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Detection of interactions between nucleosome arrays mediated by specific core histone tail domains
Methods 41 (2007) 278–285 www.elsevier.com/locate/ymeth
Detection of interactions between nucleosome arrays mediated by specific core histone tail domains Pu-Yeh Kan, Jeffrey J. Hayes
*
Department of Biochemistry and Biophysics, Box 712, University of Rochester Medical Center, Rochester, NY 14642, USA Accepted 17 August 2006
Abstract The core histone tail domains play important roles in different stages of chromatin condensation. The tails are required for folding nucleosome arrays into secondary chromatin structures such as the 30 nm diameter chromatin fiber and for mediating fiber–fiber interactions important for formation of tertiary chromatin structures. Crosslinking studies have demonstrated that inter-nucleosomal tail– DNA contacts appear in conjunction with salt-induced folding of nucleosome arrays into in higher order chromatin structures. However, since both folding of nucleosome arrays and fiber-fiber interactions take place simultaneously in >2–3 mM MgCl2 such inter-nucleosome interactions may reflect short range (intra-array) or longer range (inter-array) interactions. Here, we describe a novel technique to specifically identify inter-array interactions mediated by the histone tail domains. In addition, we describe a new method for the preparation of H3/H4 tetramers. 2006 Elsevier Inc. All rights reserved. Keywords: Histones; Chromatin; Histone tails; Nucleosomes; Nucleosome arrays; Chromatin folding
1. Introduction Chromatin in the eukaryotic nucleus is comprised of a hierarchy of structures formed by DNA, histone proteins and numerous non-histone chromosomal proteins such as HMG proteins, topoisomerases, HP1 and the Sir proteins [1,2]. Importantly, recent evidence indicates that at least the initial secondary and tertiary levels of chromatin structure can be generated by salt-induced condensation of nucleosome arrays containing only DNA and the four core histones. For example, both native and reconstituted nucleosome arrays exist as fully extended chains in low salt (TE) conditions but fold into maximally condensed secondary structures in the presence of 2 mM MgCl2 and further self-associate into oligomeric macrostructures in slightly higher MgCl2 [1,3,4]. This latter process can be detected by sedimentation in a benchtop microfuge as the arrays *
Corresponding author. Fax: +1 585 271 2683. E-mail address: [email protected] (J.J. Hayes).
1046-2023/$ - see front matter 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.ymeth.2006.08.012
undergo reversible divalent cation-dependent self-association into large (>300S) complexes [5]. On the other hand, monovalent cations such as Na+ can only induce moderate extents of chromatin folding and do not induce array self-association [4,5]. In addition to divalent cations, the core histone tail domains are also required for salt-dependent chromatin condensation. Oligonucleosome arrays lacking the tail domains are unable to form secondary chromatin structures such as the 30 nm fiber or self-associate into higherorder tertiary structures even in the presence of high MgCl2 [6–8]. Structural studies indicate that the H4 tail domain can contact a negatively charged patch on the surface of H2A/H2B dimers of neighboring nucleosomes [9,10]. Moreover, hydrodynamic studies of chromatin fibers containing selected combinations of the histone tail domains indicate that each core histone tail domain contributes to some extent to chromatin folding and oligomerization [8,11]. However, detailed mechanisms of core histone tail functions in chromatin condensation remain undefined.
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Crosslinking studies have been successful in determining the strengths and locations of histone tail interactions with DNA in chromatin complexes [12–14]. Previously our lab has characterized interactions between different histone tail domains and nucleosomal DNA [13,15–17]. Notably, we determined that the H3 tail domain participates in primarily intra-nucleosome contacts when a 13-mer nucleosome array is fully extended in low-salt conditions but rearranges to primarily inter-nucleosome interactions upon array condensation in elevated MgCl2 [17]. Interestingly, these interactions were not detected in a dinucleosome system suggesting that such systems do not recapitulate all native tail–DNA interactions [16,17]. In agreement with our results, hydrodynamic studies have also indicated H3/H4 tails play more important roles than H2A/H2B tails in both chromatin folding and inter-array self-association [8,11]. However, both chromatin folding and array self-association take place in the presence of >2 mM MgCl2 [5,18]. Thus, these inter-nucleosome contacts could represent either intra- or inter-array interactions important for either folding or self-association or both. We, therefore, designed a new experimental system that would allow the specific detection of inter-array histone tail–DNA interactions. We also describe a new method for purification and preparation of H3/H4 tetramers when the individual proteins are over-expressed in bacterial cells. 2. Methods We previously characterized intra- and inter-nucleosomal H3 tail–DNA interactions in a 13-mer nucleosome array system [16,17]. To determine whether inter-nucleosome
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interactions detected in our in vitro system represent interactions important for MgCl2-induced array folding or selfassociation or both we designed a novel method able to specifically detect inter-array interactions. This method employs two nucleosome arrays of different lengths that can be easily distinguished by gel electrophoresis: a 12mer array containing a radiolabeled core histone modified within the tail domain with a photoactivatable crosslinking agent and a 35-mer array reconstituted with native, unlabeled core histones (see below). Briefly, the arrays are mixed together and induced to undergo MgCl2-dependent array self-association. Then the arrays are briefly exposed to UV light to induce crosslinking and non-covalently bound proteins stripped from the DNA templates by SDS. Crosslinking is assessed by separation of the DNA templates on SDS–agarose gels, followed by staining and autoradiography. Detection of crosslinking to the template of the second array is taken as indicating the presence of inter-array interactions mediated by the tail domain of the radiolabeled histone (Fig. 1). Since we had previously examined inter-nucleosomal histone–DNA interactions mediated by the H3 tail domain within a nucleosomal array, we chose to investigate interarray interactions of the H3 tail. This required the generation of H3 mutants and purification of related H3/H4 tetramers (see below). However, in our hands purification of H3/H4 tetramers in the high yields required for array reconstitution has been inconsistent. Moreover, our protocol requires the preparation of radiolabeled H3 from limited amounts of bacterial culture grown in minimal media. Therefore, we developed a new method for the rapid and efficient purification of H3/H4 tetramers.
Fig. 1. Experimental design. A 35-mer nucleosome array is assembled with unlabeled native histones and mixed with 12-mer array reconstituted with three unlabeled, native histones and a 3H-labeled histone that is site-specifically modified with the bi-functional reagent 4-azidophenacyl bromide within the tail domain (APB) (H3 in the experiment shown). MgCl2 of different concentrations are then added to induce inter-array oligomerization. After UV irradiation, H3 tail crosslinking products with both DNA templates are viewed by autoradiography. The 3H signal detected on 35-mer DNA template indicates inter-array H3 tail–DNA interactions.
