The authors would like to thank Tilman Borggrefe (University of Giessen) for kindly sharing the sonication device. The work from M.L.S. is supported by grants from the Deutsche Forschungsgemeinschaft (SCHM 1417/8-3), SFB/TRR81, SFB1021, SFB1213, the Excellence Cluster Cardio-Pulmonary System (ECCPS, EXC 147/2), the Deutsche Krebshilfe (111447) and the IMPRS-HLR program of the Max-Planck Society.

1. Day 1: Cell Culture
<ol>
	<li>Culture the cells to be analyzed to the desired amount. Cell culture conditions and media depend on the cell types under investigation. 
	NOTE: In principle, any eukaryotic cell is amenable to an analysis of chromatin accessibility by FAIRE. For this experiment, 4 x 106 HEK-293T cells are seeded in a single 10 cm dish in DMEM medium supplemented with 10% (v/v) FBS and 1% (w/v) penicillin/streptomycin, and incubated at 5% CO2 and 37 &#176;C for 24 h until the cells reach 70 - 80% confluence (~ 8 x 106 cells per dish).</li>
</ol>
2. Day 2: Formaldehyde Crosslinking and Cell Harvesting
CAUTION: Formaldehyde is highly toxic and must be always used under a fume hood. Wear protective clothes (gloves and lab coat). Discard the waste appropriately.
<ol>
	<li>Add formaldehyde directly to the cell culture medium in a fume hood to reach a final concentration of 1% (v/v) (270 &#181;L of 37% (v/v) formaldehyde to 10 mL of cell culture medium). Incubate for 10 min at room temperature (20 - 25 &#176;C) and slew the plates manually every 2 min. 
	NOTE: The crosslinking time can be adjusted according to the cell type. For the majority of cell lines, 10 min of crosslinking is adequate. For tissues, longer incubations might be required to allow the fixation of all cells.</li>
	<li>Add glycine to a final concentration of 0.125 M to quench the formaldehyde (add 500 &#181;L of a 2.5 M glycine stock to 10 mL of culture medium). Incubate for 5 min at room temperature and slew every 2 min.</li>
	<li>Wash the cells 3x with ice cold PBS (8 g/L NaCl, 0.2 g/L KCl, 1.44 g/L Na2HPO4 &#183; 2H2O, 0.24 g/L KH2PO4, adjust pH to 7.4 with HCl or NaOH). Aspirate the medium and wash directly in the cell culture dish or flask by adding 10 mL of PBS. Repeat this step twice being very careful in the case of loosely adherent cells such as HEK-293T. Add PBS carefully to the side of the dish in order to avoid detachment of the cells.</li>
	<li>After the third wash, resuspend the cell in 1 mL of ice cold PBS and transfer to a 1.5 mL reaction tube on ice. Use a cell scraper to detach the cells when necessary. 
	NOTE: When starting with a limited material, use low DNA-binding tubes to avoid sample loss during the procedure.</li>
	<li>Centrifuge for 5 min at 300 x g at 4 &#176;C in a cooled tabletop centrifuge. Discard the supernatant by carefully aspirating it.</li>
</ol>
3. Cell Lysis and Chromatin Fragmentation
<ol>
	<li>Resuspend the cell pellet in 1 mL FAIRE lysis buffer (1% (w/v), 10 mM EDTA, 50 mM Tris HCl pH 8.1, 10 &#181;g/mL leupeptin, 10 &#181;g/mL aprotinin, 2 mM PMSF), and resuspend the cells by carefully pipetting up and down several times. Incubate for 20 min on ice. Store the samples at -80 &#176;C at this step if needed.</li>
	<li>Sonicate the crosslinked DNA to shear it to an average size of 200 - 300 bp. The specific settings of the sonicator must be determined for each source of samples and the used sonication device. In any case ensure that the DNA is efficiently sheared. An optimization step for DNA shearing is described under step 5.14. Cool the samples on ice during sonication to avoid heating of the sample.</li>
	<li>Centrifuge the sample for 15 min and 13,000 x g at 4 &#176;C.</li>
	<li>Transfer the supernatant to a new reaction tube. Split the samples in 100 &#181;L aliquots. Store the samples at -80 &#176;C, if needed. 
	NOTE: The protocol can be interrupted at this point.</li>
</ol>
4. De-crosslinking of Control DNA
<ol>
	<li>Take a 100 &#181;L aliquot from step 3.4 to reverse the crosslink and to be used as a reference for total DNA. Add 10 &#181;L of RNase-A (10 mg/mL) and incubate for 1 h at 37 &#176;C.</li>
	<li>Add 10 &#181;L of Proteinase K (20 mg/mL). Use a programmable thermo-block to incubate for 4 h at 37 &#176;C and then for 6 h at 65 &#176;C to cleave the proteins and to reverse the crosslinks. Finally, hold the samples at 4 &#176;C until the protocol is resumed and then proceed to step 5. 
	NOTE: Other FAIRE protocols use chromatin samples from cells which have not been crosslinked as reference, thus abolishing the need for de-crosslinking11. The fact that these samples are not crosslinked could lead to differences in the sonication efficiency or in DNA stability, therefore the use of total DNA reference samples that have undergone the same procedure as the test samples is more accurate.</li>
