Yeast homemade array CGH
Maitreya Dunham September
2007
hybrid of Brown and deRisi
lab protocols and Agilent 60-mer oligo microarray processing protocols
This labeling protocol
seems to be rather sensitive to DNA quality. I use the Winston and Hoffman prep with good results. If you're worried about quality, use
the Zymo genomic DNA kit, or a Qiagen kit. The Zymo kit can also be used to make DNA from colonies.
Measure starting DNA
concentration with a fluorometer.
Once it's been purified with the Zymo columns, the spectrophotometer
seems to be reasonably accurate.
Bring 4 µg DNA to 100µl in
a 1.5mL eppendorf tube.
Sonicate 30 pulses power
level 1, 1 second on/1 second off.
Run 2 µl on a gel to check
fragmentation. Fragments should be
a smear around 1 kb.
Purify with zymo DNA clean
and concentrator 5 kit.
Elute in 20 µl water.
Nanodrop 1 µl.
DNA can be stored -20C.
Bring 2 µg to 21µl with
water.
Add 20 µl 2.5X random
primer/reaction buffer mix:
|
STOCKS: |
1X: |
125mM Tris pH 6.8 |
0.5M Tris, pH 6.8 |
5 µl |
12.5mM MgCl2 |
50mM MgCl2 |
5 µl |
25mM 2-mercaptoethanol |
0.143M 2-mercaptoethanol
(dilute fresh) |
3.5 µl |
750µg/mL random nonamers |
5µg/µl random nonamers |
3 µl |
|
Water to 20 µl |
3.5 µl |
95¡C 5 minutes.
Ice 5 minutes.
Add 5µl 10X dNTP mix (in
TE, pH 8.0):
For dCTP Cy dyes: 1.2mM each dATP, dGTP, dTTP, and 0.6mM
dCTP
For dUTP Cy dyes: 1.2mM each dATP, dGTP, dCTP, and 0.6mM
dTTP
Add 3µl appropriate
Cy-dNTP.
Add 1µl Klenow (5U/µl).
Incubate 37C for 2 hours.
Add 5µl 0.5M EDTA pH 8.0.
Ice.
Purify using zymo columns
with 0.5 ml binding buffer.
Elute in 25 µl water.
Nanodrop 1 µl to check
yield and dye incorporation.
Dye incorporation should
be ~20 pmol/µg.
Mix 1 µg each Cy3 and Cy5 reactions. Bring to 160 ml total with water.
Add 40 ml 10X Agilent block.
Stagger the following steps if you are doing a lot of arrays.
Incubate 95C 5 minutes.
Incubate RT 5 minutes.
Add 200 ml Agilent 2X hybe buffer.
You can start adding the hybe solution while the samples are cooling.
Place a single array
backing slide, Agilent side up, in a hybe chamber.
Pipet the whole volume of
probe, avoiding bubbles, onto the center of the gasket area. Don't eject the last µl or two in order
to avoid bubbles. Spread it around
as you pipet, but not too close to the gasket.
Remove a crosslinked and
blocked array from the box. The
barcode side is the array side, and so should face down, onto the probe. Carefully lower the array over the
gasket slide, keeping it flat.
Once the array is resting
on the gasket slide, place the top of the hybe chamber, and slide the screw
over the assembly. Tighten the
screw all the way down, finger tight.
Look through the back of
the chamber and quickly rotate the slide to wet all the gaskets. There should be one big bubble that
moves freely. There may be one big
bubble and a couple of little ones stuck to the sides. If they are small and isolated, don't
worry too much about them. You
will probably do more harm than good trying to remove them. If they seem like they'll interfere,
you can try knocking the array with the heel of your hand to dislodge them.
Put the array in the hybe
oven. Make sure to balance the
rotisserie.
Hybe
65C overnight at
rotation setting 4.
Prepare
for the washes by placing a bottle with 500 ml Wash 1 in the 65C waterbath.
Washes
Prepare your wash
solutions. All solutions should be
filtered.
Wash 1 (1 L)
940 ml |
MilliQ water |
50 ml |
20X SSC |
10 ml |
10% SDS |
Wash 2 (1 L)
950 ml |
MilliQ water |
50 ml |
20X SSC |
Wash 3 (1 L)
995 ml |
MilliQ water |
5 ml |
20X SSC |
Set up a glass chamber
with enough Wash 1 to submerge a slide and a 500 ml beaker with the vertical
rack with the bent hook and enough Wash 1 to cover the racked slides.
Pour the warmed Wash 1
into a 500 ml beaker in the 65C bath.
Place it on the submergible stirplate and add a stir bar. Keep it covered with aluminum foil.
In the plastic tubs with
lids, pour ~500 ml Wash 2 plus a horizontal rack and another with ~500 ml Wash
3.
Disassemble each hybe
chamber one at a time. Use the
plastic tweezers to gently wedge open the sandwich while submerged in the
chamber of Wash 1. Transfer slide
to the rack in RT Wash 1 beaker, with a space between each slide (maximum 6 if
you place them back to back).
Once all the slides are in
the rack, hook the rack on the side of the 65C Wash 1 beaker.
Stir 10 minutes with
enough agitation that the solution is visibly turbulent.
Transfer slides to the
rack in Wash 2.
Shake 10 minutes on the
orbital shaker.
Transfer rack to Wash 3.
Shake 5 minutes on the
orbital shaker.
Spin the slides dry (~15
seconds) one at a time in the little benchtop slide spinner.
Load the slides into
scanning holders, barcode side up and barcode sticking out. The scanner scans through the back of
the slide. Don't touch anywhere
but the edges and the barcode.
Scan at 5 µm resolution.
Scan no more than 6 slides
at a time to avoid ozone in the scanner.
Stagger the washes if
you're doing multiple batches.
Grid with Genepix.
Crosslink Corning slides
at 700 energy on Donna's Stratalinker.
Make 600 ml 5X SSC, 0.1%
SDS, and 6 g Roche Blocking Reagent (1096176).
Warm 65C with occasional
agitation until dissolved.
Pour into a 500 ml beaker
and warm until 65C.
Add the crosslinked slides
in a vertical hanging rack and stir 35 minutes at 65C on submergible stirplate.
Move slides one by one
into a new rack in RT water in a tupperware.
Wash on orbital shaker for
5 minutes.
Spin dry.
Use the same day, maybe the next only if absolutely necessary.