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2.1. Preparation of the 12-mer nucleosome array containing radiolabeled, APB-modified H3 2.1.1. 12-mer DNA template We chose to reconstitute the crosslinker-containing nucleosome arrays with a DNA template containing 12 208 bp tandem repeats of a DNA fragment containing a 5S gene from sea urchin Lytechinus variegates since the 5S repeat harbors a powerful nucleosome positioning sequence [19]. Moreover, folding and self-association of this array has been well characterized by numerous physical assays and appears to be identical to that of native arrays [1,3]. Our experimental plan also requires a second array with a DNA length distinguishable from the 12-mer array template by gel electrophoresis. We tested several length arrays also based on the 5S 208 bp repeat (6-mer, 11-mer, 14-mer, 35-mer) for this purpose and found that the 35-mer template was most easily separated from the 12-mer template and free histone proteins in our electrophoresis system (see below). The 12-mer template described in this work is a slight derivative of the standard 208-12 5S array template, and was obtained from the plasmid p12-5S-C1 (Cheeptip Benyajati, unpublished results). The original 208-12 plasmid was slightly modified from the original by the addition of a DraIII site at one end of the array [17]. However, the standard 208-12 template, or any other template for the generation of nucleosome arrays of similar size may be incorporated into the methodology described here. The 208-12 template was released by complete digestion of p12-5S-C1 with the restriction enzyme HhaI. Approximately, 3 mg of plasmid is digested at a DNA/HhaI ratio of about 1 lg DNA/0.3 U HhaI overnight at 37 C. The digestion products are then isolated by preparative gel electrophoresis (1% agarose, 1/2· TBE, with 0.2 lg/ml EtBr in both gel and running buffer) at 120 V for 2 h. According to the size of 208-12 template, a DNA fragment of 2.4 kb is excised from the agarose gel and electroeluted according to standard procedures. Briefly, the gel slice is placed in a dialysis bag with 1–2 volumes of electrophoresis buffer. The bag is returned to horizontal gel box and 120 V of current is applied for 2 h to elute DNA from the gel. The EtBr and any trace proteins in the eluted DNA are removed by three phenol–chloroform extractions followed by EtOH precipitation. We typically recover the insert in about 50–60% yield by this method. The reconstituted 35-mer array was a kind gift of X. Lu and J. Hansen. 2.1.2. Preparation of core histone proteins 2.1.2.1. Design of radiolabeled, chemically modified histones. In order to detect core histone tail domain–DNA interactions between nucleosome arrays, the core histone to be studied is modified with a photo-activatable crosslinking moiety at a defined position within the tail domain to allow histone tail–DNA crosslinking. In addition, the core histone protein should be labeled in some manner so that crosslinking products can be easily detected. To achieve
site-specific incorporation of a crosslinking moiety, we first generated a histone protein in which a single amino acid residue in the tail domain is substituted for cysteine. Typically, serine is chosen as this residue is isosteric with cysteine and common within the tails; otherwise a noncharged residue such as threonine or valine is chosen. The coding sequence for the protein is prepared in a bacterial expression vector and the protein expressed and purified (see below). The cysteine-containing protein is modified with 4-azidophenacylbromide (APB), a reagent we have found to provide for efficient cysteine modification and subsequent UV-light induced crosslinking, allowing tail interactions with DNA to be accurately mapped [13,15–17]. APB modification results in site-specific attachment of the phenacylazide photo-inducible crosslinking group to cysteine residues with no detectable modification of other residues [17]. Note that the APB-modified proteins should be stored in the dark and only minimal lighting used for all benchtop manipulations. Core histone sequences normally lack cysteine residues (except for C110 of H3 which has been replaced with alanine in our constructs, see below), which makes them ideal candidates for this approach. 2.1.2.2. Preparation of H3. The inter-nucleosomal H3 tail– DNA interactions we previously detected are likely to include some inter-array interactions. Thus we choose to study inter-array interactions mediated by the portion of the H3 tail that exhibited the strongest inter-nucleosomal interactions in our previous study, the very N-terminus of H3 tail, by using the mutant H3 protein H3T6C. The amino acid sequence of H3T6C is based on H3C110A, a Xenopus H3 sequence with the native cysteine residue mutated to alanine to avoid possible inappropriate modification of this cysteine [17,20]. BL21(DE3) cells transformed with pET3a-H3T6C are grown in minimal medium to enable radiolabeling with 3H-lysine. Due to the slow-growing nature of minimal medium culture, the culture is normally grown overnight to reach the proper OD600 for protein induction. Typically, 50 ml of M9-glucose minimal media (6 mg/ml Na2HPO4, 3 mg/ml KH2PO4, 0.5 mg/ml NaCl, 0.5 mg/ml NH4Cl, 0.6% glucose, 1 mM MgSO4, 0.3 mM CaCl2) supplemented with Biotin (1 lg/ml), Thiamin (1 lg/ml), Ampicillin (100 lg/ml) and trace elements (50 mg/L EDTA, 5 mg/L FeCl3, 0.84 mg/L ZnCl2, 0.1 mg/ L CuCl2, 0.05 mg/L CoCl2, 0.1 mg/L H3BO3, 8.6 lg/L MnCl2) are inoculated with BL21(DE3) harboring H3-expressing plasmids from glycerol stocks and grown overnight in a 200 mL flask with shaking. When the OD600 reaches 0.6 (usually early morning of the next day), IPTG is added to the culture to a final concentration of 0.4 mM to induce protein expression, followed 20 min later by addition of 3Hlysine (2.5 mCi) into the 50 ml of culture (see Note 1). The culture is then grown for an additional 3 h at 37 C, then the cells spun down at 5000 RPM in a GSA rotor and either used directly for purification of H3/H4 complexes or stored frozen at 80 C (see Note 2).
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2.1.2.3. Preparation of H4. Bacterial cells expressing H4 are prepared according to a method developed in our lab [20]. Typically, 50 ml of LB are inoculated with pET3a-H4 transformed BL21(DE3) cells from a glycerol stock then incubated at 37 C overnight without shaking. The OD600 of the culture normally reaches 0.4–0.5 in the next morning due to the slow-growing nature of cells without aeration. Cells are then placed in a 37 C shaker and grown for another 0.5–1 h to reach the density (0.6 OD600) for protein expression. IPTG is then added into the LB culture to a final concentration of 0.4 mM for induction of H4 expression. After 3 h of protein expression, cells are spun down and stored in the same manner as described for H3 above. 2.1.2.4. Preparation of H3/H4 tetramers. It is important to note that method we designed for purification of H2A/H2B dimers expressed in bacteria [21] does not work well for H3/H4 as these proteins have a high tendency to form aggregates of terminally misfolded species. Previously, our laboratory has developed and utilized two methods for the preparation of H3/H4 tetramers when these proteins are expressed independently in bacterial cells. One method involves alkaline lysis of H3 and H4 expressing bacterial cells, followed by mixing both cell lysates in 1:1 ratio and gently neutralizing the pH through dialysis [20]. Presumably, the genomic DNA present in the sample serves as a chaperone for H3/H4, helping to avoid protein aggregation [20]. Though producing high quality purified protein, the final yield of purified protein is estimated to be low, about 10%. A second protocol employs acid lysis of the cells to remove soluble material, followed by dissolving insoluble inclusion bodies by urea then a second acid solubilization/precipitation to remove DNA and non-acid-soluble bacterial proteins [22]. However, we find that H4 is not as acid soluble as H3 and thus the amount of H4 remaining after the second acid precipitation step becomes limiting. In order to increase the yield of H3/H4 tetramers, we designed a new method for H3/H4 purification that combines aspects of both these protocols: 1. Cell pellets from 50 ml cultures expressing H3 and cells containing an equal amount of H4 are each re-dissolved in 5 ml of 25 mM Tris, 5 mM EDTA, 0.2% Triton X-100. 2. Then 50 ll of 10 M NaOH is added to each tube, to reach a final concentration of 0.1 M NaOH. Alkaline lysis of cells is allowed to take place at room temperature for 1 h at 25 C, as in our alkaline lysis method. 3. The cell lysates are sonicated (Bramson Sonifier 250, output 4, duty cycle 50%) for 1–2 min to fragment chromosomal DNA. We have found that this step improves the yield as the long chromosomal DNA apparently traps some protein, reducing the final yield. 4. The lysates are mixed with equal volumes (5 ml) of 8 M urea to break apart insoluble inclusion bodies and to denature the proteins. Importantly, the process of dissolving inclusion bodies is slow and normally takes overnight.
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5. Nucleic acids are then removed by addition of 100 ll of 10 M HCl, with vortexing, followed immediately by centrifugation at 13 K RPM for 30 min in an SS34 (or similar) rotor at 4 C. The supernatant is then carefully removed and neutralized by the addition of 50 ll of 10 M NaOH. The pH of the solution should be 7–8, and may need slight adjustment with dilute acid or base (Fig. 2a). 6. The H3 and H4 lysates are then mixed together (such that H3 and H4 are present in a 1:1 ratio) and dialyzed against 2 M NaCl/TE overnight to remove urea and refold the proteins. 7. The dialyzed sample is then centrifuged to remove insoluble material, diluted threefold with TE to reduce the NaCl concentration to 0.6 M and the solution applied to a Biorex-70 column for final purification. Biorex-70 is a weakly acidic cation exchange resin that has been proven to be efficient in purification of core histone proteins [21]. Typically, 2 ml of Biorex-70 resin (50–100 mesh, 50% slurry in TE buffer) is used for purification of H3/H4 tetramers from 100 ml (combined total) of H3/H4-expressing culture. The Biorex-70 resin slurry is slowly poured into a Bio-Rad 10 ml Econo-Pac disposable column to form a bed volume of 1 ml. The dialyzed, TE diluted sample is then gently added into the column. After all the sample flows through the column, 100 ml of 0.6 M NaCl/TE is slowly applied to the sample-bound
Fig. 2. Preparation of 3H-labeled, APB modified H3/H4 tetramer. (a) SDS–PAGE of acid soluble proteins from the cleared lysate prepared from bacterial cells (see Section 2.1.2.4, Step 5) expressing H3 and H4. Lane 1, H3; lane 2, H4. Gel was stained with Commassie blue. (b) H3/H4 complex purified via Biorex-70 chromatography (see Section 2.1.2.4, Step 7). Lanes 1–10, fractions 1–10 eluted with 2 M NaCl/TE from the column. Fractions 3–6 were combined and used for reconstitution. (c) Check of crosslinking activity of APB-modified H3 in the H3/H4 complex. Purified, modified H3/H4 was irradiated with UV light as described in the text then samples analyzed on SDS–PAGE. Lane 1, UV ( ). Lane 2, UV (+). (d) Autoradiograph of (c) revealing 3H-H3. Bands due to crosslinking are indicated. A small amount of H5 added as a loading/no crosslinking control is indicated by the star.