</ol>
5. Day 3: Phenol:Chloroform Purification of DNA
<ol>
	<li>Take the non-de-crosslinked aliquot from step 3.4. Add 10 &#181;L of RNase-A (10 mg/mL) and incubate for 1 h at 37 &#176;C.</li>
	<li>Take the non-de-crosslinked aliquot form step 5.1 and the de-crosslinked sample from step 4.2 and proceed in parallel. Add H2O to reach a final volume of 300 &#181;L.</li>
	<li>Add 300 &#181;L of phenol:chloroform:isoamyl alcohol (25:24:1) in a fume hood and vortex vigorously. 
	CAUTION: Phenol is corrosive and toxic and must be used always under a fume hood. Wear protective clothes (gloves and lab coat), make sure the reaction tubes are tightly closed, and dispose the waste appropriately.</li>
	<li>Centrifuge for 10 min at 13,000 x g at 4 &#176;C.</li>
	<li>Carefully transfer 280 &#181;L of the upper aqueous phase to a new reaction tube. Make sure not to take any debris from the interphase which also contains proteins. Discard the remaining lower phase as organic solvent waste.</li>
	<li>Repeat steps 5.3 to 5.5 and carefully transfer 270 &#181;L of the upper aqueous phase to a new reaction tube.</li>
	<li>Add 270 &#181;L of chloroform in a fume hood and vortex vigorously. 
	NOTE: This step is used to eliminate any remaining phenol from the sample.</li>
	<li>Centrifuge for 10 min at 13,000 x g at 4 &#176;C.</li>
	<li>Carefully transfer 250 &#181;L of the upper aqueous phase to a new reaction tube, taking care not to carry any debris from the interphase. Discard the remaining sample as organic solvent waste.</li>
	<li>Add 25 &#181;L of 5 M NaCl and 250 &#181;L of isopropanol. Add 5 &#181;L of glycogen (20 mg/mL) as the DNA carrier. Mix well by inverting the tubes and incubate for 20 min at room temperature. Centrifuge for 10 min at 13,000 x g at 4 &#176;C to precipitate the DNA.</li>
	<li>Wash the DNA pellet with 400 &#181;L of 70% (v/v) ethanol. Centrifuge for 10 min at 13,000 x g at 4 &#176;C.</li>
	<li>Aspirate the supernatant carefully and let the pellet dry for 10 min at room temperature. Resuspend in 100 &#181;L TE buffer (10 mM Tris pH 8, 1 mM EDTA).</li>
	<li>Incubate the non-de-crosslinked free DNA sample for 4 h at 65 &#176;C to remove inter-DNA crosslinks. Store the samples at -20 &#176;C.</li>
	<li>Quantify the amount of DNA using a spectrophotometer and run an aliquot to check the fragment size on a 2% (w/v) agarose gel. Adjust the shearing time and intensities, depending on the results of the DNA size analysis by agarose gel electrophoresis, to obtain an average size of 200-300 bp.</li>
</ol>
6. Determination of Free Versus Total DNA by Quantitative Real Time PCR (qRT-PCR)
<ol>
	<li>The ratio of nucleosome-free DNA (in the following referred to as free DNA) versus total DNA is determined by qRT-PCR17,18. Samples can also be quantitatively analyzed by microarray hybridization or deep sequencing for genome-wide determinations.</li>
	<li>Design qPCR primers for the regions to be tested and for positive and negative controls. 
	NOTE: Suitable positive controls are primers situated in the promoter of actively transcribed housekeeping genes such as ACTB, GAPDH, TPI1, or TUBA1A (encoding &#946;-Actin, GAPDH, TPI or &#945;-Tubulin). Appropriate negative controls are found in heterochromatin regions or gene deserts. Other suitable negative controls can be designed in genomic regions adjacent to the dynamically regulated locus under investigation to control for local copy number differences (Table 1).</li>
	<li>Dilute the DNA samples to a final concentration of 10 ng/&#181;L with TE buffer, and take the dilution factor into account for the final calculations (see step 6.5).</li>
	<li>Set up a qPCR reaction in triplicates in a 96-well plate, with the following reaction mix per well: template DNA: 20 ng (2 &#181;L), forward primer 200 nM (0.4 &#181;L of a 5 &#181;M stock), reverse primer 200 nM (0.4 &#181;L of a 5 &#181;M stock), a commercial ready to use reaction mix containing green dye (5 &#181;L from a 2x stock), and H2O to 10 &#181;L.</li>
	<li>Run in a real time qPCR device, using the absolute quantification mode, and determine the Ct of the samples with the different primer pairs. Calculate the ratio of recovery of free DNA to total DNA for each primer using the following formula, taking into account the dilution factors from step 6.3: 
	<img alt="Equation 1" src="/files/ftp_upload/57272/57272eq1.jpg" /> 
	Representative results and an example calculation are given in Table 2.</li>
	<li>To compensate for differences among samples, normalize to the positive control recovery ratio: 
	<img alt="Equation 2" src="/files/ftp_upload/57272/57272eq2.jpg" /></li>
</ol>

