Nuclei were prepared as previously described (McKittrick et al. 2004) and chromatin was extracted by modification of our previous procedure (Mito et al. 2005). Briefly, cells were suspended in TM2 (10mM Tris pH7.4, 2mM MgCl2, 0.5mM pmsf) with gentle vortexing to a concentration of 1-2x108/ml, held 1' in ice-water, and 10% NP-40 was added with gentle vortexing to 1.5%. After 2-5min on ice, crude nuclei were pelleted at 800 rpm in a Sorvall SS-34 rotor at 4oC, the supernatant was carefully removed, and the pellet was transferred with cold TM2 to a 2ml microfuge tube. Nuclei were spun in an Eppendorf microcentrifuge at 1000 rpm and the pellet was resuspended in TM2 to an estimated concentration of 0.5-1x109/ml for MNase digestion. CaCl2 was added to 1mM, the tubes were warmed to 37oC for 1', then digested by addition of MNase (Sigma 1.6U/ml) for 10'. Tubes were transferred to ice and reactions were stopped by addition of 0.2M EGTA (Na+) to 2mM. Nuclei were pelleted 10' at 2000 rpm 4oC, washed in TM2 and resuspended in 0.7ml cold 80mM or 150mM buffer: 140mM (or 70mM) NaCl, 10mM Tris 7.4, 2mM MgCl2, 2mM EGTA, 0.1% Triton X-100, 0.5mM pmsf pH7.5. After 1-2 hr rotation on a nutator at 4oC, nuclei were pelleted 10' at 2000rpm, and the pellet was gently resuspended in 0.7ml cold 600mM NaCl, 10mM Tris 7.4, 2mM MgCl2, 2mM EGTA, 0.1% Triton X-100, 0.5mM pmsf pH7.5. Supernatants from each extraction were clarified by spinning 2' at 13,200 rpm and held on ice to be used for pulldowns with aliquots taken for DNA and protein analyses (80mM, 150mM and 80-150mM extractions). The 600mM resuspended pellet was rotated overnight at 4oC, pelleted 10' at 2000rpm, and the supernatant was clarified and held on ice (80-600mM and 150-600mM extractions). For some experiments, MNase treatment was stopped by addition of EDTA (Na+ at pH 8) to 10mM, and after pelleting and saving the supernatant, nuclei were suspended in 0.6ml 10mM Tris 0.25mM EDTA pH7.4. The suspension was passed five times through a 26-gauge needle and combined with the supernatant (EDTA-needle extraction) (Jin and Felsenfeld 2007). For experiments described in Fig. 3D, DNAs extracted from samples prepared in this way were electrophoresed and the mononucleosome band was excised, and the DNA was extracted using the Qiagen Qiaquick kit. Affinity purification was performed by addition of streptavidin-Sepharose and rotating for 2hrs at 4¡C as described (Mito et al. 2005), except that the same buffer (without Triton) was used for washing the resin, which was brought up in 0.2ml 150mM buffer (without Triton) and (usually) frozen at -20oC prior to extraction. Aliquots of MNase-digested nuclei and clarified soluble nucleosomes were also frozen prior to extraction. DNA was purified from chromatin fractions and pulldown samples as described (Mito et al. 2005). Briefly, samples were RNAse-treated, deproteinized with SDS and proteinase K, extracted using a mixture of phenol/chloroform/isoamyl alcohol (25:24:1) followed by chloroform extraction, and DNA was precipitated with 2 vol ethanol. References: Jin, C. and Felsenfeld, G. 2007. Nucleosome stability mediated by histone variants H3.3 and H2A.Z. Genes Dev 21: 1519-1529. McKittrick, E., Gafken, P.R., Ahmad, K., and Henikoff, S. 2004. Histone H3.3 is enriched in covalent modifications associated with active chromatin. Proc Natl Acad Sci USA 101: 1525Ð1530. Mito, Y., Henikoff, J., and Henikoff, S. 2005. Genome-scale profiling of histone H3.3 replacement patterns. Nat Genet 37: 1090-1097.
