The author thanks Dr. Philipp Selenko (FMP-Berlin) for providing wet-lab space and infrastructure to perform experiments and Deutsche Forschungsgemeinschaft (DFG) for funding the work through a research grant (LI 2402/2-1).

1. Reconstitution of Nucleosomes with Differentially Isotope-labeled (and Asymmetrically Modified) Sister Histones
NOTE: The current protocol describes the reconstitution of nucleosomes with differentially isotope-labeled histone H3. To this end, two pools of histone H3 were used; one was 15N-labeled and contained a 6xHis-tag at the N-terminus and the second was 13C-labeled and contained the Strep peptide (WSHPQFEK) fused at the N-terminus. Both tags were separated from the native H3 sequence with a Tobacco Etch Virus (TEV) protease recognition site. To prepare additional asymmetrically modified nucleosomes, one of the two H3 pools is included in a premodified form. Premodification of a histone pool can be performed by reacting the isotope-labeled histone of interest with the respective histone-modifying enzyme in the presence of the necessary for each type of reaction cofactors/PTM donors16. The efficient placement of the respective PTM can be assessed by NMR spectroscopy or mass spectrometry. The amounts of histones/DNA used here are rough recommendations to obtain a nucleosome preparation with an approximate concentration of 10 &#181;M (measured as DNA concentration at 260 nm).This final yield is sufficient to record good quality NMR spectra with relatively short acquisition times.
<ol>
	<li>Reconstitution of a histone octamer with differentially isotope-labeled (and asymmetrically modified) histone H3 
	NOTE: Recombinant histone proteins can be produced in mg quantities in BL21(DE3)plysS cells14. To obtain isotope-labeled histones, the same expression/purification protocol is followed with the exception of using for bacterial growth a minimal medium containing 15NH4Cl and/or 13C-Glucose as nitrogen and carbon sources, for 15N- or 13C-labeling respectively. Purified histones are extensively dialyzed against 5 mM &#946;-mercaptoethanol and stored lyophilized before use.
	<ol>
		<li>Dissolve lyophilized histone aliquots, containing ~5 mg of each histone in 1 mL of unfolding buffer (7 M Guanidinium-HCl, 20 mM Tris pH 7.5, 10 mM DTT). Include both types of histone H3. Mix by pipetting and do not vortex. Keep the tubes on ice for 30 min to allow complete unfolding.</li>
		<li>Determine the exact concentration of each histone by measuring the absorbance at 280 nm against unfolding buffer and using extinction coefficients calculated with the ProtParam tool of the ExPASy portal.15</li>
		<li>Mix the core histones to equimolar ratios, keeping in mind to include 50% each of the two histone H3 pools. Start by using the whole content&#160;of the less concentrated histone and add all of the others accordingly.</li>
		<li>Adjust final protein concentration to approximately 1 mg/mL using unfolding buffer.</li>
		<li>Transfer the mixture to a 6 kDa-cutoff dialysis membrane and dialyze against 1-2 L of chilled refolding buffer (10 mM Tris pH 7.5, 2 M NaCl, 1 mM EDTA, 2 mM DTT) at 4 &#176;C. Change the buffer at least once after dialysis has proceeded for a minimum of 3 h. Perform a final dialysis O/N at 4 &#176;C.</li>
		<li>Collect dialyzed material and remove any precipitates by centrifugation in a microfuge for 5 min at 4 &#176;C on maximum speed (normally no precipitation should be observed). Determine octamer concentration by measuring the absorbance at 280 nm. In the calculations, use the added extinction coefficients of histones, keeping in mind to double each value.</li>
		<li>Concentrate to a final volume of ~2 mL using a 10 kDa-cutoff centrifugal filter unit. Recalculate octamer concentration and estimate loss. At this point, an octamer concentration between 50 - 100 &#181;M should be obtained.</li>
		<li>Connect the gel filtration column (stated in the Table of Materials) to an FPLC system and equilibrate with 2 column volumes (CV) of 0.22 &#181;m-filtered and degassed refolding buffer at a flow rate of 1 mL/min and a pressure limit of 0.5 MPa. 
		NOTE: The gel filtration run should be performed in the cold room or in a cold cabinet.</li>
		<li>Inject concentrated material using a flow rate of 1 mL/min and collect 1.0 - 1.5 mL fractions.</li>
		<li>Follow the chromatographic profile and observe histone octamer elution at ~65 - 68 mL. 
		NOTE: A small shoulder of the elution peak and a peak at ~80 mL indicates the existence of free H3-H4 tetramers and H2A-H2B dimers, respectively.</li>
		<li>Run 20 &#181;L of each collected fraction in an 18% SDS-PAGE gel. 
		NOTE: Dilute the samples by a factor of at least 2 with water before loading them on the gel to reduce the high salt concentration and thus, avoid distortions. The same applies for subsequent SDS-PAGE analyses of samples in refolding buffer.</li>
		<li>Pool relevant fractions containing pure octamers judged by the equal distribution of the 4 histones on the stained gel and determine concentration as before (step 1.1.7).</li>
		<li>Connect a 1 mL Ni-NTA column (Table of Materials) to an FPLC system and equilibrate with 10 CV of 0.22 &#181;m-filtered and degassed refolding buffer at a flow rate of 1 mL/min. 
		NOTE: The Ni-NTA chromatography should be performed in the cold room or in a cold cabinet.</li>
		<li>Pass purified octamer through the Ni-NTA using a flow rate of 1 mL/min.</li>
