Name: adaptation of hybridization capture of chromatin-associated proteins for proteomics to mammalian cells
Uploaded: 2018-06-01T13:30:00.000+00:00
Duration: 9 min 27 s
Description: Watch this Scientific Journal Video about Adaptation of Hybridization Capture of Chromatin-associated Proteins for Proteomics to Mammalian Cells at JoVE.com

This work was supported by NIH Grants P50HG004952 and R01GM109099 to MO.

1. Capture Oligonucleotide Design
<ol>
	<li>Design a panel of oligonucleotides to be used during the hybridization capture of the target region/s.
	<ol>
		<li>Aim to design 4&#8211;8 oligonucleotides per target region, but as a minimum, design at least one oligonucleotide targeting each end of the target region.</li>
		<li>If the target sequence is long (&#62;500 base pairs), design the oligonucleotides as spread out as possible to assure effective enrichment of chromatin across the entire target region. 
		NOTE: We have observed optimal captures with 4&#8211;8 oligonucleotides. Even though this process can work well with more oligonucleotides, it is not uncommon to observe a decrease in enrichment yields when too many oligonucleotides are used.</li>
	</ol>
	</li>
	<li>Design oligonucleotides with similar melting temperatures, and make sure that they do not form strong hairpins nor have strong interactions (taking into account that hybridizations will be carried out at 42 &#7506;C). 
	NOTE: If a toehold elution is chosen (4.12.3), an additional external 8 base sequence must be added to the oligonucleotide and will require complementary &#39;release&#39; oligonucleotides at the time of the elution17. Several commercial software tools can be used to select and analyze the oligonucleotides. 25 mers have shown good results, but depending on the genome size and sequence, capture oligonucleotides can contain somewhere between 20&#8211;80 bases. Typically, capture oligonucleotides have a melting temperature close to 62 &#7506;C, but that can change depending on their length. Consistent melting temperatures and/or oligonucleotide length throughout the capture mix can help avoid unwanted biases in enrichment.</li>
	<li>Design the capture oligonucleotides with a biotin moiety (preferably separated by a spacer from the capture sequence) on the 3&#39; end.</li>
</ol>
2. Cell Culture
<ol>
	<li>Grow human lymphoblastoid cells (such as GM12878 shown here) or other cell lines in RPMI 1640 media supplemented with 1% penicillin-streptomycin, 2 mM L-glutamine and 5% fetal bovine serum (FBS), at 37 &#7506;C and 5% CO2.
	<ol>
		<li>Seed initial frozen aliquots (2&#8211;5 x 105 cells) in a T-25 flask with 6 mL of RPMI 1640 media prepared in 2.1.</li>
		<li>Monitor cell density and viability.
		<ol>
			<li>Take a representative aliquot of cells and stain them with an appropriate dye such as trypan blue.</li>
			<li>Measure the cell density using a hemocytometer or an automated cell counter.</li>
		</ol>
		</li>
		<li>Before the cell reaches density of 1 x 106 cells/mL, spin the cells down at 100 x g for 5 min and transfer the cells to a T-75 flask with 25 mL of RPMI 1640 media prepared in 2.1. Grow the cells until cell density approaches 1 x 106 cells/mL.</li>
		<li>Transfer the cells to a T-150 flask in 38 mL of RPMI 1640 media (2.1) and grow the cells for an additional two days. 
		NOTE: Lymphoblastoid cells grow in suspension. General guidelines for growing lymphoblastoid cells recommend cell densities to be kept between 0.2&#8211;1 x 106 cells/mL. Because of the large amount of material required in this technology, the present protocol is purposely written to grow cells to a cell density beyond what is commonly used. Cell growth protocols can be revised and adapted by the reader depending on the cell types to be used.</li>
		<li>Spin the cells down at 100 x g for 5 min, resuspend the cells in 10 mL of RPMI 1640 media (2.1) and pour the suspension into an 850 cm2 roller bottle containing 500 mL of media. Incubate the cells under the same conditions as before but at this point start using constant rotation (~0.5 rpm).</li>
		<li>Monitor the cell growth and change the media when needed (Typically every 3&#8211;4 days). Let the cells grow to a density as close as possible to 2 x 106 cells/mL (1 x 109 cells in total).</li>
		<li>Once the desired cell count is reached, harvest the cells by spinning the cells down at 100 x g for 5 min and resuspend the cells in 36 mL of RPMI 1640 media (2.1). Transfer the cell suspension to a 50 mL conical tube. Take small aliquots for cell counting and DNA extraction.</li>
	</ol>
	</li>
	<li>Crosslink by adding 1 mL of 37 % formaldehyde to reach a final concentration of 1%. 
	CAUTION: Formaldehyde is a carcinogen and should be handled in a fume hood.
	<ol>
		<li>Incubate the cells with rotation (~30&#8211;60 rpm) for 10 min at room temperature (RT).</li>
		<li>Quench the crosslink reaction by adding 2 mL of a 2.5 M glycine solution, for a final concentration of 125 mM.</li>
		<li>Incubate the cells with rotation (~30&#8211;60 rpm) for 5 min at RT. Pellet the cells by centrifuging at 100 x g for 5 min and wash the cells twice with ice cold 1x PBS.</li>
