Probe Preparation
Plasmid Minipreps (using Promega Wizard Plus Minipreps DNA Purification
System)
1. Grow overnight culture in 3 ml of LB broth in a 14 ml Falcon tube
with a loosened cap (add bacteria with plasmid inclusion to LB broth
using a tooth pick).
2. Transfer ~1.9 ml into 2 ml eppendorf tube and spin at 4500 rpm for
4-5 min. Pour off supernatant and re-suspend the pellet in 200 ul of
Cell Re-suspension Solution.
3. Add 200 ul of Cell Lysis Solution and mix by inverting the tube several
times. The cell suspension should clear immediately.
4. Add 200 ul of Neutralization Solution and mix by inverting the tube
4 times.
5. Spin at top speed for 5 minutes.
6. Decant supernatant into1.5 ml tube, spin again if supernatant is
not clear.
7. Add 1 ml DNA Purification Resin, mix by inversion.
8. For each miniprep, prepare one Minicolumn for the Vac-Man Vacuum
Manifold.
9. Pipet 1 ml of the re-suspended resin into each barrel of the Minicolumn/Syringe
assembly. Open the stopcocks and apply a vacuum to pull the resin/lysate
mix into the Minicolumn. When all of the sample has been pulled through
the column, break the vacuum at the source.
10. Add 2 ml of the Column Wash Solution to the Syringe Barrel and reapply
the vacuum to draw the solution through the Minicolumn. Dry the resin
by continuing to draw a vacuum for 30 seconds after the solution has
been pulled through the column. Do not dry the resin for more than 30
seconds. Remove the Syringe Barrel and transfer the Minicolumn to a
1.5 ml microcentrifuge tube. Spin at top speed for 2 minutes to remove
any remaining column wash solution.
11. Transfer minicolumn to a new 1.5 ml tube. Apply 50 ul of TE to Minicolumn
and wait 1 minute. Spin at top speed for 20 seconds. Remove and discard
the Minicolumn.
Prehybing
1. Pre-wet Immobilon-N blot with Isopropanol or 95% Ethanol (blot turns
translucent gray). Rinse inddH2O for ~10 min. Let the blots remain in
ddH2O until you’re ready to put them into the pre-hyb solution.
(If you are using nylon blots, just pre-wet in ddH2O or 2X SSC.) Preheat
the hyb solution to 65 C. Boil the DNA for 5-7 minutes; add it to the
preheated solution just before pouring the solution into your boxes
or bottles.
| Pre-hyb/Hyb Solution (per 100ml) |
Final Concentration |
| 15 ml 50% Dextran Sulphate |
7.5% |
| 25 ml 20 X SSC (filter sterilized) |
5X |
| 48.5 ml ddH2O |
-- |
| 2.0 ml 50X Denhardt’s Solution |
1X |
| 5.0 ml 1 M Tris (pH 8.0) |
0.05M |
| 1.0 ml 20% SDS |
0.01M |
| 1.5 ml SSS (Salmon Sperm DNA, 10 mg/ml) |
0.15mg/ml |
Notes: Pre-hyb a new blot for 24 hours before the
first use; then for about 3-4 hours (or overnight) for each succeeding
use. It’s easier to pre-hyb in the same box (or bottle) in which
you plan to hybridize. The dextran sulfate amounts in the pre-hyb/hyb
solution cam vary (5%-10% total volume) according to your probe. If
you are using a very low amount of probe, i.e. 25 ng, you will want
to use a greater concentration of dextran sulfate. Also, be careful
not to use too much hyb solution because your probe concentration can
become too low for a good hybridization.
Random Priming
2. Remove 32P-labled nucleotide form the freezer ~40 minutes before
use. Place the pig snugly against the back of the shield.
| Notes: There’s no need
to remove the stock vial from the pig while it thaws, however, it’s
a good idea to check the fluid level of the vial when you first
remove it from the freezer. There may not be as much there as you
think. Sometimes people forget to sign out their reactions. |
3. When you’re ready to start labeling, put on a lab coat, one
pair of gloves, your ring badge, then, another pair of gloves. Check
that the Geiger counter is working correctly by testing the battery
and testing the pancake probe against the “check source”-
the Coleman lantern mantle. The cpms should read between 4000-6000 depending
on which counter you use. Check shield area with Geiger counter. Position
the open solid waste box beside you.
4. Label microcentrifuge tubes.
MISSING PAGE MISSING PAGE MISSING PAGE
10. Later, when you’re ready to add the probes to your blots,
place the reaction tubes into the 95 C heating block for 7-10 minutes.
Quick spin the tubes in the mini-centrifuge.
| Notes: You can keep your reactions
in the heating block until you’re ready to pipet them. |
Hybing
11. Retrieve blot boxes (or bottles) from the hyb ovens one or two at
a time, and immediately pipet the reactions into their respective boxes.
