Maitreya Dunham June 2006
Modified from various Brown and DeRisi lab protocols
Throughout the protocol, I combine replicate reactions where
possible. For example, I would do
all the reference labeling in 1 scaled up reaction. Split the samples back into 1X reactions for the zymo
purification. Once you elute,
recombine the samples if appropriate.
This method helps make sure the reference is as standardized as
possible.
Unless otherwise noted, all spins are at full speed. Also note that you will lose 1 ml of volume in the zymo elutions.
Use RNeasy-purified RNA for best results.
Bring 30-50 mg total RNA to 14.4 ml with water.
Add 1 ml 5 mg/ml anchored oligo dT (T20VN).
Incubate 70C 10 minutes.
Ice 5 minutes.
Add 3 ml appropriate Cy-dUTP (or Cy-dCTP, just make sure the dNTP
mix is modified appropriately).
Add 11.6 ml RT cocktail:
6 ul 5X
superscript buffer
0.6 ul 50X
dNTP mix (25 ml each 100 mM dATP, dCTP, dGTP; 10 ml 100 mM dTTP, 15 ml water)
3 ul 0.1
M DTT
2 ul 200
U/ml
Superscript II (or III)
Incubate 42C 2 hours.
Incubate 95C 5 minutes.
Ice.
Add 15 ml 0.1N NaOH and 1 ml 0.5 M EDTA pH 8.
Incubate 15 min 67C.
Add 15 ml 0.1N HCl.
Purify sample with Zymo Clean and Concentrator 5 columns:
Add 0.5 ml binding buffer.
Load on the column and spin 20 seconds. Discard flowthrough.
Add 200 ml wash buffer.
Spin 30 seconds. Discard
flowthrough.
Add 200 ml wash buffer.
Spin 1 min.
Transfer column to a new Eppendorf tube with the lid cut off.
Add 22 ml water directly to the matrix.
Incubate RT 1 minute.
Spin 1 min to elute.
Check yield with nanodrop.
You should get in the neighborhood of 1 mg cDNA at 20 pmol dye/mg cDNA.
Mix Cy3 and Cy5 reactions.
Add 120 ml 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 no more than 6 slides
at a time to avoid ozone in the scanner.
Stagger the washes if
you're doing multiple batches.
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.