	</ol>
	</li>
	<li>Continue to next step or resuspend the pellet in 5 mL of Cell Lysis Buffer (5 mM Hepes, 85 mM KCl, 0.5% Igepal, protease inhibitors (PI)) and snap-freeze the suspension drop by drop in liquid nitrogen. Store the samples at -80 &#7506;C. 
	NOTE: Prepare &#62;25 mL of Cell Lysis Buffer without Igepal and PI and add those reagents fresh when ready to use. Avoid long-term storage of Cell Lysis Buffer once Igepal and PI have been added. Store Cell Lysis Buffer at 4 &#7506;C for up to 3 months. Igepal is a nonionic, non-denaturing detergent. NP-40 can be potentially used as an alternative to Igepal but we have not used it in this context as NP-40 is not recommended when samples are eventually analyzed by mass spectrometry to characterize the DNA binding proteins.</li>
</ol>
3. Lysis and Shearing
<ol>
	<li>Prepare &#62;15 mL of Nuclei Lysis Buffer (10 mM EDTA, 50 mM Tris pH 8.0, 1 % SDS, PI). 
	NOTE: Add PI fresh, just to the amounts to be used. Avoid long-term storage of Nuclei Lysis Buffer once PI has been added. Nuclei Lysis buffer can be stored at RT for up to 1 month.</li>
	<li>Resuspend the pellet in 20 mL of Cell Lysis Buffer (containing Igepal and PI). Let the pellets thaw on ice and mix well once thawed. Let samples sit for an additional 10 min on ice. Centrifuge the cells at 400 x g at 4 &#7506;C for 5 min.</li>
	<li>Discard the supernatant and resuspend the cells in 6 mL of Nuclei Lysis Buffer with PI (volume can be increased if cells do not form a suspension).</li>
	<li>In preparation for sonication, divide the sample evenly in ~6&#8211;8 microfuge tubes (600 to 1,000 &#181;L for each tube).
	<ol>
		<li>Place the samples on ice or a cold rack and sonicate the sample using a sonicator with a microtip. Use 65% amplitude with 5 x 20 s constant bursts, allowing the suspension to cool down for at least 40 s between pulses. 
		NOTE: If volumes increase &#62;10 mL per sample, a glass cooling cell can be used instead. More and longer bursts would be required then. Sonicators differ greatly. Conditions might have to be optimized by the reader. Other sonication alternatives can be used as well.</li>
	</ol>
	</li>
	<li>Centrifuge the cells at 12,000 x g for 10 min at 4 &#7506;C, transfer the supernatant to a new tube, and discard the pellets.</li>
	<li>Estimate total recovery using a fluorometric method.</li>
	<li>Continue to next step or snap-freeze and store the sample at -80 &#7506;C.</li>
	<li>Optional: Take an aliquot, reverse crosslinks by incubating it overnight at 67 &#7506;C in 0.3 M NaCl. Clean DNA and run it in Bioanalyzer to determine fragment lengths. The bulk of the fragments should be between 500 and 1,000 bp. 
	NOTE: It is OK to stop at this point and store the samples at -80 &#7506;C.</li>
</ol>
4. Hybridization Capture
<ol>
	<li>Prepare 500 mL of 2x Hybridization (Hyb) Buffer (200 mM MES, 2M NaCl, 40 mM EDTA, 0.02 % Tween-20) and 500 mL of Wash Buffer (200 mM NaCl, 0.2 % SDS, 50 mM Tris pH 8). Store the Hyb and Wash Buffers at 4 &#7506;C and room temperature, respectively, for up to 3 months. Use half of the 2x Hyb buffer to prepare a 1x Hybridization Buffer. Also store it at 4 &#7506;C for up to 3 months. 
	NOTE: The total volume of the hybridization capture will be twice the volume of the resulting sample in the previous step (as described so far, the sample volume would be around 6 mL and thus the total hybridization volume would be 12 mL).</li>
	<li>Set aside beads equal to 5 % of the total hybridization capture volume (600 &#181;L in this case). An appropriate and suitable magnet is needed to work with the beads in the following steps.</li>
	<li>Wash the beads three times with 1x Hyb Buffer using twice the beads volume (1200 &#181;L). Resuspend the beads in 1,200 &#181;L of 2x Hyb Buffer. Place the tubes on the magnet until the solution clears and remove the buffer.</li>
	<li>Perform pre-clearing reactions in 1.5 mL microfuge tubes with a maximum of 700 &#181;L of sample per tube. Add enough beads (5%) and 2x Hyb Buffer to dilute the Hyb buffer to 1x (10 tubes with 600 &#181;L of sample, 120 &#181;L of washed beads resuspended in 2x Hyb Buffer, and 480 &#181;L of 2x Hyb Buffer). Incubate them for 10 min at 31 &#7506;C with end over end rotation (~30&#8211;60 rpm).</li>
	<li>Place the tubes on a magnet until beads are immobilized and transfer the samples to new tubes. Discard the beads. 
	NOTE: From this point forward, low-binding tubes are recommended.</li>
	<li>Resuspend the capture oligonucleotides to a working solution of 10 pmol/&#181;L and add 4 &#181;L of this solution to each tube. 
	NOTE: Different regions can act as control for each other, but it is recommended to include a capture oligonucleotide with its sequence scrambled to serve as a true negative control.</li>
	<li>Incubate the tubes for 40 min at 42 &#7506;C with end over end rotation (~30&#8211;60 rpm).</li>
	<li>During the incubation, set aside the beads needed (0.3 &#181;L/pmol of capture oligonucleotide). Wash the beads three times as described in step 4.3, but resuspend the cells in 1 x Hyb Buffer.</li>
