Agilent yeast cRNA arrays for 44k platform
Maitreya Dunham September
2007
modification of the
Agilent low RNA input fluorescent linear amplification kit and Agilent 60-mer
oligo microarray processing (SSPE wash) protocols
Use RNase free solutions
and plastics throughout.
Remember that the arrays
come 4/slide.
If starting with crude
total RNA, clean up an aliquot with a Qiagen RNeasy column.
QC on bioanalyzer. Measure concentration with nanodrop.
Make a 100 ng/µl stock of
total RNA.
The amplification/labeling
is done per Agilent instructions with half volume reactions and a quarter
amount of recommended dye.
Aliquot 3.25 µl (325 ng)
total RNA into a PCR tube.
Add 2.5 µl T7 Promoter
Primer.
65C 10 min
ice 5 min
Warm 5X first strand buffer
at 80C, with occasional vortexing, until it completely dissolves (3-4 min).
Prepare cDNA master mix,
in this order at RT:
1X
2 µl |
5X first strand buffer |
1 µl |
0.1 M DTT |
0.5 µl |
10 mM dNTP mix |
0.5 µl |
MMLV RT |
0.25 µl |
RNaseOUT |
Add 4.25 µl to each reaction.
40C 2 hours
65C 15 min
ice 5 min
Add 0.6 µl appropriate Cy
CTP to each reaction.
Warm 50% PEG at 40C until
it's resuspended and easy to pipet.
Prepare transcription
master mix, in this order at RT:
1X
8.25 µl |
water |
10 µl |
4X transcription buffer |
3 µl |
0.1 M DTT |
4 µl |
NTP mix |
3.2 µl |
50% PEG |
0.25 µl |
RNaseOUT |
0.3 µl |
inorganic
pyrophosphatase |
0.4 µl |
T7 RNA polymerase |
Mix by pipetting.
Add 29.4 µl to each
reaction. Mix by pipetting.
40C 2 hours in the dark
Purify with an RNeasy
column. All spins are full speed:
Bring reaction to 100 µl
with 60 µl water.
Add 350 µl Buffer RLT and
mix.
Add 250 µl ethanol and mix
by pipetting.
Add mix to column.
Spin 30 sec. The filter should be tinted.
Move column to new
collection tube.
Add 500 µl Buffer RPE.
Spin 30 sec. Discard flowthrough.
Add 500 µl Buffer RPE.
Spin 60 sec.
Move column to a new
eppendorf tube with the lid cut off.
Add 30 µl water directly
to the membrane. Let sit RT 1 min.
Spin 30 sec.
If any color remains on
the filter, repeat with another 30 µl water.
Nanodrop 1 ul to check
yield and dye incorporation.
Store RNA at -80C.
Find the amount of sample
that gives 2.5-5 pmol dye.
Determine how many ng are in that amount. The kit says you should get between 10-20 pmol dye/ug cRNA. Mix the red and the green reactions
such that there is at least 2.5 pmol dye in each channel and there is the same
amount of cRNA in each channel.
This means that one channel with have more than 2.5 pmol dye. Make sure that the amount of cRNA is no
more than 1000 ng per channel.
Bring total volume to 41.8
µl with water.
You may want to randomize
the arrays that are neighbors on the arrays. A simple way to do this in Excel is to list the samples in
one column and the function =RAND() into each cell in the neighboring
column. Copy the random number
column, then, with the column still selected, paste special -> values so
that the cells won't recalculate.
Then, sort both columns by the random number. List the arrays in a third column next to the sorted
list. A01 is nearest the barcode
and A04 is farthest from the barcode.
Final probe will consist
of:
Cy3-labeled DNA |
at least 2.5pmol |
Cy5-labeled DNA |
at least 2.5pmol |
Water |
to 41.8µl |
10X Agilent Blocking Agent |
11µl |
2X Hi-RPM hybridization buffer |
55µl |
TOTAL VOLUME |
110µl |
VOLUME LOADED |
100µl |
Prepare 10X Agilent
blocking agent per tube directions.
