This work was supported by NIH grants GM45916 and GM66834 to J.C.H. and a fellowship from the International Rett Syndrome Foundation to A.K. This work was also supported by NIH grant GM088409 and Howard Hughes Medical Institute contributions to K.L.

1. Assembly of Recombinant Core Histones into Octamers
Rationale: The first step in nucleosomal array reconstitution is to prepare native core histone octamers from lyophilized recombinant core histones. Histone proteins are combined in equal molar amounts and assembled into histone octamers by dialyzing the samples out of a denaturing buffer into refolding buffer.
<ol>
	<li>Purify and lyophilize recombinant core histones (H2A, H2B, H3, H4) as described13.</li>
	<li>Dissolve approximately 5 mg of each lyophilized core histone in 3 ml of unfolding buffer (6 M guanidinium HCl, 20 mM Tris pH 7.5, 5 mM DTT). Allow each aliquot to dissolve for at least one hour. Ensure that no protein remains on the sides of the container.</li>
	<li>Measure the absorbance of each histone at 276 nm using unfolding buffer as a reference.</li>
	<li>Calculate the molarity of each histone using their extinction coefficients (See Table 1 for Xenopus histone extinction coefficients). Compare the absorbance determined concentration of histone to the lyophilized dry weight, and estimate if the majority of the protein is dissolved.</li>
	<li>Determine which histone you have the fewest moles of, as this will be the limiting number of moles for all histones.</li>
	<li>Combine the histones in equal molar amounts and dilute with unfolding buffer to a final concentration of 1 mg/ml.</li>
	<li>Place the sample into 6-8 kDa MWCO dialysis tubing. Seal the tubing and place into 2 L of cold refolding buffer (2 M NaCl, 10 mM Tris pH 7.5, 1 mM EDTA, 5 mM &#946;-mercaptoethanol).</li>
	<li>Dialyze the sample for 18 hr at 4 &#176;C with stirring. Ensure the dialysis tubing can rotate freely and vigorously, or else the histones may precipitate.</li>
	<li>Change the dialysis buffer twice in the 18 hr period (i.e. change the refolding buffer about every 6 hr until done). Even if precipitate is formed do not discard the sample, some octamer may be recovered although the yield will be decreased.</li>
</ol>
Note: See step 2.1-2.2 to prepare column equilibration for the next day.
2. Purification of Histone Octamers by Size Exclusion Chromatography
Rationale: After concluding with section 1, samples will contain histone octamers as well as other histone complexes such as aggregates, H3/H4 tetramers, and H2A/H2B dimers. Histone octamers will be purified away from these other complexes using size exclusion chromatography (SEC).
<ol>
	<li>Connect a HiLoad 16/60 Superdex200 16/60 (S200) column to an FPLC system. Ensure that air bubbles do not enter the system.</li>
	<li>Clean the S200 column with 0.2 &#956;m filtered water. Use a flow rate of 0.3 ml/min and set a back pressure limit of 0.5 MPa. Ensure you have enough water and allow the column to clean overnight.</li>
	<li>Equilibrate the S200 column and sample loop of the FPLC using refolding buffer. Use 1 L of 0.2 &#956;m filtered refolding buffer. Flow refolding buffer through the column at a rate of 1 ml/min and a maximum pressure of 0.5 MPa for about 2 hr.</li>
	<li>Equilibrate a Vivaspin centrifugal concentrator (50 kDa MWCO) with 1 ml of refolding buffer. Remove the sample from dialysis tubing and centrifuge the sample at 4 &#176;C to remove any precipitates. Pipette the sample supernatant into the concentrator. Concentrate the sample to a volume of approximately 500 &#956;l.</li>
	<li>Remove the concentrated octamer sample to a new container and rinse the concentrator with 1 ml refolding buffer. Concentrate the rinse down to 500 &#956;l, and add it to the octamer sample. This helps to salvage any octamer remaining in the concentrator.</li>
	<li>Spin the sample in a 1.5 ml microfuge tube for 5 min at 10,000 rpm at 4 &#176;C. Collect the supernatant and transfer to a new tube. This helps to remove any precipitate before loading the sample on the FPLC.</li>
	<li>Load the sample onto the S200 column. Load a maximum of 1.5 ml total volume or 15 mg octamer in a single purification.</li>
	<li>Elute the sample using freshly made refolding buffer. Use a flow rate of 1 ml/min and a back pressure limit of 0.5 MPa. Use two column volumes (400 ml) refolding buffer and collect 2 ml fractions. Monitor the absorbance at 280 nm during elution. The different species will elute in order of decreasing size. Histone aggregates usually elute at approximately 45 ml, octamer at about 65 ml, and dimer at around 84 ml (Figure 1).</li>
	<li>Analyze the elution fractions from peaks of interest by running samples on an 18-20% SDS-PAGE. Fractions containing purified octamers should have equal molar amounts of the four core histone proteins (Figure 1).</li>
	<li>Pool the fractions that contain purified octamers, and concentrate to &#8804;15 mg/ml with a Vivaspin centrifugal filter. If left dilute, octamers may disassociate into H2A/H2B dimers and H3/H4 tetramers.</li>
	<li>Determine the octamer concentration by measuring the absorbance at 280 nm and calculate using the extinction coefficient from Table 1.</li>
	<li>Histone octamers can be kept short term in refolding buffer at 4 &#176;C. If being stored long term, the octamers will be more stable at -20 &#176;C and in a 50% glycerol solution. Octamers stored in glycerol should be dialyzed into fresh refolding buffer before absorbance measurements and use in nucleosomal array reconstitutions.</li>