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Biorex-70 resin to extensively wash out the contaminating DNA and bacterial proteins. The H3/H4 tetramer is then eluted by 2 M NaCl/TE (Fig. 2b). We have found that H3/H4 tetramers purified by this protocol are of high quality and can be isolated with high (>80%) yield. The cysteine group on H3 remains reduced and active toward cysteine-specific reagents such as APB (Fig. 2; see below). However, H3/H4 prepared by this method contains trace amounts of bacterial DNA, about 1 lg/ml, compared to about 1 mg/ml protein. In case of the currently described protocol, this amount of DNA is inconsequential and is ignored. However, if subsequent protocols require completely DNA-free proteins, trace DNAs may be removed by treatment with hydroxyapatite [20]. APB modification is carried out by mixing purified H3/ H4 tetramer and APB in a 1:5 molar ratio at 25 C in the dark for 1 h according to published protocols [20]. The efficiency of APB modification is determined by treatment of a small (0.5 lg) sample of the modification reaction with fluorescein 5-maleimide, which will react with any unmodified cysteines, and subsequent SDS–PAGE and visualization on a standard UV light box; completely APB-modified H3/H4 tetramers should exhibit no fluorescence. The activity of the crosslinker on the H3 tail is tested by a 30 s UV irradiation of a sample of free H3/H4 tetramer containing the modified protein followed by SDS– PAGE and Coomassie staining. Typically about 25% of the total H3 will form crosslinked products within the free tetramer if the crosslinking moiety is fully active (Fig. 2c and d). The modified H3/H4 is then used for reconstitution of 12-mer nucleosome arrays (see next section). 2.1.2.5. H2A/H2B dimers. The H2A/H2B dimers used for 12-mer array reconstitution are prepared from bacterially expressed proteins and purified by Biorex-70 cation exchange chromatography as described [21]. 2.1.3. Reconstitution of 12-mer and 35-mer nucleosome arrays Standard salt-dialysis is used for reconstitution of oligonucleosome arrays [18]. Basically core histone proteins and DNA template are mixed in 1:1 ratio in 2 M NaCl/ TE. Then the salt concentration in the mixture is gradually decreased through several rounds of dialysis. In the case of 12-mer nucleosome array reconstitution, 15 ll of 208-12mer DNA template (0.7 lg/ll) is mixed with 8 ll of 3H-labeled H3T6C/H4 (0.9 lg/ll) and 45 ll of H2A/H2B (0.15 lg/ll) in a final volume of 200 ll. The overall protein: DNA ratio is about 1.4: 1 (see Note 2). The mixture is sequentially dialyzed against 1 M NaCl/TE for 4 h, 0.75 M NaCl/TE for another 3 h, and 10 mM Tris/0.1 mM EDTA/0.1 mM EGTA overnight. The lower EDTA concentration in the final dialysis step is to avoid interference of EDTA when Mg2+ is added back to the chromatin to induce folding and selfassociation.
After reconstitution, the level of nucleosome saturation is determined by two methods. The first method involves digestion of the reconstituted templates with the restriction enzyme EcoRI, which cuts at a pair of EcoRI enzyme sites in the ‘linker’ regions between where the nucleosome cores are likely to be located on the 208 bp 5s rRNA nucleosome positioning sequence [19]. Digestions are carried out by treating 0.5 lg of reconstituted template (10 ll) with 20 U EcoRI in NEB buffer 2 in a total volume of 20 ll for 1 h at 37 C. After digestion, the digestions are separated on 0.7% agarose nucleoprotein gels and the saturation levels determined by quantitative comparison of the mononucleosome and free DNA bands [23]. Previous work by the Hansen lab has shown that the digestion should yield primarily mononucleosomes for a saturated template, with about 4–5% of the template running as naked DNA when the array templates are properly saturated with nucleosomes (Fig. 3a). A second method to check the level of nucleosome saturation is based on the propensity of nucleosome arrays to self-associate in the presence of divalent cations and the fact that arrays containing different levels of saturation tend to self-associate at different concentrations of divalent cations [5]. We therefore test all reconstituted arrays for the extent of self-association in increasing concentrations of MgCl2 to determine the level of saturation as compared to literature values [5]. We find that the arrays undergo about 40–80% self-association in 3–4 mM, MgCl2, respectively, when fully saturated. Arrays should be tested by both EcoRI digestion and self-association assay before being used for crosslinking studies (Fig. 3b and c). 2.2. Detection of inter-array crosslinking As mentioned above, our basic experimental design allows detection of inter-array interactions between the H3 tail domain on the 12-mer array and the DNA of the 35-mer array. Thus we mix 12-mer and 35-mer arrays in the presence of MgCl2 to induce inter-fiber self association of the two arrays, followed by UV light-induced crosslinking and detection of crosslinks by SDS–agarose gel electrophoresis, EtBr staining and autoradiography. Since 3H-labeled H3 is the only source of radioactive signal in the system and is only reconstituted into 12-mer array, any detectable 3 H signal co-migrating with the 35-mer DNA template likely results from inter-array H3 tail–DNA interactions. The inter-array crosslinking experiment is carried out as described. All steps are carried out at room temperature: 1. Mix 8.1 ll of unlabeled 35-mer array (0.2 lg/ll) with 8.1 ll of 3H-labeled 12-mer array (0.04 lg/ll) in a siliconized microfuge tube (see Note 4). 2. Add 1.8 ll of 10 mM Tris containing 0–80 mM MgCl2 into the sample, mix immediately and rapidly by pippeting to such that the final MgCl2 concentration is 0–8 mM.
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solution and help convert array oligomers into monomers. Two microliters of 10· gel loading dye (0.1% SDS and 0.02% EtBr) is then added into each sample and the samples incubated for 10 min at 37 C to dissociate non-covalently bound proteins from the DNA templates. Samples are then analyzed by 0.7% agarose gel electrophoresis. Both the agarose gel and electrophoresis buffer (0.5· TBE) contain the same concentrations of SDS and EtBr as samples (0.01% SDS, 0.2 lg/ml EtBr) to maximize protein removal from DNA. Consequently only covalently crosslinked proteins co-migrate with the DNA templates. After 1 h of gel electrophoresis at 120 V, the DNA templates of 12-mer and 35-mer arrays are well separated on the gel (Fig. 4a). The location of the EtBr-stained DNA templates on the gel is recorded by digital photography under UV light. The gel is then soaked in standard SDS– PAGE destain buffer I (45% methanol, 10 % acetic acid) for 15 min, washed with ddH2O, and soaked in 1 M sodium salicylate for another 15 min to provide an enhancer for the detection of 3H-labeled species [24]. The gel is then dried and exposed to autoradiographic film (Kodak BioMax XAR) at 80 C for 1–2 weeks (see Note 5). 2.4. Analysis of the results Bands corresponding to the 12-mer and 35-mer templates are apparent in the ethidium bromide-stained gel (Fig. 4a).
Fig. 3. Determination of the level of nucleosome saturation. (a) Effect of saturation on EcoRI digestion. Samples of 12-mer DNA templates reconstituted with about 12, 10, and 8 nucleosomes per template were treated with EcoRI and digestion products separated on a native 0.7% agarose nucleoprotein gel. Lane 1 shows the result expected for full saturation of the template, with about 5% of the monomer DNA appearing as free DNA. (b) MgCl2-dependent self-association assay. The same arrays analyzed in (a) were incubated with increasing concentrations of MgCl2 then centrifuged for 15 min at maximum speed in a benchtop microfuge. The supernatants were loaded onto the SDS–agarose gel and the gel briefly electrophoresed. The saturated, unsaturated and very unsaturated arrays are shown top to bottom in the gel, respectively. (c) The fraction of arrays in the supernatant was quantified from the gel in (b) and plotted vs MgCl2 concentration.