Label
Cy5
Label protocol
Strand-displacement labeling for NimbleGen arrays Steve Henikoff 5/2008. This is a modification of the NimbleGen protocol for ChIP-chip labeling (http://www.nimblegen.com/). It yields adequate amounts of end-labeled material without PCR amplification starting with ? 0.2 ug fragmented DNA. 1) In a 0.5ml PCR tube, mix up to 20uL ds DNA with 20uL either Cy3- or Cy5-labeled 9mers in random primer buffer (Cy3 and Cy5 9mer Wobble from TriLink are mixed according to the NimbleGen protocol, aliquoted and stored frozen). Use 0.2-3ug DNA, the more the better, and add water to 20uL. Protect the dye-labeled material from direct light, especially Cy5, throughout the procedure. 2) Heat 98oC 10 min. Quick chill in ice-water (this is important). 3) While the samples are denaturing, make a master mix with Klenow exo- (NEB 50,000u/ml) and dNTPs. For example, for 10 samples mix 42uL water, 10.5uL Klenow, 52.5uL 10mM dNTPs (aliquoted and stored frozen). While samples are still in the ice-water bath and chilled, add 10uL master mix to each tube with 10 or more strokes of the Pipetteman tip. 4) Following a brief spin, transfer tubes to a 37oC cycler. Incubate overnight (16 hr). 5) Prepare 1.5ml tubes with NaCl+EDTA (equivalent of 5uL 0.5M EDTA, 5.7uL 5M NaCl, which can be added together). Remove the tubes from the cycler and transfer contents from the cycler to these 1.5 ml tubes. Add 60uL 2-propanol (the volume is important). Let sit in dark 10' at room temperature. 6) Spin 13.2krpm 10 room temperature. Carefully decant the supernatant from the deeply colored pellet with a P200. Add 500uL ice-cold 80% ethanol and spin briefly in the cold. Carefully pour or pull off the ethanol - large pellets tend to dislodge. Quick spin and pull off the excess ethanol. Let air dry (~10min room temperature). 7) Add 20uL water to dissolve. Note that the deepness of the red (Cy3) or blue (Cy5) is a good measure of the yield. For samples in which there was little starting material, and/or the DNA was small, there will not be enough for an array, and the color will be faint. For samples with deep color, add an additional 30uL water. Measure OD260 and calculate yield. Store at 4oC for brief periods or at -20oC longer term in dark. 8) For sample with less than 30ug/array, do another round of labeling, using 20uL of the labeled DNA from the first round prep. The second round is much more efficient than the first round and consistently yields ? 60ug. 9) Check sizes on a 1.5% gel without ethidium. Cy3 fluoresces well enough to see 1 ug, but Cy5 is much fainter and might not be visible. DNA should be about the same average size as the starting material. Follow with ethidium staining: DNA typically smears up towards the top of the gel, but this material is evidently low in molar amount of label and does not appear to cause a problem. 10) For each 2.1M array, mix 30ug Cy3-labeled DNA with 30ug Cy5-labeled DNA and speed-vac dry on low or medium. If Cy3 is input DNA then Cy5 is pull-down DNA and vice versa. 11) Reduce volume by speedvac on low to 12.3 ML, or lyophilize samples for hybridization to NimbleGen arrays using standard NimbleGen hybridization protocol.
Strand-displacement labeling for NimbleGen arrays Steve Henikoff 5/2008. This is a modification of the NimbleGen protocol for ChIP-chip labeling (http://www.nimblegen.com/). It yields adequate amounts of end-labeled material without PCR amplification starting with ? 0.2 ug fragmented DNA. 1) In a 0.5ml PCR tube, mix up to 20uL ds DNA with 20uL either Cy3- or Cy5-labeled 9mers in random primer buffer (Cy3 and Cy5 9mer Wobble from TriLink are mixed according to the NimbleGen protocol, aliquoted and stored frozen). Use 0.2-3ug DNA, the more the better, and add water to 20uL. Protect the dye-labeled material from direct light, especially Cy5, throughout the procedure. 2) Heat 98oC 10 min. Quick chill in ice-water (this is important). 3) While the samples are denaturing, make a master mix with Klenow exo- (NEB 50,000u/ml) and dNTPs. For example, for 10 samples mix 42uL water, 10.5uL Klenow, 52.5uL 10mM dNTPs (aliquoted and stored frozen). While samples are still in the ice-water bath and chilled, add 10uL master mix to each tube with 10 or more strokes of the Pipetteman tip. 4) Following a brief spin, transfer tubes to a 37oC cycler. Incubate overnight (16 hr). 5) Prepare 1.5ml tubes with NaCl+EDTA (equivalent of 5uL 0.5M EDTA, 5.7uL 5M NaCl, which can be added together). Remove the tubes from the cycler and transfer contents from the cycler to these 1.5 ml tubes. Add 60uL 2-propanol (the volume is important). Let sit in dark 10' at room temperature. 