We thank Moustafa Saleh and Jay Tourigny for critical reading of the manuscript and Steven Hahn and Steven Henikoff for mentorship and support during the development of ChEC-seq and its application to the Mediator complex. S.G. is supported by NIH grants R01GM053451 and R01GM075114 and G.E.Z. is supported by Indiana University startup funds.

1. Generation of Yeast Strains
<ol>
	<li>Generate a yeast strain bearing the factor of interest tagged with MNase.
	<ol>
		<li>PCR amplify the MNase tagging cassette from the desired vector (Table 1) using the specified reaction mixture (Table 2) and cycling conditions (Table 3). Mix 5 &#181;L of the PCR reaction and 1 &#181;L of 6X DNA loading dye. Run each PCR aliquot on a 0.8% agarose gel at 120 V for 40 min. The expected product size is ~2.3 kb.</li>
		<li>Transform the amplified tagging cassette into the strain of choice using standard lithium acetate transformation22 and perform selection.</li>
		<li>Verify correct integration of the tagging cassette with colony PCR.
		<ol>
			<li>Prepare the specified reaction mixture (Table 4) for the number of colonies to be screened. Keep on ice until needed.</li>
			<li>Pick a portion of each colony to be tested into the bottom of a PCR tube using a sterile toothpick or pipet tip.</li>
			<li>Microwave the picked colonies at high power for 1 min.</li>
			<li>Add 50 &#181;L of PCR mix (Table 4) to each microwaved colony and pipet up and down to mix. Perform PCR using the specified cycling conditions (Table 5).</li>
			<li>Mix the 50 &#181;L of PCR reaction and 10 &#181;L of 6X DNA loading dye directly in each PCR tube. Run 10 &#181;L of each sample on a 1% agarose gel at 120 V for 40 min. The expected product size is ~ 700 bp. 
			NOTE: If desired, verify appropriate expression of the MNase fusion protein via western blotting. A ~ 20.6 kilodalton shift in the molecular weight of the tagged protein is expected.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Generate a free MNase strain by homology-based cloning of the promoter of the gene of interest into pGZ136 (Table 1) and transformation and selection as in step 1.1.2.
	<ol>
		<li>Alternatively, assemble the promoter of the gene of interest and 3xFLAG-MNase-SV40 NLS into a plasmid vector, transform and select as in step 1.1.2, and grow the strain in the appropriate selective condition for maintenance of the plasmid.</li>
	</ol>
	</li>
</ol>
2. ChEC
<ol>
	<li>In the afternoon/evening of the day prior to the ChEC experiment, inoculate 3 mL yeast-peptone-dextrose (YPD) or appropriate selective medium with a single colony of the strain bearing the MNase-tagged factor. Incubate this culture overnight (180 RPM shaking, 30 &#176;C).</li>
	<li>In the morning of the day of the ChEC experiment, dilute the overnight culture into 50 mL of medium to an optical density at 600 nm (OD600) of 0.2 - 0.3. Incubate this culture (180 RPM shaking, 30 &#176;C) until it reaches an OD600 of 0.5-0.7. As the culture approaches the appropriate OD600, set heat block or water bath to 30 &#176;C and start thawing Buffer A additives (Table 6).</li>
	<li>Prepare 5 mL of Buffer A plus additives (Table 6) per culture. Keep buffer A at RT during the procedure.</li>
	<li>Prepare microfuge tubes containing 90 &#181;L of stop solution (Table 7) and 10 &#181;L of 10% SDS for each time point to be collected. For each new factor analyzed, take samples at 0 s, 30 s, 1 min, 2.5 min, 5 min, and 10 min.</li>
	<li>Decant culture into a 50 mL conical tube and centrifuge at 1,500 x g and room temperature (RT) for 1 min.</li>
	<li>Thoroughly resuspend cells in 1 mL of Buffer A and transfer the resuspended cells to a microfuge tube. Pellet resuspended cells in a microfuge at 1,500 x g and RT for 30 s.</li>
	<li>Aspirate supernatant and thoroughly resuspend cells in 1 mL of Buffer A by pipetting up and down. Pellet resuspended cells in a microfuge at 1,500 x g and RT for 30 s. Repeat this step once.</li>
	<li>Thoroughly resuspend cells in 570 &#181;L of Buffer A. Add 30 &#181;L of 2% digitonin (0.1% final concentration) to facilitate permeabilization of the cells, and invert to mix.</li>
	<li>Permeabilize cells in heat block or water bath at 30 &#176;C for 5 min.</li>
	<li>Pipet the reaction up and down to ensure an even distribution of cells. Remove a 100 &#181;L aliquot of permeabilized cells as a negative control to a microfuge tube containing stop solution and SDS (step 2.4) and vortex briefly to mix.</li>
	<li>Add 1.1 &#181;L of 1 M CaCl2 (2 mM final concentration) to permeabilized cells to activate MNase and invert several times quickly or vortex briefly to mix. Immediately return the mixed reaction at 30 &#176;C and start a timer.</li>
