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

Summary

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.

Abstract

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.

Introduction

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 yeast^4,5,6, 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 methods^8,9,10, plasmid insertions^11,12,13,14, 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 yeast^4,5,6, 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.

Protocol

1. Capture Oligonucleotide Design

1. Design a panel of oligonucleotides to be used during the hybridization capture of the target region/s.
   1. Aim to design 4–8 oligonucleotides per target region, but as a minimum, design at least one oligonucleotide targeting each end of the target region.
   2. If the target sequence is long (>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 4–8 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.
2. 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 ᵒC).
   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 'release' 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 20–80 bases. Typically, capture oligonucleotides have a melting temperature close to 62 ᵒC, 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.
3. Design the capture oligonucleotides with a biotin moiety (preferably separated by a spacer from the capture sequence) on the 3' end.

2. Cell Culture

1. 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 ᵒC and 5% CO₂.
   1. Seed initial frozen aliquots (2–5 x 10⁵ cells) in a T-25 flask with 6 mL of RPMI 1640 media prepared in 2.1.
   2. Monitor cell density and viability.
      1. Take a representative aliquot of cells and stain them with an appropriate dye such as trypan blue.
      2. Measure the cell density using a hemocytometer or an automated cell counter.
   3. 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.
   4. 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 0.2–1 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.
   5. 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).
   6. Monitor the cell growth and change the media when needed (Typically every 3–4 days). Let the cells grow to a density as close as possible to 2 x 10⁶ cells/mL (1 x 10⁹ cells in total).
   7. 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.
2. 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.
   1. Incubate the cells with rotation (~30–60 rpm) for 10 min at room temperature (RT).
   2. Quench the crosslink reaction by adding 2 mL of a 2.5 M glycine solution, for a final concentration of 125 mM.
   3. Incubate the cells with rotation (~30–60 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.
3. 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 ᵒC.
   NOTE: Prepare >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 ᵒC 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.

3. Lysis and Shearing

1. Prepare >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.
2. 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 ᵒC for 5 min.
3. 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).
4. In preparation for sonication, divide the sample evenly in ~6–8 microfuge tubes (600 to 1,000 µL for each tube).
   1. 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 >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.
5. Centrifuge the cells at 12,000 x g for 10 min at 4 ᵒC, transfer the supernatant to a new tube, and discard the pellets.
6. Estimate total recovery using a fluorometric method.
7. Continue to next step or snap-freeze and store the sample at -80 ᵒC.
8. Optional: Take an aliquot, reverse crosslinks by incubating it overnight at 67 ᵒC 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 ᵒC.

4. Hybridization Capture

1. 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 ᵒC 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 ᵒC 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).
2. Set aside beads equal to 5 % of the total hybridization capture volume (600 µL in this case). An appropriate and suitable magnet is needed to work with the beads in the following steps.
3. Wash the beads three times with 1x Hyb Buffer using twice the beads volume (1200 µL). Resuspend the beads in 1,200 µL of 2x Hyb Buffer. Place the tubes on the magnet until the solution clears and remove the buffer.
4. Perform pre-clearing reactions in 1.5 mL microfuge tubes with a maximum of 700 µL of sample per tube. Add enough beads (5%) and 2x Hyb Buffer to dilute the Hyb buffer to 1x (10 tubes with 600 µL of sample, 120 µL of washed beads resuspended in 2x Hyb Buffer, and 480 µL of 2x Hyb Buffer). Incubate them for 10 min at 31 ᵒC with end over end rotation (~30–60 rpm).
5. 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.
6. Resuspend the capture oligonucleotides to a working solution of 10 pmol/µL and add 4 µL 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.
7. Incubate the tubes for 40 min at 42 ᵒC with end over end rotation (~30–60 rpm).
8. During the incubation, set aside the beads needed (0.3 µL/pmol of capture oligonucleotide). Wash the beads three times as described in step 4.3, but resuspend the cells in 1 x Hyb Buffer.
9. Add the washed beads to the sample and incubate the samples for 30 min at RT with end over end rotation (~30–60 rpm). Place the tubes on the magnet and remove the supernatant once the beads have been immobilized.
10. Washing the beads
    1. Add 1.2 mL of Wash Buffer and incubate the sample for 5 min with end over end rotation (~30–60 rpm) at RT. Place tubes on the magnet and wait a couple of min until solution clears. Remove the buffer and repeat this step.
    2. 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.
    3. Add 200 µL of Wash Buffer and incubate it for 15 min with end over end rotation (~30–60 rpm) at 31 ᵒC. Place the tubes on the magnet and wait a couple of min until solution clears. Remove the buffer.
    4. Add 200 µL of PBS and incubate it for 5 min 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.
11. Simple elution
    1. Add 40 µL of molecular grade water to the beads and incubate it at 94 ᵒC for 5 min.
    2. Place the tubes on the magnet and wait a couple of minutes until the solution clears.
    3. Transfer the supernatant into a new tube. Discard the beads.
    4. Proceed to the proteomic analysis or store the sample at -80 ᵒC.
       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 "cleaner" eluates.
12. DNase elution.
    1. Add 40 µL of 1x DNase Buffer and 2 U of DNase to each tube.
    2. Incubate the tubes for 15 min at 37 ᵒC.
    3. Place the tubes on a magnet and wait a couple of min until the solution clears.
    4. Remove the eluted samples from the beads and place the samples in a new low-binding tube.
    5. Incubate the tube at 94 ᵒC 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.
    6. Proceed to proteomic analysis or store the sample at -80 ᵒC.
13. Toehold elution¹⁷.
    NOTE: This method is not recommended for longer oligonucleotides (>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.
    1. Dilute 2 nanomoles of release oligonucleotides in 40 µL of SSC buffer.
    2. Incubate it for 20 min at RT.
    3. Place the tubes on a magnet and wait a couple of minutes until the solution clears.
    4. Remove the solution from the beads.
    5. Proceed to proteomic analysis or store the sample at -80 ᵒC.