</ol>
3. Reconstitution of Nucleosomal Arrays from DNA and Purified Octamers
Rationale: Reconstitution of nucleosomal arrays requires that histone octamers and template DNA be combined at specific molar ratios. Mixtures of DNA and histone octamers in 2 M NaCl are step dialyzed into buffers of decreasing ionic strength. A gradual change to lower salt ensures proper nucleosome formation. Obtaining nucleosomal arrays with the desired level of histone octamer saturation of the template DNA requires small scale test reconstitutions followed by a large scale preparative reconstitution. The appropriate conditions defined by the small scale reconstitutions are used to guide the preparative reconstitution.
<ol>
	<li>Purify tandemly repeated nucleosome positioning DNA as described 4,12. Briefly, isolate the plasmid DNA using a Qiagen GigaPrep kit or a similar product. Set-up a restriction enzyme digest in order to release the template DNA and digest the plasmid DNA to smaller sizes (&#8804;700bp). The template DNA can then be purified away from the small plasmid DNA remnants using size exclusion chromatography (SEC). A gravity fed column, with a height of about 115 cm packed with Sephacryl S-1000 beads is sufficient for purification of 601207bp x 12mer DNA.</li>
</ol>
Notes: For plasmid DNA purification at decreased cost but increased effort, one can use alkaline lysis with phenol/chloroform DNA purification in order to isolate plasmid DNA from E. coli23,24. Strategies for separating template DNA from plasmid DNA may vary with changes in template DNA size. For SEC it is important to set up the restriction enzyme digest in a fashion which maximizes the difference in size between the template and plasmid DNA.
<ol start="2">
	<li>Determine which molar ratios (r) to use for reconstitutions. r is equal to the ratio of moles of octamer to moles of DNA repeat. For initial small scale trials, r values of 0.9, 1, and 1.1 are appropriate if the intent is to obtain saturated nucleosomal arrays.</li>
	<li>Prepare the small scale samples by calculating the amount of octamer to add to approximately 18 &#956;g of DNA for each molar ratio to be tested. Mix the DNA and histone octamers. The final concentration of NaCl in the sample should be &#8805;2 M, and the final concentration of DNA should be around 0.3 &#956;g/&#956;l. The final sample buffer conditions should also include 10 mM Tris pH 7.8, and 1 mM EDTA.</li>
	<li>The samples are now ready for dialysis. Load the samples into 12k-14k MWCO dialysis tubing. Dialyze the samples against 2L of buffer of decreasing ionic strength as follows.</li>
</ol>
Components in all buffers: 10 mM Tris pH 7.8, 1 mM EDTA. Buffer 1 (add 1 M NaCl, 1 mM DTT) for 5-6 hr. Buffer 2 (add 0.75 M NaCl, 1 mM DTT) overnight. Buffer 3 (add 2.5 mM NaCl, 1 mM DTT) for 5-6 hr. Buffer 4 (add 2.5 mM NaCl, 0.1 mM PMSF) overnight.
<ol start="5">
	<li>Remove the samples from the dialysis tubing and proceed to sections 4-6. If the small-scale samples yield the desired results, repeat the steps in sections 3-6 on the large scale.</li>
</ol>
Note: In order to conserve resources, small scale samples should have the minimum volume necessary to suffice for downstream screening assays. In order to have enough sample for the sedimentation velocity and AFM experiments listed here, 50 &#956;l samples at 0.3 mg/ml DNA are appropriate. The size of large scale, preparative samples are dependent on their intended use. A large scale sample should be prepared in the same manner as the small scale samples.
4. Sedimentation Velocity Analysis of Reconstituted Nucleosomal Arrays
Rationale: Sedimentation velocity experiments in the Beckman Xl-A/I analytical ultracentrifuge yield information on the size and shape of molecules in solution. Sedimentation velocity experiments performed under low salt conditions are used to determine the extent to which the reconstituted arrays are saturated with nucleosomes.
<ol>
	<li>Dilute the sample to an absorbance around 0.5 at 260 nm using TEN buffer (10 mM Tris pH 7.8, 1 mM EDTA, 2.5 mM NaCl). Load 400 &#956;l of sample into a cell with a two sector centerpiece, with the sample on one side and the reference (TEN buffer) on the other25. Keep the reference meniscus higher than the sample meniscus (i.e. add 420 &#956;l reference solution).</li>
	<li>Load the cells into the rotor, and align the cells properly using the hash marks on the bottom of the cells and rotor25. Gently dust the lenses of the cells using compressed air.</li>
	<li>Turn on the XL-A/XL-I centrifuge, insert the rotor, and attach and secure the optics as described in the manual. Open the Proteome Lab software and select file: new.</li>
	<li>Set up a single scan run at 3,000 rpm and a temperature of 25 &#176;C. Under options, select stop after last scan, and run radial calibration (radial calibration is not necessary if the rotor was the last rotor used in the AUC).</li>
	<li>For each cell, name the samples, select a wavelength (260 nm), select absorbance, and choose a save file location on your computer. Begin the single scan run.</li>
	<li>Use the single scan to determine the appropriate radial scan length (Figure 2A). Decreasing the length of the cell that will be scanned decreases the run time. However, be sure to include the sample meniscus and extend the end very close to, or at the bottom of the cell.</li>