	<li>At each time point to be collected, pipet the reaction up and down to ensure an even distribution of cells, then remove a 100 &#181;L aliquot of permeabilized cells to a microfuge tube containing stop solution and SDS and vortex briefly to mix. For each new factor analyzed, take 0 s (no CaCl2&#160;added), 30 s, 1 min, 2.5 min, 5 min, and 10 min time points. 
	NOTE: ChEC experiments with factors that rapidly cleave DNA at 30&#176;C may be performed at lower temperatures to slow MNase cleavage kinetics.</li>
	<li>Once all time points have been collected, add 4 &#181;L of 20 mg/mL proteinase K to each sample, vortex briefly to mix, and incubate at 55 &#176;C for 30 min.</li>
	<li>Add 200 &#181;L of 25:24:1 phenol:chloroform:isoamyl alcohol to each sample, vortex vigorously to mix, and centrifuge at maximum speed and RT in a microfuge for 5 min.</li>
	<li>Remove each aqueous phase (~ 150 &#181;L) to a new tube. Add 30 &#181;g of glycogen and 500 &#181;L of 100% ethanol. Vortex vigorously to mix and precipitate on dry ice for 10 min or until solution is viscous.</li>
	<li>Pellet precipitated DNA at maximum speed and 4 &#176;C in a microfuge for 10 min.</li>
	<li>Decant supernatants and wash pellets with 1 mL of RT 70% ethanol.</li>
	<li>Decant ethanol, removing the remainder by inverting and gently tapping the tubes on a paper towel. Briefly spin samples using a benchtop centrifuge and use a pipet to remove residual ethanol, taking care not to disturb the pellet. Air-dry pellets for at RT for 5 min.</li>
	<li>While pellets are drying, make a master mix to resuspend dried pellets in 29 &#181;L of Tris, pH 8.0 + 1 &#181;L of 10 mg/mL RNase A each. Digest RNA at 37 &#176;C for 20 min.</li>
	<li>Mix 1 &#181;L of 6X DNA loading dye with 5 &#181;L of each sample and run on a 1.5% agarose gel at 120 V for 40 min.</li>
</ol>
3. Size Selection
NOTE: The goal of size selection is to remove multi-kilobase fragments of genomic DNA from the sample to be sequenced and enrich fragments of ~150 bp (approximately the size of nucleosomal DNA) or smaller. In sequencing data, fragments &#60;400 bp are enriched, with a notable peak around the size of nucleosomal DNA (~150 bp) and a peak or broad distribution of subnucleosomal fragments.
<ol>
	<li>To each sample, add 75 &#181;L of solid-phase reversible immobilization (SPRI) beads in a polyethylene glycol (PEG) solution23 and pipet up and down 10 times to mix. Incubate the bead:DNA mixture at RT for 5 min.</li>
	<li>During the SPRI bead incubation, prepare microfuge tubes containing 96 &#181;L of 10 mM Tris, pH 8.0 and 4 &#181;L of 5 M NaCl for each sample.</li>
	<li>Place tubes in a magnetic rack to collect beads on the side of the tube. Allow the beads to collect on the side of the tube for 2 min.</li>
	<li>Transfer each supernatant to a new tube containing Tris and NaCl (step 3.2).</li>
	<li>Add 200 &#181;L of 25:24:1 phenol:chloroform:isoamyl alcohol to each sample, vortex to mix, and centrifuge at maximum speed and RT in a microfuge for 5 min.</li>
	<li>Remove each aqueous phase to a new tube. Add 30 &#181;g of glycogen and 500 &#181;L of 100% ethanol. Precipitate DNA on dry ice for 10 min or until solution is viscous.</li>
	<li>Pellet precipitated DNA at maximum speed and 4 &#176;C in a microfuge for 10 min.</li>
	<li>Decant supernatants and wash pellets with 1 mL of 70% ethanol.</li>
	<li>Decant ethanol, removing the remainder by inverting and gently tapping the tubes on a paper towel. Briefly spin samples using a benchtop centrifuge and use a pipet to remove residual ethanol, taking care not to disturb the pellet. Air-dry pellets for at RT for 5 min.</li>
	<li>Resuspend dried pellets in 25 &#181;L of Tris, pH 8.0. Determine sample concentration using a high-sensitivity assay. For TF ChEC experiments, 10-100 ng DNA is routinely recovered after size selection.</li>
	<li>Prepare sequencing libraries. Library preparation protocols that do not rely upon size selection, as previously described24, are desirable to allow sequencing of a large range of fragment sizes (25-500 bp).</li>
	<li>Sequence libraries in paired-end mode. A minimum of 25 bases of sequence on each end is needed for high quality read mapping. Between 1 and 2 million reads provides sufficient coverage for a ChEC sample. 
	NOTE: Analysis of ChEC-seq data is a complex procedure and beyond the scope of this article. Detailed information on data analysis can be found in previous publications1,21, and the custom scripts used for analysis are available on GitHub (https://github.com/zentnerlab/chec-seq). HOMER25 (http://homer.salk.edu) can also be used for command-line analysis of ChEC-seq data.</li>
</ol>

