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 border="0" cellpadding="0" cellspacing="0" width="849">
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:368px;">Phenol:chloroform:isoamyl alcohol 25:24:1</td>
			<td style="width:143px;">Carl Roth</td>
			<td style="width:133px;">A156.1</td>
			<td style="width:205px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Glycogen, 20 mg/ml</td>
			<td>Thermo Fisher</td>
			<td>R0561</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">ABsolute QPCR Mix, SYBR-Green, Rox</td>
			<td>Thermo Fisher</td>
			<td>AB1162B</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Proteinase K, 20 mg/ml</td>
			<td>Thermo Fisher</td>
			<td>EO0491</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">RNase A, 10 mg/ml</td>
			<td>Thermo Fisher</td>
			<td>EN0531</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Leupeptin</td>
			<td>Sigma</td>
			<td>L8511</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Aprotinin</td>
			<td>Sigma</td>
			<td>A1603</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Phenylmethylsulfonyl fluoride (PMSF)</td>
			<td>Sigma</td>
			<td>78830</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">milliTUBE 1 ml AFA Fiber</td>
			<td>Covaris</td>
			<td>520130</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Holder milliTUBE 1ml</td>
			<td>Covaris</td>
			<td>500371</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Covaris S220 Focused-ultrasonicator</td>
			<td>Covaris</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">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

11.2K Views.  University Hospital Virgen del Rocío. 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.

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 ...

Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide RNA (sgRNA) that confers specificity, the Cas9 protein cleaves both DNA strands at the targeted locus. The DNA break can trigger either non-homologous end joining (NHEJ) or homology directed repair (HDR). NHEJ can introduce small deletions or insertions which lead to frame-shift mutations, while HDR allows for larger and more precise perturbations. Here, we present protocols for generating knockout cell lines by coupling established CRISPR/Cas9 methods with two options for downstream selection/screening. The NHEJ approach uses a single sgRNA cut site and selection-independent screening, where protein production is assessed by dot immunoblot in a high-throughput manner. The HDR approach uses two sgRNA cut sites that span the gene of interest. Together with a provided HDR template, this method can achieve deletion of tens of kb, aided by the inserted selectable resistance marker. The appropriate applications and advantages of each method are discussed.

Recent advances in the ability to genetically manipulate somatic cell lines hold great potential for basic and applied research. Here, we present two approaches for CRISPR/Cas9 generated knockout production and screening in mammalian cell lines, with and without the use of selectable markers.

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide ...

Chromatin Immunoprecipitation (ChIP) in Mouse T-cell Lines

Signaling pathways regulate gene expression programs via the modulation of the chromatin structure at different levels, such as by post-translational modifications (PTMs) of histone tails, the exchange of canonical histones with histone variants, and nucleosome eviction. Such regulation requires the binding of signal-sensitive transcription factors (TFs) that recruit chromatin-modifying enzymes at regulatory elements defined as enhancers. Understanding how signaling cascades regulate enhancer activity requires a comprehensive analysis of the binding of TFs, chromatin modifying enzymes, and the occupancy of specific histone marks and histone variants. Chromatin immunoprecipitation (ChIP) assays utilize highly specific antibodies to immunoprecipitate specific protein/DNA complexes. The subsequent analysis of the purified DNA allows for the identification the region occupied by the protein recognized by the antibody. This work describes a protocol to efficiently perform ChIP of histone proteins in a mature mouse T-cell line. The presented protocol allows for the performance of ChIP assays in a reasonable timeframe and with high reproducibility.

Chromatin Immunoprecipitation ChIP in Mouse T-cell Lines

This work describes a protocol for chromatin immunoprecipitation (ChIP) using a mature mouse T-cell line. This protocol is suitable to investigate the distribution of specific histone marks at specific promoter sites or genome-wide.

Signaling pathways regulate gene expression programs via the modulation of the chromatin structure at different levels, such as by post-translational ...

