the primer extension assay, a 32P-end-labeled DNA oligonucleotide is annealed to a complementary region in the mRNA and then extended by a reverse transcriptase to the 5′ end of the mRNA. The resulting labeled cDNA is then fractionated and detected on a polyacrylamide/urea gel. The amount of the product is a measure of the transcriptional activation by the activators. Generally, the addition of 0.1 pmol of primer to the extension reaction is sufficient because it will be in vast excess over the amount of mRNA, facilitating quantitative primer hybridization and extension.

INTRODUCTIONThe nuclear run-on assay was developed as a method for establishing that the transcription initiation rate contributes to the regulated expression of mammalian genes. The difference between monitoring gene expression by the nuclear run-on assay versus most other assays is that the nuclear run-on assay provides a measure of the frequency of transcription initiation and is largely independent of the effects of RNA stability. It can also be used to determine whether polymerase pausing or attenuation contributes to gene regulation. Briefly, the nuclear run-on assay begins with samples of cells that contain different steady-state amounts of the mRNA or protein of interest. The cells are chilled, and the plasmid membranes are permeabilized or lysed. These steps result in polymerase pausing. The nuclei are then incubated for a short time at 37°C in the presence of nucleoside triphosphates (NTPs) and radiolabeled uridine 5′-triphosphate (UTP). New transcripts are not initiated during this incubation, but the radiolabeled nucleotide becomes incorporated into transcripts that were being synthesized when the cells were first chilled and lysed. The number of nascent transcripts on the gene at the time of chilling is thought to be proportional to the frequency of transcription initiation. To determine the relative number of nascent transcripts in each sample, the radiolabeled RNA is purified and hybridized to a membrane containing immobilized DNA from the gene of interest. The amount of radioactivity that hybridizes to the membrane is approximately proportional to the number of nascent transcripts.

NF-kappaB is activated by many stimuli and NF-kappaB binding sites have been identified in a wide variety of genes. Yet, NF-kappaB-dependent gene expression must be stimulus- and cell-type-specific. In others words, the cellular response to different NF-kappaB activating stimuli, such as TNFalpha, IL-1, and LPS, must be different; and the response of different cell types, such as lymphocytes, fibroblasts, or epithelial cells, to the same NF-kappaB-inducing stimulus must also be different. Finally, kinetics of gene expression must be accounted for, so that all NF-kappaB-dependent genes are not activated simultaneously even if cell type and stimulus are constant. Here, we explore the mechanistic framework in which such regulatory aspects of NF-kappaB-dependent gene expression have been analyzed because they are likely to form the basis for physiological responses.

INTRODUCTIONGenomic footprinting and related methods can reveal important protein-DNA interactions not detected using other methods. This protocol presents procedures for DNase I genomic footprinting, micrococcal nuclease (MNase) mapping of nucleosome positioning, and monitoring nucleosome remodeling via restriction enzyme accessibility. These procedures rely on a technique known as ligation-mediated polymerase chain reaction (LM-PCR), which can be used in concert with a variety of agents that cleave or modify genomic DNA within an intact cell or nucleus, with each agent providing unique insight into the regulation of a gene. LM-PCR is a powerful technique for detecting DNA strand breaks within a complex sample because of its extreme sensitivity and specificity. This general method has proven to be invaluable for several purposes, in particular, in analysis of the properties of an endogenous DNA locus. Coupling LM-PCR to DNase I digestion results in assays that are analogous to in vitro DNase I footprinting or methylation protection. Coupling to MNase digestion allows for the high-resolution analysis of nucleosome positioning. Coupling to restriction enzyme digestion in isolated nuclei provides evidence of nucleosome remodeling. For each of these methods, nuclei must first be isolated by lysis of the cells with a non-ionic detergent, followed by centrifugation. The isolated nuclei are then treated with the appropriate nuclease, followed by purification of the genomic DNA. LM-PCR is then performed on the cleaved genomic DNA.