	<li>Add the washed beads to the sample and incubate the samples for 30 min at RT with end over end rotation (~30&#8211;60 rpm). Place the tubes on the magnet and remove the supernatant once the beads have been immobilized.</li>
	<li>Washing the beads
	<ol>
		<li>Add 1.2 mL of Wash Buffer and incubate the sample for 5 min with end over end rotation (~30&#8211;60 rpm) at RT. Place tubes on the magnet and wait a couple of min until solution clears. Remove the buffer and repeat this step.</li>
		<li>Add 1.2 mL of Wash Buffer and incubate it for 1 h with end over end rotation (~30-60 rpm) at RT. Place the tubes on the magnet and wait a couple of min until solution clears. Remove the buffer and repeat this step reducing the incubation time to 30 min the second time.</li>
		<li>Add 200 &#181;L of Wash Buffer and incubate it for 15 min with end over end rotation (~30&#8211;60 rpm) at 31 &#7506;C. Place the tubes on the magnet and wait a couple of min until solution clears. Remove the buffer.</li>
		<li>Add 200 &#181;L of PBS and incubate it for 5 min with end over end rotation (~30&#8211;60 rpm) at RT. Place the tubes on the magnet and wait a couple of min until solution clears. Remove the buffer and repeat this step.</li>
	</ol>
	</li>
	<li>Simple elution
	<ol>
		<li>Add 40 &#181;L of molecular grade water to the beads and incubate it at 94 &#7506;C for 5 min.</li>
		<li>Place the tubes on the magnet and wait a couple of minutes until the solution clears.</li>
		<li>Transfer the supernatant into a new tube. Discard the beads.</li>
		<li>Proceed to the proteomic analysis or store the sample at -80 &#7506;C. 
		NOTE: This type of elution works well for later qPCR analysis and some proteomics measurements, but in other instances two alternative approaches described below yield &#34;cleaner&#34; eluates.</li>
	</ol>
	</li>
	<li>DNase elution.
	<ol>
		<li>Add 40 &#181;L of 1x DNase Buffer and 2 U of DNase to each tube.</li>
		<li>Incubate the tubes for 15 min at 37 &#7506;C.</li>
		<li>Place the tubes on a magnet and wait a couple of min until the solution clears.</li>
		<li>Remove the eluted samples from the beads and place the samples in a new low-binding tube.</li>
		<li>Incubate the tube at 94 &#7506;C for 7 min to inactivate the enzyme and proceed to proteomic analyses. 
		NOTE: Lower DNase concentrations can be used if interference with proteomic analyses is observed. It will not be possible to calculate the capture enrichments by qPCR if this elution method is used since the DNase treatment completely destroys the DNA material in the sample.</li>
		<li>Proceed to proteomic analysis or store the sample at -80 &#7506;C.</li>
	</ol>
	</li>
	<li>Toehold elution17. 
	NOTE: This method is not recommended for longer oligonucleotides (&#62;40 bases). This method requires for the capture oligonucleotides to contain an additional 8 bases that do not overlap with the genomic sequence and complementary release oligonucleotides in order to displace the target from the capture oligonucleotides.
	<ol>
		<li>Dilute 2 nanomoles of release oligonucleotides in 40 &#181;L of SSC buffer.</li>
		<li>Incubate it for 20 min at RT.</li>
		<li>Place the tubes on a magnet and wait a couple of minutes until the solution clears.</li>
		<li>Remove the solution from the beads.</li>
		<li>Proceed to proteomic analysis or store the sample at -80 &#7506;C.</li>
	</ol>
	</li>
</ol>
5. Evaluation of Capture Yield by qPCR with Primers for the Target Capture Region of Interest
<ol>
	<li>Use an appropriate software or online tool to design primers and probes for the target region. 
	NOTE: In this experiment, the promoter region of the DUSP3 and DUSP5 were targeted. The primers and probes used are the following. DUSP3: Primer 1 - GTG TTT AGG TTC CCT GAT CC, primer 2 - CGG AAT GTC TGC CTT CG, FAM-labeled probe - TCC ATG CCC GAA TTG TGA CGT CGG; DUSP5: Primer 1 - TGA GAA AGC CCT GAT GTG TC, primer 2 - GCC TTC AGA AAC CCC AAA AG, FAM-labeled probe TGC ATT TAG AGC AGC CAG ATG AGG TT.</li>
	<li>Take an aliquot of the eluate (no more than 10 &#181;L), and dilute 1:10 with molecular grade H2O.</li>
	<li>Extract the DNA of one of the aliquots taken in step 2.1.7 and use that DNA sample to make a standard curve by doing five serial 1:10 dilutions in molecular grade H2O.</li>
	<li>Prepare a qPCR master mix (wide variety of commercial alternatives), add primers and probes and dispense appropriate amount (15 &#181;L if running a 20 &#181;L reaction) in each well.</li>
	<li>Add 5 &#181;L of either: a) diluted sample, b) the standard curve or c) molecular grade H2O (to serve as no template control) to each well.</li>
	<li>Seal the plate and centrifuge it at 100 x g for 1 min.</li>
	<li>Run in a qPCR system with the following parameters: 5 min at 95 &#7506;C followed by 40 cycles of 95 &#7506;C for 20 s, 59 &#7506;C for 20 s and 72 &#7506;C for 25 s.</li>
	<li>Use the standard curve to asses yields and scrambled capture to assess capture specificity.</li>
</ol>

