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>

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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Unfolding Buffer</td>
			<td>7M guanidinium HCl</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>20mM Tris pH7.5</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>10mM DTT</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Refolding Buffer</td>
			<td>10mM Tris pH7.5</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>2M NaCl</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>1mM EDTA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>2mM DTT</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Assay Buffer</td>
			<td>25mM NaxHxPO4 pH6.8</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>25mM NaCl</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>2mM DTT</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Guanidinium HCl</td>
			<td>Applichem</td>
			<td>A14199</td>
			<td>Use high quality Gu-HCl</td>
		</tr>
		<tr>
			<td>Tris&#160;</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>Na2HPO4</td>
			<td>Applichem</td>
			<td>A1046</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</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 200pg</td>
			<td>GE Healthcare</td>
			<td>28-9893-35</td>
			<td></td>
		</tr>
		<tr>
			<td>6-8kDa dialysis membrane</td>
			<td>Spectrumlabs</td>
			<td>spectra/por 1, 132650</td>
			<td></td>
		</tr>
		<tr>
			<td>50kDa dialysis membrane</td>
			<td>Spectrumlabs</td>
			<td>spectra/por 7, 132129</td>
			<td></td>
		</tr>
		<tr>
			<td>10kDa centrigugal filter unit</td>
			<td>Merck Millipore</td>
			<td>UFC901024</td>
			<td></td>
		</tr>
		<tr>
			<td>30kDa centrigugal filter unit</td>
			<td>Merck Millipore</td>
			<td>UFC903024</td>
			<td></td>
		</tr>
		<tr>
			<td>Solution-state NMR spectrometer&#160;</td>
			<td></td>
			<td></td>
			<td>at least 500 MHz operating frequency, equipped with a triple-resonance cryoprobe</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 ...

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Biochemistry

10.5K Views.  Leibniz Institute of Molecular Pharmacology (FMP-Berlin). 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.

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

Analysis of Histone Antibody Specificity with Peptide Microarrays

Post-translational modifications (PTMs) on histone proteins are widely studied for their roles in regulating chromatin structure and gene expression. The mass production and distribution of antibodies specific to histone PTMs has greatly facilitated research on these marks. As histone PTM antibodies are key reagents for many chromatin biochemistry applications, rigorous analysis of antibody specificity is necessary for accurate data interpretation and continued progress in the field. This protocol describes an integrated pipeline for the design, fabrication and use of peptide microarrays for profiling the specificity of histone antibodies. The design and analysis aspects of this procedure are facilitated by ArrayNinja, an open-source and interactive software package we recently developed to streamline the customization of microarray print formats. This pipeline has been used to screen a large number of commercially available and widely used histone PTM antibodies, and data generated from these experiments are freely available through an online and expanding Histone Antibody Specificity Database. Beyond histones, the general methodology described herein can be applied broadly to the analysis of PTM-specific antibodies.

This manuscript describes methods for applying peptide microarray technology to specificity profiling of antibodies that recognize histones and their post-translational modifications.

Post-translational modifications (PTMs) on histone proteins are widely studied for their roles in regulating chromatin structure and gene expression. The ...

Detergent-free Ultrafast Reconstitution of Membrane Proteins into Lipid Bilayers Using Fusogenic Complementary-charged Proteoliposomes.

Detergents are indispensable for delivery of membrane proteins into 30-100 nm small unilamellar vesicles, while more complex, larger model lipid bilayers are less compatible with detergents.
Here we describe a strategy for bypassing this fundamental limitation using fusogenic oppositely charged liposomes bearing a membrane protein of interest. Fusion between such vesicles occurs within 5 min in a low ionic strength buffer. Positively charged fusogenic liposomes can be used as simple shuttle vectors for detergent-free delivery of membrane proteins into biomimetic target lipid bilayers, which are negatively charged. We also show how to reconstitute membrane proteins into fusogenic proteoliposomes with a fast 30-min protocol.
Combining these two approaches, we demonstrate a fast assembly of an electron transport chain consisting of two membrane proteins from E. coli, a primary proton pump bo3-oxidase and F1Fo ATP synthase, in membranes of vesicles of various sizes, ranging from 0.1 to &#62;10 microns, as well as ATP production by this chain.

Here we present two ultrafast protocols for reconstitution of membrane proteins into fusogenic proteoliposomes, and fusion of such proteoliposomes with target lipid bilayers for detergent-free delivery of these membrane proteins into the postfusion bilayer. The combination of these approaches enables fast and easily controlled assembly of complex multi-component membrane systems.

Detergents are indispensable for delivery of membrane proteins into 30-100 nm small unilamellar vesicles, while more complex, larger model lipid bilayers ...