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The authors have nothing to disclose.

We have shown that ChEC can map diverse classes of yeast proteins on chromatin, and anticipate that it will be broadly applicable to different families of TFs and other chromatin-binding factors in yeast. ChEC-seq is advantageous in that it does not require crosslinking, chromatin solubilization, or antibodies. Thus, ChEC avoids artifacts potentially present in X-ChIP-seq, such as the hyper-ChIPable artifact3,4, and native ChIP, such as false negatives due to inc...

Mapping the binding sites of transcription factors (TFs), chromatin remodelers, and other chromatin-associated regulatory factors is key to understanding all chromatin-based processes. While chromatin immunoprecipitation and high-throughput sequencing (ChIP-seq) approaches have been used to gain many important insights into genome biology, they have notable limitations. We recently introduced an alternative method, termed chromatin endogenous cleavage and high-throughput sequencing (ChEC-seq)1, to circumvent these drawbacks.
ChIP-seq is most often performed with an initial formaldehyde crosslinking step (X-ChIP-seq) to preserve protein-DNA interactions. However, a number of recent studies have indicated that X-ChIP-seq captures transient or nonspecific protein-DNA interactions2,3,4,5,6,7,8, giving rise to false positive binding sites. In addition, sonication, commonly used to fragment chromatin in X-ChIP-seq experiments, preferentially shears regions of open chromatin, leading to biased recovery of fragments from these regions9,10. Sonication also yields a heterogeneous mixture of fragment lengths, ultimately limiting binding site resolution, though the addition of an exonuclease digestion step can greatly improve resolution11,12. Native ChIP methods such as occupied regions of genomes from affinity-purified naturally isolated chromatin (ORGANIC)13 do not use crosslinking and fragment chromatin with micrococcal nuclease (MNase), alleviating potential biases associated with formaldehyde crosslinking and sonication. However, the solubility of many chromatin-bound proteins under the relatively mild conditions required for native chromatin extraction is poor, potentially leading to reduced dynamic range and/or false negatives14.
While various iterations of ChIP-seq are most commonly used for genome-wide mapping of protein-DNA interactions, several mapping techniques based on fusion of proteins of interest to various DNA-modifying enzymes have also been implemented. One such approach is DNA adenine methyltransferase identification (DamID)15, wherein a chromatin-binding protein of interest is genetically fused to Dam and this fusion is expressed in cells or animals, resulting in methylation of GATC sequences proximal to binding sites for the protein. DamID is advantageous in that it does not rely upon immunoprecipitation and so avoids crosslinking, antibodies, or chromatin solubilization. It is also performed in vivo. However, the resolution of DamID is limited to the kilobase scale and the methylating activity of the Dam fusion protein is constitutive. A second method based on enzymatic fusion is Calling Card-seq16, which employs fusion of a factor of interest to a transposase, directing site-specific integration of transposons. Like DamID, Calling Card-seq is not based on immunoprecipitation and thus has similar advantages, with the added benefit of increased resolution. However, Calling Card-seq may be limited by sequence biases of transposases and is also reliant on the presence of restriction sites close to transposon insertion sites.
A third enzymatic fusion method, developed in the Laemmli lab, is chromatin endogenous cleavage (ChEC)17. In ChEC, a fusion between a chromatin-associated protein and MNase is expressed in cells, and upon calcium addition to activate MNase, DNA is cleaved proximal to binding sites for the tagged factor (Figure 1). In conjunction with Southern blotting, ChEC has been used to characterize chromatin structure and protein binding at a number of individual loci in yeast17,18, and has been combined with low-resolution microarray analysis to probe the interaction of nuclear pore components with the yeast genome19. ChEC offers benefits similar to DamID and Calling Card-seq, and its resolution is nearly single-base pair when analyzed by primer extension19. ChEC is also controllable: robust DNA cleavage by MNase depends on the addition of millimolar calcium, ensuring that MNase is inactive at the low free calcium concentrations observed in live yeast cells20.
Previously, we postulated that combining ChEC with high-throughput sequencing (ChEC-seq) would provide high-resolution maps of TF binding sites. Indeed, ChEC-seq generated high-resolution maps of the budding yeast general regulatory factors (GRFs) Abf1, Rap1, and Reb1 across the genome1. We have also successfully applied ChEC-seq to the modular Mediator complex, a conserved, essential global transcriptional coactivator21, expanding the applicability of ChEC-seq to megadalton-size complexes that do not directly contact DNA and may be difficult to map by ChIP-based methods. ChEC-seq is a powerful method both for independent validation of ChIP-seq results and generation of new insights into the regulation of chromatin-resident processes. Here, we present a step-by-step protocol for the implementation of this method in budding yeast.