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There are no conflicts of interest to disclose.

The HyCCAPP method described here has many unique features that make it a powerful approach to uncover DNA-interactions that otherwise would remain elusive. The nature of the process gives HyCCAPP the flexibility to work in different organisms and regions of the genome. It is a method, however, that has several limitations to be considered.
HyCCAPP is a method that avoids any cell modifications so that it can potentially be applied in primary cells, cell culture systems, or even tissue samples...

During the past decade, there has seen a dramatic improvement in sequencing technologies, allowing the study of a wide range of genomes in large numbers of samples, and with astonishing resolution. The Encyclopedia of DNA Elements (ENCODE) Consortium, a large-scale multi-institutional effort spearheaded by the National Human Genome Research Institute of the National Institutes of Health, has provided insights into how individual transcription factors and other regulatory proteins bind to and interact with the genome. The initial effort characterized specific DNA-protein interactions, as assessed by Chromatin immunoprecipitation (ChIP) for over 100 known DNA-binding proteins1. Alternative methods such as DNase footprinting2 and formaldehyde assisted isolation of regulatory elements (FAIRE)3 have also been used to locate specific regions of the genome interacting with proteins, but with the obvious limitation that these experimental approaches do not identify the interacting proteins. Despite the extensive efforts over the past years, no technology has emerged that efficiently allows the comprehensive characterization of protein-DNA interactions in chromatin, and the identification and quantification of chromatin-associated proteins.
To address this need, we developed a novel approach which we termed as Hybridization Capture of Chromatin-Associated Proteins for Proteomics (HyCCAPP). Initially developed in yeast4,5,6, the approach isolates crosslinked chromatin regions of interest (with bound proteins) using sequence-specific hybridization capture. After isolation of the protein-DNA complexes, approaches such as mass spectrometry can be used to characterize the set of proteins bound to the sequence of interest. Thus, HyCCAPP can be considered as a non-biased approach to uncover novel DNA-protein interactions, in the sense that it does not rely on antibodies and it is completely agnostic about the proteins that might be found. There are other approaches capable to uncover novel DNA-interacting proteins7, but most rely on ChIP-like methods8,9,10, plasmid insertions11,12,13,14, or regions with high copy numbers15. In contrast, HyCCAPP can be applied to multi- and single-copy regions, and it does not require any prior information about the proteins in the region. In addition, while some of the methods mentioned above have valuable features, notably avoiding the need for DNA-protein crosslinking reactions, the unique feature of HyCCAPP is that it can be applied to single-copy regions in unmodified cells, and without any prior knowledge about putative binding proteins, or available antibodies.
At this point, HyCCAPP has predominantly been applied to the analysis of various genomic regions in yeast4,5,6, and was recently used to analyze protein-DNA interactions in alpha-satellite DNA, a repeat region in the human genome16. As part of our ongoing work, we have adapted the hybridization capture approach initially developed for yeast chromatin to be applicable to the analysis of human cells, and present here a modified protocol that allows the selective capture of single-copy target regions in the human genome with efficiencies similar to our initial studies in yeast. This new optimized protocol now allows the adaptation and utilization of the technology to interrogate protein-DNA interactions across the human genome, using mass spectrometry or other analytical approaches.
It is important to emphasize that the HyCCAPP method is meant for the analysis of specific target regions and is not yet suitable for genome-wide analyses. The technology is especially useful when dealing with regions for which there is scarce information about interacting proteins, or when a more comprehensive in-depth analysis of interacting proteins at a specific genome locus is desired. HyCCAPP is meant to uncover DNA-binding proteins but not characterize accurately the specific protein binding sites in genomic DNA. In its current implementation, the methodology does not provide information about the DNA binding sequences or motifs for individual proteins. Therefore, it nicely complements existing technologies such as FAIRE, and may allow the identification of novel binding proteins in genomic regions identified by an initial FAIRE analysis.