		<li>Collect flow-through and keep samples for SDS-PAGE/WB analysis (20 &#181;L and 4 &#181;L, respectively). Subsequently, wash column with 10 CV of refolding buffer or until a baseline signal is reached.</li>
		<li>Elute with 10 CV of refolding buffer containing 250 mM imidazole using a flow rate of 1 mL/min. Keep samples for SDS-PAGE/WB analysis (20 &#181;L and 4 &#181;L, respectively).</li>
		<li>Equilibrate a 1 mL affinity column of a commercial streptavidin (Table of Materials) with 10 CV refolding buffer containing 250 mM imidazole using a batch/gravity flow setup. 
		NOTE: This chromatographic step should be performed in the cold room.</li>
		<li>Pass Ni-NTA elution through the commercial streptavidin affinity column.</li>
		<li>Collect flow through and keep samples for SDS-PAGE/WB analysis (20 &#181;L and 4 &#181;L, respectively). Subsequently, wash with 10 CV of refolding buffer.</li>
		<li>Elute with 10 CV of refolding buffer containing 2.5 mM desthiobiotin. Keep samples for SDS-PAGE/WB analysis (20 and 4 &#181;L respectively).</li>
		<li>Measure octamer concentration, add 10 times less TEV protease and let reaction proceed overnight at 4 &#176;C in a standard plastic centrifuge tube. 
		NOTE: The necessary amount of TEV to obtain complete digestion (removal of the affinity tags) depends on the origin (construct) and on the activity (freshly made, recombinant TEV is more active compared to the one stored for prolonged periods at - 80 &#176;C). Try initially adding 10 times less TEV compared to the octamer (estimated as protein concentrations) and adjust accordingly.</li>
		<li>Check for the efficient digestion by running 20 &#181;L of the mixture prior and after TEV addition on an 18% SDS-PAGE gel. If digestion is not complete, add more TEV protease and continue incubation for another 3-4 h.</li>
		<li>Dialyze sample against refolding buffer using a 50 kDa-cutoff dialysis membrane to remove TEV protease and adjust buffer composition.</li>
		<li>Concentrate the purified asymmetric octamer using&#160;a 10 kDa-cutoff centrifugal filter unit (the same filter unit from step 1.1.7 can be also used) to a volume of 1.0-2.0 mL. Measure the final concentration. Either use soon after for nucleosome reconstitution or adjust to 50% (v/v) glycerol to store at -20 &#176;C for longer periods. At this point and expecting at least 70% loss during the previous steps, the octamer concentration should be in the range of 20 - 50 &#181;M.</li>
	</ol>
	</li>
	<li>Nucleosome reconstitution
	<ol>
		<li>If octamer was stored in 50% glycerol, dialyze overnight at 4 &#176;C against refolding buffer before proceeding with nucleosome reconstitution.</li>
		<li>Adjust DNA (containing a nucleosome-positioning sequence) salt concentration to 2 M NaCl by using a 5 M NaCl stock solution. 
		NOTE: The purified DNA either isolated after purification of a plasmid that contains multiple copies of the desired sequence or synthesized by PCR amplification should be stored concentrated to ~50 &#181;M in water or a standard Tris-EDTA buffer.</li>
		<li>Mix octamer and DNA using the optimum ratio determined from a small-scale test. Use refolding buffer to adjust the final volume to ~3.0 mL. Always add the octamer last to prevent any possibility of mixing the octamer and the DNA at &#60;2 M salt concentrations.
		<ol>
			<li>Perform initial small-scale reconstitutions to determine the octamer:DNA ratio that leads to optimum nucleosome yields. For this, assemble three reactions of 0.9:1.0, 1.0:1.0, and 1.1:1.0 DNA:octamer ratios in a volume between 50-100 &#181;L. Typically, use a concentration of 5 &#181;M DNA.</li>
			<li>Proceed with reconstitution as described below (steps 1.2.4-1.2.8) and at the end, estimate the amount of soluble and good quality reconstituted nucleosome by measuring the DNA concentration at 260 nm and by running a 6% native-PAGE DNA gel, stained with ethidium bromide, respectively (Figure 5A).</li>
		</ol>
		</li>
		<li>Transfer the mix to a 6 kDa-cutoff dialysis membrane and dialyze against 1 L of refolding buffer overnight at 4 &#176;C.</li>
		<li>Reduce the salt concentration of the dialysis buffer in a step-wise manner from 2, to 0.85, 0.64, and 0.2 M NaCl respectively. Keep sample in each buffer for at least 3 h. Sometimes precipitate is formed after the last transition. In this case continue dialysis as planned and remove precipitate by microfuge centrifugation at the end of the next step.</li>
		<li>Transfer dialysis membrane in a beaker with 1 L of assay buffer (25 mM NaH2PO4/Na2HPO4 pH 6.8, 25 mM NaCl, 2 mM DTT) and continue dialysis O/N at 4 &#176;C.</li>
		<li>Collect sample, remove any precipitate formed at step 1.2.5 by centrifugation in a microfuge for 5 min at 4 &#176;C on maximum speed and concentrate using a 30 kDa-cutoff centrifugal filter unit to a final volume of ~300 &#181;L.</li>
		<li>Measure DNA concentration at 260 nm, run an 18% SDS-PAGE protein gel (4 &#181;L of sample) and a 6% native-PAGE DNA gel (0.2 &#181;L of sample) and store sample at 4 &#176;C for several weeks. The final concentration should be in the range of 10-20 &#181;M.</li>
	</ol>
	</li>
</ol>
2. NMR Analysis of Modification Reactions on the Two Sister Histones
NOTE: Assignment of 2D 1H-15N correlation spectra of nucleosomal histone tails can be found at references16,17,18. Additionally, assignments of CHx groups of lysines, serines and threonines on 2D 1H-13C correlation spectra can be found at reference16.
<ol>
	<li>NMR setup and recording of reference spectra 