<table border="0" cellpadding="0" cellspacing="0" width="751">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:307px;">dNTPs</td>
			<td style="width:159px;">NEB</td>
			<td style="width:119px;">N0447</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:307px;">Q5 high-fidelity DNA polymerase</td>
			<td style="width:159px;">NEB</td>
			<td style="width:119px;">M0491L</td>
			<td>Other high-fidelity DNA polymerases, such as Phusion, may be used for cassette amplification.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:307px;">TrackIt 1 Kb Plus DNA ladder</td>
			<td style="width:159px;">ThermoFisher Scientific</td>
			<td style="width:119px;">10488085</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:307px;">Taq DNA polymerase</td>
			<td style="width:159px;">NEB</td>
			<td style="width:119px;">M0273L</td>
			<td></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:307px;">cOmplete Mini EDTA-free protease inhibitor cocktail</td>
			<td style="width:159px;">Sigma-Aldrich</td>
			<td style="width:119px;">11836170001</td>
			<td>It is important that an EDTA-free protease inhibitor mix is used, so as not to inhibit Mnase cleavage by chelation of Ca2+.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:307px;">PMSF</td>
			<td style="width:159px;">ACROS Organics</td>
			<td style="width:119px;">AC215740010</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:307px;">Digitonin, High Purity</td>
			<td style="width:159px;">EMD Millipore</td>
			<td style="width:119px;">300410-250MG</td>
			<td>Make a 2% stock by dissolving 20 mg digitonin in 1 mL DMSO with vigorous vortexing.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:307px;">Proteinase K, 20 mg/mL</td>
			<td style="width:159px;">Invitrogen</td>
			<td style="width:119px;">25530049</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:307px;">RNase A, 10 mg/mL</td>
			<td style="width:159px;">ThermoFisher Scientific</td>
			<td style="width:119px;">EN0531</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:307px;">Ampure XP beads</td>
			<td style="width:159px;">Beckman Coulter</td>
			<td style="width:119px;">A63880</td>
			<td>Ampure-like beads can be generated using a published protocol (ref 24).</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:307px;">MagneSphere magnetic rack</td>
			<td style="width:159px;">Promega</td>
			<td style="width:119px;">Z5342</td>
			<td></td>
		</tr>
	</tbody>
</table>

genome-wide mapping of protein-dna interactions with chec-seq in saccharomyces cerevisiae

Genome-wide mapping of protein-DNA interactions is critical for understanding gene regulation, chromatin remodeling, and other chromatin-resident processes. Formaldehyde crosslinking followed by chromatin immunoprecipitation and high-throughput sequencing (X-ChIP-seq) has been used to gain many valuable insights into genome biology. However, X-ChIP-seq has notable limitations linked to crosslinking and sonication. Native ChIP avoids these drawbacks by omitting crosslinking, but often results in poor recovery of chromatin-bound proteins. In addition, all ChIP-based methods are subject to antibody quality considerations. Enzymatic methods for mapping protein-DNA interactions, which involve fusion of a protein of interest to a DNA-modifying enzyme, have also been used to map protein-DNA interactions. We recently combined one such method, chromatin endogenous cleavage (ChEC), with high-throughput sequencing as ChEC-seq. ChEC-seq relies on fusion of a chromatin-associated protein of interest to micrococcal nuclease (MNase) to generate targeted DNA cleavage in the presence of calcium in living cells. ChEC-seq is not based on immunoprecipitation and so circumvents potential concerns with crosslinking, sonication, chromatin solubilization, and antibody quality while providing high resolution mapping with minimal background signal. We envision that ChEC-seq will be a powerful counterpart to ChIP, providing an independent means by which to both validate ChIP-seq findings and discover new insights into genomic regulation.