Determining if DNA Stained with a Cyanine Dye Can Be Digested with Restriction Enzymes

Visualization of DNA for fluorescence microscopy utilizes a variety of dyes such as cyanine dyes. These dyes are utilized due to their high affinity and sensitivity for DNA. In order to determine if the DNA molecules are full length after the completion of the experiment, a method is required to determine if the stained molecules are full length by digesting DNA with restriction enzymes. However, stained DNA may inhibit the enzymes, so a method is needed to determine what enzymes one could use for fluorochrome stained DNA. In this method, DNA is stained with a cyanine dye overnight to allow the dye and DNA to equilibrate. Next, stained DNA is digested with a restriction enzyme, loaded into a gel and electrophoresed. The experimental DNA digest bands are compared to an in silico digest to determine the restriction enzyme activity. If there is the same number of bands as expected, then the reaction is complete. More bands than expected indicate partial digestion and less bands indicate incomplete digestion. The advantage of this method is its simplicity and it uses equipment that a scientist would need for a restriction enzyme assay and gel electrophoresis. A limitation of this method is that the enzymes available to most scientists are commercially available enzymes; however, any restriction enzymes could be used.

Staining DNA molecules for fluorescence microscopy allows a scientist to view them during an experiment. In the method presented here, DNA molecules are pre-stained with fluorescent dyes and digested with methylation and non-methylation sensitive restriction enzymes.

Visualization of DNA for fluorescence microscopy utilizes a variety of dyes such as cyanine dyes. These dyes are utilized due to their high affinity and ...

Purification and Reconstitution of TRPV1 for Spectroscopic Analysis

Polymodal ion channels transduce multiple stimuli of different natures into allosteric changes; these dynamic conformations are challenging to determine and remain largely unknown. With recent advances in single-particle cryo-electron microscopy (cryo-EM) shedding light on the structural features of agonist binding sites and the activation mechanism of several ion channels, the stage is set for an in-depth dynamic analysis of their gating mechanisms using spectroscopic approaches. Spectroscopic techniques such as electron paramagnetic resonance (EPR) and double electron-electron resonance (DEER) have been mainly restricted to the study of prokaryotic ion channels that can be purified in large quantities. The requirement for large amounts of functional and stable membrane proteins has hampered the study of mammalian ion channels using these approaches. EPR and DEER offer many advantages, including determination of the structure and dynamic changes of mobile protein regions, albeit at low resolution, that might be difficult to obtain by X-ray crystallography or cryo-EM, and monitoring reversible gating transition (i.e., closed, open, sensitized, and desensitized). Here, we provide protocols for obtaining milligrams of functional detergent-solubilized transient receptor potential cation channel subfamily V member 1 (TRPV1) that can be labeled for EPR and DEER spectroscopy.

This article describes specific methods to obtain biochemical quantities of detergent-solubilized TRPV1 for spectroscopic analysis. The combined protocols provide biochemical and biophysical tools that can be adapted to facilitate structural and functional studies for mammalian ion channels in a membrane-controlled environment.

Polymodal ion channels transduce multiple stimuli of different natures into allosteric changes; these dynamic conformations are challenging to determine ...

Benchtop Immobilized Metal Affinity Chromatography, Reconstitution and Assay of a Polyhistidine Tagged Metalloenzyme for the Undergraduate Laboratory

Benchtop immobilized metal affinity chromatography (IMAC), of polyhistidine tagged proteins is easily mastered by undergraduate students and has become the most widely used protein purification method in the modern literature. But, the application of affinity chromatography to metal binding proteins, especially those with redox sensitive metals such as iron, is often limited to laboratories with access to a glove box - equipment that is not routinely available in the undergraduate laboratory. In this article, we demonstrate our benchtop methods for isolation, IMAC purification and metal-ion reconstitution of a poly-histidine tagged, redox-active, non-heme iron binding extradiol dioxygenase and the assay of the dioxygenase with varied substrate concentrations and saturating oxygen. These methods are executed by undergraduate students and implemented in the undergraduate teaching and research laboratory with instrumentation that is accessible and affordable at primarily undergraduate institutions.

Here we present a protocol for the benchtop immobilized metal affinity chromatography purification and subsequent reconstitution of a polyhistidine tagged, non-heme iron binding dioxygenase suitable for the undergraduate teaching laboratory.

Benchtop immobilized metal affinity chromatography (IMAC), of polyhistidine tagged proteins is easily mastered by undergraduate students and has become ...