<table><tbody><tr><td>PrimerQuest tool</td><td>IDT</td><td></td><td></td></tr><tr><td>OligoAnalyzer 3.1</td><td>IDT</td><td></td><td>Analyze capture oligonucleotids</td></tr><tr><td>PairFold</td><td>UBC</td><td></td><td>Analyze interactions</td></tr><tr><td>RPMI 1640 media</td><td>Thermo Fisher</td><td>11875093</td><td></td></tr><tr><td>Penicillin-streptomycin</td><td>Thermo Fisher</td><td>15140122</td><td></td></tr><tr><td>L-Glutamine&amp;nbsp;</td><td>Thermo Fisher</td><td>25030081</td><td></td></tr><tr><td>Fetal bovine serum</td><td>Thermo Fisher</td><td>26140079</td><td></td></tr><tr><td>Countess automated cell counter</td><td>Thermo Fisher</td><td></td><td></td></tr><tr><td>850 cm&lt;sup&gt;2&lt;/sup&gt; roller bottle&amp;nbsp;</td><td>Greiner Bio-one</td><td>680058</td><td></td></tr><tr><td>Roller system</td><td>Wheaton</td><td>22-288-525</td><td></td></tr><tr><td>37 % formaldehyde&amp;nbsp;</td><td>Sigma-Aldrich</td><td>F8775</td><td></td></tr><tr><td>Glycine</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>Igepal CA-630</td><td>Sigma-Aldrich</td><td>I3021</td><td></td></tr><tr><td>Protease inhibitor cocktail</td><td>Sigma-Aldrich</td><td>P4380</td><td></td></tr><tr><td>HEPES</td><td>Thermo Fisher</td><td>15630080</td><td></td></tr><tr><td>Branson digital sonifier SFX 150</td><td>Emerson</td><td></td><td></td></tr><tr><td>Qubit 3.0 fluorometer</td><td>Thermo Fisher</td><td></td><td></td></tr><tr><td>Qubit dsDNA BR assay kit</td><td>Thermo Fisher</td><td>Q32850</td><td></td></tr><tr><td>Bioanalyzer 2100</td><td>Agilent</td><td></td><td></td></tr><tr><td>Agilent DNA 1000 kit</td><td>Agilent</td><td>5067-1504</td><td></td></tr><tr><td>MES sodium salt</td><td>Sigma-Aldrich</td><td>M3885</td><td></td></tr><tr><td>NaCl 5M</td><td>Thermo Fisher</td><td>AM9760G</td><td></td></tr><tr><td>EDTA</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>DynaMag-2 magnet</td><td>Thermo Fisher</td><td>12321D</td><td></td></tr><tr><td>DynaMag-15 magnet</td><td>Thermo Fisher</td><td>12301D</td><td></td></tr><tr><td>Dynabeads M-280 atreptavidin</td><td>Thermo Fisher</td><td>60210</td><td></td></tr><tr><td>Low binding tubes</td><td>Eppendorf</td><td>22431081</td><td></td></tr><tr><td>Hybridization oven</td><td>SciGene</td><td></td><td></td></tr><tr><td>Tube shaker and rotator</td><td>Thermo Fisher</td><td>415110Q</td><td></td></tr><tr><td>DNase I (RNase-free)</td><td>New England BioLabs</td><td>M0303</td><td></td></tr><tr><td>SSC buffer 20&amp;times; concentrate</td><td>Sigma-Aldrich</td><td>S6639</td><td></td></tr></tbody></table>

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.