	NOTE: Enzymatic assays were performed using 140 &#181;L of the nucleosome sample in assay buffer into a 3-mm NMR tube at 300 K. Different NMR tubes with other sample volumes can be also used. The aforementioned temperature is appropriate to obtain good quality 1H-15N correlation spectra of nucleosomal histone tails and good enzymatic activities. 2D-SOFAST-HMQC 1H-15N and 2D-HSQC 1H-13C spectra were recorded respectively with a setup that confers similar acquisition times. Typical acquisition parameters are 128 transients with 1.024 (1H) x 128 (15N) complex points and 32 transients with 1.024 (1H) x 64 (13C) complex points for the two different types of correlation spectra. For both 2D spectra, the acquisition time was ~30 min.
	<ol>
		<li>Set 300 K as the sample temperature in the spectrometer. Add 10% D2O (v/v) to the sample to be used for the spectrometer frequency lock.</li>
		<li>Place the sample in the spectrometer and perform basic operations (locking, tuning, shimming). Additionally, determine optimal pulse lengths and general acquisition parameters.</li>
		<li>Record 1D and 2D 1H-15N and 1H-13C reference spectra.</li>
		<li>Process reference spectra and ensure that the signals to be used for mapping the modification reactions are well resolved and have good intensities.</li>
		<li>Recover sample and transfer it to a microfuge tube.</li>
	</ol>
	</li>
	<li>Setup of the enzymatic reaction, NMR monitoring and quantitative analysis
	<ol>
		<li>Copy and arrange a series of interleaved 2D 1H-15N and 1H-13C spectra. Aim for a total acquisition time of several hours and adjust accordingly in a new run based on the enzymatic rates obtained from the first reaction.</li>
		<li>Add to the sample the necessary cofactors for the respective type of modification reaction (1 mM ATP/2 mM MgCl2 for phosphorylation, 1 mM acetyl-CoA for acetylation, 1 mM SAM for methylation, etc.).</li>
		<li>Add the enzyme of interest, mix by pipetting, transfer the sample back in the NMR tube and subsequently in the spectrometer (perform these operations quickly). 
		NOTE: If there is no data about the expected enzymatic activity, adjust the substrate-to-enzyme molar ratio to 10:1. Perform a first trial run and adjust accordingly for the real run.</li>
		<li>Perform a fast automatic shimming and use the rest of parameters from the reference spectra.</li>
		<li>Initiate the series of the interleaved 2D 1H-15N and 1H-13C spectra.</li>
		<li>Follow the progression of the reaction in real-time and stop acquisition when the reaction reaches completion or a desired level.</li>
		<li>Process spectra as before with exactly the same parameters.</li>
		<li>Repeat the whole run using a different acquisition order for the two types of correlation spectra. If in the first experiment a 1H-15N correlation was first in the series, start now with a 1H-13C correlation. By this and having same acquisition time for both spectra, it is possible to plot and compare PTM reactions on both differentially isotope-labeled histones on the same time scale. Additionally, it is possible to calculate averages and plot error bars. 
		NOTE: A thorough description of the methodologies for NMR signal analysis and subsequent quantification of enzymatic reactions can be found here19.</li>
		<li>Select NMR signals from each time-course NMR spectrum that report the progression of the reaction and calculate their relative signal intensities using a visualization and analysis software for NMR spectra. Do this for signals that report the unmodified as well as the modified state. Extract modification levels.</li>
		<li>Plot the calculated values of modification levels against the reaction time.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Hansen, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Conformational+Dynamics+of+the+Chromatin+Fiber+in+Solution:+Determinants,+Mechanisms,+and+Functions.">Conformational Dynamics of the Chromatin Fiber in Solution: Determinants, Mechanisms, and Functions.</a> Annu. Rev. Biophys. Biomol. Struct. 31, 361-392 (2002).</li><li>Clapier, C. R., Cairns, B. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Biology+of+Chromatin+Remodeling+Complexes.">The Biology of Chromatin Remodeling Complexes.</a> Annu. Rev. Biochem. 78, 273-304 (2009).</li><li>Musselman, C. A., Lalonde, M., Cote, J., Kutateladze, T. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perceiving+the+epigenetic+landscape+through+histone+readers.">Perceiving the epigenetic landscape through histone readers.</a> Nat. Struct. Mol. Biol. 19 (12), 1218-1227 (2012).</li><li>Suganuma, T., Workman, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Signals+and+Combinatorial+Functions+of+Histone+Modifications.">Signals and Combinatorial Functions of Histone Modifications.</a> Annu. Rev. Biochem. 80, 473-499 (2011).</li><li>Lee, J. S., Smith, E., Shilatifard, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Language+of+Histone+Crosstalk.">The Language of Histone Crosstalk.</a> Cell. 142 (5), 682-685 (2010).</li><li>Liokatis, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phosphorylation+of+histone+H3+Ser10+establishes+a+hierarchy+for+subsequent+intramolecular+modification+events.">Phosphorylation of histone H3 Ser10 establishes a hierarchy for subsequent intramolecular modification events.</a> Nat. Struct. Mol. Biol. 19 (8), 819-823 (2012).</li><li>Voigt, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Asymmetrically+Modified+Nucleosomes.">Asymmetrically Modified Nucleosomes.