PFGE arrays
Maitreya Dunham September
2007
modification of the
Invitrogen BioPrime array CGH kit and Agilent 60-mer oligo microarray
processing
You can also use the
labeling protocol with a homemade array hybe protocol, but use the entire
amount of labeled band DNA in the probe.
Run PFGE as usual. Using double-wide lanes will give
higher yields.
Excise band from gel.
Incubate in 1 ml water for
1 hour twice.
Replace water with 250 µl
1X HaeIII buffer (may need more depending on gel slice size). Add 5 µl HaeIII and digest at 37C
overnight.
Gel purify using QIAQuick
kit.
Nanodrop 1 ul
(concentration is generally very low).
DNA can be stored -20C.
DNA labeling is done with
the Bio-Prime array CGH kit plus zymo columns. For reference, either use a wt PFGE plug that has not been
run on a gel but has been processed as above, or use 1 µg HaeIII-digested wt
genomic DNA.
Aliquot 21 µl gel-purified
DNA into a new tube.
Add 20 µl 2.5X random
primers solution.
Incubate 95C 5 min.
Ice 5 min.
Make a master mix for each
dye:
1X
5 µl |
10X dCTP nucleotide mix |
3 µl |
Cy dCTP |
1 µl |
Klenow |
Add 9 µl to each reaction.
Mix gently.
Incubate 37C 2 hours.
Add 5 µl stop buffer.
Ice. The reaction can be stored -20C
overnight if necessary.
Purify using the Zymo
columns according to their instructions, except use 500 µl binding buffer for
the first step.
Elute in 25 µl water.
Nanodrop 1 µl to check
yield and dye incorporation.
Yield should be a few
hundred ng with dye incorporation >20 pmol/µg.
Mix 100ng Cy3-labeled DNA
and 100ng Cy5-labeled DNA.
Add water to bring the
total volume to 44µl.
Add 11µl Agilent 10X
Blocking Agent (prepare 10X Blocking Agent according to instructions that came
with the tube).
For the next steps, only
process 4 samples at a time so they don't rehybridize before assembling the
array.
95C for 5 minutes.
Cool @ room temperature
for 5 minutes.
Add 55µl 2X Hi-RPM
hybridization buffer, mix by pipetting.
Spin down.
Load 100µl onto the gasket
slide:
Place a backing slide,
Agilent side up, in a hybe chamber.
Pipet the whole volume of
probe, avoiding bubbles, onto the center of one gasket area. Don't eject the last µl or two in order
to avoid bubbles (only spare a couple of µl for this, otherwise the volume
won't be enough and you'll get a dark spot in the middle of the array). Spread it around as you pipet, but not
too close to the gasket.
Do the same for the other
gasket areas.
Remove the array from the
box. The Agilent 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 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
with the heel of your hand to dislodge them.
Put the array in the hybe
oven. Make sure to balance the
rotisserie.
Hybe 17 hrs. @ 65C, 20RPM
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, covering when not in use and 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.
Process according to 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.