Dear Nadiza,

The following are text from two relevant papers. I hope you will find an answer to your question after reading the text.

1-Histone Modification Research Methods

 Judith Erkmann (judith_erkmann at yahoo dot com)

University of Massachusetts Medical School, United States

DOI http://dx.doi.org/10.13070/mm.en.1.92

Date last modified : 2014-12-11; original version : 2011-09-28

Cite as MATER METHODS 2011;1:92

Abstract

A detailed review of methods and reagents in histone modification research.

 BACKGROUND

Histone proteins are subject to a variety of posttranslational modifications (PTMs), many of which (i.e., methylation, acetylation, ubiquitylation and SUMOylation) occur at lysines [1]. While only a handful of modification sites have been identified in the buried histone-fold domains, PTMs on the flexible N-terminal tail regions are quite common. Many histone modifications mediate functionally significant changes in chromatin structure and do so either by altering chromatin structure/dynamics directly or through the recruitment of histone modifiers and/or nucleosome remodeling complexes [1, 2].

Histone PTMs have been found to affect a host of chromatin-based reactions, including transcription, heterochromatic gene silencing and genome stability [3-6]. Modifications implicated in gene expression are especially significant as they have the potential to influence whole transcription programs. Defects in histone PTM metabolism have been linked to misregulated gene expression in various in vitro models and, in some cases, also correlate with human disease, as has been demonstrated for immunodeficiency disorders and a variety of human cancers [7-12]. Thus, how histone markers are regulated and influence the association of PTM-specific binding proteins will continue to be an area of significant investigation [2, 13-15].

Determining the function of histone PTMs often involves investigating the modification's abundance and interacting partners. The methods described here address these areas and include summaries and references for protocols detailing histone purification methods, the production of recombinant site-specifically modified histones, peptide-based systems to characterize PTM-binding proteins, and different methods for analyzing chromatin immunoprecipitation samples.

METHODS FOR HISTONE MODIFICATION RESEARCH

Cell lysate

For detection of histone modifications by Western blotting, it often is possible to use whole-cell lysates made by extraction with SDS Laemmli sample buffer. In the case of animal cell lines, cells collected by centrifugation can be directly resuspended in sample buffer and boiled for loading [16, 17] ; note, however, that some protocols additionally recommend sonicating the samples after the extraction step. Protein extracts of fungi can be prepared the same way with the exception of an optional alkali pre-treatment step [18]. This step can be omitted, however, if it is experimentally necessary to minimize sample handling time, so long as the omission is noted and samples for follow-up experiments qare treated the same way. Following extraction, the sample's insoluble fraction is removed by centrifugation, leaving behind the soluble whole cell extract in the supernatant.

Enriched histone fraction

For some applications, it is necessary to examine either histone-enriched fractions or purified histone proteins; examples of an enriched sample would be isolated nuclei or a crude chromatin preparation. The isolation of nuclei from cultured metazoan cells or yeast is very straightforward and requires essentially three steps: hypotonic swelling (after prior cell wall digestion in the case of yeast), cell membrane lysis by mechanical shearing (i.e., breakage with a dounce homogenizer or mild agitation on a rotator), and isolation of nuclei by centrifugation [19,20] (Figure 1A). The isolation of crude chromatin is also quite simple and includes only a detergent lysis step followed by the sedimentation of chromatin by centrifugation [21-23] (Figure 1B).

Histone purification

Several of the available histone purification protocols are excellent and easy to follow [20]. In the method described here, histones are extracted from nuclei using a dilute sulfuric acid solution and then purified by column chromatography (Figure 1C). The beauty of this protocol is that nucleic acids and many of the non-histone proteins can be readily removed by centrifugation because of their insolubility at acidic pH. The soluble histone fraction is then precipitated with trichloroacetic acid (TCA) and, if desired, purified over a reversed-phase HPLC column. The histones at this point can be used in a variety of applications, including Western blotting and mass spectrometry. Please note that another publication [20] also describes an alternate, high-salt histone extraction method.

[enlarge]

Figure 1. Protocols for isolating intact nuclei (A), preparing curde chromatin fractions (B), and purifying histones (C).