</ol>
Note: Single scans allow the measurement area to be shortened by starting measurements just before the sample meniscus (Figure 2A). Most of the scans generated during the AUC run should have boundary fractions past the meniscus as well as stable plateaus (Figure 2B). Dirty lenses on the AUC cells can generate scans with large spikes, these may affect analysis of the data. The UltraScan software includes a manual which is a great resource for information regarding the analysis of analytical ultracentrifugation data26.
<ol start="7">
	<li>Using the XL-A/XL-I control panel, enter a speed of 0 and press start. This will prompt the centrifuge chamber to pull a vacuum and allow the temperature of the chamber to equilibrate. Wait 1 hr to allow for temperature equilibration before beginning a run.</li>
	<li>Use the software to set the run up for the desired speed and number of scans. Higher speeds increase resolution, but one should collect at least 20 scans (preferably more) before the sample has sedimented to the bottom of the cell. It is convenient to overlay the last several scans (accomplished in the options menu) in order to monitor the progress of the run and check for potential problems such as leaking cells. Monitor the first couple of scans to ensure proper operation.</li>
</ol>
5. Editing and Analyzing Sedimentation Velocity Data
Rationale: Raw sedimentation velocity data should be analyzed by a program that yields a diffusion-corrected sedimentation coefficient distribution. This information in turn indicates the fraction of a reconstituted sample that contains a given saturation level, e.g. if using a 12-mer DNA template it will be possible to determine the fraction of reconstituted arrays that has 10, 11 and 12 nucleosomes/DNA.
<ol>
	<li>Use AUC data analysis software such as UltraScan to edit the data26. Editing of scan data must be done for each cell and creates a data set for subsequent analysis. Proper editing of scans involves defining the sample meniscus, reducing the data to a region of interest, and defining baseline as well as plateau regions (Figure 2). Unwanted scans can also be removed from the data set at this point. However, since it is possible to remove unwanted scans during analysis, it is recommended that all scans be kept at this point.</li>
	<li>We strongly recommend that the enhanced van Holde-Weischet method be used to analyze the data27. This analysis method corrects for the effects of diffusion over the course of the run and yields the integral distribution of sedimentation coefficients. In particular, the enhanced van Holde-Weischet method (as implemented in UltraScan) will allow for the inclusion of scans which lack stable plateaus, or which contain boundary fractions which have not cleared the meniscus in the analysis28.</li>
	<li>Determine the distribution of sedimentation coefficients for the assembled arrays. Representative results for 601-12 (207bp, 12-mer) nucleosomal arrays are shown in Figure 3.</li>
	<li>If one of the small scale samples contains arrays with the desired range of sedimentation coefficients, then repeat the process at an increased scale starting with section 3. If none of the small scale samples has a proper distribution of sedimentation coefficients then repeat the small scale tests with a new r range starting at section 3.</li>
</ol>
6. Visualizing Nucleosomal Arrays Using Atomic Force Microscopy (AFM)
Rationale: AFM allows visualization of the level of saturation of nucleosomal arrays. This technique complements sedimentation velocity by AUC and restriction enzyme digestion as a quality control assay.
<ol>
	<li>Using methods described above, obtain a well saturated nucleosome array (Figure 5A).</li>
	<li>Prepare APTES treated mica slides by placing ~30 &#956;l 1:1,000 dilution of commercial (Sigma-Aldrich (3-Aminopropyl)triethoxysilane A3648-100ML) in 0.22 &#956;m filtered nanopure water for 30 min.</li>
	<li>After 30 min rinse the APTES off with filtered water and gently dry them in a Nitrogen flow.</li>
	<li>Place appropriately diluted nucleosome array sample (~1.5 ng/&#956;l) on the slide, and incubate covered at room temperature for 15 min.</li>
	<li>Rinse sample off with filtered sample buffer, dry as before, and place slide on the stage of AFM (in this case an Asylum Research MFP-3D Atomic Force Microscope).</li>
	<li>Start by imaging 2 x 2 &#956;m scans with 512 scan lines. Count the number of nucleosomes to ascertain level of saturation (Figure 5B).</li>
	<li>Ideally you should image areas on the slide with well separated nucleosome arrays.</li>
	<li>For higher resolution images, a 500 x 500 nm scan can be obtained (Figure 5C).</li>
	<li>Images can be flattened and analyzed using the MFP-3D software provided by Asylum.</li>
	<li>Each image can be divided into four quadrants and zoomed in digitally to obtain a clearer view of well-separated arrays and multiple free hand lines can be drawn through the arrays and height profiles are obtained for the nucleosomes (Figure 5C, right panel-height trace and 5D).</li>
	<li>Several such images should be analyzed for each type of array encompassing several hundreds of nucleosomes. Height profiles are recorded, categorized and plotted in MS Excel.</li>
</ol>