Due to the need for large input amounts of chromatin for HyCCAPP to succeed, cells are grown to relatively high levels of confluency. Trypan blue staining is used to confirm that cell death rates are moderate (&#60;10 %). In single copy experiments, chromatin content prior to hybridization capture needs to be in the femtomolar range, which usually requires at least 109 cells as starting material. Prior to full scale experiments, it is recommended to test capture oligonucleotide...

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

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 ...

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Research

JoVE Journal

Genetics

6.2K Views.  Texas Biomedical Research Institute. 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.

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

Protein Complex Affinity Capture from Cryomilled Mammalian Cells

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this technique is also frequently referred to as immunoprecipitation. Affinity capture can be applied in a bench-scale and in a high-throughput context. When coupled with protein mass spectrometry, affinity capture has proven to be a workhorse of interactome analysis. Although there are potentially many ways to execute the numerous steps involved, the following protocols implement our favored methods. Two features are distinctive: the use of cryomilled cell powder to produce cell extracts, and antibody-coupled paramagnetic beads as the affinity medium. In many cases, we have obtained superior results to those obtained with more conventional affinity capture practices. Cryomilling avoids numerous problems associated with other forms of cell breakage. It provides efficient breakage of the material, while avoiding denaturation issues associated with heating or foaming. It retains the native protein concentration up to the point of extraction, mitigating macromolecular dissociation. It reduces the time extracted proteins spend in solution, limiting deleterious enzymatic activities, and it may reduce the non-specific adsorption of proteins by the affinity medium. Micron-scale magnetic affinity media have become more commonplace over the last several years, increasingly replacing the traditional agarose- and Sepharose-based media. Primary benefits of magnetic media include typically lower non-specific protein adsorption; no size exclusion limit because protein complex binding occurs on the bead surface rather than within pores; and ease of manipulation and handling using magnets.

Here we describe protocols to disrupt mammalian cells by solid-state milling at a cryogenic temperature, produce a cell extract from the resulting cell powder, and isolate protein complexes of interest by affinity capture upon antibody-coupled micron-scale paramagnetic beads.

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this ...

Biochemistry

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 ...

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

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

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

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

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

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 ...

Quantitative Analysis of Chromatin Proteomes in Disease

In the nucleus reside the proteomes whose functions are most intimately linked with gene regulation. Adult mammalian cardiomyocyte nuclei are unique due to the high percentage of binucleated cells,1 the predominantly heterochromatic state of the DNA, and the non-dividing nature of the cardiomyocyte which renders adult nuclei in a permanent state of interphase.2 Transcriptional regulation during development and disease have been well studied in this organ,3-5 but what remains relatively unexplored is the role played by the nuclear proteins responsible for DNA packaging and expression, and how these proteins control changes in transcriptional programs that occur during disease.6 In the developed world, heart disease is the number one cause of mortality for both men and women.7 Insight on how nuclear proteins cooperate to regulate the progression of this disease is critical for advancing the current treatment options.
Mass spectrometry is the ideal tool for addressing these questions as it allows for an unbiased annotation of the nuclear proteome and relative quantification for how the abundance of these proteins changes with disease. While there have been several proteomic studies for mammalian nuclear protein complexes,8-13 until recently14 there has been only one study examining the cardiac nuclear proteome, and it considered the entire nucleus, rather than exploring the proteome at the level of nuclear sub compartments.15 In large part, this shortage of work is due to the difficulty of isolating cardiac nuclei. Cardiac nuclei occur within a rigid and dense actin-myosin apparatus to which they are connected via multiple extensions from the endoplasmic reticulum, to the extent that myocyte contraction alters their overall shape.16 Additionally, cardiomyocytes are 40% mitochondria by volume17 which necessitates enrichment of the nucleus apart from the other organelles. Here we describe a protocol for cardiac nuclear enrichment and further fractionation into biologically-relevant compartments. Furthermore, we detail methods for label-free quantitative mass spectrometric dissection of these fractions-techniques amenable to in vivo experimentation in various animal models and organ systems where metabolic labeling is not feasible.

Advances in mass spectrometry have allowed the high throughput analysis of protein expression and modification in a host of tissues. Combined with subcellular fractionation and disease models, quantitative mass spectrometry and bioinformatics can reveal new properties in biological systems. The method described herein analyzes chromatin-associated proteins in the setting of heart disease and is readily applicable to other in vivo models of human disease.

In the nucleus reside the proteomes whose functions are most intimately linked with gene regulation. Adult mammalian cardiomyocyte nuclei are unique due ...