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The authors have nothing to disclose.

FAIRE is a robust and inexpensive method allowing the identification of nucleosome-free regions. It is based on the principle that DNA wrapped around nucleosomes can easily be crosslinked to the histones. A phenol:chloroform extraction is used to separate the non-crosslinked and protein-free DNA fragments from the protein-bound DNA, which is found in the lower phenol phase and to some extent in the interphase (Figure 1). Therefore, the nucleosome-free regions are overrepresented in the fraction of free DNA in the aqueous phase. A number of publications describe detailed protocols of the FAIRE method23,24,25,26, which can be consulted to complement the procedure described here.
Similar to other methods used to determine the chromatin structure, the FAIRE method also critically depends on optimal shearing of the DNA. If the fragments are too long, any nucleosome-free region will be masked by the adjacent chromatin-bound DNA. If the fragments are too small, the detection by qPCR or deep sequencing becomes very difficult. For this reason, the sonication of the DNA is a crucial parameter that must be controlled and optimized in order to obtain fragments around 200 - 300 bp length, which represent 1 - 2 nucleosomes (Figure 4).
Another parameter that can affect the final outcome is the crosslinking of the sample. Although the fixation procedure described here is valid for most cell lines, the researcher must carefully evaluate this step for the particular sample under study, especially when using tissues, where longer fixation times might be necessary to allow the formaldehyde to reach the cells in all the tissue sample.
Unlike other methods, such as DNase I or micrococcal nuclease hypersensitivity, ChIP, or ATAC, FAIRE does not rely on an enzymatic activity or specific antibodies, and therefore it does not need titration or optimization of the reaction conditions. This makes FAIRE an ideal method to measure chromatin accessibility for labs with little experience in the chromatin field. In addition, pilot experiments are only required in order to optimize the DNA shearing step and the crosslinking time, when necessary.
The method was initially developed in yeast6, but it has also been extended to mammalian cells. The DNA purified with the FAIRE method can be used for deep sequencing, allowing the genome-wide characterization of the chromatin status. This has already proven useful in a number of studies8,9,10,11,12,13,19,27,28.
Results from FAIRE-seq experiments show a large overlap with results obtained by other methods such as DNase I-seq14, although some differences arise. This is due to methodological differences as for example relaxed or remodeled nucleosomes could still hinder the cleavage by DNase I, while these DNA regions would be less prone to produce DNA/protein crosslinks and thus would be determined as nucleosome-free regions using FAIRE9. Also, the presence of non-histone proteins such as RNA polymerases or reader proteins for epigenetic marks could result in DNA/protein crosslinks and thus would be interpreted as protein-packed regions in FAIRE experiments. These limitations are inherent to every method used for the determination of the chromatin state, thus raising the need to use a combination of different methods to obtain a comprehensive insight.

The genomic DNA of eukaryotic cells is tightly packed into nucleosomes, which needs to be removed or remodeled in order to allow the interaction of other proteins with the DNA. The transcriptional activation of a gene typically leads to a more open configuration of its nucleosomal structure, particularly in the +1 nucleosome1, to facilitate transcription by RNA polymerase. Also, regulatory regions such as promoters and enhancers undergo dynamic changes in their chromatin accessibility to allow the interaction with chromatin modifiers or transcription factors2. Several approaches have been developed to identify these nucleosome-free regions. These include the sensitivity to nucleases such as DNase I3 or micrococcal nuclease4, which can only access the DNA when it is not tightly wrapped around nucleosomes. Other methods for the identification of open chromatin or free DNA include the transposase-mediated integration of retrotransposable elements (Assay for Transposase-Accessible Chromatin, ATAC)5, or chromatin immunoprecipitation (ChIP) using antibodies against histones. The FAIRE method is not based on the capacity of an enzyme to reach the DNA or the binding of an antibody to a particular protein, but instead relies on the purification of nucleosome-free regions by a phenol:chloroform extraction6,7. In this method, the DNA is covalently bound to chromatin proteins by the crosslinking agent formaldehyde and is sheared afterwards to small pieces by sonication. Tightly packed chromatin regions will have abundant DNA/protein crosslinks, while DNA regions with no or few nucleosomes will have little or no crosslinked DNA/protein complexes. A phenol:chloroform extraction allows purification of the non-crosslinked free DNA in the aqueous phase, while the DNA crosslinked to proteins is captured in the organic phenol phase or aqueous-organic interphase together with the proteins. The quantification of this free DNA compared to a reference of total DNA sample allows the identification of the chromatin free regions. The FAIRE method can be used for the characterization of individual genomic regions as described here, but is also suitable for the identification of genome-wide chromatin accessibility when coupled to deep sequencing as described in different models8,9,10,11,12,13. Results obtained by FAIRE-seq and other methods such as ATAC-seq are largely overlapping14. The FAIRE method is a very robust, cheap, and easy to perform method to identify nucleosome-free regions. Unlike other methods, it does not require pilot experiments to determine the appropriate level of digestion as required for nucleases (DNase-seq, MNase-seq) or transposases (ATAC-seq) and discussed in detail in a recent review15. The only parameters to be adjusted are the sonication conditions and in some cases the fixation time. This paper describes a straightforward protocol that is schematically displayed in Figure 1 and that does not require any special equipment other than a sonicator. The protocol can be performed in a reasonably short time and makes FAIRE an easy and reliable method to test chromatin accessibility in a number of different cell types ranging from lung cells11 to primary cells obtained from mouse livers16.