We have studied transcriptional control of the murine terminal deoxynucleotidyltransferase (TdT) gene, which is activated specifically in immature B and T lymphocytes. This analysis has led to the identification and purification of a 50-kDa sequence-specific DNA-binding protein, LyF-1, that interacts with the approximate consensus sequence PyPyTGGGAGPu and is enriched in cells at most stages of B- and T-cell differentiation. LyF-1 binds tightly to an element in the TdT promoter that we show is required for transcription in lymphocytes. LyF-1 also interacts with an element in the immunoglobulin mu enhancer, called microB, that was recently shown to be important for lymphocyte-specific enhancer activity. Moreover, LyF-1 binds to the promoters for the lymphocyte-specific genes lambda 5, VpreB, and lck, all of which we speculate have additional features in common with the TdT promoter. Thus, LyF-1 may be a general transcriptional activator for genes whose expression is restricted to the B- and/or T-lymphocyte lineages.

Transcription of mammalian genes by RNA polymerase II often begins at a specific nucleotide, whose location is determined either by an upstream DNA element known as a TATA box or by an element positioned at the transcription start site called an initiator (Inr). By in vitro analysis of synthetic promoters, we demonstrate here that the TATA and Inr elements are functionally similar and that the Inr is contained between nucleotides -3 and +5 relative to the initiation site. Moreover, we found that a mammalian transcription factor IID (TFIID) protein fraction is required for transcriptional stimulation by an Sp1-dependent activating element placed upstream of either TATA or Inr elements. However, in these assays, the yeast TATA-binding protein, which previously was shown to function similarly to mammalian TFIID, could not efficiently substitute for the mammalian TFIID fraction. These results demonstrate that mammalian TFIID is functionally distinct from the yeast TATA-binding protein and may contain additional subunits or domains that are important for transcriptional activation from some promoters.

INTRODUCTIONThe hydroxyl-radical footprinting methodology has wide applications to studying protein-DNA interactions, as well as structural perturbations (e.g., bending) that occur in DNA during protein binding. Hydroxyl radicals cleave DNA by abstracting a hydrogen atom from C4 of the sugar in the minor groove. Protein binding over the minor groove generally protects the sugar from cleavage. Because a hydroxyl radical molecule is small and thus is not subject to the same steric restrictions as other agents (such as DNase I), hydroxyl-radical footprinting can give detailed information on protein binding to the minor groove. In this protocol, the radical is generated by Fe(II) EDTA, which cleaves hydrogen peroxide into a hydroxyl radical and a hydroxide ion. The radical then cleaves the sugar in a diffusion limited reaction. Ascorbic acid is included to regenerate the active Fe(II).

INTRODUCTIONMethylation interference (and methylation protection) is the major technique for studying the interaction of proteins with major-groove guanines, although it also detects adenines in the minor groove and cytosines in single-stranded DNA. It is one of the highest-resolution methods for measuring the bases involved in sequence-specific recognition by proteins. A 32P-end-labeled DNA containing a recognition site is methylated (on average once per DNA molecule) by dimethyl sulfate (DMS) and then protein is allowed to bind. Certain methylation modifications prevent the protein from binding because they are at positions where the protein comes into close proximity with the DNA. Most modifications, however, have no effect on binding. The bound and unbound DNA molecules are separated using the electrophoretic mobility shift assay (or filter binding, immunoprecipitation, or any other suitable technique). The unbound fraction will be enriched in DNA molecules containing modifications at positions that interfere with protein binding. In contrast, the bound fraction will contain modifications that do not interfere with binding of the protein. Methylation weakens the nucleotide and makes it susceptible to piperidine-induced depurination. This leads, in turn, to scission of the DNA backbone. The cleaved fragments are fractionated on a sequencing gel alongside chemical sequencing ladders to identify the affected positions.