Add 11 µl 10X blocking
agent to each tube. Mix.
Add 2.2 µl 25X
fragmentation buffer. Mix.
60C 30 min in the dark
You may want to stagger
the fragmentation step if you are doing multiple samples. Do at most 4 samples/1 slide at a time.
Add 55 µl 2X Hi-RPM
hybridization buffer to stop the reaction. Mix by pipetting.
Try to spin out any bubbles that form.
Place a backing slide,
Agilent side up, in a hybe chamber.
Pipet 100 µl of probe,
avoiding bubbles, onto the center of one gasket area. Don't eject the last µl or two in order to avoid bubbles,
but don't skimp on the volume or you'll get a hole in the center of the
array. Spread the probe around as
you pipet, but not too close to the gasket.
Do the same for the next 3
samples.
Remove the array from the
box. The Agilent side is the Array
side. 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 rotate the slide.
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 with the array, you can try knocking the array
on the bench to dislodge them.
Put the array in the hybe
oven. Make sure to balance the
rotisserie.
Hybe 65C for 17 hours at
10 RPM.
Prepare your wash
solutions. Be aware of array
materials that may be for RNA only use.
Wash A (1 L)
add in this order:
700 ml |
Water |
300 ml |
20X SSPE |
0.25 ml |
20% N-lauroylsarcosine |
Filter. Shake to mix.
Wash B (1 L)
add in this order:
997 ml |
Water |
3 ml |
20X SSPE |
0.25 ml |
20% N-lauroylsarcosine |
Filter. Shake to mix.
Rinse the wash chambers, racks,
and stirbars with water.
Set up:
two Wash A chambers, one
with a rack and a stirbar on a stirplate.
one Wash B chamber with a
stirbar on a stirplate.
one acetonitrile chamber
with a stirbar on a stirplate.
For all stirring steps,
the wash liquid should be visibly turbulent. Make sure the entire slide is submerged at all times.
Disassemble each hybe
chamber one at a time. Use the
plastic tweezers to gently wedge open the sandwich while submerged in Wash
A. Transfer slide to the rack in
the other Wash A chamber. Leave a
gap between each slide and between the slides and the wall.
Once all the slides are in
the rack, stir for 1 min.
Start stirring Wash B.
Transfer the rack into
Wash B and stir for exactly 1 min.
Don't worry about transferring some Wash A into Wash B.
Start stirring the
acetonitrile.
Quickly transfer the rack
into the acetonitrile, draining off some of the Wash B as you go.
Let stir 30 sec. Slowly and evenly pull the rack out of
the acetonitrile. If you see
droplets remaining on the slides, submerge them and try again.
Set the rack on a kimwipe.
Load the slides into
scanning holders, Agilent side up and barcode sticking out, blotting excess
acetonitrile if necessary. The
scanner scans through the back of the slide. Don't touch anywhere but the edges and the barcode.
Scan no more than 5 slides
at a time to avoid ozone in the scanner.
You can reuse the wash
buffers for more slides, replacing the first Wash A for every batch.
Open the Agilent scan
control program. If the lasers
refuse to warm up, power cycle the scanner.
Place the slides in the
scanner, noting the slot numbers.
Select the appropriate
slot numbers from the pulldown menus on the upper left.
Select the directory
column and click edit values.
Browse to find the directory you want to save in. Hit set. The column values should change.
Check the default
preferences for the correct scanning area (61 x 21.6 mm), resolution (5 µm), laser power (100% each) and with the split and
rotate box not checked.
Scan.
Open the scanned tif with
the Agilent feature extraction software.
Run the appropriate
protocol.
Check the visual results to make sure it looks ok. I usually check that it's aligned properly, and that the flagged spots make some sense. If you get a larger than usual number of outlier spots, make a note of it