</a> Cell. 151 (1), 181-193 (2012).</li><li>Egelhofer, T. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+assessment+of+histone-modification+antibody+quality.">An assessment of histone-modification antibody quality.</a> Nat. Struct. Mol. Biol. 18 (1), 91-93 (2011).</li><li>Huang, H., Lin, S., Garcia, B. A., Zhao, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+Proteomic+Analysis+of+Histone+Modifications.">Quantitative Proteomic Analysis of Histone Modifications.</a> Chem. Rev. 115 (6), 2376-2418 (2015).</li><li>Liokatis, S., Dose, A., Schwarzer, D., Selenko, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+Detection,+of+Protein+Phosphorylation+and+Acetylation+by+High-Resolution+NMR+Spectroscopy.">Simultaneous Detection, of Protein Phosphorylation and Acetylation by High-Resolution NMR Spectroscopy.</a> J. Am. Chem. Soc. 132 (42), 14704-14705 (2010).</li><li>Binolfi, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracellular+repair+of+oxidation-damaged+&#945;-synuclein+fails+to+target+C-terminal+modification+sites.">Intracellular repair of oxidation-damaged &#945;-synuclein fails to target C-terminal modification sites.</a> Nat. Commun. 7, 10251 (2016).</li><li>Schanda, P., Kupce, E., Brutscher, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SOFAST-HMQC+experiments+for+recording+two-dimensional+heteronuclear+correlation+spectra+of+proteins+within+a+few+seconds.">SOFAST-HMQC experiments for recording two-dimensional heteronuclear correlation spectra of proteins within a few seconds.</a> J. Biomol. NMR. 33 (4), 199-211 (2005).</li><li>Theillet, F. X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+signaling,+post-translational+protein+modifications+and+NMR+spectroscopy.">Cell signaling, post-translational protein modifications and NMR spectroscopy.</a> J. Biomol. NMR. 54 (3), 217-236 (2012).</li><li>Dyer, P. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reconstitution+of+Nucleosome+Core+Particles+from+Recombinant+Histones+and+DNA.">Reconstitution of Nucleosome Core Particles from Recombinant Histones and DNA.</a> Methods Enzymol. 375, 23-43 (2004).</li><li>Artimo, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ExPASy+:SIB+bioinformatics+resource+portal.">ExPASy :SIB bioinformatics resource portal.</a> Nucleic Acids Res. 40 (W1), 597-603 (2012).</li><li>Liokatis, S., klingberg, R., Tan, S., Schwarzer, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differentially+Isotope-Labeled+Nucleosomes+to+Study+Asymmetric+Histone+Modification+Crosstalk+by+Time-Resolved+NMR+Spectroscopy.">Differentially Isotope-Labeled Nucleosomes to Study Asymmetric Histone Modification Crosstalk by Time-Resolved NMR Spectroscopy.</a> Angew. Chem. Int. Ed. 55 (29), 8262-8265 (2016).</li><li>St&#252;tzer, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulations+of+DNA+Contacts+by+Linker+Histones+and+Post-translational+Modifications+Determine+the+Mobility+and+Modifiability+of+Nucleosomal+H3+Tails.">Modulations of DNA Contacts by Linker Histones and Post-translational Modifications Determine the Mobility and Modifiability of Nucleosomal H3 Tails.</a> Mol. Cell. 61 (2), 247-259 (2016).</li><li>Zhou, B. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Histone+H4+K16Q+Mutation,+an+Acetylation+Mimic,+Causes+Structural+Disorder+of+Its+N-terminal+Basic+Patch+in+the+Nucleosome.">Histone H4 K16Q Mutation, an Acetylation Mimic, Causes Structural Disorder of Its N-terminal Basic Patch in the Nucleosome.</a> J. Mol. Biol. 421 (1), 30-37 (2012).</li><li>Theillet, F. X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Site-specific+NMR+mapping+and+time-resolved+monitoring+of+serine+and+threonine+phosphorylation+in+reconstituted+kinase+reactions+and+mammalian+cell+extracts.">Site-specific NMR mapping and time-resolved monitoring of serine and threonine phosphorylation in reconstituted kinase reactions and mammalian cell extracts.</a> Nat. Protoc. 8 (7), 1416-1432 (2013).</li><li>Lowary, P. T., Widom, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+DNA+sequence+rules+for+high+affinity+binding+to+histone+octamer+and+sequence-directed+nucleosome+positioning.">New DNA sequence rules for high affinity binding to histone octamer and sequence-directed nucleosome positioning.</a> J. Mol. Biol. 276 (1), 19-42 (1998).</li><li>Hackenberger, C. P., Schwarzer, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemoselective+ligation+and+modification+strategies+for+peptides+and+proteins.">Chemoselective ligation and modification strategies for peptides and proteins.</a> Angew. Chem. Int. Ed. 47 (52), 10030-10074 (2008).</li><li>Theillet, F. X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Site-specific+mapping+and+time-resolved+monitoring+of+lysine+methylation+by+high-resolution+NMR+spectroscopy.">Site-specific mapping and time-resolved monitoring of lysine methylation by high-resolution NMR spectroscopy.</a> J. Am. Chem. Soc. 134 (18), 7616-7619 (2012).</li><li>Lechner, C. C., Agashe, N. D., Fierz, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Traceless+Synthesis+of+Asymmetrically+Modified+Bivalent+Nucleosomes.">Traceless Synthesis of Asymmetrically Modified Bivalent Nucleosomes.</a> Angew. Chem. Int. Ed. 55 (8), 2903-2906 (2016).</li><li>Bernstein, B. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+bivalent+chromatin+structure+marks+key+developmental+genes+in+embryonic+stem+cells.">A bivalent chromatin structure marks key developmental genes in embryonic stem cells.</a> Cell. 125 (2), 315-326 (2006).</li></ol>