3. Transfer the sample into a Falcon 5-ml polystyrene tube, and then place the tube into a 15-ml Pyrex 9820 glass tube. This two-layer protection excludes shorter wavelength UV light that may be directly absorbed and damage the DNA templates. 4. Samples are then exposed to 365 nm UV light emitted from a standard VMR LM-20E light box for 1 min to trigger the crosslinking reaction [20]. 2.3. Analysis of crosslinking by agarose gel electrophoresis/ autoradiography After UV irradiation, 0.5 ll of 100 mM EDTA is immediately added into each sample as to chelate Mg2+ from the
Fig. 4. Inter-array crosslinking between H3 tail domains and the 35-mer array. (a) Inter-array crosslinking is dependent upon MgCl2 and UV irradiation. A sample containing the 12-mer and 35-mer arrays was prepared as described in the text, irradiated with UV light and products separated on an SDS–agarose gel. The concentration of MgCl2 in each sample is indicated above the gels. (a) EtBr staining. (b) Autoradiograph. A band corresponding to crosslinking of radiolabeled H3 to the 35-mer template is indicated as an inter-array crosslinking while crosslinking to the 12-mer array is indicated as intra-array crosslinking. The location of the two templates in the top gel and free H3 in the bottom gel is indicated. Note that the sample in lane 1 was not subjected to irradiation, as indicated.
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The autoradiograph of the gels reveals several bands. In the absence of irradiation, free H3 migrates as a diffuse band, below the 12-mer template (Fig. 4b). When samples are irradiated in the absence of MgCl2, some amount of radioactive H3 co-migrates with the 12-mer template as expected since the nucleosomes associated with this template contain the labeled, APB-modified protein. This crosslinking results in a small shift in the position of the 12-mer template on the ethidium-stained gel (Fig. 4a, compare lanes 1 and 2). Note that no crosslinking is detected to the 35-mer template in the absence of MgCl2. However, in the presence of MgCl2, some radioactivity now co-migrates with the 35-mer template, indicating that some inter-array crosslinking has taken place. The gel can be scanned on a standard densitometer and the fraction of the total crosslinked protein that co-migrates with the 35-mer template determined (not shown). About 10% of the crosslinked H3 co-migrated with the 35mer array template when the crosslinking was carried out in 4 mM MgCl2 (Fig. 4). According to the MgCl2 self-association assay, about 80% of 12-mer array and about 60% of 35-mer array are assembled into large, self-associated oligomers in the presence of 4 mM MgCl2 (Fig. 3). Self-association increases to 100% of both templates at higher MgCl2 concentrations and may involve assembly into even large structures [5]. Accordingly, inter-array crosslinks formed by the H3 tail constitute 20% of total histone–DNA crosslinks in 6–8 mM MgCl2 (Fig. 4). 3. Summary Our method allows probable inter-array interactions mediated by a histone tail domain to be detected via crosslinking. These interactions are thought to be important for formation of higher-order tertiary chromatin structures modeled by the MgCl2-dependent oligomerization of model nucleosome arrays [5,11]. The excess 35-mer serves as a sink for presumably the majority of such interactions by selected tail domains located on the 12-mer array. We estimate that our method can detect inter-array crosslinks if they represent minimally 1–2% of total crosslinks. Previously, we determined that the H3 tail domain participates in primarily inter-nucleosomal interactions when the array is condensed in MgCl2 [16,17]. Using our new method, we find that about 20% of these inter-nucleosome interactions occur between arrays and about 80% are intra-array. Thus the H3 tail domain contributes to both folding of the oligonucleosome array and to array oligomerization. Our inter-array crosslinking experiment is easily extended to examine contacts made by other positions in the H3 tail and other tail domains by moving the location of the APB-derived crosslinker to other positions of H3 tail or other histone tail domains. In addition, the effect of posttranslational modifications and the binding of linker histones on these interactions can be examined. For example, lysine fi glutamine substitutions can be used to approximate lysine acetylation or such modifications can be introduced chemically [25] or enzymatically.
4. Notes Note 1: In order to obtain the highest efficiency of 3H incorporation in the expressed protein, we have found that the timing for the addition of the 3H-lysine is critical. Thus, in trial runs, the expression of the target protein expression is carefully monitored after IPTG addition, and 3H-lysine is added just before H3T6C expression is apparent on by SDS–PAGE of whole cells. Typically for H3 expression the optimal addition is 20 min after IPTG addition. Note 2: Retain the supernatant after spinning down cells expressing radiolabeled protein and dispose of properly in the radioactive waste. Note 3: In practice the actual ratio of histones–DNA template must be empirically determined to avoid underor over-saturating the templates with histones. Thus, several trial reconstitutions are typically set up with histone:DNA ratios covering the range 0.9:1 to 1.5:1 were examined. The level of saturation is determined as described. Note 4: An excess of the unlabeled 35-mer array is used to maximize the chance that inter-array interactions by the radiolabled and APB-modified histone occur to this template since interarray interactions between 12-mer templates will not be detected in our experiment. Note 5: Make sure that the film is pressed tightly up against the gel during the exposure. We use paper cassettes and place a weight on top of the cassette to ensure that a tight fit. Also, given the very long half-life of 3H, exposure times may be adjusted over a wide range of times depending on the specific activity of the protein. Acknowledgments We thank Drs. Jeffrey Hansen and Xu Lu for kindly providing the 35-mer nucleosome array. This work was supported by NIH Grant GM52426. References [1] J.C. Hansen, Annu. Rev. Biophys. Biomol. Struct. 31 (2002) 361–392. [2] C.L. Woodcock, S. Dimitrov, Curr. Opin. Genet. Dev. 11 (2001) 130–135. [3] M. Garcia-Ramirez, F. Dong, J. Ausio, J. Biol. Chem. 267 (1992) 19587–19595. [4] J.C. Hansen, J. Ausio, V.H. Stanik, K.E. van Holde, Biochemistry 28 (1989) 9129–9136. [5] P.M. Schwarz, A. Felthauser, T.M. Fletcher, J.C. Hansen, Biochemistry 35 (1996) 4009–4015. [6] J. Allan, N. Harborne, D.C. Rau, H. Gould, J. Cell Biol. 93 (1982) 285–297. [7] T.M. Fletcher, J.C. Hansen, J. Biol. Chem. 270 (1995) 25359–25362. [8] C. Tse, J.C. Hansen, Biochemistry 36 (1997) 11381–11388. [9] K. Luger, A.W. Mader, R.K. Richmond, D.F. Sargent, T.J. Richmond, Nature 389 (1997) 251–260. [10] T. Schalch, S. Duda, D.F. Sargent, T.J. Richmond, Nature 436 (2005) 138–141. [11] F. Gordon, K. Luger, J.C. Hansen, J. Biol. Chem. 280 (2005) 33701–33706.
P.-Y. Kan, J.J. Hayes / Methods 41 (2007) 278–285 [12] D. Angelov, V. Stefanovsky, S.I. Dimitrov, V.R. Russanova, E. Keskinova, I.G. Pashev, Nucleic Acids Res. 16 (1988) 4525–4538. [13] D. Angelov, J.M. Vitolo, V. Mutskov, S. Dimitrov, J.J. Hayes, Proc. Natl. Acad. Sci. USA 98 (2001) 6599–6604. [14] S.I. Usachenko, S.G. Bavykin, I.M. Gavin, E.M. Bradbury, Proc. Natl. Acad. Sci. USA 91 (1994) 6845–6849. [15] K.M. Lee, J.J. Hayes, Biochemistry 37 (1998) 8622–8628. [16] C. Zheng, J.J. Hayes, J. Biol. Chem. 278 (2003) 24217–24224. [17] C. Zheng, X. Lu, J.C. Hansen, J.J. Hayes, J. Biol. Chem. 280 (2005) 33552–33557.