6) Spin 13.2krpm 10 room temperature. Carefully decant the supernatant from the deeply colored pellet with a P200. Add 500uL ice-cold 80% ethanol and spin briefly in the cold. Carefully pour or pull off the ethanol - large pellets tend to dislodge. Quick spin and pull off the excess ethanol. Let air dry (~10min room temperature). 7) Add 20uL water to dissolve. Note that the deepness of the red (Cy3) or blue (Cy5) is a good measure of the yield. For samples in which there was little starting material, and/or the DNA was small, there will not be enough for an array, and the color will be faint. For samples with deep color, add an additional 30uL water. Measure OD260 and calculate yield. Store at 4oC for brief periods or at -20oC longer term in dark. 8) For sample with less than 30ug/array, do another round of labeling, using 20uL of the labeled DNA from the first round prep. The second round is much more efficient than the first round and consistently yields ? 60ug. 9) Check sizes on a 1.5% gel without ethidium. Cy3 fluoresces well enough to see 1 ug, but Cy5 is much fainter and might not be visible. DNA should be about the same average size as the starting material. Follow with ethidium staining: DNA typically smears up towards the top of the gel, but this material is evidently low in molar amount of label and does not appear to cause a problem. 10) For each 2.1M array, mix 30ug Cy3-labeled DNA with 30ug Cy5-labeled DNA and speed-vac dry on low or medium. If Cy3 is input DNA then Cy5 is pull-down DNA and vice versa. 11) Reduce volume by speedvac on low to 12.3 ML, or lyophilize samples for hybridization to NimbleGen arrays using standard NimbleGen hybridization protocol.
Hybridization protocol
Hybridization protocol provided and performed by NimbleGen (http://www.nimblegen.com/products/chip/index.html). Set MAUI hybridization unit to 42 deg C and allow time for the temperature to stabilize. Based on the A_260 measurement, combine 13 micrograms each of the Test and Reference Samples into a single 1.5 ml microcentrifuge tube. Protect tube from light during handling to prevent photobleaching of the light-sensitive Cy dyes. Dry the combined contents in a Speed-Vac on low heat. Resuspend the sample in 10.9microliter VWR water and vortex to completely dissolve the sample. Spin the tube down briefly to collect the contents in the bottom. Using the NimbleGen Array Reuse Kit, add the following to the resuspended sample: 19.5 microliter 2X Hybridization Buffer, 7.8 microliter Hybridization Component A, 0.4 microliter Cy3 CPK6 50mer Oligo (50nM), 0.4 microliter Cy5 CPK6 50mer Oligo (100nM). CPK6 oligos are included in the hybridization as controls that hybridize to alignment features on the NimbleGen arrays. They are required for proper extraction of array data from the scanned image. Mix the tube briefly then spin down to collect the contents in the bottom and place at 95 deg C for 5 minutes. Immediately transfer the tube to the MAUI 42 deg C sample block and hold at this temperature until you are are ready for sample loading. Place the MAUI Mixer SL Hybridization Chamber on the array using the provided assembly/disassembly jig and carefully follow MAUI setup instructions. Use the braying tool to remove all air bubbles from the adhesive gasket around the outside of the hybridization chamber. Put the array and hybridization chamber on the MAUI and allow 30 seconds for the chips to come up to temperature. Load the sample using the pipet supplied with the MAUI Station and following manufacturer instructions. During the loading, a small amount (3-7 microliter) of the sample may flow out of the outlet port. Confirm that there are no bubbles in the chamber. If there are, very gently massage any bubbles to either of the ends, away from the center of the array. Avoid applying too much pressure since this will force liquid out of the ports. Place the loaded array into one of the four MAUI bays and let equilibrate for 30 seconds. Wipe off any sample leakage at the ports with a Kim-Wipe, and adhere MAUI stickers to both ports. Close the bay clamp and select mix mode B. Hold down the mix button to start mixing. Confirm that the mixing is in progress before closing the cover. Hybridize the sample 16-20 hours. Prior to removing the array from the MAUI Hybridization Station, prepare the following solutions. You will need two 250ml dishes of Wash I, and one each for Wash II and Wash III. One dish for Wash I should be shallow and be wide enough to accomodate the array and mixer loaded in the MAUI assembly/dissassembly jig. The lid from a 1000 microliter pipet tip box works well. Also, the buffer in the first Wash I dish should be heated to 42 deg C to help soften the adhesive on the hybridization chamber; this will help prevent braking the array. Place