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Competitive hybridization of labeled nucleic acids to a microarray is conceptually similar to other hybridization methods, such as Southern blotting. For massively multiplexed microarrays, the adoption of two-color hybridization schemes has been a significant advance. The use of two colors-typically Cy3- and Cy5-labeled nucleic acids-makes it possible to control for factors that affect hybridization intensity, including the number of labeled nucleotides and the T m of each oligonucleotide. Thus, the difference in intensity among spots on a microarray can be quantified and analyzed to assess biological phenomena, like changes in gene expression or details of transcript structure. This protocol for hybridization is conceptually straightforward-"cold" (nonfluorescent) blocking nucleotide is added to the mixed nucleic acid material, Hybridization buffer is added, and the mixture is applied to the microarray surface. Hybridization occurs overnight, after which the microarray is washed and scanned., (© 2019 Cold Spring Harbor Laboratory Press.)

Most laboratories will choose to order microarrays from a commercial vendor; however, microarray printing is sufficiently straightforward that even small laboratories with extensive microarray needs may find it cost-effective and worth the effort to produce their own microarrays. Printing microarrays requires a spotting robot. For typical microarrays, oligonucleotides are distributed into 384-well plates and printed to polylysine-coated microarray slides. The details of a printing protocol vary depending on the robot being used; thus, follow the manufacturer's instructions. However, a typical workflow using a robot equipped with contact steel quill pins is provided in this protocol., (© 2019 Cold Spring Harbor Laboratory Press.)

Every microarray experiment is based on a common format. First, a large number of nucleotide "spots" are arrayed onto a substrate, typically a glass slide, a silicon chip, or microbeads. Second, a complex population of nucleic acids (isolated from cells, selected from in vitro-synthesized libraries, or obtained from another source) is labeled, typically with fluorescent dyes. Third, the labeled nucleic acids are allowed to hybridize to their complementary spot(s) on the microarray. Fourth, the hybridized microarray is washed, allowing the amount of hybridized label to then be quantified. Analysis of the raw data generates a readout of the levels of each species of RNA in the original complex population. This introduction includes several examples of microarray applications and provides a discussion of the basic steps of most microarray experiments., (© 2019 Cold Spring Harbor Laboratory Press.)

Homemade microarrays are printed on polylysine-coated slides. The lysines form a positively charged surface that can bind nonspecifically to the acidic nucleic acids during hybridization, resulting in significant background fluorescence. Thus, a key step in microarray processing is blocking all of the surface lysines not associated with the oligonucleotides in the microarray spots. In this protocol, the ε-amino group of lysine is succinylated by reacting with succinic anhydride. The procedure is straightforward. Microarrays are, if necessary, rehydrated; excess liquid is removed by drying at a moderate temperature; and the succinylation reaction is performed. After the reaction is complete, the slides are washed and dried with ethanol, at which point they are ready for hybridization or they can be stored in a desiccator., (© 2019 Cold Spring Harbor Laboratory Press.)

The goal of this procedure is to randomly amplify a sample of DNA to achieve the best possible sequence representation for hybridization to microarrays. This protocol has been used successfully to amplify genomic representations starting with <10 ng of DNA. The protocol consists of three sets of enzymatic reactions. In Round A, Sequenase is used to extend randomly annealed primers to generate templates for subsequent PCR. During Round B, a specific primer is used to amplify the previously generated templates. Amplified material can be labeled by adding fluorescent moieties to the nucleic acids. Alternatively, Round C in this protocol can be used to incorporate either aminoallyl-dUTP or Cy-dye-coupled nucleotides during additional PCR cycles. This protocol may be unsuitable for amplifying material smaller than 250 bp because such material will not be amplified uniformly., (© 2019 Cold Spring Harbor Laboratory Press.)

Round A/Round B amplification has been used extensively in genomic localization analysis and appears to work quite well for typical microarray applications in which DNA is sheared by sonication to ∼500 bp. For amplification of material <500 bp in size, however, T7 linear amplification of DNA (TLAD) is preferred because it more accurately maintains uniform representation of short DNA fragments. Amplification of double-stranded DNA by TLAD begins with the addition of a 3' tail of poly-thymidine to DNA by TdT. Second, the Klenow fragment of Escherichia coli DNA polymerase is used, along with a T7-poly(A) primer, to generate a complementary strand that carries a 5' T7 primer. Finally, extension of the original T-tailed DNA strand yields a template suitable for T7-based transcription, which generates amplified RNA (aRNA). This technique avoids the "jackpotting" issues observed with PCR, in which an early amplification event leads to disproportionate representation of a particular sequence, because PCR follows exponential kinetics, whereas transcription is a linear amplification method., (© 2019 Cold Spring Harbor Laboratory Press.)

As with RNA labeling protocols, the main difference between direct and indirect DNA labeling protocols is a trade-off of cost and time. Indirect labeling of DNA, described here, takes ∼2 h longer than direct labeling but is hundreds of dollars cheaper., (© 2019 Cold Spring Harbor Laboratory Press.)