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P.M. Schwarz, J.C. Hansen, J. Biol. Chem. 269 (1994) 16284–16289. R.T. Simpson, F. Thoma, J.M. Brubaker, Cell 42 (1985) 799–808. C. Zheng, J.J. Hayes, Methods Enzymol. 375 (2004) 179–193. J.J. Hayes, K.M. Lee, Methods 12 (1997) 2–9. C. Thiriet, Methods 33 (2004) 86–92. C. Tse, T. Sera, A.P. Wolffe, J.C. Hansen, Mol. Cell. Biol. 18 (1998) 4629–4638. [24] Z. Yang, C. Zheng, C. Thiriet, J.J. Hayes, Mol. Cell. Biol. 25 (2005) 241–249. [25] M. Shogren-Knaak, H. Ishii, J.M. Sun, M.J. Pazin, J.R. Davie, C.L. Peterson, Science 311 (2006) 844–847.
Detection of interactions between nucleosome arrays mediated by specific core histone tail domains Pu-Yeh Kan, Jeffrey J. Hayes
*
Department of Biochemistry and Biophysics, Box 712, University of Rochester Medical Center, Rochester, NY 14642, USA Accepted 17 August 2006
Abstract The core histone tail domains play important roles in different stages of chromatin condensation. The tails are required for folding nucleosome arrays into secondary chromatin structures such as the 30 nm diameter chromatin fiber and for mediating fiber–fiber interactions important for formation of tertiary chromatin structures. Crosslinking studies have demonstrated that inter-nucleosomal tail– DNA contacts appear in conjunction with salt-induced folding of nucleosome arrays into in higher order chromatin structures. However, since both folding of nucleosome arrays and fiber-fiber interactions take place simultaneously in >2–3 mM MgCl2 such inter-nucleosome interactions may reflect short range (intra-array) or longer range (inter-array) interactions. Here, we describe a novel technique to specifically identify inter-array interactions mediated by the histone tail domains. In addition, we describe a new method for the preparation of H3/H4 tetramers. 2006 Elsevier Inc. All rights reserved. Keywords: Histones; Chromatin; Histone tails; Nucleosomes; Nucleosome arrays; Chromatin folding
1. Introduction Chromatin in the eukaryotic nucleus is comprised of a hierarchy of structures formed by DNA, histone proteins and numerous non-histone chromosomal proteins such as HMG proteins, topoisomerases, HP1 and the Sir proteins [1,2]. Importantly, recent evidence indicates that at least the initial secondary and tertiary levels of chromatin structure can be generated by salt-induced condensation of nucleosome arrays containing only DNA and the four core histones. For example, both native and reconstituted nucleosome arrays exist as fully extended chains in low salt (TE) conditions but fold into maximally condensed secondary structures in the presence of 2 mM MgCl2 and further self-associate into oligomeric macrostructures in slightly higher MgCl2 [1,3,4]. This latter process can be detected by sedimentation in a benchtop microfuge as the arrays *
Corresponding author. Fax: +1 585 271 2683. E-mail address: [email protected] (J.J. Hayes).
1046-2023/$ - see front matter 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.ymeth.2006.08.012
undergo reversible divalent cation-dependent self-association into large (>300S) complexes [5]. On the other hand, monovalent cations such as Na+ can only induce moderate extents of chromatin folding and do not induce array self-association [4,5]. In addition to divalent cations, the core histone tail domains are also required for salt-dependent chromatin condensation. Oligonucleosome arrays lacking the tail domains are unable to form secondary chromatin structures such as the 30 nm fiber or self-associate into higherorder tertiary structures even in the presence of high MgCl2 [6–8]. Structural studies indicate that the H4 tail domain can contact a negatively charged patch on the surface of H2A/H2B dimers of neighboring nucleosomes [9,10]. Moreover, hydrodynamic studies of chromatin fibers containing selected combinations of the histone tail domains indicate that each core histone tail domain contributes to some extent to chromatin folding and oligomerization [8,11]. However, detailed mechanisms of core histone tail functions in chromatin condensation remain undefined.
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Crosslinking studies have been successful in determining the strengths and locations of histone tail interactions with DNA in chromatin complexes [12–14]. Previously our lab has characterized interactions between different histone tail domains and nucleosomal DNA [13,15–17]. Notably, we determined that the H3 tail domain participates in primarily intra-nucleosome contacts when a 13-mer nucleosome array is fully extended in low-salt conditions but rearranges to primarily inter-nucleosome interactions upon array condensation in elevated MgCl2 [17]. Interestingly, these interactions were not detected in a dinucleosome system suggesting that such systems do not recapitulate all native tail–DNA interactions [16,17]. In agreement with our results, hydrodynamic studies have also indicated H3/H4 tails play more important roles than H2A/H2B tails in both chromatin folding and inter-array self-association [8,11]. However, both chromatin folding and array self-association take place in the presence of >2 mM MgCl2 [5,18]. Thus, these inter-nucleosome contacts could represent either intra- or inter-array interactions important for either folding or self-association or both. We, therefore, designed a new experimental system that would allow the specific detection of inter-array histone tail–DNA interactions. We also describe a new method for purification and preparation of H3/H4 tetramers when the individual proteins are over-expressed in bacterial cells. 2. Methods We previously characterized intra- and inter-nucleosomal H3 tail–DNA interactions in a 13-mer nucleosome array system [16,17]. To determine whether inter-nucleosome
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interactions detected in our in vitro system represent interactions important for MgCl2-induced array folding or selfassociation or both we designed a novel method able to specifically detect inter-array interactions. This method employs two nucleosome arrays of different lengths that can be easily distinguished by gel electrophoresis: a 12mer array containing a radiolabeled core histone modified within the tail domain with a photoactivatable crosslinking agent and a 35-mer array reconstituted with native, unlabeled core histones (see below). Briefly, the arrays are mixed together and induced to undergo MgCl2-dependent array self-association. Then the arrays are briefly exposed to UV light to induce crosslinking and non-covalently bound proteins stripped from the DNA templates by SDS. Crosslinking is assessed by separation of the DNA templates on SDS–agarose gels, followed by staining and autoradiography. Detection of crosslinking to the template of the second array is taken as indicating the presence of inter-array interactions mediated by the tail domain of the radiolabeled histone (Fig. 1). Since we had previously examined inter-nucleosomal histone–DNA interactions mediated by the H3 tail domain within a nucleosomal array, we chose to investigate interarray interactions of the H3 tail. This required the generation of H3 mutants and purification of related H3/H4 tetramers (see below). However, in our hands purification of H3/H4 tetramers in the high yields required for array reconstitution has been inconsistent. Moreover, our protocol requires the preparation of radiolabeled H3 from limited amounts of bacterial culture grown in minimal media. Therefore, we developed a new method for the rapid and efficient purification of H3/H4 tetramers.
Fig. 1. Experimental design. A 35-mer nucleosome array is assembled with unlabeled native histones and mixed with 12-mer array reconstituted with three unlabeled, native histones and a 3H-labeled histone that is site-specifically modified with the bi-functional reagent 4-azidophenacyl bromide within the tail domain (APB) (H3 in the experiment shown). MgCl2 of different concentrations are then added to induce inter-array oligomerization. After UV irradiation, H3 tail crosslinking products with both DNA templates are viewed by autoradiography. The 3H signal detected on 35-mer DNA template indicates inter-array H3 tail–DNA interactions.