5. Evaluation of Capture Yield by qPCR with Primers for the Target Capture Region of Interest

1. 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.
2. Take an aliquot of the eluate (no more than 10 µL), and dilute 1:10 with molecular grade H₂O.
3. 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.
4. Prepare a qPCR master mix (wide variety of commercial alternatives), add primers and probes and dispense appropriate amount (15 µL if running a 20 µL reaction) in each well.
5. Add 5 µL of either: a) diluted sample, b) the standard curve or c) molecular grade H₂O (to serve as no template control) to each well.
6. Seal the plate and centrifuge it at 100 x g for 1 min.
7. Run in a qPCR system with the following parameters: 5 min at 95 ᵒC followed by 40 cycles of 95 ᵒC for 20 s, 59 ᵒC for 20 s and 72 ᵒC for 25 s.
8. Use the standard curve to asses yields and scrambled capture to assess capture specificity.

Results

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 (<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 ...

Discussion

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...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
PrimerQuest tool	IDT
OligoAnalyzer 3.1	IDT		Analyze capture oligonucleotids
PairFold	UBC		Analyze interactions
RPMI 1640 media	Thermo Fisher	11875093
Penicillin-streptomycin	Thermo Fisher	15140122
L-Glutamine	Thermo Fisher	25030081
Fetal bovine serum	Thermo Fisher	26140079
Countess automated cell counter	Thermo Fisher
850 cm2 roller bottle	Greiner Bio-one	680058
Roller system	Wheaton	22-288-525
37 % formaldehyde	Sigma-Aldrich	F8775
Glycine	Sigma-Aldrich
Igepal CA-630	Sigma-Aldrich	I3021
Protease inhibitor cocktail	Sigma-Aldrich	P4380
HEPES	Thermo Fisher	15630080
Branson digital sonifier SFX 150	Emerson
Qubit 3.0 fluorometer	Thermo Fisher
Qubit dsDNA BR assay kit	Thermo Fisher	Q32850
Bioanalyzer 2100	Agilent
Agilent DNA 1000 kit	Agilent	5067-1504
MES sodium salt	Sigma-Aldrich	M3885
NaCl 5M	Thermo Fisher	AM9760G
EDTA	Sigma-Aldrich
DynaMag-2 magnet	Thermo Fisher	12321D
DynaMag-15 magnet	Thermo Fisher	12301D
Dynabeads M-280 atreptavidin	Thermo Fisher	60210
Low binding tubes	Eppendorf	22431081
Hybridization oven	SciGene
Tube shaker and rotator	Thermo Fisher	415110Q
DNase I (RNase-free)	New England BioLabs	M0303
SSC buffer 20× concentrate	Sigma-Aldrich	S6639