In Vitro Reconstitution of Self-Organizing Protein Patterns on Supported Lipid Bilayers

Many aspects of the fundamental spatiotemporal organization of cells are governed by reaction-diffusion type systems. In vitro reconstitution of such systems allows for detailed studies of their underlying mechanisms which would not be feasible in vivo. Here, we provide a protocol for the in vitro reconstitution of the MinCDE system of Escherichia coli, which positions the cell division septum in the cell middle. The assay is designed to supply only the components necessary for self-organization, namely a membrane, the two proteins MinD and MinE and energy in the form of ATP. We therefore fabricate an open reaction chamber on a coverslip, on which a supported lipid bilayer is formed. The open design of the chamber allows for optimal preparation of the lipid bilayer and controlled manipulation of the bulk content. The two proteins, MinD and MinE, as well as ATP, are then added into the bulk volume above the membrane. Imaging is possible by many optical microscopies, as the design supports confocal, wide-field and TIRF microscopy alike. In a variation of the protocol, the lipid bilayer is formed on a patterned support, on cell-shaped PDMS microstructures, instead of glass. Lowering the bulk solution to the rim of these compartments encloses the reaction in a smaller compartment and provides boundaries that allow mimicking of in vivo oscillatory behavior. Taken together, we describe protocols to reconstitute the MinCDE system both with and without spatial confinement, allowing researchers to precisely control all aspects influencing pattern formation, such as concentration ranges and addition of other factors or proteins, and to systematically increase system complexity in a relatively simple experimental setup.

In Vitro Reconstitution of Self-Organizing Protein Patterns on Supported Lipid Bilayers

We provide a protocol for in vitro self-organization assays of MinD and MinE on a supported lipid bilayer in an open chamber. Additionally, we describe how to enclose the assay in lipid-clad PDMS microcompartments to mimic in vivo conditions by reaction confinement.

Many aspects of the fundamental spatiotemporal organization of cells are governed by reaction-diffusion type systems. In vitro reconstitution of such ...

Reconstitution of Cell-cycle Oscillations in Microemulsions of Cell-free Xenopus Egg Extracts

Real-time measurement of oscillations at the single-cell level is important to uncover the mechanisms of biological clocks. Although bulk extracts prepared from Xenopus laevis eggs have been powerful in dissecting biochemical networks underlying the cell-cycle progression, their ensemble average measurement typically leads to a damped oscillation, despite each individual oscillator being sustained. This is due to the difficulty of perfect synchronization among individual oscillators in noisy biological systems. To retrieve the single-cell dynamics of the oscillator, we developed a droplet-based artificial cell system that can reconstitute mitotic cycles in cell-like compartments encapsulating cycling cytoplasmic extracts of Xenopus laevis eggs. These simple cytoplasmic-only cells exhibit sustained oscillations for over 30 cycles. To build more complicated cells with nuclei, we added demembranated sperm chromatin to trigger nuclei self-assembly in the system. We observed a periodic progression of chromosome condensation/decondensation and nuclei envelop breakdown/reformation, like in real cells. This indicates that the mitotic oscillator functions faithfully to drive multiple downstream mitotic events. We simultaneously tracked the dynamics of the mitotic oscillator and downstream processes in individual droplets using multi-channel time-lapse fluorescence microscopy. The artificial cell-cycle system provides a high-throughput framework for quantitative manipulation and analysis of mitotic oscillations with single-cell resolution, which likely provides important insights into the regulatory machinery and functions of the clock.

Reconstitution of Cell-cycle Oscillations in Microemulsions of Cell-free Xenopus Egg Extracts

We present a method for the generation of in vitro self-sustained mitotic oscillations at the single-cell level by encapsulating egg extracts of Xenopus laevis in water-in-oil microemulsions.

Real-time measurement of oscillations at the single-cell level is important to uncover the mechanisms of biological clocks. Although bulk extracts ...

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 ...

Detergent-assisted Reconstitution of Recombinant Drosophila Atlastin into Liposomes for Lipid-mixing Assays

Membrane fusion is a crucial process in the eukaryotic cell. Specialized proteins are necessary to catalyze fusion. Atlastins are endoplasmic reticulum (ER) resident proteins implicated in homotypic fusion of the ER. We detail here a method for purifying a glutathione S-transferase (GST) and poly-histidine tagged Drosophila atlastin by two rounds of affinity chromatography. Studying fusion reactions in vitro requires purified fusion proteins to be inserted into a lipid bilayer. Liposomes are ideal model membranes, as lipid composition and size may be adjusted. To this end, we describe a reconstitution method by detergent removal for Drosophila atlastin into preformed liposomes. While several reconstitution methods are available, reconstitution by detergent removal has several advantages that make it suitable for atlastins and other similar proteins. The advantage of this method includes a high reconstitution yield and correct orientation of the reconstituted protein. This method can be extended to other membrane proteins and for other applications that require proteoliposomes. Additionally, we describe a FRET based lipid mixing assay of proteoliposomes used as a measurement of membrane fusion.

Detergent-assisted Reconstitution of Recombinant Drosophila Atlastin into Liposomes for Lipid-mixing Assays

Biological membrane fusion is catalyzed by specialized fusion proteins. Measuring the fusogenic properties of proteins can be achieved by lipid mixing assays. We present a method for purifying recombinant Drosophila atlastin, a protein that mediates homotypic fusion of the ER, reconstituting it to preformed liposomes, and testing for fusion capacity.

Membrane fusion is a crucial process in the eukaryotic cell. Specialized proteins are necessary to catalyze fusion. Atlastins are endoplasmic reticulum ...