Histone PTM detection

Histone PTM are generally detected through the antibodies. The quality and specificity of anti-PTM antibodies should be carefully evaluated prior to experimental application. Concerns to be addressed include cross-reactivity with alternate histone modification sites, recognition of the unmodified (recombinant) protein and cross-reactivity with other nuclear species. The procedures for this type of evaluation are very straightforward and involve either Western blotting of nuclear extract preparations against recombinant histones or immunoblotting an array of modified and unmodified peptides spotted on nitrocellulose membrane.

anti-PTM histone antibody quality

Recent initiatives to address anti-PTM histone antibody quality issue have resulted in the characterization of more than 200 antibodies against 57 different histone modifications [24]. About 20% antibodies were deemed not effective. The supplementary data to this report catalogues the performance of the different antibodies tested in dot blots, Western blots (across different species), and chromatin immunoprecipitation (ChIP) assays. This is an excellent resource for the field that the authors encourage others to contribute to by logging new test results with the Antibody Validation Database [25]. A description of the dot blot protocol and images of the dot blots from this study can be found at [26].

Recombinant site-specifically modified histones

One method for producing site-specifically modified histones involves the ligation of protein fragment in vitro [27-31] (Figure 2A). The premise of this method is to chemically ligate a synthetic, modified peptide (corresponding to either the very N- or C-terminal end of the protein) to a recombinant fragment containing the complementary part of the protein, followed by purification of the full-length ligation product [32]. A related method, published more recently, describes the use of native chemical ligation to produce a fully synthetic modified histone by the sequential addition of synthetic peptides corresponding to consecutive parts of the protein [33].

[enlarge]

Figure 2. Production of site-specifically modified histones by expressed protein ligation (A) and direct incorporation in vivo (B).

Besides by chemical ligation, lysine-acetylated histones can also be produced by direct incorporation in vivo using E.coli that have been genetically engineered to incorporate acetyl-lysine at UAG codons [34] (Figure 2B). The premise of this system is to introduce into E. coli an M. barkeri pyrrolysyl tRNA synthetase variant that has been sequence-optimized to charge its cognate tRNACUA with acetyl-lysine. Thus, to produce a site-specifically acetylated histone, one would need only to add to the strain a construct containing a UAG codon at the desired modification site.

Characterization of histone PTM binding partners - co-immunoprecipitation (CoIP)/pulldown experiments

In cases where it is of interest to study the interaction of a protein with endogenous histones, it is necessary to first convert the chromatin to mononucleosome-sized pieces by digestion with micrococcal nuclease (MNase). The protein of interest can then be immunoprecipitated from the soluble chromatin fraction and evaluated for association with the core histones and different histone modifications by Western blotting [35, 36]. This experiment can also be done as a pulldown assay using a recombinant bait protein that has been incubated in the solubilized nucleosome fraction and then purified [37]. If it is obvious that a particular modification is enriched in complexes containing the target protein, it may then be of interest to characterize the interaction in binding assays using modified peptides (see below). In cases where no such preference is determined, yet it is still of interest to know whether an interaction is modification dependent, relative affinities for modified versus unmodified histones can be determined in binding assays using native and recombinant histones [38].

One common method of purifying native histones from tissue culture cells is to produce oligonucleosome-sized pieces of chromatin by either limited digestion with MNase or mechanical shearing, followed by chromatography of the fragments on a hydroxyapatite column and elution at high salt [39, 40] (Figure 3A). Recombinant histones are insoluble when overproduced as monomers; however, it is possible to recover the overexpressed proteins from inclusion bodies [41, 42] (Figure 3B). To start, preparations of inclusion bodies are made from bacterial cultures individually overexpressing each histone. The histones are then extracted from the inclusion bodies, purified over sequential ion-exchange resins, and eluted using a linear salt gradient. To generate histone octamers, equimolar amounts of H3, H4, H2A and H2B are first unfolded, combined, refolded by dialysis, and then purified over a sizing column.

[enlarge]

Figure 3. Purification schemes for isolating native (A) and recombinant histones (B).

Characterization of histone PTM binding partners - peptide pulldown assays

The modification preferences of a protein can be determined by comparing the relative binding affinities of the protein for differentially modified peptides. This can be accomplished in a few different ways. In one format, bead-immobilized peptides are used to pull down a test protein - using either recombinant protein or a nuclear extract – and the relative recoveries of the protein are determined by Western blotting [43-45] (Figure 4A). The principle of short peptides as bait molecules has also been applied in unbiased experiments to discover previously unknown histone PTM-binding proteins [36, 37, 46, 47]. In this assay, binding proteins from a nuclear extract are identified on the basis of enrichment on the modified peptide relative to an unmodified peptide or a peptide carrying the same modification at a different amino acid.