Similar to direct RNA labeling, direct DNA labeling with Cy-dCTP is the simplest and fastest method for labeling DNA. This is a standard Klenow labeling protocol in which Cy-dCTP is incorporated during the labeling reaction. After stopping the reaction, labeled nucleotide is separated from unreacted Cy-dCTP, and Cy3- and Cy5-labeled materials are combined for hybridization. This protocol is suitable for many applications, including detection of copy-number variation, nucleosome mapping, and other location analysis (e.g., chromatin immunoprecipitation [ChIP]-chip)., (© 2019 Cold Spring Harbor Laboratory Press.)

The labeling procedure described here is called indirect because the fluorescent moiety is not incorporated during the reverse transcription reaction. Instead, a reactive nucleotide analog (aminoallyl-dUTP) is incorporated during reverse transcription, the cDNA is isolated, and then the cyanine dyes are incorporated by binding with the aminoallyl group to produce the desired fluorescent cDNA., (© 2019 Cold Spring Harbor Laboratory Press.)

Gene expression profiling typically requires microgram quantities of mRNA, which can be difficult to obtain. In such cases, RNA must be amplified to have enough material for microarray labeling and hybridization. This protocol generates amplified antisense RNA (aRNA) from limited quantities of total RNA. It is designed around maximizing yield and product length while minimizing template-independent side reactions. Template-independent reactions compete with the desired template-dependent reaction, an undesirable situation that becomes more severe as less RNA template is used. Amplification products dominated by template-independent product result in greatly reduced sensitivity and compression of differences in microarray hybridization experiments. Most notably, the oligo(dT) primer used in reverse transcription (RT) yields a high-molecular-weight product in the in vitro transcription (IVT) reaction independent of any cDNA template. This protocol therefore limits the amount of primer used by using small cDNA synthesis volumes. In addition, high-molecular-weight, template-independent product is generated in the presence of biotinylated NTPs and the absence of any polymer when excessive amounts of T7 RNA polymerase activity are used. Template-dependent product of questionable molecular weight and limited functionality in downstream reactions can also be produced with excessive T7 RNA polymerase activity. Essentially, more yield is not always better. Here, a single round of amplification typically results in a fivefold to 20-fold mass conversion of starting material. If the first-round aRNA is used as a template for a second round of amplification, 200- to 400-fold amplification is typical., (© 2019 Cold Spring Harbor Laboratory Press.)

This is the simplest method to label RNA for use in expression analysis. RNA is reverse-transcribed using both oligo(dT) and random hexamers as primers. The random hexamers improve overall efficiency of labeling, especially at the 5' end of the RNA. Fluorescently labeled dUTP is incorporated into the cDNA. After reverse transcription, the RNA is degraded, and the labeled cDNA is purified from unincorporated Cy dyes. Finally, samples labeled with Cy3 and Cy5 dyes are mixed and combined with blocking nucleotides and used for hybridization., (© 2019 Cold Spring Harbor Laboratory Press.)

Significant insight about biological networks arises from the study of network motifs--overly abundant network subgraphs--but such wiring patterns do not specify when and how potential routes within a cellular network are used. To address this limitation, we introduce activity motifs, which capture patterns in the dynamic use of a network. Using this framework to analyze transcription in Saccharomyces cerevisiae metabolism, we find that cells use different timing activity motifs to optimize transcription timing in response to changing conditions: forward activation to produce metabolic compounds efficiently, backward shutoff to rapidly stop production of a detrimental product and synchronized activation for co-production of metabolites required for the same reaction. Measuring protein abundance over a time course reveals that mRNA timing motifs also occur at the protein level. Timing motifs significantly overlap with binding activity motifs, where genes in a linear chain have ordered binding affinity to a transcription factor, suggesting a mechanism for ordered transcription. Finely timed transcriptional regulation is therefore abundant in yeast metabolism, optimizing the organism's adaptation to new environmental conditions.

Lymphocyte activation is accompanied by visible changes in chromatin structure. We find that antigen receptor signaling induces the rapid association of the BAF complex with chromatin. PIP2, which is regulated by activation stimuli, is sufficient in vitro to target the BAF complex to chromatin, but it has no effect on related chromatin remodeling complexes containing SNF2L or hISWI. Purification and peptide sequencing of the subunits of the complex revealed beta-actin as well as a novel actin-related protein, BAF53. beta-actin and BAF53 are required for maximal ATPase activity of BRG1 and are also required with BRG1 for association of the complex with chromatin/matrix. This work indicates that membrane signals control the activity of the mammalian SWI/SNF or BAF complex and demonstrates a direct interface between signaling and chromatin regulation.

We demonstrate an essential role for the proteasome complex in two proteolytic processes required for activation of the transcription factor NF-kappa B. The p105 precursor of the p50 subunit of NF-kappa B is processed in vitro by an ATP-dependent process that requires proteasomes and ubiquitin conjugation. The C-terminal region of p105 is rapidly degraded, leaving the N-terminal p50 domain. p105 processing can be blocked in intact cells with inhibitors of the proteasome or in yeast with proteasome mutants. These inhibitors also block the activation of NF-kappa B and the rapid degradation of I kappa B alpha induced by tumor necrosis factor alpha. Thus, the ubiquitin-proteasome pathway functions not only in the complete degradation of polypeptides, but also in the regulated processing of precursors into active proteins.