In the case of a successful ChEC experiment, analysis of DNA by agarose gel electrophoresis will reveal a calcium-dependent increase in DNA fragmentation over-time, as indicated by smearing and eventual complete digestion of genomic DNA. In some cases, a ladder of bands similar to that seen with a traditional MNase digestion is observed after extended digestion. This is the case for ChEC analysis of Reb1, a general regulatory factor that binds nucleosome-depleted regions (NDRs) (<strong c...

Watch this Scientific Journal Video about Genome-wide Mapping of Protein-DNA Interactions with ChEC-seq in Saccharomyces cerevisiae at JoVE.com

Genome-wide Mapping of Protein-DNA Interactions with ChEC-seq in Saccharomyces cerevisiae

We describe chromatin endogenous cleavage coupled with high-throughput sequencing (ChEC-seq), a chromatin immunoprecipitation (ChIP)-orthogonal method for mapping protein binding sites genome-wide with micrococcal nuclease (MNase) fusion proteins.

Genome-wide Mapping of Protein-DNA Interactions with ChEC-seq in Saccharomyces cerevisiae

Genome-wide mapping of protein-DNA interactions is critical for understanding gene regulation, chromatin remodeling, and other chromatin-resident ...

genome-wide-mapping-protein-dna-interactions-with-chec-seq

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JoVE Journal

Genetics

11.0K Views.  Fred Hutchinson Cancer Research Center. We describe chromatin endogenous cleavage coupled with high-throughput sequencing (ChEC-seq), a chromatin immunoprecipitation (ChIP)-orthogonal method for mapping protein binding sites genome-wide with micrococcal nuclease (MNase) fusion proteins.

Video: Genome-wide Mapping of Protein-DNA Interactions with ChEC-seq in Saccharomyces cerevisiae

Genome-wide Determination of Mammalian Replication Timing by DNA Content Measurement

Replication of the genome occurs during S phase of the cell cycle in a highly regulated process that ensures the fidelity of DNA duplication. Each genomic region is replicated at a distinct time during S phase through the simultaneous activation of multiple origins of replication. Time of replication (ToR) correlates with many genomic and epigenetic features and is linked to mutation rates and cancer. Comprehending the full genomic view of the replication program, in health and disease is a major future goal and challenge.
This article describes in detail the &#34;Copy Number Ratio of S/G1 for mapping genomic Time of Replication&#34; method (herein called: CNR-ToR), a simple approach to map the genome wide ToR of mammalian cells. The method is based on the copy number differences between S phase cells and G1 phase cells. The CNR-ToR method is performed in 6 steps: 1. Preparation of cells and staining with propidium iodide (PI); 2. Sorting G1 and S phase cells using fluorescence-activated cell sorting (FACS); 3. DNA purification; 4. Sonication; 5. Library preparation and sequencing; and 6. Bioinformatic analysis. The CNR-ToR method is a fast and easy approach that results in detailed replication maps.

We describe here a relatively fast and simple approach for mapping genome-wide mammalian replication timing, from cell isolation to the basic analysis of the sequencing results. A genomic map of a representative replication program will be provided following the protocol.

Replication of the genome occurs during S phase of the cell cycle in a highly regulated process that ensures the fidelity of DNA duplication. Each genomic ...

Mapping RNA-RNA Interactions Globally Using Biotinylated Psoralen

Knowing how RNAs interact with themselves and with others is key to understanding RNA based gene regulation in the cell. While examples of RNA-RNA interactions such as microRNA-mRNA interactions have been shown to regulate gene expression, the full extent to which RNA interactions occur in the cell is still unknown. Previous methods to study RNA interactions have primarily focused on subsets of RNAs that are interacting with a particular protein or RNA species. Here, we detail a method named Sequencing of Psoralen crosslinked, Ligated, and Selected Hybrids (SPLASH) that allows genome-wide capture of RNA interactions in vivo in an unbiased manner. SPLASH utilizes in vivo crosslinking, proximity ligation, and high throughput sequencing to identify intramolecular and intermolecular RNA base-pairing partners globally. SPLASH can be applied to different organisms including bacteria, yeast and human cells, as well as diverse cellular conditions to facilitate the understanding of the dynamics of RNA organization under diverse cellular contexts. The entire experimental SPLASH protocol takes about 5 days to complete and the computational workflow takes about 7 days to complete.