the remaining three wash solutions in 300 microliter Tissue-Tek slide staining dishes. Wash I: 225 ml VWR Water, 25ml 10X Wash Buffer I, 25microliter 1M DTT. Wash II: 225ml VWR Water, 25ml 10X Wash Buffer II, 25microliter 1M DTT. Wash III: 225ml VWR Water, 25microliter 10X Wash Buffer III, 25microliter 1M DTT. Remove chip from MAUI Hybridization Station, load it back into the MAUI assembly/disassembly jig, and immerse in the shallow 250ml Wash I at 42 deg C. Peel the hybridization chamber off very slowly to prevent the slide from cracking. Do not let the surface of the slide dry out at any point during washing. While the chip is submerged, carefully peel off the mixer. Gently agitate the chip in Wash I for 10-15 seconds. Transfer the slide into a slide rack in the second dish of Wash I and incubate 2 minutes with agitation. Transfer to Wash II and incubate 1 minute with agitation. Rock the disk to move the wash over the tops of the arrays. Transfer to Wash III and incubate for 15 sec with agitation. Remove array and spin dry in array-drying unit for 1 minute. Store the dried array in a dark desiccator.
Scan protocol
Sample scan protocol provided and performed by NimbleGen (http://www.nimblegen.com/products/chip/index.html). Axon Scanner (model 4000B). Turn on the power of the scanner. The switch is on the back right hand side. Launch GenePix Software 10 minutes before scanning to allow lasers to warm. Open the scanner door and open the side carriage. Place chip in carriage so that array is face down and barcode end is closest to you. Move the black lever on the left side of the carriage left until the array is lying flat in the carriage. Release the lever so that the arry is pushed over gently to the right side of the carriage and held firmly. Close the carriage (you should hear a click) and slide the scanner door shut. Open the hardware settings. Select the following settings for scanning: Wavelength (532=Cy3, 635=Cy5). Set the 532 laser PMT Gain to 650 and the 635 laser PMT Gain to 750. Power=100%. Pixel Size=5m. Lines to average=1. Focus position=0micrometer. Select Image/Ratio to view both channels simultaneously. Perform a preview scan by clicking on the fast-forward icon. Stop scan once the full arry is in view by pressing the stop icon. Switch to zoom mode by clicking on the array image and pressing Z. Click and drag the mouse on the area in which you want to zoom. Center the image of the entire array area within the viewing window. Under Tools on the left side of the screen, click the Scan Area icon. Click and drag to define a box that bounds the visibile array area. The box dimensions should be stretched to create a box just slightly larger than the array area. It is critical to include all of the corner probes within the scan image. Scan chip by clicking the play icon. While scanning, set the zoom level to view the whole image and adjust the brightness and contrast of the displayed image to eliminate visible saturation. Based on the appearance of the array features, adjust the PMT setting as appropriate. The features should be mostly yellow. If your features are mostly green, either decrease the 532 PMT or raise the 635 PMT. If the features are mostly red or orange, either raise the 532 PMT or decrease the 635 PMT. Zoom the view to a region scanned under the most recent PMT settings and click on the Histogram tab at the top of the left side of the screen to check global intensity of the features. On the top left side of the screen make sure you have the 532 and 635 wavelength boxes checked so both wavelength histograms are displayed. Make sure the Log Axis box is checked. You want the red and green curves to be as close as possible to one another. If the red curve is above the green, lower the red PMT setting or raise the green PMT setting. You want the curve to have 1e-5 normalized counts at the 65,000 intensity level (saturation). This means that you have about 1% of the features saturated. The position of the two curves on top of each other is more important than hitting the 1% saturation mark. The histogram only graphs the area of the image that is viewable in the screen on the image tab. If the histogram is no longer changing then either the chip is done scanning or the area that you can see on the image screen has been scanned. After the PMT settings are properly adjusted, stop the current scan, do not save this image. Restart the scan under the new settings and wait until the scan has completed. Save the images. Make sure you save both the 532nm and 635nm images as separate single image tif files.
Description
Biotin-H2A pulldown of EDTA-needle-extracted chromatin
Data processing
Log base 2 (Cy5/Cy3) ratios were bi-weight mean centered