The author has nothing to disclose.

For nucleosome reconstitution, the current protocol utilizes a 165 bp long DNA template containing the 601-Widom nucleosome positioning sequence20, but similar performance is expected using various lengths of DNA templates. The protocol was designed and employed using asymmetric types of histone H3. With the same principle, the method can be applied also for the other core histones and additionally can be used to reconstitute complexes carrying two distinct core histones with different isotope lab...

Eukaryotic DNA is tightly packaged within cell nuclei into chromatin. The fundamental building block of chromatin is the nucleosome core particle that contains ~147 bp of DNA wrapped around an octameric complex made up of two copies each of the four core histones (H3, H4, H2A, H2B). Histone proteins harbor a plethora of Post-translational Modifications (PTMs). These covalent substitutions induce alterations in chromatin structure, both directly by affecting the physical chemistry of the system and indirectly by recruiting chromatin-remodeling activities1,2,3. By those means, histone PTMs control chromatin accessibility and, hence, regulate all DNA-based cellular functions4.
PTMs are installed by histone-modifying enzyme systems mainly on the unstructured N-terminal segments (tails) of nucleosome-incorporated core histones. Due to the many modification sites on the relatively short sequence of histone tails, PTMs influence each other by inducing or blocking subsequent modification reactions, an effect known as modification cross-talk5. Because of the overall symmetric architecture of the nucleosome, modification reactions and crosstalk mechanisms were thought to occur similarly onto the two copies of each nucleosomal histone (sister histones). This concept was recently challenged and subsequently disproved. Particularly, in vitro enzymatic assays on free histone H3 tail peptides and on nucleosomes demonstrated that a set of H3 kinases introduced phosphorylation in an asymmetric manner6. Additionally, affinity-purification-based LC-MS/MS analysis revealed the existence of asymmetrically H3-methylated nucleosomes in several types of eukaryotic cells7. Thus, asymmetrically modified nucleosomes constitute novel species, and tools are needed to uncover the mechanisms that control their formation and to analyze the crosstalk effects that this asymmetry might exert.
Commonly, Western Blotting (WB) and Mass Spectrometry (MS) analysis have been used to detect histone PTMs. Despite its easy application, WB suffers from specificity/cross-reactivity problems. On top of that, it is incapable of performing simultaneous multi-PTM analysis and direct quantification of the modification reactions8. On the other hand, MS analysis employs sophisticated instrumentation that requires high-level training, but provides high specificity as well as simultaneous mapping and quantification of multiple PTMs9.&#160;However, both methods are disruptive and the nucleosomal complexes are dissociated before analysis, giving rise to a mixture of histones and/or histone-derived peptides. This manipulation removes the ability to distinguish independent modification reactions that occur on each of the two sister histones and to report the copy-specific modification status of the nucleosomal histones.
Nuclear Magnetic Resonance (NMR) spectroscopy evolved as an alternative method to map PTM reactions. NMR is nondisruptive and thus allows the monitoring of PTM events in a real-time manner in reconstituted mixtures, and even in intact cells10,11. The development of routines for fast data acquisition as well as for high-resolution mapping based on 2D hetero-nuclear correlation methods of isotope-labeled (15N and/or 13C) samples12 allowed the simultaneous mapping of different types of PTMs, such as serine/threonine/tyrosine phosphorylation, lysine acetylation/methylation, and arginine methylation13. Depending on the PTM under investigation, 15N- or 13C-labeling protocols can be employed to mark the protein functional group that serves as a modification reporter. Consequently, PTM mapping can be performed by following the characteristic chemical shift displacement of the corresponding functional group &#39;sensing&#39; the alteration on the chemical environment. In most cases, both N-H and C-H chemical groups can be used to report the evolution of the PTM of interest.
The current protocol describes the generation of nucleosomes containing differentially isotope-labeled sister histones. It combines the flexibility of NMR spectroscopy to map PTMs using both 1H-15N and 1H-13C correlation spectra with the utilization of different protein affinity tags for purification of the selected reconstituted histone complexes. Notably, the protocol employs two different pools of a particular histone for nucleosome reconstitution. These pools are differentially isotope-labeled (one with 15N, the other with 13C), and they are fused to a polyhistidine and a streptavidin affinity tag, respectively. A tandem affinity purification scheme with Ni-NTA and streptavidin-based chromatography initially used by Voigt et al.7&#160;is&#160;employed to purify asymmetric species from symmetric counterparts (Figure 1A). Asymmetric histone octamers are used subsequently to reconstitute equivalent nucleosomal complexes (Figure 1B), using the standard salt dialysis method14. Additionally, through the same procedure and by having one of the histone pools pre-modified, a PTM can be incorporated asymmetrically onto the resulting nucleosomes. The reaction of these substrates with histone-modifying enzymes and subsequent NMR-mapping of modification events enable the characterization of crosstalk mechanisms both in-cis (premodified histone copy) and in-trans (unmodified histone copy) (Figure 1C).

<table><tbody><tr><td>Guanidinium HCl</td><td>Applichem</td><td>A14199</td><td>Use high quality Gu-HCl</td></tr><tr><td>Tris&amp;nbsp;</td><td>Roth</td><td>4855.2</td><td></td></tr><tr><td>DTT</td><td>Applichem</td><td>A1101</td><td></td></tr><tr><td>NaCl</td><td>VWR chemicals</td><td>27810.364</td><td></td></tr><tr><td>EDTA</td><td>Roth</td><td>8043.2</td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;H&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Applichem</td><td>A1046</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Applichem</td><td>A3902</td><td></td></tr><tr><td>Imidazole</td><td>Applichem</td><td>A1073</td><td></td></tr><tr><td>d-Desthiobiotin</td><td>Sigma</td><td>D1411</td><td></td></tr><tr><td>Ni-NTA Superflow</td><td>Qiagen</td><td>1034557</td><td></td></tr><tr><td>Strep-Tactin Superflow</td><td>IBA</td><td>2-1207-001</td><td></td></tr><tr><td>His-probe antibody</td><td>Santa Cruz</td><td>sc-8036</td><td></td></tr><tr><td>Strep-tactin conjugated HRP</td><td>IBA</td><td>2-1502-001</td><td></td></tr><tr><td>Hi-Load 16/600 Superdex 200 pg</td><td>GE Healthcare</td><td>28-9893-35</td><td></td></tr><tr><td>6 - 8 kDa dialysis membrane</td><td>Spectrumlabs</td><td>spectra/por 1, 132650</td><td></td></tr><tr><td>50 kDa dialysis membrane</td><td>Spectrumlabs</td><td>spectra/por 7, 132129</td><td></td></tr><tr><td>10 kDa centrigugal filter unit</td><td>Merck Millipore</td><td>UFC901024</td><td></td></tr><tr><td>30 kDa centrigugal filter unit</td><td>Merck Millipore</td><td>UFC903024</td><td></td></tr><tr><td>Solution-state NMR spectrometer&amp;nbsp;</td><td></td><td></td><td>at least 500 MHz operating frequency, equipped with a triple-resonance cryoprobe</td></tr><tr><td>&lt;strong&gt;Buffer name&lt;/strong&gt;</td><td>&lt;strong&gt;Component&lt;/strong&gt;</td><td></td><td></td></tr><tr><td>Unfolding Buffer</td><td>7 M Guanidinium HCl</td><td></td><td></td></tr><tr><td></td><td>20 mM Tris, pH 7.5</td><td></td><td></td></tr><tr><td></td><td>10 mM DTT</td><td></td><td></td></tr><tr><td>Refolding Buffer</td><td>10 mM Tris, pH 7.5</td><td></td><td></td></tr><tr><td></td><td>2 M NaCl</td><td></td><td></td></tr><tr><td></td><td>1 mM EDTA</td><td></td><td></td></tr><tr><td></td><td>2 mM DTT</td><td></td><td></td></tr><tr><td>Assay Buffer</td><td>25 mM Na&lt;sub&gt;2&lt;/sub&gt;H&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, pH 6.8</td><td></td><td></td></tr><tr><td></td><td>25 mM NaCl</td><td></td><td></td></tr><tr><td></td><td>2 mM DTT</td><td></td><td></td></tr></tbody></table>

reconstitution of nucleosomes with differentially isotope-labeled sister histones

Asymmetrically modified nucleosomes contain the two copies of a histone (sister histones) decorated with distinct sets of Post-translational Modifications (PTMs). They are newly identified species with unknown means of establishment and functional implications. Current analytical methods are inadequate to detect the copy-specific occurrence of PTMs on the nucleosomal sister histones. This protocol presents a biochemical method for the in vitro reconstitution of nucleosomes containing differentially isotope-labeled sister histones. The generated complex can be also asymmetrically modified, after including a premodified histone pool during refolding of histone subcomplexes. These asymmetric nucleosome preparations can be readily reacted with histone-modifying enzymes to study modification cross-talk mechanisms imposed by the asymmetrically pre-incorporated PTM using nuclear magnetic resonance (NMR) spectroscopy. Particularly, the modification reactions in real-time can be mapped independently on the two sister histones by performing different types of NMR correlation experiments, tailored for the respective isotope type. This methodology provides the means to study crosstalk mechanisms that contribute to the formation and propagation of asymmetric PTM patterns on nucleosomal complexes.