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(33), e1530 (2009).</li><li>Demeler, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=UltraScan:+a+comprehensive+data+analysis+software+package+for+analytical+ultracentrifugation+experiments.">UltraScan: a comprehensive data analysis software package for analytical ultracentrifugation experiments.</a> Modern Analytical Ultracentrifugation: Techniques. , 210-230 (2005).</li><li>Holde, K. V., Weischet, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Boundary+analysis+of+sedimentation+velocity+experiments+with+monodisperse+and+paucidisperse+solutes.">Boundary analysis of sedimentation velocity experiments with monodisperse and paucidisperse solutes.</a> Biopolymers. 17 (6), 1387-1403 (1978).</li><li>Demeler, B., van Holde, K. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sedimentation+velocity+analysis+of+highly+heterogeneous+systems.">Sedimentation velocity analysis of highly heterogeneous systems.</a> Analytical Biochemistry. 335, 279-288 (2004).</li><li>Hansen, J., Lohr, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assembly+and+structural+properties+of+subsaturated+chromatin+arrays.">Assembly and structural properties of subsaturated chromatin arrays.</a> Journal of Biological Chemistry. 8, 5840-5848 (1993).</li><li>Routh, A., Sandin, S., Rhodes, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nucleosome+repeat+length+and+linker+histone+stoichiometry+determine+chromatin+fiber+structure.">Nucleosome repeat length and linker histone stoichiometry determine chromatin fiber structure.</a> Proceedings of the National Academy of Sciences of the U S A. 105 (26), 8872-8877 (2008).</li><li>Zhou, J., Fan, J. Y., Rangasamy, D., Tremethick, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+nucleosome+surface+regulates+chromatin+compaction+and+couples+it+with+transcriptional+repression.">The nucleosome surface regulates chromatin compaction and couples it with transcriptional repression.</a> Nature Structural &amp; Molecular Biology. 14 (11), 1070-1076 (2007).</li><li>Dorigo, B., Schalch, T., Kulangara, A., Duda, S., Schroeder, R. R., Richmond, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nucleosome+arrays+reveal+the+two-start+organization+of+the+chromatin+fiber.">Nucleosome arrays reveal the two-start organization of the chromatin fiber.</a> Science (New York, N.Y.). 306 (5701), 1571-1573 (2004).</li><li>Correll, S. J., Schubert, M. H., Grigoryev, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=a+Short+nucleosome+repeats+impose+rotational+modulations+on+chromatin+fibre+folding.">a Short nucleosome repeats impose rotational modulations on chromatin fibre folding.</a> The EMBO Journal. 31 (10), 2416-2426 (2012).</li><li>Mcbryant, S. J., Krause, C., Woodcock, C. L., 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=The+Silent+Information+Regulator+3+Protein+,+SIR3p+,+Binds+to+Chromatin+Fibers+and+Assembles+a+Hypercondensed+Chromatin+Architecture+in+the+Presence+of+Salt.">The Silent Information Regulator 3 Protein , SIR3p , Binds to Chromatin Fibers and Assembles a Hypercondensed Chromatin Architecture in the Presence of Salt.</a> Molecular and Cellular Biology. 28 (11), 3563-3572 (2008).</li><li>Schalch, T., Duda, S., Sargent, D. F., Richmond, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=X-ray+structure+of+a+tetranucleosome+and+its+implications+for+the+chromatin+fibre.">X-ray structure of a tetranucleosome and its implications for the chromatin fibre.</a> Nature. 436 (7047), 138-1341 (2005).</li><li>Fan, J. Y., Gordon, F., Luger, K., Hansen, J. C., Tremethick, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+essential+histone+variant+H2A.Z+regulates+the+equilibrium+between+different+chromatin+conformational+states.">The essential histone variant H2A.Z regulates the equilibrium between different chromatin conformational states.</a> Nature Structural Biology. 9 (3), 172-176 (2002).</li><li>Carruthers, L. M., Bednar, J., Woodcock, C. L., 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=Linker+histones+stabilize+the+intrinsic+salt-dependent+folding+of+nucleosomal+arrays:+mechanistic+ramifications+for+higher-order+chromatin+folding.">Linker histones stabilize the intrinsic salt-dependent folding of nucleosomal arrays: mechanistic ramifications for higher-order chromatin folding.</a> Biochemistry. 37 (42), 14776-14787 (1998).</li><li>Huynh, V. A. T., Robinson, P. J. J., Rhodes, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Method+for+the+In+Vitro+Reconstitution+of+a+Defined+&amp;#34;30+nm&amp;#34;+Chromatin+Fibre+Containing+Stoichiometric+Amounts+of+the+Linker+Histone.">A Method for the In Vitro Reconstitution of a Defined &amp;#34;30 nm&amp;#34; Chromatin Fibre Containing Stoichiometric Amounts of the Linker Histone.</a> Journal of Molecular Biology. 345 (5), 957-968 (2005).</li><li>Dorigo, B., Schalch, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chromatin+fiber+folding:+requirement+for+the+histone+H4+N-terminal+tail.">Chromatin fiber folding: requirement for the histone H4 N-terminal tail.</a> J. Mol. Biol. 2836 (03), 85-96 (2003).</li><li>Qian, R. L., Liu, Z. 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=Visualization+of+chromatin+folding+patterns+in+chicken+erythrocytes+by+atomic+force+microscopy+(AFM.">Visualization of chromatin folding patterns in chicken erythrocytes by atomic force microscopy (AFM.</a> Cell Research. 7 (2), 143-150 (1997).</li></ol>