<table><tbody><tr><td>Phenol:chloroform:isoamyl alcohol 25:24:1</td><td>Carl Roth</td><td>A156.1</td><td></td></tr><tr><td>Glycogen, 20 mg/ml</td><td>Thermo Fisher</td><td>R0561</td><td></td></tr><tr><td>ABsolute QPCR Mix, SYBR-Green, Rox</td><td>Thermo Fisher</td><td>AB1162B</td><td></td></tr><tr><td>Proteinase K, 20 mg/ml</td><td>Thermo Fisher</td><td>EO0491</td><td></td></tr><tr><td>RNase A, 10 mg/ml</td><td>Thermo Fisher</td><td>EN0531</td><td></td></tr><tr><td>Leupeptin</td><td>Sigma</td><td>L8511</td><td></td></tr><tr><td>Aprotinin</td><td>Sigma</td><td>A1603</td><td></td></tr><tr><td>Phenylmethylsulfonyl fluoride (PMSF)</td><td>Sigma</td><td>78830</td><td></td></tr><tr><td>milliTUBE 1 ml AFA Fiber</td><td>Covaris</td><td>520130</td><td></td></tr><tr><td>Holder milliTUBE 1ml</td><td>Covaris</td><td>500371</td><td></td></tr><tr><td>Covaris S220 Focused-ultrasonicator</td><td>Covaris</td><td></td><td></td></tr><tr><td>StepOnePlus Real-Time PCR System</td><td>Applied Biosistems</td><td>4376600</td><td></td></tr></tbody></table>

formaldehyde-assisted isolation of regulatory elements to measure chromatin accessibility in mammalian cells

Appropriate gene expression in response to extracellular cues, that is, tissue- and lineage-specific gene transcription, critically depends on highly defined states of chromatin organization. The dynamic architecture of the nucleus is controlled by multiple mechanisms and shapes the transcriptional output programs. It is, therefore, important to determine locus-specific chromatin accessibility in a reliable fashion that is preferably independent from antibodies, which can be a potentially confounding source of experimental variability. Chromatin accessibility can be measured by various methods, including the Formaldehyde-Assisted Isolation of Regulatory Elements (FAIRE) assay, that allow the determination of general chromatin accessibility in a relatively low number of cells. Here we describe a FAIRE protocol that allows simple, reliable, and fast identification of genomic regions with a low protein occupancy. In this method, the DNA is covalently bound to the chromatin proteins using formaldehyde as a crosslinking agent and sheared to small pieces. The free DNA is afterwards enriched using phenol:chloroform extraction. The ratio of free DNA is determined by quantitative polymerase chain reaction (qPCR) or DNA sequencing (DNA-seq) compared to a control sample representing total DNA. The regions with a looser chromatin structure are enriched in the free DNA sample, thus allowing the identification of genomic regions with lower chromatin compaction.