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2.1. Preparation of the 12-mer nucleosome array containing radiolabeled, APB-modified H3 2.1.1. 12-mer DNA template We chose to reconstitute the crosslinker-containing nucleosome arrays with a DNA template containing 12 208 bp tandem repeats of a DNA fragment containing a 5S gene from sea urchin Lytechinus variegates since the 5S repeat harbors a powerful nucleosome positioning sequence [19]. Moreover, folding and self-association of this array has been well characterized by numerous physical assays and appears to be identical to that of native arrays [1,3]. Our experimental plan also requires a second array with a DNA length distinguishable from the 12-mer array template by gel electrophoresis. We tested several length arrays also based on the 5S 208 bp repeat (6-mer, 11-mer, 14-mer, 35-mer) for this purpose and found that the 35-mer template was most easily separated from the 12-mer template and free histone proteins in our electrophoresis system (see below). The 12-mer template described in this work is a slight derivative of the standard 208-12 5S array template, and was obtained from the plasmid p12-5S-C1 (Cheeptip Benyajati, unpublished results). The original 208-12 plasmid was slightly modified from the original by the addition of a DraIII site at one end of the array [17]. However, the standard 208-12 template, or any other template for the generation of nucleosome arrays of similar size may be incorporated into the methodology described here. The 208-12 template was released by complete digestion of p12-5S-C1 with the restriction enzyme HhaI. Approximately, 3 mg of plasmid is digested at a DNA/HhaI ratio of about 1 lg DNA/0.3 U HhaI overnight at 37 C. The digestion products are then isolated by preparative gel electrophoresis (1% agarose, 1/2· TBE, with 0.2 lg/ml EtBr in both gel and running buffer) at 120 V for 2 h. According to the size of 208-12 template, a DNA fragment of 2.4 kb is excised from the agarose gel and electroeluted according to standard procedures. Briefly, the gel slice is placed in a dialysis bag with 1–2 volumes of electrophoresis buffer. The bag is returned to horizontal gel box and 120 V of current is applied for 2 h to elute DNA from the gel. The EtBr and any trace proteins in the eluted DNA are removed by three phenol–chloroform extractions followed by EtOH precipitation. We typically recover the insert in about 50–60% yield by this method. The reconstituted 35-mer array was a kind gift of X. Lu and J. Hansen. 2.1.2. Preparation of core histone proteins 2.1.2.1. Design of radiolabeled, chemically modified histones. In order to detect core histone tail domain–DNA interactions between nucleosome arrays, the core histone to be studied is modified with a photo-activatable crosslinking moiety at a defined position within the tail domain to allow histone tail–DNA crosslinking. In addition, the core histone protein should be labeled in some manner so that crosslinking products can be easily detected. To achieve
site-specific incorporation of a crosslinking moiety, we first generated a histone protein in which a single amino acid residue in the tail domain is substituted for cysteine. Typically, serine is chosen as this residue is isosteric with cysteine and common within the tails; otherwise a noncharged residue such as threonine or valine is chosen. The coding sequence for the protein is prepared in a bacterial expression vector and the protein expressed and purified (see below). The cysteine-containing protein is modified with 4-azidophenacylbromide (APB), a reagent we have found to provide for efficient cysteine modification and subsequent UV-light induced crosslinking, allowing tail interactions with DNA to be accurately mapped [13,15–17]. APB modification results in site-specific attachment of the phenacylazide photo-inducible crosslinking group to cysteine residues with no detectable modification of other residues [17]. Note that the APB-modified proteins should be stored in the dark and only minimal lighting used for all benchtop manipulations. Core histone sequences normally lack cysteine residues (except for C110 of H3 which has been replaced with alanine in our constructs, see below), which makes them ideal candidates for this approach. 2.1.2.2. Preparation of H3. The inter-nucleosomal H3 tail– DNA interactions we previously detected are likely to include some inter-array interactions. Thus we choose to study inter-array interactions mediated by the portion of the H3 tail that exhibited the strongest inter-nucleosomal interactions in our previous study, the very N-terminus of H3 tail, by using the mutant H3 protein H3T6C. The amino acid sequence of H3T6C is based on H3C110A, a Xenopus H3 sequence with the native cysteine residue mutated to alanine to avoid possible inappropriate modification of this cysteine [17,20]. BL21(DE3) cells transformed with pET3a-H3T6C are grown in minimal medium to enable radiolabeling with 3H-lysine. Due to the slow-growing nature of minimal medium culture, the culture is normally grown overnight to reach the proper OD600 for protein induction. Typically, 50 ml of M9-glucose minimal media (6 mg/ml Na2HPO4, 3 mg/ml KH2PO4, 0.5 mg/ml NaCl, 0.5 mg/ml NH4Cl, 0.6% glucose, 1 mM MgSO4, 0.3 mM CaCl2) supplemented with Biotin (1 lg/ml), Thiamin (1 lg/ml), Ampicillin (100 lg/ml) and trace elements (50 mg/L EDTA, 5 mg/L FeCl3, 0.84 mg/L ZnCl2, 0.1 mg/ L CuCl2, 0.05 mg/L CoCl2, 0.1 mg/L H3BO3, 8.6 lg/L MnCl2) are inoculated with BL21(DE3) harboring H3-expressing plasmids from glycerol stocks and grown overnight in a 200 mL flask with shaking. When the OD600 reaches 0.6 (usually early morning of the next day), IPTG is added to the culture to a final concentration of 0.4 mM to induce protein expression, followed 20 min later by addition of 3Hlysine (2.5 mCi) into the 50 ml of culture (see Note 1). The culture is then grown for an additional 3 h at 37 C, then the cells spun down at 5000 RPM in a GSA rotor and either used directly for purification of H3/H4 complexes or stored frozen at 80 C (see Note 2).
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2.1.2.3. Preparation of H4. Bacterial cells expressing H4 are prepared according to a method developed in our lab [20]. Typically, 50 ml of LB are inoculated with pET3a-H4 transformed BL21(DE3) cells from a glycerol stock then incubated at 37 C overnight without shaking. The OD600 of the culture normally reaches 0.4–0.5 in the next morning due to the slow-growing nature of cells without aeration. Cells are then placed in a 37 C shaker and grown for another 0.5–1 h to reach the density (0.6 OD600) for protein expression. IPTG is then added into the LB culture to a final concentration of 0.4 mM for induction of H4 expression. After 3 h of protein expression, cells are spun down and stored in the same manner as described for H3 above. 2.1.2.4. Preparation of H3/H4 tetramers. It is important to note that method we designed for purification of H2A/H2B dimers expressed in bacteria [21] does not work well for H3/H4 as these proteins have a high tendency to form aggregates of terminally misfolded species. Previously, our laboratory has developed and utilized two methods for the preparation of H3/H4 tetramers when these proteins are expressed independently in bacterial cells. One method involves alkaline lysis of H3 and H4 expressing bacterial cells, followed by mixing both cell lysates in 1:1 ratio and gently neutralizing the pH through dialysis [20]. Presumably, the genomic DNA present in the sample serves as a chaperone for H3/H4, helping to avoid protein aggregation [20]. Though producing high quality purified protein, the final yield of purified protein is estimated to be low, about 10%. A second protocol employs acid lysis of the cells to remove soluble material, followed by dissolving insoluble inclusion bodies by urea then a second acid solubilization/precipitation to remove DNA and non-acid-soluble bacterial proteins [22]. However, we find that H4 is not as acid soluble as H3 and thus the amount of H4 remaining after the second acid precipitation step becomes limiting. In order to increase the yield of H3/H4 tetramers, we designed a new method for H3/H4 purification that combines aspects of both these protocols: 1. Cell pellets from 50 ml cultures expressing H3 and cells containing an equal amount of H4 are each re-dissolved in 5 ml of 25 mM Tris, 5 mM EDTA, 0.2% Triton X-100. 2. Then 50 ll of 10 M NaOH is added to each tube, to reach a final concentration of 0.1 M NaOH. Alkaline lysis of cells is allowed to take place at room temperature for 1 h at 25 C, as in our alkaline lysis method. 3. The cell lysates are sonicated (Bramson Sonifier 250, output 4, duty cycle 50%) for 1–2 min to fragment chromosomal DNA. We have found that this step improves the yield as the long chromosomal DNA apparently traps some protein, reducing the final yield. 4. The lysates are mixed with equal volumes (5 ml) of 8 M urea to break apart insoluble inclusion bodies and to denature the proteins. Importantly, the process of dissolving inclusion bodies is slow and normally takes overnight.
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5. Nucleic acids are then removed by addition of 100 ll of 10 M HCl, with vortexing, followed immediately by centrifugation at 13 K RPM for 30 min in an SS34 (or similar) rotor at 4 C. The supernatant is then carefully removed and neutralized by the addition of 50 ll of 10 M NaOH. The pH of the solution should be 7–8, and may need slight adjustment with dilute acid or base (Fig. 2a). 6. The H3 and H4 lysates are then mixed together (such that H3 and H4 are present in a 1:1 ratio) and dialyzed against 2 M NaCl/TE overnight to remove urea and refold the proteins. 7. The dialyzed sample is then centrifuged to remove insoluble material, diluted threefold with TE to reduce the NaCl concentration to 0.6 M and the solution applied to a Biorex-70 column for final purification. Biorex-70 is a weakly acidic cation exchange resin that has been proven to be efficient in purification of core histone proteins [21]. Typically, 2 ml of Biorex-70 resin (50–100 mesh, 50% slurry in TE buffer) is used for purification of H3/H4 tetramers from 100 ml (combined total) of H3/H4-expressing culture. The Biorex-70 resin slurry is slowly poured into a Bio-Rad 10 ml Econo-Pac disposable column to form a bed volume of 1 ml. The dialyzed, TE diluted sample is then gently added into the column. After all the sample flows through the column, 100 ml of 0.6 M NaCl/TE is slowly applied to the sample-bound
Fig. 2. Preparation of 3H-labeled, APB modified H3/H4 tetramer. (a) SDS–PAGE of acid soluble proteins from the cleared lysate prepared from bacterial cells (see Section 2.1.2.4, Step 5) expressing H3 and H4. Lane 1, H3; lane 2, H4. Gel was stained with Commassie blue. (b) H3/H4 complex purified via Biorex-70 chromatography (see Section 2.1.2.4, Step 7). Lanes 1–10, fractions 1–10 eluted with 2 M NaCl/TE from the column. Fractions 3–6 were combined and used for reconstitution. (c) Check of crosslinking activity of APB-modified H3 in the H3/H4 complex. Purified, modified H3/H4 was irradiated with UV light as described in the text then samples analyzed on SDS–PAGE. Lane 1, UV ( ). Lane 2, UV (+). (d) Autoradiograph of (c) revealing 3H-H3. Bands due to crosslinking are indicated. A small amount of H5 added as a loading/no crosslinking control is indicated by the star.
