Genomic ssDNA mapping by microarray

Part I.  Sample collection and Preparation of Yeast DNA Embedded in Agarose Plugs
(the preparation of agarose plug embedded DNA is adapted from Iadonato, S. P., and A. Gnirke. 1996.  RARE-cleavage analysis of YACs. Methods Mol Biol 54: 75-85.)

Yeast cultures are grown in synthetic complete media at 25 oC or 30oC.  Log phase cells (at OD660 of 0.25) are synchronized in G1 by the addition of alpha factor at 200 nM or 3 mM (for bar1 and BAR1 strains respectively), followed by incubation for approximately one and a half generations, or until the unbudded cell population reaches at least 90%.  HU is added at 200 mM, and cells are released from the G1 arrest by the addition of pronase at 0.02 mg/ml or 0.3 mg/ml (for bar1 and BAR1 strains respectively).
Harvest the cells by adding them to test tubes containing NaN3 on ice such that the final concentration of NaN3 will be 0.1%.  I typically collect 50 ml of each time point sample to make 3 plugs of ~100 ml each.  Scale up for the 0 time point control sample if multiple S phase samples are collected.
Spin down cells at 3000 x rpm in the floor centrifuge for 5 min. Wash the cell pellet twice with 50 mM EDTA, spinning 5 min at 3000 x rpm as above.
Resuspend the cells in 50 mM EDTA to a final concentration of 2 x 109 cells/ml and warm the cell suspension at 45°C for 5 min. Add an equal volume of 1% low-melt InCert (or SeaPlaque) agarose in 50 mM EDTA, also prewarmed to 45°C . (This procedure will make 0.5% plugs, which are fairly fragile, but are better if further manipulations like in gelo digest are required. If not, use 2% agarose to make 1% plugs, which are less fragile). Mix the suspension thoroughly by vortexing and pipette 100 µl aliquots into each plug mold to harden.  Plugs may be allowed to set at room temperature or placed at 4°C (for 15 minutes).
Extrude each plug from the plug mold into a six-well dish, up to 3 plugs per well. To each well add 6 ml of freshly prepared spheroplasting solution. Incubate at 37°C for 4 h with gentle shaking.
Aspirate off the spheroplast solution from each well and add 6 ml of LDS solution. Incubate with gentle shaking at 37°C for at least 15 min. Remove and add fresh LDS solution. Incubate with gentle shaking at 37°C overnight.
Wash the plugs 2 x 30 min with 6 ml of 0.2x NDS solution with gentle shaking at room temperature.
Wash the plugs 3 x 30 min with 6 ml of TE, pH 8.0 with gentle shaking at room temperature. Plugs may be stored at 4°C in six-well dishes with TE, pH 8.0, covered with Saran wrap to prevent excessive evaporation. High-molecular-mass DNA will usually remain undegraded for at least six months.
Refer to the CHEF gel apparatus manual for suggested parameters. To visualize all the yeast chromosomes, we use a switch time ramped from 60-120s, 200V, 24 hours. To obtain better separation of the smaller chromosomes, we use a switch time ramped from 35-70s, 200V, 22 hrs.
Spheroplasting Solution
40 ml 1M Sorbitol (approx. 1M final conc.)
1.6 ml 0.5M EDTA, pH 8.0 (20mM final)
0.4 ml 1M Tris-HCl, pH7.5 (10mM final)
40 µl 2-mercaptoethanol (14mM final)
0.5 mg/ml Zymolyase 20-T

NDS Solution
0.5 M EDTA
10 mM Tris base
1% Sarkosyl
(pH 9.5)
To 350 ml H2O add 93 g Na2EDTA•2H2O and 0.6g Tris base
Adjust the pH to greater than 8.0 with 100 to 200 pellets of solid NaOH 
Add 50 ml of 10% N-lauryl sarcosine
Adjust the pH to 9.5 with concentrated NaOH
Bring the final volume to 500 ml with H2O
Filter-sterilize and store at room temperature.

LDS Solution
1% lithium dodecyl sulfate
100 mM EDTA
10 mM Tris-HCl, pH 8.0
Per 1 liter:
10 g lithium dodecyl sulfate (Sigma Chemical Cat #L-4632)
200 ml 0.5M EDTA, pH8.0
10 ml 1M Tris-HCl, pH8.0
Bring volume to 1 liter with H2O, filter-sterilize, and store at room temperature

Part II.  Random-primed in-gel labeling of ssDNA by Klenow
The steps from here on involve working with Cy dyes which are light-sensitive.  Try to minimize exposure to light throughout the procedures.