Properly refolded octameric species are isolated after a run of the reconstitution mixture through a gel filtration column (Figure 2). The reconstituted octameric pool containing the three different types of octamers is subjected to the tandem affinity purification scheme. Samples are collected from all the steps and analyzed by SDS-PAGE and subsequently WB. Verification of the correct execution of the protocol that results in the isolation of asymmetric species is achiev...

Watch this Scientific Journal Video about Reconstitution of Nucleosomes with Differentially Isotope-labeled Sister Histones at JoVE.com

Reconstitution of Nucleosomes with Differentially Isotope-labeled Sister Histones

This protocol describes the reconstitution of nucleosomes containing differentially isotope-labeled sister histones. At the same time, asymmetrically post-translationally modified nucleosomes can be generated after using a premodified histone copy. These preparations can be readily used to study modification crosstalk mechanisms, simultaneously on both sister histones, by using high-resolution NMR spectroscopy.

Asymmetrically modified nucleosomes contain the two copies of a histone (sister histones) decorated with distinct sets of Post-translational Modifications ...

reconstitution-nucleosomes-with-differentially-isotope-labeled-sister

Research

JoVE Journal

Biochemistry

10.9K Views.  Leibniz Institute of Molecular Pharmacology (FMP-Berlin).  Dit protocol beschrijft de reconstructie van nucleosomen die differentieel-isotoop gemerkte zus histonen. Tegelijkertijd asymmetrisch post-translationeel gemodificeerde nucleosomen worden gegenereerd na het gebruik van een histon premodified kopie. Deze preparaten kunnen gemakkelijk worden gebruikt om wijzigingen crosstalk mechanismen bestuderen, gelijktijdig op beide zus histonen, met behulp van hoge-resolutie NMR-spectroscopie. 

Video: Reconstitution of Nucleosomes with Differentially Isotope-labeled Sister Histones

Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription to take place in eukaryotic cells. The dynamics of nucleosomes provides access to the DNA by replication and transcription machineries, and critically contributes to the molecular mechanisms underlying chromatin functions. Single-molecule studies such as atomic force microscopy (AFM) imaging have contributed significantly to our current understanding of the role of nucleosome structure and dynamics. The current protocol describes the steps enabling high-resolution AFM imaging techniques to study the structural and dynamic properties of nucleosomes. The protocol is illustrated by AFM data obtained for the centromere nucleosomes in which H3 histone is replaced with its counterpart centromere protein A (CENP-A). The protocol starts with the assembly of mono-nucleosomes using a continuous dilution method. The preparation of the mica substrate functionalized with aminopropyl silatrane (APS-mica) that is used for the nucleosome imaging is critical for the AFM visualization of nucleosomes described and the procedure to prepare the substrate is provided. Nucleosomes deposited on the APS-mica surface are first imaged using static AFM, which captures a snapshot of the nucleosome population. From analyses of these images, such parameters as the size of DNA wrapped around the nucleosomes can be measured and this process is also detailed. The time-lapse AFM imaging procedure in the liquid is described for the high-speed time-lapse AFM that can capture several frames of nucleosome dynamics per second. Finally, the analysis of nucleosome dynamics enabling the quantitative characterization of the dynamic processes is described and illustrated.

Here, we present a protocol to characterize nucleosome particles at the single-molecule level using static and time-lapse atomic force microscopy (AFM) imaging techniques. The surface functionalization method described allows for the capture of the structure and dynamics of nucleosomes in high-resolution at the nanoscale.

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription ...

In Vitro Ubiquitination and Deubiquitination Assays of Nucleosomal Histones

Ubiquitination is a post-translational modification that plays important roles in various signaling pathways and is notably involved in the coordination of chromatin function and DNA-associated processes. This modification involves a sequential action of several enzymes including E1 ubiquitin-activating, E2 ubiquitin-conjugating and E3 ubiquitin-ligase and is reversed by deubiquitinases (DUBs). Ubiquitination induces degradation of proteins or alteration of protein function including modulation of enzymatic activity, protein-protein interaction and subcellular localization. A critical step in demonstrating protein ubiquitination or deubiquitination is to perform in vitro reactions with purified components. Effective ubiquitination and deubiquitination reactions could be greatly impacted by the different components used, enzyme co-factors, buffer conditions, and the nature of the substrate.&#160; Here, we provide step-by-step protocols for conducting ubiquitination and deubiquitination reactions. We illustrate these reactions using minimal components of the mouse Polycomb Repressive Complex 1 (PRC1), BMI1, and RING1B, an E3 ubiquitin ligase that monoubiquitinates histone H2A on lysine 119. Deubiquitination of nucleosomal H2A is performed using a minimal Polycomb Repressive Deubiquitinase (PR-DUB) complex formed by the human deubiquitinase BAP1 and the DEUBiquitinase ADaptor (DEUBAD) domain of its co-factor ASXL2. These ubiquitination/deubiquitination assays can be conducted in the context of either recombinant nucleosomes reconstituted with bacteria-purified proteins or native nucleosomes purified from mammalian cells. We highlight the intricacies that can have a significant impact on these reactions and we propose that the general principles of these protocols can be swiftly adapted to other E3 ubiquitin ligases and deubiquitinases.

Ubiquitination is a post-translational modification that plays important roles in cellular processes and is tightly coordinated by deubiquitination. Defects in both reactions underlie human pathologies. We provide protocols for conducting ubiquitination and deubiquitination reaction in vitro using purified components.

Ubiquitination is a post-translational modification that plays important roles in various signaling pathways and is notably involved in the coordination ...