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Biorex-70 resin to extensively wash out the contaminating DNA and bacterial proteins. The H3/H4 tetramer is then eluted by 2 M NaCl/TE (Fig. 2b). We have found that H3/H4 tetramers purified by this protocol are of high quality and can be isolated with high (>80%) yield. The cysteine group on H3 remains reduced and active toward cysteine-specific reagents such as APB (Fig. 2; see below). However, H3/H4 prepared by this method contains trace amounts of bacterial DNA, about 1 lg/ml, compared to about 1 mg/ml protein. In case of the currently described protocol, this amount of DNA is inconsequential and is ignored. However, if subsequent protocols require completely DNA-free proteins, trace DNAs may be removed by treatment with hydroxyapatite [20]. APB modification is carried out by mixing purified H3/ H4 tetramer and APB in a 1:5 molar ratio at 25 C in the dark for 1 h according to published protocols [20]. The efficiency of APB modification is determined by treatment of a small (0.5 lg) sample of the modification reaction with fluorescein 5-maleimide, which will react with any unmodified cysteines, and subsequent SDS–PAGE and visualization on a standard UV light box; completely APB-modified H3/H4 tetramers should exhibit no fluorescence. The activity of the crosslinker on the H3 tail is tested by a 30 s UV irradiation of a sample of free H3/H4 tetramer containing the modified protein followed by SDS– PAGE and Coomassie staining. Typically about 25% of the total H3 will form crosslinked products within the free tetramer if the crosslinking moiety is fully active (Fig. 2c and d). The modified H3/H4 is then used for reconstitution of 12-mer nucleosome arrays (see next section). 2.1.2.5. H2A/H2B dimers. The H2A/H2B dimers used for 12-mer array reconstitution are prepared from bacterially expressed proteins and purified by Biorex-70 cation exchange chromatography as described [21]. 2.1.3. Reconstitution of 12-mer and 35-mer nucleosome arrays Standard salt-dialysis is used for reconstitution of oligonucleosome arrays [18]. Basically core histone proteins and DNA template are mixed in 1:1 ratio in 2 M NaCl/ TE. Then the salt concentration in the mixture is gradually decreased through several rounds of dialysis. In the case of 12-mer nucleosome array reconstitution, 15 ll of 208-12mer DNA template (0.7 lg/ll) is mixed with 8 ll of 3H-labeled H3T6C/H4 (0.9 lg/ll) and 45 ll of H2A/H2B (0.15 lg/ll) in a final volume of 200 ll. The overall protein: DNA ratio is about 1.4: 1 (see Note 2). The mixture is sequentially dialyzed against 1 M NaCl/TE for 4 h, 0.75 M NaCl/TE for another 3 h, and 10 mM Tris/0.1 mM EDTA/0.1 mM EGTA overnight. The lower EDTA concentration in the final dialysis step is to avoid interference of EDTA when Mg2+ is added back to the chromatin to induce folding and selfassociation.
After reconstitution, the level of nucleosome saturation is determined by two methods. The first method involves digestion of the reconstituted templates with the restriction enzyme EcoRI, which cuts at a pair of EcoRI enzyme sites in the ‘linker’ regions between where the nucleosome cores are likely to be located on the 208 bp 5s rRNA nucleosome positioning sequence [19]. Digestions are carried out by treating 0.5 lg of reconstituted template (10 ll) with 20 U EcoRI in NEB buffer 2 in a total volume of 20 ll for 1 h at 37 C. After digestion, the digestions are separated on 0.7% agarose nucleoprotein gels and the saturation levels determined by quantitative comparison of the mononucleosome and free DNA bands [23]. Previous work by the Hansen lab has shown that the digestion should yield primarily mononucleosomes for a saturated template, with about 4–5% of the template running as naked DNA when the array templates are properly saturated with nucleosomes (Fig. 3a). A second method to check the level of nucleosome saturation is based on the propensity of nucleosome arrays to self-associate in the presence of divalent cations and the fact that arrays containing different levels of saturation tend to self-associate at different concentrations of divalent cations [5]. We therefore test all reconstituted arrays for the extent of self-association in increasing concentrations of MgCl2 to determine the level of saturation as compared to literature values [5]. We find that the arrays undergo about 40–80% self-association in 3–4 mM, MgCl2, respectively, when fully saturated. Arrays should be tested by both EcoRI digestion and self-association assay before being used for crosslinking studies (Fig. 3b and c). 2.2. Detection of inter-array crosslinking As mentioned above, our basic experimental design allows detection of inter-array interactions between the H3 tail domain on the 12-mer array and the DNA of the 35-mer array. Thus we mix 12-mer and 35-mer arrays in the presence of MgCl2 to induce inter-fiber self association of the two arrays, followed by UV light-induced crosslinking and detection of crosslinks by SDS–agarose gel electrophoresis, EtBr staining and autoradiography. Since 3H-labeled H3 is the only source of radioactive signal in the system and is only reconstituted into 12-mer array, any detectable 3 H signal co-migrating with the 35-mer DNA template likely results from inter-array H3 tail–DNA interactions. The inter-array crosslinking experiment is carried out as described. All steps are carried out at room temperature: 1. Mix 8.1 ll of unlabeled 35-mer array (0.2 lg/ll) with 8.1 ll of 3H-labeled 12-mer array (0.04 lg/ll) in a siliconized microfuge tube (see Note 4). 2. Add 1.8 ll of 10 mM Tris containing 0–80 mM MgCl2 into the sample, mix immediately and rapidly by pippeting to such that the final MgCl2 concentration is 0–8 mM.
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solution and help convert array oligomers into monomers. Two microliters of 10· gel loading dye (0.1% SDS and 0.02% EtBr) is then added into each sample and the samples incubated for 10 min at 37 C to dissociate non-covalently bound proteins from the DNA templates. Samples are then analyzed by 0.7% agarose gel electrophoresis. Both the agarose gel and electrophoresis buffer (0.5· TBE) contain the same concentrations of SDS and EtBr as samples (0.01% SDS, 0.2 lg/ml EtBr) to maximize protein removal from DNA. Consequently only covalently crosslinked proteins co-migrate with the DNA templates. After 1 h of gel electrophoresis at 120 V, the DNA templates of 12-mer and 35-mer arrays are well separated on the gel (Fig. 4a). The location of the EtBr-stained DNA templates on the gel is recorded by digital photography under UV light. The gel is then soaked in standard SDS– PAGE destain buffer I (45% methanol, 10 % acetic acid) for 15 min, washed with ddH2O, and soaked in 1 M sodium salicylate for another 15 min to provide an enhancer for the detection of 3H-labeled species [24]. The gel is then dried and exposed to autoradiographic film (Kodak BioMax XAR) at 80 C for 1–2 weeks (see Note 5). 2.4. Analysis of the results Bands corresponding to the 12-mer and 35-mer templates are apparent in the ethidium bromide-stained gel (Fig. 4a).