In-gel labeling of ssDNA by Klenow:
a.        Make 1X reaction buffer (I usually make 40 – 50 ml, 5 ml per wash per agarose plug of 100 ul volume):
50 mM Tris-HCl pH 6.8
5 mM MgCl2
10 mM BME
(do not include hexamers in this step, too expensive)
b.       Remove agarose plugs to new dishes (6-well plates, one plug per well)
c.        Add 5 ml TE 0.1 to each well and incubate on rotator at RT for 15 min
d.       Aspirate buffer.  Repeat c.
e.        Add 5 ml 1X reaction buffer and incubate on rotator at RT for 30 min
f.         Aspirate buffer.  Repeat e.
g.        During the last wash step, prepare labeling reaction mix:
Estimate gel volume approx. 50 ul
Mix (50 ml) contains:
10 μl     10X dNTPs (from the ArrayBox, this mix contains 1.2 mM dATP, dCTP and dGTP but 0.6 mM dTTP, IMPORTANT!!!)
20 μl    2.5X reaction buffer (from the ArrayBox)
6 μl       Cy5 or Cy3 (Perkin Elmer)
3 μl       Klenow (10,000units/ml, NEB M0212M Klenow Fragment, exo-)
11 μl    H2O
h.       Assemble a small water bath.  Use a 1ml Blue tip box, remove insert, place a CLEAN small gel tray inside, add approx. 50 ml of agd H2O or enough to submerge the gel tray legs halfway.  Cut a piece of parafilm and use gloved hands to place it on the gel tray.  You could mark the positions/sample names on the edges of the parafilm to indicate where the plugs go.
i.         Use a CLEAN spatula, move the plugs onto the parafilm according to the arranged positions.
j.         Pipet 50 μl of reaction mix onto each agarose plug slowly.  I try to let the liquild hover on top of the agarose but you may get some overflow on the side.  However, surface tension will hold the liquid near the plugs. 
k.        Incubate at 37oC (in the dark) for 2-3 hours.
AgarAce digestion to recover DNA:
a.        Transfer band to empty 1.5ml tube and weigh
b.       Melt at 70°C for 10 min in water bath
c.        Spin max speed for 1 min at RT
d.       Transfer to 47°C heat block or water bath for 1min
e.        Add ½U (2λ) Promega agarase (AgarAceTM) per 200 mg molten agarose and mix
f.         Incubate at 47°C for 1hr
g.        Split into 500 μl aliquots, ice for 5 min, and spin at max speed for 20 min at 4°C
h.       Recover supernatant
Sonicate and Clean-up DNA:
a.       Estimate volume of the liquid.  If a sample volume is greater than 300 μl, aliquot into multiple tubes to have maximum volume of 300 μl.  Try to aliquot into equal volumes for a given sample.
b.       Sonicate sample to shear DNA fragments using the Bioruptor in the Hanes lab.  Use 20 cycles, 30 sec on and 30 sec off, on “H” (high) setting.
c.        Use Qiagen PCR cleanup kit to clean the samples (follow instructions).  Pool all aliquots of the same sample into one column.  You might have to load and spin the column multiple times to get all the volume in.
d.       Elute each sample with 44 μl of EB.
e.       Samples can be stored at 4oC or proceed to microarray hybridization.

Solutions needed:
2.5x reaction buffer for Klenow
            125 mM         Tris-HCl pH 6.8
            12.5 mM         MgCl2
            25 mM            beta-mercaptoethanol
            750 mg/ml     random hexamers
 10x dNTP mix
            1.2 mM each dATP, dCTP, dGTP
            0.6 mM           dTTP
            10 mM            Tris-HCl pH 8.0
 random hexamers (IDT)
MW = 1,791.7
1 mM stock = 1.79 mg/ml

Part III.  Agilen microarray hybridization and scanning

This protocol is adapted from the Dunham lab protocol which in turn was adapted from the Brown and deRisi lab protocols and various Agilent 60-mer oligo microarray processing (SSPE wash) protocols.  The arrays we use are 4x44K platform yeast ChIP to chip arrays.  There are 4 genomes per slide and therefore can hold 4 experiments per slide.  Array 1 is the nearest to the barcode and array 4 is the farthest from the barcode.

Microarray Slide Hybridization

Final hybridization reaction is composed of:
            Mixed Cy5- and Cy3-labeled DNA and H2O          44 μl
            10x Agilent Blocking Agent (warm to RT)              11 μl
            2x Hi-RPM hybridization buffer                              55 μl
            total volume                                                            110 μl
            volume to load on slide                                          100 μl