We have used the FAIRE method to measure differences in the chromatin accessibility of MHC class II genes in HEK293T cells as published19. MHC class II encoding genes are expressed in antigen presenting cells (APCs), either constitutively or in response to cytokine signaling20. The expression of MHCII genes is driven by an activator complex which binds to specific DNA elements, termed XY boxes, located in the promoter and enhancers of these genes21,22. This is reflected at the level of chromatin, as revealed by our FAIRE analysis of selected regions in the MHC class II genes HLA-DPB1 and HLA-DMA. These experiments showed that chromatin is open at the regions encompassing the XY boxes, while it is more compact in the gene bodies (Figure 2).
As an example, for the calculations, in this particular experiment, we diluted the total DNA sample 10 times (dilution factor 10), and the free DNA sample was not diluted (dilution factor 1) for the qRT-PCR. The Cts obtained for the different regions tested and the calculations to obtain the final recovery ratios and relative recovery ratios are listed in Table 2. These relative recovery ratios were obtained using the ACTB promoter as the positive control. Note that these calculations are for one of the replicates used to generate the results shown in Figure 2.
In a different context, chromatin accessibility was tested in a model for inducible gene expression. In this case, rapid changes of chromatin compaction in response to the pro-inflammatory stimulus tumor necrosis factor (TNF) were measured. HeLa cells were left untreated or stimulated with TNF for 1 h, followed by FAIRE to measure chromatin compaction at the promoter region controlling expression of the gene encoding the TNF-responsive cytokine interleukin 8 (IL8). These results showed a strongly increased chromatin accessibility at the IL8 promoter of TNF-stimulated cells (Figure 3). The accessibility at the IL-8 promoter after TNF stimulation is even higher than that found in the ACTB promoter showing a dramatic chromatin remodeling in this regulatory region. As the recovery ratio obtained in this case is higher than that obtained in the region used as the positive control (ACTB promoter) the relative recovery ratio is higher than one.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57272/57272fig1.jpg" /> 
Figure 1: The FAIRE workflow. Cells are crosslinked directly in the cell culture flask or dish by adding formaldehyde (A). After quenching of the formaldehyde, cells are washed three times with ice cold PBS (B), and transferred to a reaction tube where they are resuspended in FAIRE lysis buffer and sonicated to shear the DNA (C). One aliquot of the extracted chromatin is de-crosslinked by heating at 65 &#176;C, to get the total DNA reference (D), and afterwards the free DNA is extracted using phenol:chloroform both from the crosslinked and the de-crosslinked aliquots (E). Finally, the DNA is quantified, and the ratio of free vs. total DNA is calculated (F). <a href="https://cloudflare.jove.com/files/ftp_upload/57272/57272fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57272/57272fig2.jpg" /> 
Figure 2: Determination of chromatin accessibility in the MHC class II gene cluster in HEK293T cells. HEK293T cells were analyzed by FAIRE. The ratio between free versus total DNA (i.e., the relative recovery ratio) was determined for the promoter of the ACTB gene as a positive control, a heterochromatin region on Chr. 12 as negative control, and the regulatory regions and gene bodies of the MHC class II genes HLA-DMA and HLA-DPB1. The upper part shows a schematic representation of the locus and the positions of the primers. Error bars represent standard deviations from two biological replicates measured in triplicates. The figure was modified from a published report19. <a href="https://cloudflare.jove.com/files/ftp_upload/57272/57272fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57272/57272fig3.jpg" /> 
Figure 3: Changes of DNA accessibility at the IL8 promoter in response to TNF treatment. HeLa cells were stimulated with TNF (20 ng/mL) for 1 h and analyzed by FAIRE. The ratio between free versus total DNA was determined for the promoter of ACTB (positive control), a heterochromatin region on Chr. 12 (negative control), and the IL8 promoter. Error bars represent standard errors of the mean (SEM) from three biological replicates measured in duplicates. <a href="https://cloudflare.jove.com/files/ftp_upload/57272/57272fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57272/57272fig4.jpg" /> 
Figure 4: Optimization of the sonication conditions. Chromatin extracted from 293T cells was sonified using a focused sonicator with appropriate tubes (Table of Materials). The chromatin was sonicated for the indicated time with repetitions of 30 s with the following settings: peak power 150, duty factor 15, cycles per burst 500, followed by 30 s: peak power 2.5, duty factor 15, cycles per burst 500. The resulting chromatin was de-crosslinked, purified by phenol:chloroform extraction and loaded in a 2% agarose gel. An ethidium-bromide stained gel is shown, and the positions of the molecular weight markers are indicated in bp. In this experiment, the optimal chromatin size (200 - 300 bp) was obtained after 20 min of sonication. <a href="https://cloudflare.jove.com/files/ftp_upload/57272/57272fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Primer</td>
			<td>Sequence</td>
		</tr>
		<tr>
			<td>ACTB-Promoter-F</td>
			<td>AAAGGCAACTTTCGGAACGG</td>
		</tr>
		<tr>
			<td>ACTB-Promoter-R</td>
			<td>TTCCTCAATCTCGCTCTCGC</td>
		</tr>
		<tr>
			<td>Chr. 12 negative control-F</td>
			<td>ATGGTTGCCACTGGGGATCT</td>
		</tr>
		<tr>
			<td>Chr. 12 negative control- R</td>
			<td>TGCCAAAGCCTAGGGGAAGA</td>
		</tr>
		<tr>
			<td>HLA-DMA-XY-Box-F</td>
			<td>CATCAGTCACTGGGGAGACG</td>
		</tr>
		<tr>
			<td>HLA-DMA-XY-Box-R</td>
			<td>GCTTCCCAGCCCAGTTACAT</td>
		</tr>
		<tr>
			<td>HLA-DMA-5&#8217;-F</td>
			<td>GGAGAGAACAATCTCCGCTTCA</td>
		</tr>
		<tr>
			<td>HLA-DMA-5&#8217;-R</td>
			<td>AGCTGCTATGTGTGGTTGGT</td>
		</tr>
		<tr>
			<td>HLA-DMA-3&#8217;-F</td>
			<td>TGGGGACCTAGTTAGGGAGC</td>
		</tr>
		<tr>
			<td>HLA-DMA-3&#8217;-R</td>
			<td>AGATCCATGGGAGGAGGCTT</td>
		</tr>
		<tr>
			<td>HLA-DPB1-XY-Box-F</td>
			<td>GTCCAATCCCAGGGTCACAG</td>
		</tr>
		<tr>
			<td>HLA-DPB1-XY-Box-R</td>
			<td>TGAAAAGAGCTGCAGTCAGGA</td>
		</tr>
		<tr>
			<td>HLA-DPB1-5&#8217;-F</td>
			<td>GCGTGTTCATGTCTGCATCC</td>
		</tr>
		<tr>
			<td>HLA-DPB1-5&#8217;-R</td>
			<td>TGATCCTCAGAGCCTGGACA</td>
		</tr>
		<tr>
			<td>HLA-DPB1-3&#8217;-F</td>
			<td>CCAGCCTAGGGTGAATGTTT</td>
		</tr>
		<tr>
			<td>HLA-DPB1-3&#8217;-R</td>
			<td>GCCTGGGTAGAAATCCGTCA</td>
		</tr>
		<tr>
			<td>IL8 Promoter-F</td>
			<td>GTGATGACTCAGGTTTGCCCT</td>
		</tr>
		<tr>
			<td>IL8 Promoter-R</td>
			<td>CTTATGGAGTGCTCCGGTGG</td>
		</tr>
	</tbody>
</table>
Table 1: Primers for qPCR.
<img alt="Table 2" src="/files/ftp_upload/57272/57272table2.jpg" /> 
Table 2: Example calculations of FAIRE Relative Recovery Ratios. <a href="http://cloudflare.jove.com/files/ftp_upload/57272/Table_2.xlsx">Please click here to download this file.</a>

Watch this Scientific Journal Video about Formaldehyde-assisted Isolation of Regulatory Elements to Measure Chromatin Accessibility in Mammalian Cells at JoVE.com

Formaldehyde-assisted Isolation of Regulatory Elements to Measure Chromatin Accessibility in Mammalian Cells

The plasticity of nuclear architecture is controlled by dynamic epigenetic mechanisms including non-coding RNAs, DNA methylation, nucleosome repositioning as well as histone composition and modification. Here we describe a Formaldehyde-Assisted Isolation of Regulatory Elements (FAIRE) protocol which allows the determination of chromatin accessibility in a reproducible, inexpensive, and straightforward manner.