Retroviral Scanning: Mapping MLV Integration Sites to Define Cell-specific Regulatory Regions

Moloney murine leukemia (MLV) virus-based retroviral vectors integrate predominantly in acetylated enhancers and promoters. For this reason, mLV integration sites can be used as functional markers of active regulatory elements. Here, we present a retroviral scanning tool, which allows the genome-wide identification of cell-specific enhancers and promoters. Briefly, the target cell population is transduced with an mLV-derived vector and genomic DNA is digested with a frequently cutting restriction enzyme. After ligation of genomic fragments with a compatible DNA linker, linker-mediated polymerase chain reaction (LM-PCR) allows the amplification of the virus-host genome junctions. Massive sequencing of the amplicons is used to define the mLV integration profile genome-wide. Finally, clusters of recurrent integrations are defined to identify cell-specific regulatory regions, responsible for the activation of cell-type specific transcriptional programs.
The retroviral scanning tool allows the genome-wide identification of cell-specific promoters and enhancers in prospectively isolated target cell populations. Notably, retroviral scanning represents an instrumental technique for the retrospective identification of rare populations (e.g. somatic stem cells) that lack robust markers for prospective isolation.

Here, we describe a protocol for genome-wide mapping of the integration sites of Moloney murine leukemia virus-based retroviral vectors in human cells.

Moloney murine leukemia (MLV) virus-based retroviral vectors integrate predominantly in acetylated enhancers and promoters. For this reason, mLV ...

Chromatin Immunoprecipitation (ChIP) Protocol for Low-abundance Embryonic Samples

Chromatin immunoprecipitation (ChIP) is a widely-used technique for mapping the localization of post-translationally modified histones, histone variants, transcription factors, or chromatin-modifying enzymes at a given locus or on a genome-wide scale. The combination of ChIP assays with next-generation sequencing (i.e., ChIP-Seq) is a powerful approach to globally uncover gene regulatory networks and to improve the functional annotation of genomes, especially of non-coding regulatory sequences. ChIP protocols normally require large amounts of cellular material, thus precluding the applicability of this method to investigating rare cell types or small tissue biopsies. In order to make the ChIP assay compatible with the amount of biological material that can typically be obtained in vivo during early vertebrate embryogenesis, we describe here a simplified ChIP protocol in which the number of steps required to complete the assay were reduced to minimize sample loss. This ChIP protocol has been successfully used to investigate different histone modifications in various embryonic chicken and adult mouse tissues using low to medium cell numbers (5 x 104 - 5 x 105 cells). Importantly, this protocol is compatible with ChIP-seq technology using standard library preparation methods, thus providing global epigenomic maps in highly relevant embryonic tissues.

Chromatin Immunoprecipitation ChIP Protocol for Low-abundance Embryonic Samples

Here, we describe a chromatin immunoprecipitation (ChIP) and ChIP-seq library preparation protocol to generate global epigenomic profiles from low-abundance chicken embryonic samples.

Chromatin immunoprecipitation (ChIP) is a widely-used technique for mapping the localization of post-translationally modified histones, histone variants, ...

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 ...

A Simple, Rapid, and Quantitative Assay to Measure Repair of DNA-protein Crosslinks on Plasmids Transfected into Mammalian Cells

The purpose of this method is to provide a flexible, rapid, and quantitative technique to examine the kinetics of DNA-protein crosslink (DPC) repair in mammalian cell lines. Rather than globally assaying removal of xenobiotic-induced or spontaneous chromosomal DPC removal, this assay examines the repair of a homogeneous, chemically defined lesion specifically introduced at one site within a plasmid DNA substrate. Importantly, this approach avoids the use of radioactive materials and is not dependent on expensive or highly-specialized technology. Instead, it relies on standard recombinant DNA procedures and widely available real-time, quantitative polymerase chain reaction (qPCR) instrumentation. Given the inherent flexibility of the strategy utilized, the size of the crosslinked protein, as well as the nature of the chemical linkage and the precise DNA sequence context of the attachment site can be varied to address the respective contributions of these parameters to the overall efficiency of DPC repair. Using this method, plasmids containing a site-specific DPC were transfected into cells and low molecular weight DNA recovered at various times post-transfection. Recovered DNA is then subjected to strand-specific primer extension (SSPE) using a primer complementary to the damaged strand of the plasmid. Since the DPC lesion blocks Taq DNA polymerase, the ratio of repaired to un-repaired DNA can be quantitatively assessed using qPCR. Cycle threshold (CT) values are used to calculate percent repair at various time points in the respective cell lines. This SSPE-qPCR method can also be used to quantitatively assess the repair kinetics of any DNA adduct that blocks Taq polymerase.