Characterization of histone PTM binding partners - peptide microarrays

Also described in the literature are array-based methods capable of screening the interaction of probe proteins with multiple peptides simultaneously (Figure 4B). For such experiments, the protein of interest is incubated over the surface of a peptide microarray, which is essentially a streptavidin-coated slide spotted with different biotinylated peptides, and the protein peptide complexes are visualized by detection with a fluorophore-conjugated antibody and an array scanner [48-50]. Studies using an inverse setup, i.e., protein arrays incubated with fluorescently-labeled peptide, have been described as well [51].

[enlarge]

Figure 4. Characterization of histone modifications using peptide-based pulldown assays (A) and peptide microarrays (B).

Determination of the genomic loci for specific PTM - chromatin immunoprecipitation

Genomic sites that are enriched for a particular PTM can be characterized by immunoprecipitating chromatin fragments containing the mark of interest and then quantifying the relative proportion of different loci that the PTM is associated with (Figure 5). Many labs have produced detailed ChIP protocols that can be found either online [52, 53] or in the literature [54-56]. The first step of most ChIP protocols is to treat cells with formaldehyde to "freeze" the position of chromatin-associated proteins by crosslinking. Whereas cells of metazoan origin are then lysed directly, yeast cells must first undergo cell wall breakage, which can be accomplished by either mechanical shearing or enzymatic digestion. Depending on the nucleotide resolution desired at detection, chromatin is cut into short pieces in one of two ways: shearing by sonication to yield fragments of between 200 and 500 bp or digestion with MNase down to individual nucleosomes. Immunoprecipitation of the extracts is performed the same way as in a conventional IP experiment except that after the elution step, the eluate is incubated overnight at 65°C to reverse cross-links. On the following day, the samples are treated with Proteinase K and then phenol/chloroform extracted to recover the co-precipitating DNA fragments.

[enlarge]

Figure 5. Chromatin immunoprecipitation (A) results in the isolation of co-purifying DNA fragments, whose sequence and enrichment can be analyzed by PCR (B), ChIP-chip (C), and/or Chip-seq (D).

Determination of the genomic loci for specific PTM - CHiP detection - PCR

If it is of interest to analyze PTM levels at only a handful of loci, this can be accomplished by either real-time PCR or quantification of PCR products on an ethidium stained gel (Figure 5B). Briefly, the loci to be analyzed are amplified from dilutions of the IP and input fractions and compared, ideally, against these fractions from a parallel IP of the histone the PTM is attached to. To determine whether the PTM is enriched at a particular location, the normalized PTM/histone ratios should be compared with those at a region nearby that the PTM is not predicted to be associated with.

Determination of the genomic loci for specific PTM - CHiP detection - ChIP-chip

Microarray-based methods enable the analysis of histone modification enrichment at large numbers of loci simultaneously. The microarray itself is a coated glass slide affixed with different oligonucleotides – ranging in number from the tens of thousands to the tens of millions. Sites where a protein is enriched are determined by co-hybridizing to the array fluorescently labeled DNAs derived from the IP and input fractions and comparing the normalized IP/input intensities across the array (Figure 5C).

The first steps of preparing DNAs for microarray analysis are to amplify the DNA from each test (IP) and reference (input) sample and differentially label the amplification products with either of two fluorophores. DNA amplification can be accomplished in several ways, and the methods that are PCR based begin with the addition of primer binding sites to the DNA ends. This part can be done by either ligating the DNAs to short linkers of known sequence [57] or performing an initial two rounds of annealing and extension using primers that have degenerate 3' sequence and known 5' sequence [54, 58]. The end-tagged products are then PCR amplified for several rounds using primers that recognize the added linker sequence.

Another widely used method for DNA amplification involves the conversion of DNAs into transcription templates, followed by linear amplification of the products by transcription with T7 RNA polymerase [59]. To start, short polyT tails are added to the DNA 3' ends to create a priming site for first-strand synthesis by Klenow. The oligo used for priming contains a stretch of A's at the 3' end of the minimal T7 RNA polymerase promoter sequence, which enables subsequent amplification of the first-strand products by in vitro transcription.

Samples that have been amplified by PCR can be fluorescently labeled in one of two ways: (1) by incorporating a fluorophore-modified dNTP during the final set of PCR cycles or (2) indirectly, using an amine-modified dNTP that can be dye coupled later [53]. The same labeling principles apply to products amplified by transcription, except that the modified nucleotides are incorporated during a final reverse transcription step [53]. When preparing samples for labeling, it is important to also label an appropriate reference sample that the test-sample can be co-hybridized with. Thus, the labelings are set up such that one of the two samples is labeled with Cy3 and the other with Cy5. After the final amplification or post-labeling step, the samples are purified, and the labeled test and reference samples are mixed and prepared for hybridization.