Site Specific Lysine Acetylation of Histones for Nucleosome Reconstitution using Genetic Code Expansion in Escherichia coli

Acetylated histone proteins can be easily expressed in Escherichia coli encoding a mutant, N&#949;-acetyl-lysine (AcK)-specific Methanosarcina mazi pyrrolysine tRNA-synthetase (MmAcKRS1) and its cognate tRNA (tRNAPyl) to assemble reconstituted mononucleosomes with site specific acetylated histones. MmAcKRS1 and tRNAPyl deliver AcK at an amber mutation site in the mRNA of choice during translation in Escherichia coli. This technique has been used extensively to incorporate AcK at H3 lysine sites. Pyrrolysyl-tRNA synthetase (PylRS) can also be easily evolved to incorporate other noncanonical amino acids (ncAAs) for site specific protein modification or functionalization. Here we detail a method to incorporate AcK using the MmAcKRS1 system into histone H3 and integrate acetylated H3 proteins into reconstituted mononucleosomes. Acetylated reconstituted mononucleosomes can be used in biochemical and binding assays, structure determination, and more. Obtaining modified mononucleosomes is crucial for designing experiments related to discovering new interactions and understanding epigenetics.

Site Specific Lysine Acetylation of Histones for Nucleosome Reconstitution using Genetic Code Expansion in Escherichia coli

Here we present a method to express acetylated histone proteins using genetic code expansion and assemble reconstituted nucleosomes in vitro.

Acetylated histone proteins can be easily expressed in Escherichia coli encoding a mutant, N&#949;-acetyl-lysine (AcK)-specific Methanosarcina mazi ...

Preparation of Nucleosome Core Particles Complexed with DNA Repair Factors for Cryo-Electron Microscopy Structural Determination

DNA repair in the context of chromatin is poorly understood. Biochemical studies using nucleosome core particles, the fundamental repeating unit of chromatin, show most DNA repair enzymes remove DNA damage at reduced rates as compared to free DNA. The molecular details on how base excision repair (BER) enzymes recognize and remove DNA damage in nucleosomes have not been elucidated. However, biochemical BER data of nucleosomal substrates suggest the nucleosome presents different structural barriers dependent on the location of the DNA lesion and the enzyme. This indicates the mechanisms employed by these enzymes to remove DNA damage in free DNA may be different than those employed in nucleosomes. Given that the majority of genomic DNA is assembled into nucleosomes, structural information of these complexes is needed. To date, the scientific community lacks detailed protocols to perform technically feasible structural studies of these complexes. Here, we provide two methods to prepare a complex of two genetically fused BER enzymes (Polymerase &#946; and AP Endonuclease1) bound to a single-nucleotide gap near the entry-exit of the nucleosome for cryo-electron microscopy (cryo-EM) structural determination. Both methods of sample preparation are compatible for vitrifying quality grids via plunge freezing. This protocol can be used as a starting point to prepare other nucleosomal complexes with different BER factors, pioneer transcription factors, and chromatin-modifying enzymes.

This protocol outlines in detail the preparation of nucleosomal complexes using two methods of sample preparation for freezing TEM grids.

DNA repair in the context of chromatin is poorly understood. Biochemical studies using nucleosome core particles, the fundamental repeating unit of ...

In Situ Nucleosome Reconstitution for Single-Molecule Studies

In Situ Nucleosome Assembly for Single-Molecule Correlative Force and Fluorescence Microscopy

Nucleosomes constitute the primary unit of eukaryotic chromatin and have been the focus of numerous informative single-molecule investigations regarding their biophysical properties and interactions with chromatin-binding proteins. Nucleosome reconstitution on DNA for these studies typically involves a salt dialysis procedure that provides precise control over the placement and number of nucleosomes formed along a DNA tether. However, this protocol is time-consuming and requires a substantial amount of DNA and histone octamers as inputs. To offer an alternative strategy, an in situ nucleosome reconstitution method for single-molecule force and fluorescence microscopy that utilizes the histone chaperone Nap1 is described. This method enables users to assemble nucleosomes on any DNA template without the need for strong nucleosome positioning sequences, adjust nucleosome density on demand, and use fewer reagents. In situ nucleosome formation occurs within seconds, offering a simpler experimental workflow and a convenient transition into single-molecule measurements. Examples of two downstream assays for probing nucleosome mechanics and visualizing the behavior of individual proteins on chromatin are further described.

Author Spotlight: Efficient Nucleosome Reconstitution for Single-Molecule Techniques

This article presents a detailed experimental procedure for reconstituting nucleosome-containing DNA tethers for single-molecule correlative force and fluorescence microscopy. It further describes several downstream experiments that can be conducted to visualize the binding behavior of chromatin-interacting proteins and analyze changes in the physical properties of nucleosomes.

Nucleosomes constitute the primary unit of eukaryotic chromatin and have been the focus of numerous informative single-molecule investigations regarding ...

2D Gel Separation for Histone Isoform Enrichment and PTM Characterization

Unveiling Histone Proteoforms using 2D-TAU Gel Electrophoresis

Histones undergo various post-translational modifications (PTMs) such as methylation, acetylation, phosphorylation, acylation, and ubiquitination, which control nucleosome dynamics and determine cell fate. The nucleosome, which is the functional unit of chromatin, comprises DNA, four pairs of histones (H3, H4, H2A, and H2B) making up the globular core, and the linker histone H1, which stabilizes the chromatin structure. The amino (N)-terminal tails of the histones protrude from the globular core domains and undergo distinct PTMs that influence the chromatin landscape. Some evidence suggests that histone PTM homeostasis is crucial for preserving all physiological activities. The deregulation of histone PTMs is the primary cause of abnormal cellular proliferation, invasion, and metastasis. Therefore, developing methods for characterizing histone PTMs is crucial. Here, we describe an effective technique for isolating and analyzing histone isoforms. The method, based on the combination of two orthogonal separations, allows the enrichment of histone isoforms and the following mass spectrometry identification. The technique, originally described by Shechter et al., combines acid-urea polyacrylamide gels (TAU-GEL), which can separate basic histone proteins based on size and charge, and Sodium dodecyl sulfate-polyacrylamide gels (SDS-PAGE), which can separate proteins by molecular weight. The result is a two-dimensional map of histone isoforms, suitable for in-gel digestion followed by mass spectrometry identification and western blot analysis. The result is a two-dimensional map of histone isoforms, suitable for both in-gel digestion followed by mass spectrometry identification and western blot analysis. This proteomic approach is a robust method that allows the enrichment of a single histone isoform and the characterization of new histone PTMs.