1.     Add H2O to DNA to bring the total volume to 44 μl
2.     Add 11 μl Agilent 10x Blocking Agent (prepare 10x Blocing Agent according to Agilent instructions, warm to RT before use)
3.     Incubate at 95 to 100°C in heating block for 5 min (process only 4 samples at a time)
4.     Cool at RT for 5 min
5.     Add 55 μl 2x Hi-RPM hybridization buffer, mix by pipetting, avoid making bubbles
6.     Pulse spin
7.     Load 100 μl onto the gasket slide: Place a backing slide, Agilent side up, in a hybridization chamber; pipet 110 μl of DNA, avoiding bubbles, onto the center of one gasket area.  Do not eject the last 1 or 2 μl to avoid bubbles.  Spread it around as you pipet but do not touch the slide with tip.  Repeat for the other 3 gasket areas.
8.     Remove the array slide from the storage box.  The Agilent side is the array side, so it should face down and make contact with the DNA solution.  Carefully lower the array over the gasket slide keeping it flat.
9.     Once the array is resting on the gasket slide, place the top of the hybridization chamber on top and slide the screw over the assembly.  Tighten the screw all the way down, finger tight.
10. Look through the back of the chamber and rotate the assembled slide.  There should be a single big bubble that moves freely.  You might get some small bubbles too but there’s nothing you can do at this point.  Just make sure they are all freely moving.
11. Put the hybridization chamber on the rotisserie in the 65°C oven, making sure to balance.
12. Hybridize for 17 hrs at 65°C at 20 rpm.

Slide Washing
(Turn on scanner to warm up lasers first)

Prepare wash solutions (filter before use):
Wash A (1 L):
Add in this order:
            700 ml            agd H2O
            300 ml            20x SSPE (Invitrogen)
            0.25 ml           20% N-Lauroylsarcosine

Wash B (1 L):
Add in this order:
            997 ml            agd H2O
            3 ml                20x SSPE
            0.25 ml           20% N-Lauroylsarcosine

20x SSPE:
3M NaCl
200mM sodium phosphate
in glass-distilled H2O, pH 7.4
Solution is 0.2 µm filtered

1.     Rinse the glass dishes, racks and stir bars with water.  Need 3 glass dishes (for washes) and 1 glass jar (for disassembling the hybridization chamber and releasing the slides).
2.     Set up two stir plates near the hood and one inside the hood, each with a glass dish and a stir bar inside the dish.  In the first dish, place a rack.
3.     Add Wash A to the 1st dish, Wash B to the 2nd dish and acetonitrile to the 3rd dish. Add enough liquid so that the entire slide can be submerged.  The stirring liquid must be visibly turbulent.
4.     Disassemble the hybridization chambers one at a time.  Use a pair of plastic tweezers to gently wedge open the sandwich while submerging in Wash A in the glass jar.
5.     Transfer the array slide to the rack in the 1st dish with Wash A.  Add other slides to rack and leave ample space between slides.
6.     Once all the slides are in the rack, time the wash for 1 min.
7.     Transfer the rack immediately into the 2nd dish with Wash B, time the wash for 1 min.
8.     Transfer the rack into the 3rd dish with acetonitrile, dabbing on some kimwipes to dry just before transfer.  Time the rinse for 30 sec.
9.     Slowly and evenly pull the rack out of the acetonitrile.  The slides should be clean and dry.  If you see droplets on the slides then submerge them and try again.
10. Set the rack on some kimwipes.
11. Load the slides into scanning holders with Agilent side up (the scanner scans through the back of the slide) and barcode sticking out.  Blot excess acetonitrile with kimwipe if necessary.  Do not touch anywhere but the edges and the barcode.
12. Scan no more than 5 slides at a time to avoid ozone in the scanner.  You can reuse the wash buffers for more slides.  Just replace the first Wash A in the glass jar for every batch.

1.     Open the Agilent scan control program.
2.     Remove the caroussel lid gently.
3.     Place the slide holders in the caroussel, noting the slot numbers.
4.     Select the appropriate slot numbers from the pulldown menus on the upper left.
5.     Select the directory column and click Edit Values.  Browse to find the directory you want to save the files in.  Click “set”.  The values should change.
6.     Check the default preferences for the correct scanning area (61x21.6 mm), resolution (5 mm), laser power (100% PMT) and with the split and rotate boxes unchecked.
7.     Scan.
Setting used at UW:
Profile: Agilent HD_CGH
Resolution: 5 mm
TIFF: 16 bit
G PMT: 100
R PMT: 100
XDR: 0.33

Feature Extraction
1.     Open the Feature Extraction software.
2.     Add the tif image to the project.
3.     Find the grid template file for the slide and run the appropriate protocol.  For the ChIP to chip 4x44K slides (G4493A) the template file is 014810_D_20060718.  To check which template file to use, go to Agilent website at and enter the array barcode. 
Note: The custom designed Crick array (G4497A) uses the template file 034056_D_F_20110503
4.     Check the visual results to make sure it looks good.  Check alignment and make sure that the flagging is sensible.  If you get a larger than usual number of outliers, make a note of it.
5.     If the alignment was incorrect, need to manually align the template and re-extract data.
-       Open a project
-       Open the image file
-       Click on the Grid template icon on far right
-       Adjust Main Grid by moving the grid so that it sits on top of the spots
-       Click on the Adjust Sub Grid icon
-       Adjust Sub Grid as well
-       Save the grid template file as “****_grid.csv”.  Do not add letters between “grid” and “csv”.
-       Go back to the Extraction Browser and under the “Template”, select the newly created grid file
-       Select the protocol file “ChIP_v1_95_May07” 
-     Start extraction

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