Here, we detail the method of Sequencing of Psoralen crosslinked, Ligated, and Selected Hybrids (SPLASH), which enables genome-wide mapping of intramolecular and intermolecular RNA-RNA interactions in vivo. SPLASH can be applied to study RNA interactomes of organisms including yeast, bacteria and humans.

Knowing how RNAs interact with themselves and with others is key to understanding RNA based gene regulation in the cell. While examples of RNA-RNA ...

Mapping Genome-wide Accessible Chromatin in Primary Human T Lymphocytes by ATAC-Seq

Assay for Transposase-Accessible Chromatin with high-throughput sequencing (ATAC-seq) is a method used for the identification of open (accessible) regions of chromatin. These regions represent regulatory DNA elements (e.g., promoters, enhancers, locus control regions, insulators) to which transcription factors bind. Mapping the accessible chromatin landscape is a powerful approach for uncovering active regulatory elements across the genome. This information serves as an unbiased approach for discovering the network of relevant transcription factors and mechanisms of chromatin structure that govern gene expression programs. ATAC-seq is a robust and sensitive alternative to DNase I hypersensitivity analysis coupled with next-generation sequencing (DNase-seq) and formaldehyde-assisted isolation of regulatory elements (FAIRE-seq) for genome-wide analysis of chromatin accessibility and to the sequencing of micrococcal nuclease-sensitive sites (MNase-seq) to determine nucleosome positioning. We present a detailed ATAC-seq protocol optimized for human primary immune cells i.e. CD4+ lymphocytes (T helper 1 (Th1) and Th2 cells). This comprehensive protocol begins with cell harvest, then describes the molecular procedure of chromatin tagmentation, sample preparation for next-generation sequencing, and also includes methods and considerations for the computational analyses used to interpret the results. Moreover, to save time and money, we introduced quality control measures to assess the ATAC-seq library prior to sequencing. Importantly, the principles presented in this protocol allow its adaptation to other human immune and non-immune primary cells and cell lines. These guidelines will also be useful for laboratories which are not proficient with next-generation sequencing methods.

Assay for Transposase-Accessible Chromatin coupled with high-throughput sequencing (ATAC-seq) is a genome-wide method to uncover accessible chromatin. This is a step-by-step ATAC-seq protocol, from molecular to the final computational analysis, optimized for human lymphocytes (Th1/Th2). This protocol can be adopted by researchers without prior experience in next-generation sequencing methods.

Assay for Transposase-Accessible Chromatin with high-throughput sequencing (ATAC-seq) is a method used for the identification of open (accessible) regions ...

Determining Genome-wide Transcript Decay Rates in Proliferating and Quiescent Human Fibroblasts

Quiescence is a temporary, reversible state in which cells have ceased cell division, but retain the capacity to proliferate. Multiple studies, including ours, have demonstrated that quiescence is associated with widespread changes in gene expression. Some of these changes occur through changes in the level or activity of proliferation-associated transcription factors, such as E2F and MYC. We have demonstrated that mRNA decay can also contribute to changes in gene expression between proliferating and quiescent cells. In this protocol, we describe the procedure for establishing proliferating and quiescent cultures of human dermal foreskin fibroblasts. We then describe the procedures for inhibiting new transcription in proliferating and quiescent cells with Actinomycin D (ActD). ActD treatment represents a straightforward and reproducible approach to dissociating new transcription from transcript decay. A disadvantage of ActD treatment is that the time course must be limited to a short time frame because ActD affects cell viability. Transcript levels are monitored over time to determine transcript decay rates. This procedure allows for the identification of genes and isoforms that exhibit differential decay in proliferating versus quiescent fibroblasts.

We describe a protocol for generating proliferating and quiescent primary human dermal fibroblasts, monitoring transcript decay rates, and identifying differentially decaying genes.

Quiescence is a temporary, reversible state in which cells have ceased cell division, but retain the capacity to proliferate. Multiple studies, including ...

Generation of Genome-wide Chromatin Conformation Capture Libraries from Tightly Staged Early Drosophila Embryos

Investigating the three-dimensional architecture of chromatin offers invaluable insight into the mechanisms of gene regulation. Here, we describe a protocol for performing the chromatin conformation capture technique in situ Hi-C on staged Drosophila melanogaster embryo populations. The result is a sequencing library that allows the mapping of all chromatin interactions that occur in the nucleus in a single experiment. Embryo sorting is done manually using a fluorescent stereo microscope and a transgenic fly line containing a nuclear marker. Using this technique, embryo populations from each nuclear division cycle, and with defined cell cycle status, can be obtained with very high purity. The protocol may also be adapted to sort older embryos beyond gastrulation. Sorted embryos are used as inputs for in situ Hi-C. All experiments, including sequencing library preparation, can be completed in five days. The protocol has low input requirements and works reliably using 20 blastoderm stage embryos as input material. The end result is a sequencing library for next generation sequencing. After sequencing, the data can be processed into genome-wide chromatin interaction maps that can be analyzed using a wide range of available tools to gain information about topologically associating domain (TAD) structure, chromatin loops, and chromatin compartments during Drosophila development.