Determination of the genomic loci for specific PTM - CHiP detection - ChIP-seq

Large-scale enrichment analysis can also be performed using a variety of massive parallel DNA sequencing methods. Such methods make possible the parallel sequencing of millions of DNA molecules in real time. The majority of ChIP-seq studies published so far were done using the "sequencing by synthesis" platform by Illumina [60]. The premise of this platform is to perform parallel sequencing of millions of clonal DNA clusters on the surface of a flow-cell (Figure 5D).

ChIP-seq samples are prepared by ligating the IP'd DNAs to oligonucleotide adapter molecules, after which the ligation products (in some protocols) are amplified for several rounds by PCR and then purified [61, 62]. Samples are then injected into a flow cell whose surface is coated with oligonucleotides complementary to the ligation product adapter sequences. The density of the tethered oligonucleotides is such that, during the amplification steps, the DNAs remain spatially close to the parent template, as the newly synthesized molecules originate from primers that are attached to the flow-cell surface adjacently. The sequencing phase of the protocol is accomplished by single-base extension using nucleotides that are fluorescently labeled and can terminate elongation reversibly. The sequencing reaction therefore proceeds as follows: a labeled nucleotide is added to the free 3' end, after which elongation pauses to enable detection of the incorporated nucleotide. Subsequently, the terminator group is cleaved to permit addition of the next nucleotide. Nucleotide extension is iterated for several more cycles, which typically results in read-lengths in the 40-bp range. For in depth review of next-generation sequencing methods, and comparison with microarray-based approaches, please see [63-66].

A recent publication "Systematic evaluation of factors influencing ChIP-seq fidelity" indicates that bias can be introduced due to the chromatin state [67]. Open chromatin regions often lead to false positive, and algorithmic adjustment should be made.

Determination of specific gene loci for specific PTM - PLA assay

A method has been developed to identify specific histone PTM for specific gene in a particular cell type in histological sections [68]. The cell type is marked through cell-specific marker with an antibody. The gene of interest is detected through in situ hybridization with a biotin-conjugated probe, and histone with specific PTM is labeled through a histone modification-specific antibody. Proximity ligation assay [69] is used to identify the co-location of the biotin label and histone antibody. The method enables detection at single cell level.

Common questions

I want to obtain Hela whole cell lysate without destroying its histone structure. What kind of cell lysis method should be used?

Ultrasonic cell disrupter (200W, 4 min for twice) or mechanical microfluidics. Add some glycerin in the buffer, absolutely no SDS in the buffer.

Which kind of Western blot internal control should I use?

If it is the whole cell lysates, then the regular cytoplasmic controls such as beta actin can be used. If it is the nuclear fraction, then the nuclear controls such as lamin B1 or PCNA can be used. If it is the purified histone fraction, than other members of the histone family, such as H2A can be used.

No histone positive bands are detected after Western blot?

One of the important considerations is that histone proteins have low molecular weights and can easily be transferred through the membrane during Western blotting transfer. Therefore, it's important to reduce the transfer time, and also make sure the markers with similar molecular weights show up on the membrane.

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ISSN : 2329-5139

2- Protocol

Nature Protocols 2, 1445 - 1457 (2007)

Published online: 7 June 2007 | doi:10.1038/nprot.2007.202

 Subject Categories: Isolation, purification and separation | Biochemistry and protein analysis

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David Shechter

Holger L Dormann

C David Allis

Sandra B Hake

Extraction, purification and analysis of histones

David Shechter1, Holger L Dormann1, C David Allis1 & Sandra B Hake1,2

Abstract

Histone proteins are the major protein components of chromatin, the physiologically relevant form of the genome (or epigenome) in all eukaryotic cells. Chromatin is the substrate of many biological processes, such as gene regulation and transcription, replication, mitosis and apoptosis. Since histones are extensively post-translationally modified, the identification of these covalent marks on canonical and variant histones is crucial for the understanding of their biological significance. Many different biochemical techniques have been developed to purify and separate histone proteins. Here, we present standard protocols for acid extraction and salt extraction of histones from chromatin; separation of extracted histones by reversed-phase HPLC; analysis of histones and their specific post-translational modification profiles by acid urea (AU) gel electrophoresis and the additional separation of non-canonical histone variants by triton AU(TAU) and 2D TAU electrophoresis; and immunoblotting of isolated histone proteins with modification-specific antibodies.

Good luck,

Rafik