Methods are described in publication Zilberman, et al. Nature Genetics 2007 (GSE5974).
Label
Cy3
Label protocol
Sample labeling protocol provided and performed by NimbleGen (http://www.nimblegen.com/products/chip/index.html). Prepare the following three solutions: 50X dNTP Mix, Random 9mer Buffer, 10X TE. Dilute Cy5 dye-labeled 9mers to 1 O.D./42 microliter Random 9mer Buffer. Aliquot to 40 microliter individual reaction volumes in 0.2 milliliter thin-walled PCR tubes and store at -20 degrees C. Assemble the following components in a 0.2 milliliter thin-walled PCR tube: 1 microgram LM-PCR Sample and 40 microliter Cy5-9mer Primers. Heat denature samples in thermocycler at 98 degrees C for 10 minutes. Quick chill in ice water bath. Snap-chilling step after denaturation is critical for efficiency labeling. Add the following: 10 microliters 50X dNTPmix, 8 microliters VWR deionized water, 2 microliters Kienow (50 U/microliter). Do not vortex after addition of Kienow. Mix well by pipetting 10 times. Spin down at low RPM to force contents to bottom of tube. Incubate at 37 degrees C for 2 hours in a thermocycler protected from light. Stop the reaction by addition of EDTA: 10 microliter Stop Solution (0.5 M EDTA). Transfer the reaction to a 1.5 milliliter tube. Precipitate the labeled samples by adding NaCl and isopropanol to each tube as indicated: 11.5 microliter 5M NaCl, 110 microliter Isopropanol. Vortex briefly after addition of 5M NaCl prior to addition of isopropanol. Vortex. Incubate 10 minutes at room temperature in the dark. Centrifuge at maximum speed for 10 minutes. Remove supernatant with pipette. Rinse pellet with 500 microliter 80% ice-cold ethanol. Dislodge pellet from tube wall. Centrifuge at maximum speed for 2 minutes. Remove supernatant with pipette. Speed-Vac on low heat for 5 minutes until dry, protected from light. Store labeled samples at -20 degrees C, protected from light. Rehydrate dried pellets in 25 microliter VWR deionized water. Spin tubes down briefly prior to opening. Vortex for 30 seconds and quick spin to collect contents at bottom of tube. Continue to vortex or let site in the dark at room temp until the pellet is completely rehydrated, then vortex again and quickly spin. Measure the A_260 in each sample. It is important to consume as little sample as possible when performing this measurement. Typical yields range from 10 to 30 micrograms per reaction.
Methods are described in publication Zilberman, et al. Nature Genetics 2007 (GSE5974).
Label
Cy5
Label protocol
Sample labeling protocol provided and performed by NimbleGen (http://www.nimblegen.com/products/chip/index.html). Prepare the following three solutions: 50X dNTP Mix, Random 9mer Buffer, 10X TE. Dilute Cy3 dye-labeled 9mers to 1 O.D./42 microliter Random 9mer Buffer. Aliquot to 40 microliter individual reaction volumes in 0.2 milliliter thin-walled PCR tubes and store at -20 degrees C. Assemble the following components in a 0.2 milliliter thin-walled PCR tube: 1 microgram LM-PCR Sample and 40 microliter Cy3-9mer Primers. Heat denature samples in thermocycler at 98 degrees C for 10 minutes. Quick chill in ice water bath. Snap-chilling step after denaturation is critical for efficiency labeling. Add the following: 10 microliters 50X dNTPmix, 8 microliters VWR deionized water, 2 microliters Kienow (50 U/microliter). Do not vortex after addition of Kienow. Mix well by pipetting 10 times. Spin down at low RPM to force contents to bottom of tube. Incubate at 37 degrees C for 2 hours in a thermocycler protected from light. Stop the reaction by addition of EDTA: 10 microliter Stop Solution (0.5 M EDTA). Transfer the reaction to a 1.5 milliliter tube. Precipitate the labeled samples by adding NaCl and isopropanol to each tube as indicated: 11.5 microliter 5M NaCl, 110 microliter Isopropanol. Vortex briefly after addition of 5M NaCl prior to addition of isopropanol. Vortex. Incubate 10 minutes at room temperature in the dark. Centrifuge at maximum speed for 10 minutes. Remove supernatant with pipette. Rinse pellet with 500 microliter 80% ice-cold ethanol. Dislodge pellet from tube wall. Centrifuge at maximum speed for 2 minutes. Remove supernatant with pipette. Speed-Vac on low heat for 5 minutes until dry, protected from light. Store labeled samples at -20 degrees C, protected from light. Rehydrate dried pellets in 25 microliter VWR deionized water. Spin tubes down briefly prior to opening. Vortex for 30 seconds and quick spin to collect contents at bottom of tube. Continue to vortex or let site in the dark at room temp until the pellet is completely rehydrated, then vortex again and quickly spin. Measure the A_260 in each sample. It is important to consume as little sample as possible when performing this measurement. Typical yields range from 10 to 30 micrograms per reaction.
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
Antibody-based methylation mapping of the Arabidopsis genome. Genomic DNA was extracted from wild type, met1-6 or pie1-5 lines in the Columbia background. The values are log2 signal ratios of immunoprecipitated DNA /input DNA.
Data processing
Log base 2 ratios were bi-weight mean centered by NimbleGen