Medicine

Histone Proteomic Analysis and Post-Translational Modification Characterization

Histone Modification Screening using Liquid Chromatography, Trapped Ion Mobility Spectrometry, and Time-Of-Flight Mass Spectrometry

Histone proteins are highly abundant and conserved among eukaryotes and play a large role in gene regulation as a result of structures known as posttranslational modifications (PTMs). Identifying the position and nature of each PTM or pattern of PTMs in reference to external or genetic factors allows this information to be statistically correlated with biological responses such as DNA transcription, replication, or repair. In the present work, a high-throughput analytical protocol for the detection of histone PTMs from biological samples is described. The use of complementary liquid chromatography, trapped ion mobility spectrometry, and time-of-flight mass spectrometry (LC-TIMS-ToF MS/MS) enables the separation and PTM assignment of the most biologically relevant modifications in a single analysis. The described approach takes advantage of recent developments in dependent data acquisition (DDA) using parallel accumulation in the mobility trap, followed by sequential fragmentation and collision-induced dissociation. Histone PTMs are confidently assigned based on their retention time, mobility, and fragmentation pattern.

Author Spotlight: Enhanced Histone PTM Isomer Identification Through LC-TIMS-ToF MS/MS and PASEF

An analytical workflow based on liquid chromatography, trapped ion mobility spectrometry, and time-of-flight mass spectrometry (LC-TIMS-ToF MS/MS) for high confidence and highly reproducible "bottom-up" analysis of histone modifications and identification based on principal parameters (retention time [RT], collision cross section [CCS], and accurate mass-to-charge [m/z] ratio).

Histone proteins are highly abundant and conserved among eukaryotes and play a large role in gene regulation as a result of structures known as ...

Chemistry

The ChroP Approach Combines ChIP and Mass Spectrometry to Dissect Locus-specific Proteomic Landscapes of Chromatin

Chromatin is a highly dynamic nucleoprotein complex made of DNA and proteins that controls various DNA-dependent processes. Chromatin structure and function at specific regions is regulated by the local enrichment of histone post-translational modifications (hPTMs) and variants, chromatin-binding proteins, including transcription factors, and DNA methylation. The proteomic characterization of chromatin composition at distinct functional regions has been so far hampered by the lack of efficient protocols to enrich such domains at the appropriate purity and amount for the subsequent in-depth analysis by Mass Spectrometry (MS). We describe here a newly designed chromatin proteomics strategy, named ChroP (Chromatin Proteomics), whereby a preparative chromatin immunoprecipitation is used to isolate distinct chromatin regions whose features, in terms of hPTMs, variants and co-associated non-histonic proteins, are analyzed by MS. We illustrate here the setting up of ChroP for the enrichment and analysis of transcriptionally silent heterochromatic regions, marked by the presence of tri-methylation of lysine 9 on histone H3. The results achieved demonstrate the potential of ChroP in thoroughly characterizing the heterochromatin proteome and prove it as a powerful analytical strategy for understanding how the distinct protein determinants of chromatin interact and synergize to establish locus-specific structural and functional configurations.

By combining native and crosslinking chromatin immunoprecipitation with high-resolution Mass Spectrometry, ChroP approach enables to dissect the composite proteomic architecture of histone modifications, variants and non-histonic proteins synergizing at functionally distinct chromatin domains.

Chromatin is a highly dynamic nucleoprotein complex made of DNA and proteins that controls various DNA-dependent processes. Chromatin structure and ...

Biology

Discovering Protein Interactions and Characterizing Protein Function Using HaloTag Technology

Research in proteomics has exploded in recent years with advances in mass spectrometry capabilities that have led to the characterization of numerous proteomes, including those from viruses, bacteria, and yeast.&#160; In comparison, analysis of the human proteome lags behind, partially due to the sheer number of proteins which must be studied, but also the complexity of networks and interactions these present. To specifically address the challenges of understanding the human proteome, we have developed HaloTag technology for protein isolation, particularly strong for isolation of multiprotein complexes and allowing more efficient capture of weak or transient interactions and/or proteins in low abundance. &#160;HaloTag is a genetically encoded protein fusion tag, designed for covalent, specific, and rapid immobilization or labelling of proteins with various ligands. Leveraging these properties, numerous applications for mammalian cells were developed to characterize protein function and here we present methodologies including: protein pull-downs used for discovery of novel interactions or functional assays, and cellular localization. We find significant advantages in the speed, specificity, and covalent capture of fusion proteins to surfaces for proteomic analysis as compared to other traditional non-covalent approaches. We demonstrate these and the broad utility of the technology using two important epigenetic proteins as examples, the human bromodomain protein BRD4, and histone deacetylase HDAC1.&#160; These examples demonstrate the power of this technology in enabling &#160;the discovery of novel interactions and characterizing cellular localization in eukaryotes, which will together further understanding of human functional proteomics. &#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;

HaloTag technology is a multifunctional technology which has shown significant success in isolation of both small and large protein complexes from mammalian cells.&#160; Here we highlight the advantages of this technology compared to existing alternatives and demonstrate its utility to study numerous aspects of protein function inside eukaryotic cells.