The authors have no conflicts of interest.

Model nucleosomal arrays are a very useful tool for the in vitro study of chromatin structure and function. For example, they have been widely used to study the mechanism of chromatin fiber condensation in solution 30-34, and made it possible to obtain an x-ray structure of a tetranucleosome 35. More recently they have proven useful in deciphering the structural effects of specific core histone variants, mutants and posttranslational modifications14-16,36. Here we describe a gene...

Eukaryotic genomes do not exist as naked DNA, but rather are compacted and organized by bound proteins. These complexes of DNA and protein are known as chromatin. The basic repeating unit of chromatin is the nucleosome. A nucleosome consists of histone octamer and 146 base pairs of DNA wrapped around the histone octamer about 1.6 times1. The histone octamer is composed of two copies each of the core histones H2A, H2B, H3, and H4. Core histone octamers that are repetitively spaced along a DNA molecule are called nucleosomal arrays. The extended structure of nucleosomal arrays has been referred to as the 10 nm fiber or the "beads on a string" structure and is present in vitro under low salt conditions2. The 10 nm fiber is capable of condensing into higher order structures through intra-array compaction and/or inter-array oligomerization2. These higher order structures can be induced in the presence of salts, or can be influenced through the binding of chromatin architectural proteins to the nucleosomal array3,4. Levels of chromatin compaction are inversely correlated with rate of transcription in vivo5,6. Recent research highlights the importance of the structural organization of genomes in processes such as differentiation, cancer development, and others7,8. The use of nucleosomal arrays to study chromatin structure and function has become widespread. Here we describe a method for the assembly of nucleosomal arrays from recombinant core histones and nucleosome positioning DNA.
Using recombinant DNA with tandem repeats of nucleosome positioning sequences allows for the reconstitution of arrays that contain regularly spaced nucleosomes. Two of the more popular positioning sequences are the 5S rRNA gene sequence and the "601" sequence9,10. The 601 sequence was derived from SELEX experiments and more strongly positions nucleosomes than the 5S sequence11. Consequently, the linker DNA length of the 601 arrays is more homogeneous. Tandemly repeated nucleosome positioning DNA is obtained by gel filtration4,12. Recombinant histones are purified from E. coli under denaturing conditions13. The use of recombinant histones allows one to carefully control the histone composition of the nucleosomal arrays. For example, core histones bearing specific mutations14 or post-translational modifications15,16 can be substituted for wild type core histones.
Sedimentation velocity experiments monitor the rate of sedimentation of macromolecules in solution under an applied centrifugal force17. This yields information about the size and shape of macromolecules in a sample. Sedimentation velocity experiments are thus an appropriate tool for studying solution state changes in chromatin fiber structure due to chromatin condensation18. Importantly, it is first necessary to use sedimentation velocity experiments as a quality control step in nucleosomal array reconstitution. If DNA and nucleosomes are not combined at the proper molar ratio, the arrays may be under- or over-saturated with core histones. Thus, the information gained from sedimentation velocity experiments is used to ensure that the DNA is properly saturated with nucleosomes. It is important to use alternative methods to estimate the saturation of DNA with nucleosomes, especially if working with a previously uncharacterized DNA template. Therefore, we also describe a method for analysis of nucleosomal arrays using atomic force microscopy (AFM). AFM is a powerful technique that allows visualization of the effects of a number of parameters, such as the level of saturation, effect of the presence of histone variants or the effects of MgCl2 19,20. AFM has also been applied to study nucleosome dynamics using time lapse imaging 21. In vitro assembled nucleosomal 12-mer arrays are particularly amenable to AFM studies because they belong in the right size range for AFM imaging 22. In the present study we have used AFM of nucleosomal arrays as a quality control as well as a means of affirming the data from AUC ("seeing is believing"). In addition to simple visualization, AFM allows measurement of height profiles of samples as an additional metric.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>(3-Aminopropyl)triethoxysilane</td>
 <td>Sigma-Aldrich </td>
 <td>A3648-100ML</td>
 <td></td>
 </tr>
 <tr>
 <td>6-8 kDa MWCO Dialysis Tubing</td>
 <td>Fisher</td>
 <td>21-152-5</td>
 <td></td>
 </tr>
 <tr>
 <td>HiLoad Superdex 200 16/60 Column</td>
 <td>GE</td>
 <td>17-1069-01</td>
 <td></td>
 </tr>
 <tr>
 <td>Vivaspin 50 kDa MWCO Centrifugal Concentrator</td>
 <td>Sartorius</td>
 <td>VS2031</td>
 <td></td>
 </tr>
 <tr>
 <td>12-14 kDa MWCO Dialysis Tubing</td>
 <td>Fisher</td>
 <td>08-667A</td>
 <td></td>
 </tr>
 <tr>
 <td>Illustra Sephacryl S-1000 Superfine</td>
 <td>GE</td>
 <td>17-0476-01 </td>
 <td></td>
 </tr>
 <tr>
 <td>XL-A/I Analytical Ultracentrifuge</td>
 <td>Beckman-Coulter</td>
 <td></td>
 <td></td>
 </tr>
 </tbody>
</table>

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.

To illustrate the protocol we reconstituted nucleosomal arrays from recombinant Xenopus core histones and DNA consisting of 12 tandem 207 bp repeats of the 601 positioning sequence (601207 x 12). We first assembled native octamers from lyophilized core histones and then purified the octamers by FPLC using an S200 column (Figure 1A). Larger complexes elute earlier from the S200 column. Histones generally elute in this order: non-specific histone aggregates, histone octamer, H3/H4 tetra...

Watch this Scientific Journal Video about Assembly of Nucleosomal Arrays from Recombinant Core Histones and Nucleosome Positioning DNA at JoVE.com

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

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  ...

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Unit 1

Cells and their Components

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22.5K vues.  Colorado State University.  Une méthode est présentée pour la reconstitution du modèle nucléosomiques tableaux de histones recombinantes et d'ADN répétées en tandem de positionnement des nucléosomes. Nous décrivons également comment les expériences de la vitesse de sédimentation en ultracentrifugation analytique, et la microscopie à force atomique (AFM) sont utilisés pour mesurer l'ampleur de nucléosomique saturation de tableau après reconstitution. 

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

Isolation of Genomic DNA from Mouse Tails

Combining QD-FRET and Microfluidics to Monitor DNA Nanocomplex Self-Assembly in Real-Time