Generation of Genome-wide Chromatin Conformation Capture Libraries from Tightly Staged Early Drosophila Embryos

This work describes a protocol for the generation of high resolution in situ Hi-C libraries from tightly staged pre-gastrulation Drosophila melanogaster embryos.

Investigating the three-dimensional architecture of chromatin offers invaluable insight into the mechanisms of gene regulation. Here, we describe a ...

Chromatin Immunoprecipitation (ChIP) of Histone Modifications from Saccharomyces cerevisiae

Histone post-translational modifications (PTMs), such as acetylation, methylation and phosphorylation, are dynamically regulated by a series of enzymes that add or remove these marks in response to signals received by the cell. These PTMS are key contributors to the regulation of processes such as gene expression control and DNA repair. Chromatin immunoprecipitation (chIP) has been an instrumental approach for dissecting the abundance and localization of many histone PTMs throughout the genome in response to diverse perturbations to the cell. Here, a versatile method for performing chIP of post-translationally modified histones from the budding yeast Saccharomyces cerevisiae (S. cerevisiae) is described. This method relies on crosslinking of proteins and DNA using formaldehyde treatment of yeast cultures, generation of yeast lysates by bead beating, solubilization of chromatin fragments by micrococcal nuclease, and immunoprecipitation of histone-DNA complexes. DNA associated with the histone mark of interest is purified and subjected to quantitative PCR analysis to evaluate its enrichment at multiple loci throughout the genome. Representative experiments probing the localization of the histone marks H3K4me2 and H4K16ac in wildtype and mutant yeast are discussed to demonstrate data analysis and interpretation. This method is suitable for a variety of histone PTMs and can be performed with different mutant strains or in the presence of diverse environmental stresses, making it an excellent tool for investigating changes in chromatin dynamics under different conditions.

Chromatin Immunoprecipitation ChIP of Histone Modifications from Saccharomyces cerevisiae

Here, we describe a protocol for chromatin immunoprecipitation of modified histones from the budding yeast Saccharomyces cerevisiae. Immunoprecipitated DNA is subsequently used for quantitative PCR to interrogate the abundance and localization of histone post-translational modifications throughout the genome.

Histone post-translational modifications (PTMs), such as acetylation, methylation and phosphorylation, are dynamically regulated by a series of enzymes ...

Promoter Capture Hi-C: High-resolution, Genome-wide Profiling of Promoter Interactions

The three-dimensional organization of the genome is linked to its function. For example, regulatory elements such as transcriptional enhancers control the spatio-temporal expression of their target genes through physical contact, often bridging considerable (in some cases hundreds of kilobases) genomic distances and bypassing nearby genes. The human genome harbors an estimated one million enhancers, the vast majority of which have unknown gene targets. Assigning distal regulatory regions to their target genes is thus crucial to understand gene expression control. We developed Promoter Capture Hi-C (PCHi-C) to enable the genome-wide detection of distal promoter-interacting regions (PIRs), for all promoters in a single experiment. In PCHi-C, highly complex Hi-C libraries are specifically enriched for promoter sequences through in-solution hybrid selection with thousands of biotinylated RNA baits complementary to the ends of all promoter-containing restriction fragments. The aim is to then pull-down promoter sequences and their frequent interaction partners such as enhancers and other potential regulatory elements. After high-throughput paired-end sequencing, a statistical test is applied to each promoter-ligated restriction fragment to identify significant PIRs at the restriction fragment level. We have used PCHi-C to generate an atlas of long-range promoter interactions in dozens of human and mouse cell types. These promoter interactome maps have contributed to a greater understanding of mammalian gene expression control by assigning putative regulatory regions to their target genes and revealing preferential spatial promoter-promoter interaction networks. This information also has high relevance to understanding human genetic disease and the identification of potential disease genes, by linking non-coding disease-associated sequence variants in or near control sequences to their target genes.

DNA regulatory elements, such as enhancers, control gene expression by physically contacting target gene promoters, often through long-range chromosomal interactions spanning large genomic distances. Promoter Capture Hi-C (PCHi-C) identifies significant interactions between promoters and distal regions, enabling the assignment of potential regulatory sequences to their target genes.