Fig. 3. Determination of the level of nucleosome saturation. (a) Effect of saturation on EcoRI digestion. Samples of 12-mer DNA templates reconstituted with about 12, 10, and 8 nucleosomes per template were treated with EcoRI and digestion products separated on a native 0.7% agarose nucleoprotein gel. Lane 1 shows the result expected for full saturation of the template, with about 5% of the monomer DNA appearing as free DNA. (b) MgCl2-dependent self-association assay. The same arrays analyzed in (a) were incubated with increasing concentrations of MgCl2 then centrifuged for 15 min at maximum speed in a benchtop microfuge. The supernatants were loaded onto the SDS–agarose gel and the gel briefly electrophoresed. The saturated, unsaturated and very unsaturated arrays are shown top to bottom in the gel, respectively. (c) The fraction of arrays in the supernatant was quantified from the gel in (b) and plotted vs MgCl2 concentration.
3. Transfer the sample into a Falcon 5-ml polystyrene tube, and then place the tube into a 15-ml Pyrex 9820 glass tube. This two-layer protection excludes shorter wavelength UV light that may be directly absorbed and damage the DNA templates. 4. Samples are then exposed to 365 nm UV light emitted from a standard VMR LM-20E light box for 1 min to trigger the crosslinking reaction [20]. 2.3. Analysis of crosslinking by agarose gel electrophoresis/ autoradiography After UV irradiation, 0.5 ll of 100 mM EDTA is immediately added into each sample as to chelate Mg2+ from the
Fig. 4. Inter-array crosslinking between H3 tail domains and the 35-mer array. (a) Inter-array crosslinking is dependent upon MgCl2 and UV irradiation. A sample containing the 12-mer and 35-mer arrays was prepared as described in the text, irradiated with UV light and products separated on an SDS–agarose gel. The concentration of MgCl2 in each sample is indicated above the gels. (a) EtBr staining. (b) Autoradiograph. A band corresponding to crosslinking of radiolabeled H3 to the 35-mer template is indicated as an inter-array crosslinking while crosslinking to the 12-mer array is indicated as intra-array crosslinking. The location of the two templates in the top gel and free H3 in the bottom gel is indicated. Note that the sample in lane 1 was not subjected to irradiation, as indicated.
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The autoradiograph of the gels reveals several bands. In the absence of irradiation, free H3 migrates as a diffuse band, below the 12-mer template (Fig. 4b). When samples are irradiated in the absence of MgCl2, some amount of radioactive H3 co-migrates with the 12-mer template as expected since the nucleosomes associated with this template contain the labeled, APB-modified protein. This crosslinking results in a small shift in the position of the 12-mer template on the ethidium-stained gel (Fig. 4a, compare lanes 1 and 2). Note that no crosslinking is detected to the 35-mer template in the absence of MgCl2. However, in the presence of MgCl2, some radioactivity now co-migrates with the 35-mer template, indicating that some inter-array crosslinking has taken place. The gel can be scanned on a standard densitometer and the fraction of the total crosslinked protein that co-migrates with the 35-mer template determined (not shown). About 10% of the crosslinked H3 co-migrated with the 35mer array template when the crosslinking was carried out in 4 mM MgCl2 (Fig. 4). According to the MgCl2 self-association assay, about 80% of 12-mer array and about 60% of 35-mer array are assembled into large, self-associated oligomers in the presence of 4 mM MgCl2 (Fig. 3). Self-association increases to 100% of both templates at higher MgCl2 concentrations and may involve assembly into even large structures [5]. Accordingly, inter-array crosslinks formed by the H3 tail constitute 20% of total histone–DNA crosslinks in 6–8 mM MgCl2 (Fig. 4). 3. Summary Our method allows probable inter-array interactions mediated by a histone tail domain to be detected via crosslinking. These interactions are thought to be important for formation of higher-order tertiary chromatin structures modeled by the MgCl2-dependent oligomerization of model nucleosome arrays [5,11]. The excess 35-mer serves as a sink for presumably the majority of such interactions by selected tail domains located on the 12-mer array. We estimate that our method can detect inter-array crosslinks if they represent minimally 1–2% of total crosslinks. Previously, we determined that the H3 tail domain participates in primarily inter-nucleosomal interactions when the array is condensed in MgCl2 [16,17]. Using our new method, we find that about 20% of these inter-nucleosome interactions occur between arrays and about 80% are intra-array. Thus the H3 tail domain contributes to both folding of the oligonucleosome array and to array oligomerization. Our inter-array crosslinking experiment is easily extended to examine contacts made by other positions in the H3 tail and other tail domains by moving the location of the APB-derived crosslinker to other positions of H3 tail or other histone tail domains. In addition, the effect of posttranslational modifications and the binding of linker histones on these interactions can be examined. For example, lysine fi glutamine substitutions can be used to approximate lysine acetylation or such modifications can be introduced chemically [25] or enzymatically.
4. Notes Note 1: In order to obtain the highest efficiency of 3H incorporation in the expressed protein, we have found that the timing for the addition of the 3H-lysine is critical. Thus, in trial runs, the expression of the target protein expression is carefully monitored after IPTG addition, and 3H-lysine is added just before H3T6C expression is apparent on by SDS–PAGE of whole cells. Typically for H3 expression the optimal addition is 20 min after IPTG addition. Note 2: Retain the supernatant after spinning down cells expressing radiolabeled protein and dispose of properly in the radioactive waste. Note 3: In practice the actual ratio of histones–DNA template must be empirically determined to avoid underor over-saturating the templates with histones. Thus, several trial reconstitutions are typically set up with histone:DNA ratios covering the range 0.9:1 to 1.5:1 were examined. The level of saturation is determined as described. Note 4: An excess of the unlabeled 35-mer array is used to maximize the chance that inter-array interactions by the radiolabled and APB-modified histone occur to this template since interarray interactions between 12-mer templates will not be detected in our experiment. Note 5: Make sure that the film is pressed tightly up against the gel during the exposure. We use paper cassettes and place a weight on top of the cassette to ensure that a tight fit. Also, given the very long half-life of 3H, exposure times may be adjusted over a wide range of times depending on the specific activity of the protein. Acknowledgments We thank Drs. Jeffrey Hansen and Xu Lu for kindly providing the 35-mer nucleosome array. This work was supported by NIH Grant GM52426. References [1] J.C. Hansen, Annu. Rev. Biophys. Biomol. Struct. 31 (2002) 361–392. [2] C.L. Woodcock, S. Dimitrov, Curr. Opin. Genet. Dev. 11 (2001) 130–135. [3] M. Garcia-Ramirez, F. Dong, J. Ausio, J. Biol. Chem. 267 (1992) 19587–19595. [4] J.C. Hansen, J. Ausio, V.H. Stanik, K.E. van Holde, Biochemistry 28 (1989) 9129–9136. [5] P.M. Schwarz, A. Felthauser, T.M. Fletcher, J.C. Hansen, Biochemistry 35 (1996) 4009–4015. [6] J. Allan, N. Harborne, D.C. Rau, H. Gould, J. Cell Biol. 93 (1982) 285–297. [7] T.M. Fletcher, J.C. Hansen, J. Biol. Chem. 270 (1995) 25359–25362. [8] C. Tse, J.C. Hansen, Biochemistry 36 (1997) 11381–11388. [9] K. Luger, A.W. Mader, R.K. Richmond, D.F. Sargent, T.J. Richmond, Nature 389 (1997) 251–260. [10] T. Schalch, S. Duda, D.F. Sargent, T.J. Richmond, Nature 436 (2005) 138–141. [11] F. Gordon, K. Luger, J.C. Hansen, J. Biol. Chem. 280 (2005) 33701–33706.
P.-Y. Kan, J.J. Hayes / Methods 41 (2007) 278–285 [12] D. Angelov, V. Stefanovsky, S.I. Dimitrov, V.R. Russanova, E. Keskinova, I.G. Pashev, Nucleic Acids Res. 16 (1988) 4525–4538. [13] D. Angelov, J.M. Vitolo, V. Mutskov, S. Dimitrov, J.J. Hayes, Proc. Natl. Acad. Sci. USA 98 (2001) 6599–6604. [14] S.I. Usachenko, S.G. Bavykin, I.M. Gavin, E.M. Bradbury, Proc. Natl. Acad. Sci. USA 91 (1994) 6845–6849. [15] K.M. Lee, J.J. Hayes, Biochemistry 37 (1998) 8622–8628. [16] C. Zheng, J.J. Hayes, J. Biol. Chem. 278 (2003) 24217–24224. [17] C. Zheng, X. Lu, J.C. Hansen, J.J. Hayes, J. Biol. Chem. 280 (2005) 33552–33557.
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