Advances in genomics continue to fuel the development of therapeutics that can target pathogenesis at the cellular and molecular level. Typically functional inside the cell, nucleic acid-based therapeutics require an efficient intracellular delivery system. One widely adopted approach is to complex DNA with a gene carrier to form nanocomplexes via electrostatic self-assembly, facilitating cellular uptake of DNA while protecting it against degradation. The challenge lies in the rational design of efficient gene carriers, since premature dissociation or overly stable binding would be detrimental to the cellular uptake and therapeutic efficacy. Nanocomplexes synthesized by bulk mixing showed a diverse range of intracellular unpacking and trafficking behavior, which was attributed to the heterogeneity in size and stability of nanocomplexes. Such heterogeneity hinders the accurate assessment of the self-assembly kinetics and adds to the difficulty in correlating their physical properties to transfection efficiencies or bioactivities. We present a novel convergence of nanophotonics (i.e. QD-FRET) and microfluidics to characterize the real-time kinetics of the nanocomplex self-assembly under laminar flow. QD-FRET provides a highly sensitive indication of the onset of molecular interactions and quantitative measure throughout the synthesis process, whereas microfluidics offers a well-controlled microenvironment to spatially analyze the process with high temporal resolution (~milliseconds). For the model system of polymeric nanocomplexes, two distinct stages in the self-assembly process were captured by this analytic platform. The kinetic aspect of the self-assembly process obtained at the microscale would be particularly valuable for microreactor-based reactions which are relevant to many micro- and nano-scale applications. Further, nanocomplexes may be customized through proper design of microfludic devices, and the resulting QD-FRET polymeric DNA nanocomplexes could be readily applied for establishing structure-function relationships. 

We present a novel and powerful integration of nanophotonics (QD-FRET) and microfluidics to investigate the formation of polyelectrolyte polyplexes, which is expected to provide better control and synthesis of uniform and customizable polyplexes for future nucleic acid-based therapeutics.

Advances in genomics continue to fuel the development of therapeutics that can target pathogenesis at the cellular and molecular level. Typically ...

Assembly, Loading, and Alignment of an Analytical Ultracentrifuge Sample Cell

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed centrifugation. When properly planned and executed, an AUC sedimentation velocity or sedimentation equilibrium experiment can reveal a great deal about a protein in regards to size and shape, sample purity, sedimentation coefficient, oligomerization states and protein-protein interactions.
This technique, however, requires a rigorous level of technical attention. Sample cells hold a sectored center piece sandwiched between two window assemblies. They are sealed with a torque pressure of around 120-140 in/lbs. Reference buffer and sample are loaded into the centerpiece sectors and then after sealing, the cells are precisely aligned into a titanium rotor so that the optical detection systems scan both sample and reference buffer in the same radial path midline through each centerpiece sector while rotating at speeds of up to 60, 000 rpm and under very high vacuum
Not only is proper sample cell assembly critical, sample cell components are very expensive and must be properly cared for to ensure they are in optimum working condition in order to avoid leaks and breakage during experiments. Handle windows carefully, for even the slightest crack or scratch can lead to breakage in the centrifuge. The contact between centerpiece and windows must be as tight as possible; i.e. no Newton s rings should be visible after torque pressure is applied. Dust, lint, scratches and oils on either the windows or the centerpiece all compromise this contact and can very easily lead to leaking of solutions from one sector to another or leaking out of the centerpiece all together. Not only are precious samples lost, leaking of solutions during an experiment will cause an imbalance of pressure in the cell that often leads to broken windows and centerpieces. In addition, plug gaskets and housing plugs must be securely in place to avoid solutions being pulled out of the centerpiece sector through the loading holes by the high vacuum in the centrifuge chamber. Window liners and gaskets must be free of breaks and cracks that could cause movement resulting in broken windows.
This video will demonstrate our procedures of sample cell assembly, torque, loading and rotor alignment to help minimize component damage, solution leaking and breakage during the perfect AUC experiment.

The analytical ultracentrifuge (AUC) sample cell holds sample and reference buffer and during experiments and is exposed to high vacuum and rotor speeds up to 60,000 rpm. This video will demonstrate the rigorous attention to detail necessary for assembly, loading and alignment of this very important component of an AUC experiment.

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed ...

Extraction of High Molecular Weight Genomic DNA from Soils and Sediments

The soil microbiome is a vast and relatively unexplored reservoir of genomic diversity and metabolic innovation that is intimately associated with nutrient and energy flow within terrestrial ecosystems. Cultivation-independent environmental genomic, also known as metagenomic, approaches promise unprecedented access to this genetic information with respect to pathway reconstruction and functional screening for high value therapeutic and biomass conversion processes. However, the soil microbiome still remains a challenge largely due to the difficulty in obtaining high molecular weight DNA of sufficient quality for large insert library production. Here we introduce a protocol for extracting high molecular weight, microbial community genomic DNA from soils and sediments. The quality of isolated genomic DNA is ideal for constructing large insert environmental genomic libraries for downstream sequencing and screening applications. 
The procedure starts with cell lysis. Cell walls and membranes of microbes are lysed by both mechanical (grinding) and chemical forces (&beta;-mercaptoethanol). Genomic DNA is then isolated using extraction buffer, chloroform-isoamyl alcohol and isopropyl alcohol. The buffers employed for the lysis and extraction steps include guanidine isothiocyanate and hexadecyltrimethylammonium bromide (CTAB) to preserve the integrity of the high molecular weight genomic DNA. Depending on your downstream application, the isolated genomic DNA can be further purified using cesium chloride (CsCl) gradient ultracentrifugation, which reduces impurities including humic acids. The first procedure, extraction, takes approximately 8 hours, excluding DNA quantification step. The CsCl gradient ultracentrifugation, is a two days process. During the entire procedure, genomic DNA should be treated gently to prevent shearing, avoid severe vortexing, and repetitive harsh pipetting.

A methodology to isolate high molecular weight and high quality genomic DNA from soil microbial community is described.

The soil microbiome is a vast and relatively unexplored reservoir of genomic diversity and metabolic innovation that is intimately associated with ...