INTRODUCTIONIn the ethylation interference assay, ethyl nitrosourea is used to ethylate phosphates within 32P-end-labeled DNA molecules that contain a protein recognition site; the reaction is optimized to yield approximately one ethyl group per DNA molecule. The DNA is then incubated with barely saturating concentrations of a DNA-binding protein, but the ethylated phosphates interfere with proteins coming into close proximity to the minor groove or phosphate backbone of the binding site on the DNA molecule. Because ethylation makes the DNA susceptible to piperidine-induced cleavage, the protein recognition site can be identified by analyzing the cleavage products (from modified DNA molecules with bound protein versus modified DNA molecules without bound protein) on a polyacrylamide/urea gel alongside a sequencing ladder. Two points are critical for success with this assay. First, only about 10% of the molecules should be ethylated, which avoids having a significant number of molecules with multiple ethylations. This can be determined by modifying the DNA until ~10% of the starting probe is converted to a ladder of bands. It is critical that each DNA molecule be modified only once, so that it is clear that the modification being detected--the one nearest to the 32P-labeled end--is the one responsible for interference. Second, only barely saturating amounts of protein should be bound to the DNA. Barely saturated amounts are used because although the ethylation decreases binding, it does not always abolish it. High concentrations of protein may overcome the deleterious effect of the modification.

INTRODUCTIONThis is the first phase of a two-part assay for exploring whether an endogenous control region of interest is assembled into nucleosomes or is devoid of nucleosomes. This assay can also be used to determine if nucleosomes are similarly positioned at the locus in every cell within a population. Micrococcal nuclease (MNase) is unique among nucleases in its relative ability to induce double-strand breaks within nucleosomal linker regions, but only single-stranded nicks within the nucleosome itself. Because of this property, micrococcal nuclease can be used to determine whether a DNA fragment of interest is nucleosomal. In this phase of the assay, the cells are lysed and nuclei are isolated. Limiting concentrations of micrococcal nuclease are added to the nuclei, resulting in cleavage at nucleosome linker regions (preferably cleavage at two sites per DNA molecule). The cleavage reactions are stopped, and then the genomic DNA is purified and digested with restriction enzymes.

INTRODUCTIONThis is the second phase of a two-part assay for exploring whether an endogenous control region of interest is assembled into nucleosomes or is devoid of nucleosomes. This assay can also be used to determine if nucleosomes are similarly positioned at the locus in every cell within a population. In this second part of the assay, the restricted genomic DNA is separated by agarose gel electrophoresis and stained with ethidium bromide, resulting in a ladder of bands corresponding in size to multiples of the nucleosome core plus linker (~200 bp). To determine whether a DNA fragment of interest is nucleosomal, the genomic DNA is subjected to Southern blot analysis. If a probe derived from the DNA fragment hybridizes to the ladder of nucleosomal bands, the fragment may indeed be assembled into nucleosomes.

Summary Ikaros DNA-binding proteins are critical for the development of lymphocytes and other hematopoietic lineages, but it remains unclear how they cooperate with other regulators of signaling and transcription to achieve ordered gene expression during development. Here, we show that Ikaros proteins regulate the pre-BCR component λ5 in a stage-specific manner. In pre-BI cells, Ikaros modulated λ5 expression in competition with the transcriptional activator EBF. This required Ikaros binding to the Igll1 (λ5) promoter and was abolished either by mutation of the Ikaros DNA-binding domain or by deletion of a single Ikaros site from the Igll1 promoter. At the transition from the pre-BI to pre-BII stage, the expression of the Ikaros family member Aiolos was upregulated and required for the efficient silencing of Igll1 . Aiolos expression was controlled by pre-BCR signals via the adaptor protein SLP-65. Thus, pre-BCR signaling regulates Aiolos and the silencing of Igll1 via a developmental-stage-specific feedback loop.