The three-dimensional organization of the genome is linked to its function. For example, regulatory elements such as transcriptional enhancers control the ...

Genome-wide Surveillance of Transcription Errors in Eukaryotic Organisms

Accurate transcription is required for the faithful expression of genetic information. Surprisingly though, little is known about the mechanisms that control the fidelity of transcription. To fill this gap in scientific knowledge, we recently optimized the circle-sequencing assay to detect transcription errors throughout the transcriptome of Saccharomyces cerevisiae, Drosophila melanogaster, and Caenorhabditis elegans. This protocol will provide researchers with a powerful new tool to map the landscape of transcription errors in eukaryotic cells so that the mechanisms that control the fidelity of transcription can be elucidated in unprecedented detail.

This protocol provides researchers with a new tool to monitor the fidelity of transcription in multiple model organisms.

Accurate transcription is required for the faithful expression of genetic information. Surprisingly though, little is known about the mechanisms that ...

Saccharomyces cerevisiae Metabolic Labeling with 4-thiouracil and the Quantification of Newly Synthesized mRNA As a Proxy for RNA Polymerase II Activity

Global defects in RNA polymerase II transcription might be overlooked by transcriptomic studies analyzing steady-state RNA. Indeed, the global decrease in mRNA synthesis has been shown to be compensated by a simultaneous decrease in mRNA degradation to restore normal steady-state levels. Hence, the genome-wide quantification of mRNA synthesis, independently from mRNA decay, is the best direct reflection of RNA polymerase II transcriptional activity. Here, we discuss a method using non-perturbing metabolic labeling of nascent RNAs in Saccharomyces cerevisiae (S. cerevisiae). Specifically, the cells are cultured for 6 min with a uracil analog, 4-thiouracil, and the labeled newly transcribed RNAs are purified and quantified to determine the synthesis rates of all individual mRNA. Moreover, using labeled Schizosaccharomyces pombe cells as internal standard allows comparing mRNA synthesis in different S. cerevisiae strains. Using this protocol and fitting the data with a dynamic kinetic model, the corresponding mRNA decay rates can be determined.

Saccharomyces cerevisiae Metabolic Labeling with 4-thiouracil and the Quantification of Newly Synthesized mRNA As a Proxy for RNA Polymerase II Activity

The protocol described here is based on the genome-wide quantification of newly synthesized mRNA purified from yeast cells labeled with 4-thiouracil. This method allows to measure mRNA synthesis uncoupled from mRNA decay and, thus, provides an accurate measurement of RNA polymerase II transcription.

Global defects in RNA polymerase II transcription might be overlooked by transcriptomic studies analyzing steady-state RNA. Indeed, the global decrease in ...

Genetic Mapping of Thermotolerance Differences Between Species of Saccharomyces Yeast via Genome-Wide Reciprocal Hemizygosity Analysis

A central goal of modern genetics is to understand how and why organisms in the wild differ in phenotype. To date, the field has advanced largely on the strength of linkage and association mapping methods, which trace the relationship between DNA sequence variants and phenotype across recombinant progeny from matings between individuals of a species. These approaches, although powerful, are not well suited to trait differences between reproductively isolated species. Here we describe a new method for genome-wide dissection of natural trait variation that can be readily applied to incompatible species. Our strategy, RH-seq, is a genome-wide implementation of the reciprocal hemizygote test. We harnessed it to identify the genes responsible for the striking high temperature growth of the yeast Saccharomyces cerevisiae relative to its sister species S. paradoxus. RH-seq utilizes transposon mutagenesis to create a pool of reciprocal hemizygotes, which are then tracked through a high-temperature competition via high-throughput sequencing. Our RH-seq workflow as laid out here provides a rigorous, unbiased way to dissect ancient, complex traits in the budding yeast clade, with the caveat that resource-intensive deep sequencing is needed to ensure genomic coverage for genetic mapping. As sequencing costs drop, this approach holds great promise for future use across eukaryotes.

Genetic Mapping of Thermotolerance Differences Between Species of Saccharomyces Yeast via Genome-Wide Reciprocal Hemizygosity Analysis

Reciprocal hemizygosity via sequencing (RH-seq) is a powerful new method to map the genetic basis of a trait difference between species. Pools of hemizygotes are generated by transposon mutagenesis and their fitness is tracked through competitive growth using high-throughout sequencing. Analysis of the resulting data pinpoints genes underlying the trait.

A central goal of modern genetics is to understand how and why organisms in the wild differ in phenotype. To date, the field has advanced largely on the ...