Appropriate gene expression in response to extracellular cues, that is, tissue- and lineage-specific gene transcription, critically depends on highly ...

formaldehyde-assisted-isolation-regulatory-elements-to-measure

Research

JoVE Journal

Genetics

11.3K Views.  University Hospital Virgen del Rocío. De plasticiteit van nucleaire het platform wordt beheerd door dynamische Epigenetische mechanismen, met inbegrip van niet-coderende RNAs, DNA methylering, nucleosoom herpositionering evenals Histon samenstelling en wijziging. Hier beschrijven we een Formaldehyde-Assisted isolatie van regelgevende elementen (FAIRE)-protocol, waarmee de bepaling van de chromatine toegankelijkheid in een reproduceerbare, goedkope en eenvoudige wijze.

Video: Formaldehyde-assisted Isolation of Regulatory Elements to Measure Chromatin Accessibility in Mammalian Cells

Sequential Salt Extractions for the Analysis of Bulk Chromatin Binding Properties of Chromatin Modifying Complexes

Elucidation of the binding properties of chromatin-targeting proteins can be very challenging due to the complex nature of chromatin and the heterogeneous nature of most mammalian chromatin-modifying complexes. In order to overcome these hurdles, we have adapted a sequential salt extraction (SSE) assay for evaluating the relative binding affinities of chromatin-bound complexes. This easy and straightforward assay can be used by non-experts to evaluate the relative difference in binding affinity of two related complexes, the changes in affinity of a complex when a subunit is lost or an individual domain is inactivated, and the change in binding affinity after alterations to the chromatin landscape. By sequentially re-suspending bulk chromatin in increasing amounts of salt, we are able to profile the elution of a particular protein from chromatin. Using these profiles, we are able to determine how alterations in a chromatin-modifying complex or alterations to the chromatin environment affect binding interactions. Coupling SSE with other in vitro and in vivo assays, we can determine the roles of individual domains and proteins on the functionality of a complex in a variety of chromatin environments.

Sequential salt extraction of chromatin bound proteins is a useful tool for determining the binding properties of large protein complexes. This method can be employed to evaluate the role of individual subunits or domains in the overall affinity of a protein complex to bulk chromatin.

Elucidation of the binding properties of chromatin-targeting proteins can be very challenging due to the complex nature of chromatin and the heterogeneous ...

Adaptation of Hybridization Capture of Chromatin-associated Proteins for Proteomics to Mammalian Cells

The hybridization capture of chromatin-associated proteins for proteomics (HyCCAPP) technology was initially developed to uncover novel DNA-protein interactions in yeast. It allows analysis of a target region of interest without the need for prior knowledge about likely proteins bound to the target region. This, in theory, allows HyCCAPP to be used to analyze any genomic region of interest, and it provides sufficient flexibility to work in different cell systems. This method is not meant to study binding sites of known transcription factors, a task better suited for Chromatin Immunoprecipitation (ChIP) and ChIP-like methods. The strength of HyCCAPP lies in its ability to explore DNA regions for which there is limited or no knowledge about the proteins bound to it. It can also be a convenient method to avoid biases (present in ChIP-like methods) introduced by protein-based chromatin enrichment using antibodies. Potentially, HyCCAPP can be a powerful tool to uncover truly novel DNA-protein interactions. To date, the technology has been predominantly applied to yeast cells or to high copy repeat sequences in mammalian cells. In order to become the powerful tool we envision, HyCCAPP approaches need to be optimized to efficiently capture single-copy loci in mammalian cells. Here, we present our adaptation of the initial yeast HyCCAPP capture protocol to human cell lines, and show that single-copy chromatin regions can be efficiently isolated with this modified protocol.

This is a method to identify novel DNA-interacting proteins at specific target loci, relying on sequence-specific capture of crosslinked chromatin for subsequent proteomic analyses. No prior knowledge about potential binding proteins, nor cell modifications are required. Initially developed for yeast, the technology has now been adapted for mammalian cells.

The hybridization capture of chromatin-associated proteins for proteomics (HyCCAPP) technology was initially developed to uncover novel DNA-protein ...

Improved Protocol for Chromatin Immunoprecipitation from Mouse Skeletal Muscle

We describe an efficient and reproducible protocol for the preparation of chromatin from adult mouse skeletal muscle, a physically resistant tissue with a high content of structural proteins. Dissected limb muscles from adult mice are physically disrupted by mechanical homogenisation, or a combination of mincing and douncing, in a hypotonic buffer before formaldehyde fixation of the cell lysate. The fixed nuclei are purified by further cycles of mechanical homogenisation or douncing and sequential filtrations to remove cell debris. The purified nuclei can be sonicated immediately or at a later stage after freezing. The chromatin can be efficiently sonicated and is suitable for chromatin immunoprecipitation experiments, as illustrated by the profiles obtained for transcription factors, RNA polymerase II, and covalent histone modifications. The binding events detected using chromatin prepared by this protocol are predominantly those taking place in the muscle fiber nuclei despite the presence of chromatin from other fiber-associated satellite and endothelial cells. This protocol is therefore adapted to study gene regulation in the adult mouse skeletal muscle.

A novel protocol for the preparation of chromatin from adult mouse skeletal muscle adapted to the study of gene regulation in muscle fibers by chromatin immunoprecipitation is presented.

We describe an efficient and reproducible protocol for the preparation of chromatin from adult mouse skeletal muscle, a physically resistant tissue with a ...