Addition of potassium cyanide to 2-deoxy-2-fluoro- d -arabinose ( 1 ) at pH 7.8 followed by catalytic hydrogenation over palladium—barium sulfate (5%) produced 3-deoxy-3-fluoro- d -glucose ( 4 ) and 3-deoxy-3-fluoro- d -mannose ( 5 ) in 25 and 40% isolated yields, respectively. The epimeric sugars were purified by passage through a column of Dowex-50W × 8 (Ca 2+ ). In a similar manner, 3-deoxy-3-fluoro- d -arabinose ( 6 ) was converted into 4-deoxy-4-fluoro- d -glucose ( 9 ) and 4-deoxy-4- fluoro- d -mannose ( 10 ) in 27 and 45% isolated yields, respectively. Deoxyfluorohexoses 4, 5, 9 and 10 enriched with carbon-13 and carbon-14 at C-1 have been prepared by this procedure.

Accessibility of the genomic regulatory information is largely controlled by the nucleosome-organizing activity of transcription factors (TFs). While stimulus-induced TFs bind to genomic regions that are maintained accessible by lineage-determining TFs, they also increase accessibility of thousands of cis-regulatory elements. Nucleosome remodeling events underlying such changes and their interplay with basal positioning are unknown. Here, we devised a novel quantitative framework discriminating different types of nucleosome remodeling events in micrococcal nuclease ChIP-seq (chromatin immunoprecipitation [ChIP] combined with high-throughput sequencing) data sets and used it to analyze nucleosome dynamics at stimulus-regulated cis-regulatory elements. At enhancers, remodeling preferentially affected poorly positioned nucleosomes while sparing well-positioned nucleosomes flanking the enhancer core, indicating that inducible TFs do not suffice to overrule basal nucleosomal organization maintained by lineage-determining TFs. Remodeling events appeared to be combinatorially driven by multiple TFs, with distinct TFs showing, however, different remodeling efficiencies. Overall, these data provide a systematic view of the impact of stimulation on nucleosome organization and genome accessibility in mammalian cells.

In this issue of Immunity, Ghisletti et al. (2010) examined lipopolysaccharide (LPS)-induced binding of the p300 transcription coactivator to identify inducible enhancers throughout the genome of mouse macrophages, thereby revealing common features of enhancers that contribute to the LPS response and a critical role for PU.1 in enhancer marking.

Transcription of the lymphocyte-specific terminal deoxynucleotidyltransferase gene begins at a single nucleotide, but no TATA box is present. We have identified a 17 bp element that is sufficient for accurate basal transcription of this gene both in vitro and in vivo. This motif, the initiator (Inr), contains within itself the transcription start site. Homology to the Inr is found in many TATA-containing genes, and specific mutagenesis influences both the efficiency and accuracy of initiation. Moreover, in the presence of either a TATA box or the SV40 21 bp repeats, a greatly increased level of transcription initiates specifically at the Inr. Thus, the Inr constitutes the simplest functional promoter that has been identified and provides one explanation for how promoters that lack TATA elements direct transcription initiation.

Background Signaling via B cell receptor (BCR) and Toll-like receptors (TLRs) results in activation of B cells with distinct physiological outcomes, but transcriptional regulatory mechanisms that drive activation and distinguish these pathways remain unknown. Results Two hours after ligand exposure RNA-seq, ChIP-seq and computational methods reveal that BCR- or TLR-mediated activation of primary resting B cells proceeds via a large set of shared and a smaller subset of distinct signal-selective transcriptional responses. BCR stimulation resulted in increased global recruitment of RNA Pol II to promoters that appear to transit slowly to downstream regions. Conversely, lipopolysaccharide (LPS) stimulation involved an enhanced RNA Pol II transition from initiating to elongating mode accompanied by greater H3K4me3 activation markings compared to BCR stimulation. These rapidly diverging transcriptomic landscapes also show distinct repressing (H3K27me3) histone signatures, mutually exclusive transcription factor binding in promoters, and unique miRNA profiles. Conclusions Upon examination of genome-wide transcription and regulatory elements, we conclude that the B cell commitment to different activation states occurs much earlier than previously thought and involves a multi-faceted receptor-specific transcriptional landscape. Electronic supplementary material The online version of this article (doi:10.1186/s13072-015-0012-x) contains supplementary material, which is available to authorized users.