Reseal and return boxes (or bottles) to the hyb ovens for overnight
hybing at 65 C.
| Notes: Open tubes with a microfuge
tube opener (it looks like a small bottle opener). This prevents
contamination of your fingertips. Use the 200 ul Pipetman set on
“~70 ul” and aerosol pipet tips to pipet the radioactive
probe into your hyb boxes or bottles. Try to pipet the reactions
into the hyb solutions, not directly onto the blots themselves.
If you don’t have a choice, try to rinse the blot with the
hyb solution to dilute the probe quickly. Make sure boxes or bottles
are sealed very tightly, otherwise, the hyb solution evaporates.
The bottles must have Teflon inserts in the caps, otherwise they
leak. |
Washing
12. Make up your wash solutions and preheat Wash #3 ahd of time. Be
sure to make enough for the number of blots you have. You can usually
wash all your blots in one box. For a dozen ½ size blots, make
2 liters Wash #1, 1 liter Wash #2, and 2 liters Wash #3. Be sure to
check the shield and washing area for contamination before you start.
General blot washing regimen after overnight hybridization:
#1 2.0X SSC, 0.5% SDS…wash 3 times for 5 minutes at room temperature.
#2 0.1X SSC, 0.1% SDS…wash once for 10 minutes at room temperature.
#3 0.1X SSC, 0.1% SDS…wash twice for 30 minutes at 65 C (pre-heat
solution)
13. Pour about 500-600 ml Wash #1 into one of the large plastic Rubbermaid
“wash” boxes; place it on a shaker. Unscrew the cap of the
liquid waste jar and lay it beside the jar’s neck inside the metal
container. Take the top off the solid waste box. Position the waste
containers beside you for easy access. Pour off the probe from the hyb
box (or bottle) into the liquid waste jar and dap with a kimwipe at
the lip of the box to catch drips; add a little Wash #1 to the Hyb box,
rinse gently and pour off into the jar, again dabbing with a kimwipe.
Use forceps to place blots into the “wash” box, then snap
the lid on. Wash with slow shaking, using the shaker in the hood. The
first five minute wash should be put into the waste jar; the rest of
the washes can be poured down the sink. Rinse sink with lots of cold
water.
| Note: You can wash your blots
for various time periods depending on how “hot” they
are. (Use a forceps to remove a blot from the wash solution; lay
it on the inner side of the lid of the wash box. Check radioactivity
levels with the Geiger counter.) If there is a lot of background,
you can increase the stringency and/or wash for longer at 65 C.
If you don’t have a strong signal (>3k cpm), you might
not want to use wash #3. If it is a very weak signal (<1k cpm),
don’t use Wash #2 either. Fill a waste jar only to the shoulder,
not to the very top. When the jar is full, put a piece of tape on
the lid labled “full”, and replace the jar on the waste
cart with an empty one. |
14. When you‘ve finished washing, place each blot into a pre-cut
plastic sheet holder, then into a cassette with an intensifying screen
for exposure to x-ray film.
| Note: You can either air dry
your blot, or place it into the sheet holder still damp. |
15. Check shield area, counters, sink, flooraround waste bottle, boxes
you’ve washed, yourself (including shoes), front of lab coat,
arms, etc. for contamination. Sign out on the shield.
Exposing to X-ray Film
16. In the darkroom, turn off all lights except for those rated as safe
to use with x-ray film. Open cassettes containing the plastic sheet
holders/bolts. Open x-ray film box and remove as many pieces of film
as you will need, then close the box. (This will prevent an entire box
of film being destroyed if someone walks in and turns the light on.)
Write directly on the film in an “out of the way” corner.
(Use pencil; ink can be removed during the development process.)
Place the film on top of the plastic sheet holder, then place an intensifying
screen (shiny side towards the blot) on top of the film. Close the cassette
and place it between thin lead sheets. Wrap the cassette/lead sheets
in plastic wrap, then in aluminum foil. To assure there are no pinhole
light leaks, you can use 2 pieces of foil, or a black nylon x-ray film
holder. If you have more than one cassette, put lead sheets between
each one, as well as one on top and on the bottom of the stack.
| Note: X-ray film should be
stored with the open end of the inner safety envelope folded over
the film, and pushed to the bottom of the box. This protects the
film should the top of the box be accidentally removed. |
17. Store the film/blot package in the -80 C freezer.
General exposure times:
If blot is >3k cpm, expose overnight
Between 1-3k cpm, 2-3 days
Between 0.6-1k cpm, 4-7 days
Between 0.4-0.6k cpm, 7-10 days
Between 0.2-0.4k cpm, 10-14 days
|