Research in proteomics has exploded in recent years with advances in mass spectrometry capabilities that have led to the characterization of numerous ...

Identification of Protein Interaction Partners in Mammalian Cells Using SILAC-immunoprecipitation Quantitative Proteomics

Quantitative proteomics combined with immuno-affinity purification, SILAC immunoprecipitation, represent a powerful means for the discovery of novel protein:protein interactions. By allowing the accurate relative quantification of protein abundance in both control and test samples, true interactions may be easily distinguished from experimental contaminants. Low affinity interactions can be preserved through the use of less-stringent buffer conditions and remain readily identifiable. This protocol discusses the labeling of tissue culture cells with stable isotope labeled amino acids, transfection and immunoprecipitation of an affinity tagged protein of interest, followed by the preparation for submission to a mass spectrometry facility. This protocol then discusses how to analyze and interpret the data returned from the mass spectrometer in order to identify cellular partners interacting with a protein of interest. As an example this technique is applied to identify proteins binding to the eukaryotic translation initiation factors: eIF4AI and eIF4AII.

SILAC immunoprecipitation experiments represent a powerful means for discovering novel protein:protein interactions. By allowing the accurate relative quantification of protein abundance in both control and test samples, true interactions may be easily distinguished from experimental contaminants, and low affinity interactions preserved through use of less-stringent buffer conditions.

Quantitative proteomics combined with immuno-affinity purification, SILAC immunoprecipitation, represent a powerful means for the discovery of novel ...

Chromatin Extraction from Frozen Chimeric Liver Tissue for Chromatin Immunoprecipitation Analysis

Crosslinking Chromatin Immunoprecipitation (X-ChIP) is a widely used technique to assess levels of histone marks and occupancy of transcription factors on host and/or pathogen chromatin. Chromatin preparation from tissues creates additional challenges that need to be overcome to obtain reproducible and reliable protocols comparable to those used for cell culture. Tissue disruption and fixation are critical steps to achieve efficient shearing of chromatin. Coexistence of different cell types and clusters may also require different shearing times to reach optimal fragment size and hinders shearing reproducibility. The purpose of this method is to achieve reliable and reproducible host chromatin preparations from frozen tissue (liver) suitable for both ChIP-qPCR and next generation sequencing (NGS) applications. We observed that the combination of liquid nitrogen tissue pulverization followed by homogenization leads to increased reproducibility compared to homogenization only, since it provides a suspension consisting mostly of dissociated single cells that can be efficiently sheared. Moreover, the fixation step should be performed under mild rotation to provide homogeneous crosslinking. The fixed material is then suitable for buffer-based nuclei isolation, to reduce contamination of cytoplasmic protein and pathogen DNAs and RNAs (when applicable), avoiding time-consuming centrifugation gradients. Subsequent sonication will complete nuclear lysis and shear the chromatin, producing a specific size range according to the chosen shearing conditions. The size range should fall between 100 and 300 nt for NGS applications, while it could be higher (300-700 nt) for ChIP-qPCR analysis. Such protocol adaptations can greatly improve chromatin analyses from frozen tissue specimens.

This protocol focuses on chromatin preparation from snap frozen tissues and it is suitable for Crosslinking Chromatin Immunoprecipitation (X-ChIP) followed by either quantitative PCR analysis (X-ChIP-qPCR) or next generation sequencing approaches (X-ChIP-seq).

Crosslinking Chromatin Immunoprecipitation (X-ChIP) is a widely used technique to assess levels of histone marks and occupancy of transcription factors on ...

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

Summary

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Chapters in this video

Protein Complex Affinity Capture from Cryomilled Mammalian Cells

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

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

Chromatin Immunoprecipitation Assay Using Micrococcal Nucleases in Mammalian Cells

Quantitative Analysis of Chromatin Proteomes in Disease

Histone Modification Screening using Liquid Chromatography, Trapped Ion Mobility Spectrometry, and Time-Of-Flight Mass Spectrometry

The ChroP Approach Combines ChIP and Mass Spectrometry to Dissect Locus-specific Proteomic Landscapes of Chromatin

Discovering Protein Interactions and Characterizing Protein Function Using HaloTag Technology

Identification of Protein Interaction Partners in Mammalian Cells Using SILAC-immunoprecipitation Quantitative Proteomics

Chromatin Extraction from Frozen Chimeric Liver Tissue for Chromatin Immunoprecipitation Analysis