Author Spotlight: Epigenetic Modifications and Metabolic Rewiring as Targets for Cancer Therapy

This method describes a simple and fast strategy for histone proteoform characterization and investigation.

Histones undergo various post-translational modifications (PTMs) such as methylation, acetylation, phosphorylation, acylation, and ubiquitination, which ...

Expression Analysis of Mammalian Linker-histone Subtypes

Linker histone H1 binds to the nucleosome core particle and linker DNA, facilitating folding of chromatin into higher order structure. H1 is essential for mammalian development1 and regulates specific gene expression in vivo2-4. Among the highly conserved histone proteins, the family of H1 linker histones is the most heterogeneous group. There are 11 H1 subtypes in mammals that are differentially regulated during development and in different cell types. These H1 subtypes include 5 somatic H1s (H1a-e), the replacement H10, 4 germ cell specific H1 subtypes, and H1x5. The presence of multiple H1 subtypes that differ in DNA binding affinity and chromatin compaction ability6-9 provides an additional level of modulation of chromatin function. Thus, quantitative expression analysis of individual H1 subtypes, both of mRNA and proteins, is necessary for better understanding of the regulation of higher order chromatin structure and function.
Here we describe a set of assays designed for analyzing the expression levels of individual H1 subtypes (Figure 1). mRNA expression of various H1 variant genes is measured by a set of highly sensitive and quantitative reverse transcription-PCR (qRT-PCR) assays, which are faster, more accurate and require much less samples compared with the alternative approach of Northern blot analysis. Unlike most other cellular mRNA messages, mRNAs for most histone genes, including the majority of H1 genes, lack a long polyA tail, but contain a stem-loop structure at the 3' untranslated region (UTR)10. Therefore, cDNAs are prepared from total RNA by reverse transcription using random primers instead of oligo-dT primers. Realtime PCR assays with primers specific to each H1 subtypes (Table 1) are performed to obtain highly quantitative measurement of mRNA levels of individual H1 subtypes. Expression of housekeeping genes are analyzed as controls for normalization. 
The relative abundance of proteins of each H1 subtype and core histones is obtained through reverse phase high-performance liquid chromatography (RP-HPLC) analysis of total histones extracted from mammalian cells11-13. The HPLC method and elution conditions described here give optimum separations of mouse H1 subtypes. By quantifying the HPLC profile, we calculate the relative proportion of individual H1 subtypes within H1 family, as well as determine the H1 to nucleosome ratio in the cells.

We describe a set of assays to analyze expression levels of H1 linker histones. mRNA of individual H1 genes are quantitatively measured by random primer based reverse transcription followed by real-time PCR, whereas protein quantification of H1 histones is achieved by HPLC analysis.

Linker histone H1 binds to the nucleosome core particle and linker DNA, facilitating folding of chromatin into higher order structure.  H1 is essential ...

Biology

Assembly of Nucleosomal Arrays from Recombinant Core Histones and Nucleosome Positioning DNA

Core histone octamers that are repetitively spaced along a DNA molecule are called nucleosomal arrays. Nucleosomal arrays are obtained in one of two ways: purification from in vivo sources, or reconstitution in vitro from recombinant core histones and tandemly repeated nucleosome positioning DNA. The latter method has the benefit of allowing for the assembly of a more compositionally uniform and precisely positioned nucleosomal array. Sedimentation velocity experiments in the analytical ultracentrifuge yield information about the size and shape of macromolecules by analyzing the rate at which they migrate through solution under centrifugal force. This technique, along with atomic force microscopy, can be used for quality control, ensuring that the majority of DNA templates are saturated with nucleosomes after reconstitution. Here we describe the protocols necessary to reconstitute milligram quantities of length and compositionally defined nucleosomal arrays suitable for biochemical and biophysical studies of chromatin structure and function.

A method is presented for the reconstitution of model nucleosomal arrays from recombinant core histones and tandemly repeated nucleosome positioning DNA. We also describe how sedimentation velocity experiments in the analytical ultracentrifuge, and atomic force microscopy (AFM) are used to monitor the extent of nucleosomal array saturation after reconstitution.

Core histone octamers that are repetitively spaced along a DNA molecule  are called nucleosomal arrays. Nucleosomal arrays are obtained in one of two  ...

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

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

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

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

Biochemical Assays for Analyzing Activities of ATP-dependent Chromatin Remodeling Enzymes

Members of the SNF2 family of ATPases often function as components of multi-subunit chromatin remodeling complexes that regulate nucleosome dynamics and DNA accessibility by catalyzing ATP-dependent nucleosome remodeling. Biochemically dissecting the contributions of individual subunits of such complexes to the multi-step ATP-dependent chromatin remodeling reaction requires the use of assays that monitor the production of reaction products and measure the formation of reaction intermediates. This JOVE protocol describes assays that allow one to measure the biochemical activities of chromatin remodeling complexes or subcomplexes containing various combinations of subunits. Chromatin remodeling is measured using an ATP-dependent nucleosome sliding assay, which monitors the movement of a nucleosome on a DNA molecule using an electrophoretic mobility shift assay (EMSA)-based method. Nucleosome binding activity is measured by monitoring the formation of remodeling complex-bound mononucleosomes using a similar EMSA-based method, and DNA- or nucleosome-dependent ATPase activity is assayed using thin layer chromatography (TLC) to measure the rate of conversion of ATP to ADP and phosphate in the presence of either DNA or nucleosomes. Using these assays, one can examine the functions of subunits of a chromatin remodeling complex by comparing the activities of the complete complex to those lacking one or more subunits. The human INO80 chromatin remodeling complex is used as an example; however, the methods described here can be adapted to the study of other chromatin remodeling complexes.

Here we describe biochemical assays that can be used to characterize ATP-dependent chromatin remodeling enzymes for their abilities to 1) catalyze ATP-dependent nucleosome sliding, 2) engage with nucleosome substrates, and 3) hydrolyze ATP in a nucleosome- or DNA-dependent manner.

Members of the SNF2 family of ATPases often function as components of multi-subunit chromatin remodeling complexes that regulate nucleosome dynamics and ...

Reconstitution of Nucleosomes with Differentially Isotope-labeled Sister Histones

Samenvatting

Meer video's verkennen