Selective Capture of 5-hydroxymethylcytosine from Genomic DNA

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene expression, maintenance of genome integrity, parental imprinting, X-chromosome inactivation, regulation of development, aging, and cancer1. Recently, the presence of an oxidized 5-mC, 5-hydroxymethylcytosine (5-hmC), was discovered in mammalian cells, in particular in embryonic stem (ES) cells and neuronal cells2-4. 5-hmC is generated by oxidation of 5-mC catalyzed by TET family iron (II)/&alpha;-ketoglutarate-dependent dioxygenases2, 3. 5-hmC is proposed to be involved in the maintenance of embryonic stem (mES) cell, normal hematopoiesis and malignancies, and zygote development2, 5-10. To better understand the function of 5-hmC, a reliable and straightforward sequencing system is essential. Traditional bisulfite sequencing cannot distinguish 5-hmC from 5-mC11. To unravel the biology of 5-hmC, we have developed a highly efficient and selective chemical approach to label and capture 5-hmC, taking advantage of a bacteriophage enzyme that adds a glucose moiety to 5-hmC specifically12.
Here we describe a straightforward two-step procedure for selective chemical labeling of 5-hmC. In the first labeling step, 5-hmC in genomic DNA is labeled with a 6-azide-glucose catalyzed by &beta;-GT, a glucosyltransferase from T4 bacteriophage, in a way that transfers the 6-azide-glucose to 5-hmC from the modified cofactor, UDP-6-N3-Glc (6-N3UDPG). In the second step, biotinylation, a disulfide biotin linker is attached to the azide group by click chemistry. Both steps are highly specific and efficient, leading to complete labeling regardless of the abundance of 5-hmC in genomic regions and giving extremely low background. Following biotinylation of 5-hmC, the 5-hmC-containing DNA fragments are then selectively captured using streptavidin beads in a density-independent manner. The resulting 5-hmC-enriched DNA fragments could be used for downstream analyses, including next-generation sequencing.
Our selective labeling and capture protocol confers high sensitivity, applicable to any source of genomic DNA with variable/diverse 5-hmC abundances. Although the main purpose of this protocol is its downstream application (i.e., next-generation sequencing to map out the 5-hmC distribution in genome), it is compatible with single-molecule, real-time SMRT (DNA) sequencing, which is capable of delivering single-base resolution sequencing of 5-hmC.

Described is a two-step labeling process using &beta;-glucosyltransferase (&beta;-GT) to transfer an azide-glucose to 5-hmC, followed by click chemistry to transfer a biotin linker for easy and density-independent enrichment. This efficient and specific labeling method enables enrichment of 5-hmC with extremely low background and high-throughput epigenomic mapping via next-generation sequencing.

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene ...

Design and Use of Multiplexed Chemostat Arrays

Chemostats are continuous culture systems in which cells are grown in a tightly controlled, chemically constant environment where culture density is constrained by limiting specific nutrients.1,2 Data from chemostats are highly reproducible for the measurement of quantitative phenotypes as they provide a constant growth rate and environment at steady state. For these reasons, chemostats have become useful tools for fine-scale characterization of physiology through analysis of gene expression3-6 and other characteristics of cultures at steady-state equilibrium.7 Long-term experiments in chemostats can highlight specific trajectories that microbial populations adopt during adaptive evolution in a controlled environment. In fact, chemostats have been used for experimental evolution since their invention.8 A common result in evolution experiments is for each biological replicate to acquire a unique repertoire of mutations.9-13 This diversity suggests that there is much left to be discovered by performing evolution experiments with far greater throughput. 
We present here the design and operation of a relatively simple, low cost array of miniature chemostats&mdash;or ministats&mdash;and validate their use in determination of physiology and in evolution experiments with yeast. This approach entails growth of tens of chemostats run off a single multiplexed peristaltic pump. The cultures are maintained at a 20 ml working volume, which is practical for a variety of applications. It is our hope that increasing throughput, decreasing expense, and providing detailed building and operation instructions may also motivate research and industrial application of this design as a general platform for functionally characterizing large numbers of strains, species, and growth parameters, as well as genetic or drug libraries.

We developed and validated a small-footprint array of miniature chemostats built from readily available parts for low cost. Physiological and experimental evolution results were similar to larger volume chemostats. The ministat array provides a compact, inexpensive, and accessible platform for traditional chemostat experiments, functional genomics, and chemical screening applications.

Chemostats are continuous culture systems in which cells are grown in a tightly controlled, chemically constant environment where culture density is ...

Efficient Production and Purification of Recombinant Murine Kindlin-3 from Insect Cells for Biophysical Studies

Kindlins are essential coactivators, with talin, of the cell surface receptors&#160;integrins and also participate in integrin outside-in signalling, and the control of gene transcription in the cell nucleus. The kindlins are ~75 kDa multidomain proteins and bind to an NPxY motif and upstream T/S cluster of the integrin &#946;-subunit cytoplasmic tail. The hematopoietically-important kindlin isoform, kindlin-3, is critical for platelet aggregation during thrombus formation, leukocyte rolling in response to infection and inflammation and osteoclast podocyte formation in bone resorption. Kindlin-3&#39;s role in these processes has resulted in extensive cellular and physiological studies. However, there is a need for an efficient method of acquiring high quality milligram quantities of the protein for further studies. We have developed a protocol, here described, for the efficient expression and purification of recombinant murine kindlin-3 by use of a baculovirus-driven expression system in Sf9 cells yielding sufficient amounts of high purity full-length protein to allow its biophysical characterization. The same approach could be taken in the study of the other mammalian kindlin isoforms.

Kindlins are fundamental to cell adhesion through integrins but studies of them have been hampered by the difficulty encountered in expressing them recombinantly in bacterial hosts. We describe here methods for their efficient production in baculovirus-infected insect cells.

Kindlins are essential coactivators, with talin, of the cell surface receptors&#160;integrins and also participate in integrin outside-in signalling, and ...