The goal of this protocol is to quantify the repair of defined DNA-protein crosslinks on plasmid DNA. Lesioned plasmids are transfected into recipient mammalian cell lines and low-molecular weight harvested at multiple time points post-transfection. DNA repair kinetics are quantified using strand-specific primer extension followed by qPCR.

The purpose of this method is to provide a flexible, rapid, and quantitative technique to examine the kinetics of DNA-protein crosslink (DPC) repair in ...

CRISPR-Mediated Reorganization of Chromatin Loop Structure

Recent studies have clearly shown that long-range, three-dimensional chromatin looping interactions play a significant role in the regulation of gene expression, but whether looping is responsible for or a result of alterations in gene expression is still unknown. Until recently, how chromatin looping affects the regulation of gene activity and cellular function has been relatively ambiguous, and limitations in existing methods to manipulate these structures prevented in-depth exploration of these interactions. To resolve this uncertainty, we engineered a method for selective and reversible chromatin loop re-organization using CRISPR-dCas9 (CLOuD9). The dynamism of the CLOuD9 system has been demonstrated by successful localization of CLOuD9 constructs to target genomic loci to modulate local chromatin conformation. Importantly, the ability to reverse the induced contact and restore the endogenous chromatin conformation has also been confirmed. Modulation of gene expression with this method establishes the capacity to regulate cellular gene expression and underscores the great potential for applications of this technology in creating stable de novo chromatin loops that markedly affect gene expression in the contexts of cancer and development.

Chromatin looping plays a significant role in gene regulation; however, there have been no technological advances that allow for selective and reversible modification of chromatin loops. Here we describe a powerful system for chromatin loop re-organization using CRISPR-dCas9 (CLOuD9), demonstrated to selectively and reversibly modulate gene expression at targeted loci.

Recent studies have clearly shown that long-range, three-dimensional chromatin looping interactions play a significant role in the regulation of gene ...

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 ...

Exploring Sequence Space to Identify Binding Sites for Regulatory RNA-Binding Proteins

Gene regulation plays an important role in all cells. Transcriptional, post-transcriptional (or RNA processing), translational, and post-translational steps are used to regulate specific genes. Sequence-specific nucleic acid-binding proteins target specific sequences to control spatial or temporal gene expression. The binding sites in nucleic acids are typically characterized by mutational analysis. However, numerous proteins of interest have no known binding site for such characterization. Here we describe an approach to identify previously unknown binding sites for RNA-binding proteins. It involves iterative selection and amplification of sequences starting with a randomized sequence pool. Following several rounds of these steps-transcription, binding, and amplification-the enriched sequences are sequenced to identify a preferred binding site(s). Success of this approach is monitored using in vitro binding assays. Subsequently, in vitro and in vivo functional assays can be used to assess the biological relevance of the selected sequences. This approach allows identification and characterization of a previously unknown binding site(s) for any RNA-binding protein for which an assay to separate protein-bound and unbound RNAs exists.

Sequence specificity is critical for gene regulation. Regulatory proteins that recognize specific sequences are important for gene regulation. Defining functional binding sites for such proteins is a challenging biological problem. An iterative approach for identification of a binding site for an RNA-binding protein is described here and is applicable to all RNA-binding proteins.

Gene regulation plays an important role in all cells. Transcriptional, post-transcriptional (or RNA processing), translational, and post-translational ...