Biochemistry

Chromatin Immunoprecipitation Assay Using Micrococcal Nucleases in Mammalian Cells

To express cellular phenotypes in organisms, living cells execute gene expression accordingly, and transcriptional programs play a central role in gene expression. The cellular transcriptional machinery and its chromatin modification proteins coordinate to regulate transcription. To analyze transcriptional regulation at the molecular level, several experimental methods such as electrophoretic mobility shift, transient reporter and chromatin immunoprecipitation (ChIP) assays are available. We describe a modified ChIP assay in detail in this article because of its advantages in directly showing histone modifications and the interactions between proteins and DNA in cells. One of the key steps in a successful ChIP assay is chromatin shearing. Although sonication is commonly used for shearing chromatin, it is difficult to identify reproducible conditions. Instead of shearing chromatin by sonication, we utilized enzymatic digestion with micrococcal nuclease (MNase) to obtain more reproducible results. In this article, we provide a straightforward ChIP assay protocol using MNase.

Chromatin immunoprecipitation (ChIP) is a powerful tool for understanding the molecular mechanisms of gene regulation. However, the method involves difficulties in obtaining reproducible chromatin fragmentation by mechanical shearing. Here, we provide an improved protocol for a ChIP assay using enzymatic digestion.

To express cellular phenotypes in organisms, living cells execute gene expression accordingly, and transcriptional programs play a central role in gene ...

Locus-Specific Chromatin Isolation from Yeast Cells

Single-Copy Gene Locus Chromatin Purification in Saccharomyces cerevisiae

The basic organizational unit of eukaryotic chromatin is the nucleosome core particle (NCP), which comprises DNA wrapped ~1.7 times around a histone octamer. Chromatin is defined as the entity of NCPs and numerous other protein complexes, including transcription factors, chromatin remodeling and modifying enzymes. It is still unclear how these protein-DNA interactions are orchestrated at the level of specific genomic loci during different stages of the cell cycle. This is mainly due to the current technical limitations, which make it challenging to obtain precise measurements of such dynamic interactions. Here, we describe an improved method combining site-specific recombination with an efficient single-step affinity purification protocol to isolate a single-copy gene locus of interest in its native chromatin state. The method allows for the robust enrichment of the target locus over genomic chromatin, making this technique an effective strategy for identifying and quantifying protein interactions in an unbiased and systematic manner, for example by mass spectrometry. Further to such compositional analyses, native chromatin purified by this method likely reflects the in vivo situation regarding nucleosome positioning and histone modifications and is, therefore, amenable to further structural and biochemical analyses of chromatin derived from virtually any genomic locus in yeast.

Author Spotlight: Advancing Chromatin Research and Overcoming Limitations with a High-Enrichment Locus-Specific Chromatin Isolation Protocol 

This protocol presents a locus-specific chromatin isolation method based on site-specific recombination to purify a single-copy gene locus of interest in its native chromatin context from budding yeast, Saccharomyces cerevisiae.

The basic organizational unit of eukaryotic chromatin is the nucleosome core particle (NCP), which comprises DNA wrapped ~1.7 times around a histone ...

Repressing Gene Transcription by Redirecting Cellular Machinery with Chemical Epigenetic Modifiers

Regulation of chromatin compaction is an important process that governs gene expression in higher eukaryotes. Although chromatin compaction and gene expression regulation are commonly disrupted in many diseases, a locus-specific, endogenous, and reversible method to study and control these mechanisms of action has been lacking. To address this issue, we have developed and characterized novel gene-regulating bifunctional molecules. One component of the bifunctional molecule binds to a DNA-protein anchor so that it will be recruited to an allele-specific locus. The other component engages endogenous cellular chromatin-modifying machinery, recruiting these proteins to a gene of interest. These small molecules, called chemical epigenetic modifiers (CEMs), are capable of controlling gene expression and the chromatin environment in a dose-dependent and reversible manner. Here, we detail a CEM approach and its application to decrease gene expression and histone tail acetylation at a Green Fluorescent Protein (GFP) reporter located at the Oct4 locus in mouse embryonic stem cells (mESCs). We characterize the lead CEM (CEM23) using fluorescent microscopy, flow cytometry, and chromatin immunoprecipitation (ChIP), followed by a quantitative polymerase chain reaction (qPCR). While the power of this system is demonstrated at the Oct4 locus, conceptually, the CEM technology is modular and can be applied in other cell types and at other genomic loci.

Regulation of the chromatin environment is an essential process required for proper gene expression. Here, we describe a method for controlling gene expression through the recruitment of chromatin-modifying machinery in a gene-specific and reversible manner.

Regulation of chromatin compaction is an important process that governs gene expression in higher eukaryotes. Although chromatin compaction and gene ...

Bioengineering

Isolation and Cultivation of Neural Progenitors Followed by Chromatin-Immunoprecipitation of Histone 3 Lysine 79 Dimethylation Mark

Brain development is a complex process, which is controlled in a temporo-spatial manner by gradients of morphogens and different transcriptional programs. Additionally, epigenetic chromatin modifications, like histone methylation, have an important role for establishing and maintaining specific cell fates within this process. The vast majority of histone methylation occurs on the flexible histone tail, which is accessible to histone modifiers, erasers, and histone reader proteins. In contrast, H3K79 methylation is located in the globular domain of histone 3 and is implicated in different developmental functions. H3K79 methylation is evolutionarily conserved and can be found in a wide range of species from Homo sapiens to Saccharomyces cerevisiae. The modification occurs in different cell populations within organisms, including neural progenitors. The location of H3K79 methylation in the globular domain of histone 3 makes it difficult to assess. Here, we present methods to isolate and culture cortical progenitor cells (CPCs) from embryonic cortical brain tissue (E11.5-E14.5) or cerebellar granular neuron progenitors (CGNPs) from postnatal tissue (P5-P7), and to efficiently immunoprecipitate H3K79me2 for quantitative PCR (qPCR) and genome-wide sequencing.

We present an effective and reproducible method to isolate and culture neural progenitor cells from embryonic and postnatal brain tissue for chromatin immunoprecipitation (ChIP) of histone 3 lysine 79 dimethylation (H3K79me2) - a histone mark located within the globular domain of histone 3.

Brain development is a complex process, which is controlled in a temporo-spatial manner by gradients of morphogens and different transcriptional programs. ...

Neuroscience

Efficient Chromatin Immunoprecipitation using Limiting Amounts of Biomass

Chromatin immunoprecipitation (ChIP) is a widely-used method for determining the interactions of different proteins with DNA in chromatin of living cells. Examples include sequence-specific DNA binding transcription factors, histones and their different modification states, enzymes such as RNA polymerases and ancillary factors, and DNA repair components. Despite its ubiquity, there is a lack of up-to-date, detailed methodologies for both bench preparation of material and for accurate analysis allowing quantitative metrics of interaction. Due to this lack of information, and also because, like any immunoprecipitation, conditions must be re-optimized for new sets of experimental conditions, the ChIP assay is susceptible to inaccurate or poorly quantitative results.
Our protocol is ultimately derived from seminal work on transcription factor:DNA interactions1,2 , but incorporates a number of improvements to sensitivity and reproducibility for difficult-to-obtain cell types. The protocol has been used successfully3,4 , both using qPCR to quantify DNA enrichment, or using a semi-quantitative variant of the below protocol.
This quantitative analysis of PCR-amplified material is performed computationally, and represents a limiting factor in the assay. Important controls and other considerations include the use of an isotype-matched antibody, as well as evaluation of a control region of genomic DNA, such as an intergenic region predicted not to be bound by the protein under study (or anticipated not to show changes under the experimental conditions). In addition, a standard curve of input material for every ChIP sample is used to derive absolute levels of enrichment in the experimental material. Use of standard curves helps to take into account differences between primer sets, regardless of how carefully they are designed, and also efficiency differences throughout the range of template concentrations for a single primer set. Our protocol is different from others that are available5-8 in that we extensively cover the later, analysis phase.

We describe a robust method for chromatin immunoprecipitation using primary T cells. The method is founded on standard approaches, but uses a specific set of conditions and reagents that improve efficiency for limited a quantities of cells. Importantly, a detailed description of the data analysis phase is presented.

Chromatin immunoprecipitation (ChIP) is a  widely-used method for determining the interactions of different proteins with  DNA in chromatin of living ...

Biology

A Cell Free Assay to Study Chromatin Decondensation at the End of Mitosis

During the vertebrate cell cycle chromatin undergoes extensive structural and functional changes. Upon mitotic entry, it massively condenses into rod shaped chromosomes which are moved individually by the mitotic spindle apparatus. Mitotic chromatin condensation yields chromosomes compacted fifty-fold denser as in interphase. During exit from mitosis, chromosomes have to re-establish their functional interphase state, which is enclosed by a nuclear envelope and is competent for replication and transcription. The decondensation process is morphologically well described, but in molecular terms poorly understood: We lack knowledge about the underlying molecular events and to a large extent the factors involved as well as their regulation. We describe here a cell-free system that faithfully recapitulates chromatin decondensation in vitro, based on mitotic chromatin clusters purified from synchronized HeLa cells and X. laevis egg extract. Our cell-free system provides an important tool for further molecular characterization of chromatin decondensation and its co-ordination with processes simultaneously occurring during mitotic exit such as nuclear envelope and pore complex re-assembly.

The molecular mechanisms of the decondensation of highly compacted mitotic chromatin are ill-defined. We present a cell-free assay based on mitotic chromatin clusters isolated from HeLa cells and Xenopus laevis egg extract that faithfully reconstitutes the decondensation process in vitro.

During the vertebrate cell cycle chromatin undergoes extensive structural and functional changes. Upon mitotic entry, it massively condenses into rod ...

Isolation of Specific Genomic Regions and Identification of Associated Molecules by enChIP

The identification of molecules associated with specific genomic regions of interest is required to understand the mechanisms of regulation of the functions of these regions. To enable the non-biased identification of molecules interacting with a specific genomic region of interest, we recently developed the engineered DNA-binding molecule-mediated chromatin immunoprecipitation (enChIP) technique. Here, we describe how to use enChIP to isolate specific genomic regions and identify the associated proteins and RNAs. First, a genomic region of interest is tagged with a transcription activator-like (TAL) protein or a clustered regularly interspaced short palindromic repeats (CRISPR) complex consisting of a catalytically inactive form of Cas9 and a guide RNA. Subsequently, the chromatin is crosslinked and fragmented by sonication. The tagged locus is then immunoprecipitated and the crosslinking is reversed. Finally, the proteins or RNAs that are associated with the isolated chromatin are subjected to mass spectrometric or RNA sequencing analyses, respectively. This approach allows the successful identification of proteins and RNAs associated with a genomic region of interest.

The identification of molecules associated with specific genomic regions of interest is required to understand the mechanisms of regulation of the functions of these regions. This protocol describes procedures to perform engineered DNA-binding molecule-mediated chromatin imunoprecipitation (enChIP) for identification of proteins and RNAs associated with a specific genomic region.

Formaldehyde-assisted Isolation of Regulatory Elements to Measure Chromatin Accessibility in Mammalian Cells

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