Non-chromatographic Purification of Recombinant Elastin-like Polypeptides and their Fusions with Peptides and Proteins from Escherichia coli

Elastin-like polypeptides are repetitive biopolymers that exhibit a lower critical solution temperature phase transition behavior, existing as soluble unimers below a characteristic transition temperature and aggregating into micron-scale coacervates above their transition temperature. The design of elastin-like polypeptides at the genetic level permits precise control of their sequence and length, which dictates their thermal properties. Elastin-like polypeptides are used in a variety of applications including biosensing, tissue engineering, and drug delivery, where the transition temperature and biopolymer architecture of the ELP can be tuned for the specific application of interest. Furthermore, the lower critical solution temperature phase transition behavior of elastin-like polypeptides allows their purification by their thermal response, such that their selective coacervation and resolubilization allows the removal of both soluble and insoluble contaminants following expression in Escherichia coli. This approach can be used for the purification of elastin-like polypeptides alone or as a purification tool for peptide or protein fusions where recombinant peptides or proteins genetically appended to elastin-like polypeptide tags can be purified without chromatography. This protocol describes the purification of elastin-like polypeptides and their peptide or protein fusions and discusses basic characterization techniques to assess the thermal behavior of pure elastin-like polypeptide products.

Non-chromatographic Purification of Recombinant Elastin-like Polypeptides and their Fusions with Peptides and Proteins from Escherichia coli

Elastin-like polypeptides are stimulus-responsive biopolymers with applications ranging from recombinant protein purification to drug delivery. This protocol describes the purification and characterization of elastin-like polypeptides and their peptide or protein fusions from Escherichia coli using their lower critical solution temperature phase transition behavior as a simple alternative to chromatography.

Elastin-like polypeptides are repetitive biopolymers that exhibit a lower critical solution temperature phase transition behavior, existing as soluble ...

Amplification, Next-generation Sequencing, and Genomic DNA Mapping of Retroviral Integration Sites

Retroviruses exhibit signature integration preferences on both the local and global scales. Here, we present a detailed protocol for (1) generation of diverse libraries of retroviral integration sites using ligation-mediated PCR (LM-PCR) amplification and next-generation sequencing (NGS), (2) mapping the genomic location of each virus-host junction using BEDTools, and (3) analyzing the data for statistical relevance. Genomic DNA extracted from infected cells is fragmented by digestion with restriction enzymes or by sonication. After suitable DNA end-repair, double-stranded linkers are ligated onto the DNA ends, and semi-nested PCR is conducted using primers complementary to both the long terminal repeat (LTR) end of the virus and the ligated linker DNA. The PCR primers carry sequences required for DNA clustering during NGS, negating the requirement for separate adapter ligation. Quality control (QC) is conducted to assess DNA fragment size distribution and adapter DNA incorporation prior to NGS. Sequence output files are filtered for LTR-containing reads, and the sequences defining the LTR and the linker are cropped away. Trimmed host cell sequences are mapped to a reference genome using BLAT and are filtered for minimally 97% identity to a unique point in the reference genome. Unique integration sites are scrutinized for adjacent nucleotide (nt) sequence and distribution relative to various genomic features. Using this protocol, integration site libraries of high complexity can be constructed from genomic DNA in three days. The entire protocol that encompasses exogenous viral infection of susceptible tissue culture cells to integration site analysis can therefore be conducted in approximately one to two weeks. Recent applications of this technology pertain to longitudinal analysis of integration sites from HIV-infected patients.

We describe a protocol for amplifying retroviral integration sites from the genomic DNA of infected cells, sequencing the amplified virus-host junctions, and then mapping these sequences to a reference genome. We also describe techniques to quantify the distribution of integration sites relative to various genomic annotations using BEDTools.

Retroviruses exhibit signature integration preferences on both the local and global scales. Here, we present a detailed protocol for (1) generation of ...

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules derived from the parental cells. EVs function as intra- and inter-species communication vectors while also contributing to the interaction between bacteria and host organisms in the context of infection and colonization. Given the multitude of functions attributed to EVs in health and disease, there is a growing interest in isolating EVs for in vitro and in vivo studies. It was hypothesized that the separation of EVs based on physical properties, namely size, would facilitate the isolation of vesicles from diverse bacterial cultures. 
The isolation workflow consists of centrifugation, filtration, ultrafiltration, and size-exclusion chromatography (SEC) for the isolation of EVs from bacterial cultures. A pump-driven tangential flow filtration (TFF) step was incorporated to enhance scalability, enabling the isolation of material from liters of starting cell culture. Escherichia coli was used as a model system expressing EV-associated nanoluciferase and non-EV-associated mCherry as reporter proteins. The nanoluciferase was targeted to the EVs by fusing its N-terminus with cytolysin A. Early chromatography fractions containing 20-100 nm EVs with associated cytolysin A - nanoLuc were distinct from the later fractions containing the free proteins. The presence of EV-associated nanoluciferase was confirmed by immunogold labeling and transmission electron microscopy. This EV isolation workflow is applicable to other human gut-associated gram-negative and gram-positive bacterial species. In conclusion, combining centrifugation, filtration, ultrafiltration/TFF, and SEC enables scalable isolation of EVs from diverse bacterial species. Employing a standardized isolation workflow will facilitate comparative studies of microbial EVs across species.

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Bacteria secrete nanometer-sized extracellular vesicles (EVs) carrying bioactive biological molecules. EV research focuses on understanding their biogenesis, role in microbe-microbe and host-microbe interactions and disease, as well as their potential therapeutic applications. A workflow for scalable isolation of EVs from various bacteria is presented to facilitate standardization of EV research.

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules ...