Do 10 ul reactions containing 2 ul of miniscreen/miniprep DNA where 20 ul DNA is the
yield from ~1.5 ml saturated cells by standard lab miniscreen protocol. (See alkaline
lysis miniprep method).
Choose digest such that you can use one or two enzymes, preferably of
compatible salt optima (see NEB or MBI catalogue), of low cost,
and known to cleave miniscreen DNA well (ask around).
Important considerations in choosing sites include size of relevant fragments
(preferably greater than 400 base pairs (bp)), and number of fragments generated
(preferably few sites so that digest will go to completion and not generate a confusing
pattern of partial and irrelevant fragments).
Where a new site has been created, one strategy is to demonstrate that the
parent plasmid is linearized while the desired construction results in a 500 - 1500
bp drop out. The reverse where a site has been destroyed. If comparison is being
made between size of the restriction fragment of the parent and of the construct choose
sites to maximize contrast, e.g. if difference is less than 100 bps, fragments being
compared should not be greater than 500 bp since the sizing accuracy using the 1 kb
ladder as markers is poor above this.
Including a control digest of parent to contrast with miniscreens is a good idea.
For example, if mapping 12 miniscreens and with 1 control digest of parent
plasmid (total 13 samples):
Label tubes 1 - 13, put on ice.
Prepare a "master mix" containing everything except DNA (i.e. includes salts,
buffer, divalent ions, BSA and enzymes). If a double digest is being done with
comparable salt optima and temp optima, mix both enzymes together in the
master mix. If incompatible, the digests must be done sequentially I usually
start with the lower temp or lower salt enzyme, adding the other subsequently,
as will be described. By making a master mix you can deliver all components
except DNA in a single pipetting step. After preparing the master mix
vortex it and spin
down for 1 second in a microcentrifuge. The master mix also makes it convenient to dish out
miniscule amounts of enzyme since you measure out not .2 ul of enzyme, but
rather the amount for (in this case) 14 tubes = 2.8 ul, and you dispense a larger
volume to each tube since all components are mixed together (except the DNA
which is the variable).
Dispense master mix to all tubes on ice first (you can use the same pipette tip
since all the tubes are empty at that point and you are not going into the enzyme
stock again. Absolutely never reuse a pipette tip to get enzyme or buffer, etc.
from the stock tube, even if you don't think it touched any other components -- if
you are wrong, everyone is screwed.
To continue our example:
14 x 1 ul CB 10 X (10-fold concentrate of buffer salts; this is usually provided with the enzyme by the company that sells it. Most enzymes in our lab are from either NEB or MBI Fermentas. Always check with the catalogue first before planning a digest. Both catalogues have convenient charts showing the percentage activity of each enzyme in each buffer. Some enzymes require special "unique" buffers.)
MBI Fermentas provides a 'Tango' buffer for double digests that works reasonably
well with most of their enzymes. NOTE: The way companies get a bunch of
different enzymes to all work in the same buffer is that they include 'special'
things that the enzyme needs in concentrated form in the enzyme storage
buffer. This means that the optimal buffer for the same enzyme from two
different companies may be different. If you are using enzymes from two
different companies look up the composition of the buffer that they list as
'optimal' and compare it with the composition of the buffer for the other enzyme
you are planning to use.
14 x 1 ul 10 X BSA (check the catalogue to see if the enzyme you are using requires
BSA). I don't know any enzymes that are inhibited by BSA. None of
the Fermentas enzymes require added BSA (because they include it in the storage
buffer if you need it in your reaction).
14 X 0.2 ul enzyme(s) (volume may vary from 0.05 ul per reaction to 1 ul maximum,
typically 0.2 ul enzyme per reaction, depending on enzyme
concentration, shooting for approx 1 unit per digest.
Especially in the case of double digests, remember that
enzyme -- sold in 50% glycerol -- must not make up more
than 10% of the total reaction volume or
it may inhibit the enzyme or
alter its specificity.
81.2 ul H2O
Total master mix volume is now approx. 109.2 ul, enough to dispense the correct
8 ul to all 13 tubes on ice.
Now add 2 ul dissolved, vortexed and spun down miniscreen DNA changing tips
after each addition and mixing the DNA directly into the 8 ul droplet of master
mix in each tube.
For the control digest (tube 13) use 0.25 ul of 1 mg/ml stock plasmid DNA and
make up the rest of the volume by adding 1.75 ul sterile distilled water.
Place tubes to incubate at desired temp. For 37°C incubations, digest for 2 hrs,
for 50 to 60°C digestions, stop after 1 hr since the reaction volume has just
about evaporated at that point, anyway. For this reason it is usually a good idea
to spin down high temperature digests every 15 minutes.
If a second digest at different salt or temp is to be done, don't forget to add
either the additional enzyme or combination of salt and enzyme and continue the
incubation at the appropriate temp. You can transfer the tubes to ice while you
are getting your second "master mix" ready. This addition of salt and/or enzyme
should be done in less than 1 ul to keep close to the correct total volume, and
the tube should be spun down for one second in the cold, vortexed and put to
incubate. If only enzyme is being added, 0.2 ul is too small to be practical for a
pipette, so use either the P2 pipette or simply take a very small amount
at the end of the yellow tip using the pipette person.
When finally through with digests, add 5-10 ul of DNA loading buffer.
Load 10 ul of sample onto gel. 1% agarose in TAE is good for large fragments
and linearization. 110 volts constant voltage. Remember to use 1 KB markers.(aliquots of these can be found in the DNA freezer in the "buffers and markers" box.)
Run until lower blue (bromophenol blue) is at least half way to 3/4 of the way down the gel. Dial
down voltage, turn off power, cut out your piece of gel, (wear gloves), and place it in the ethidium bromide solution for at least 15 min. Read the MSDS sheet for ethidium bromide! Visualize bands on transilluminator while
wearing safety glasses. Take picture if desired, and dispose of gel
immediately in the garbage can near the transilluminator.
If you need to dispose of any ethidium bromide solution - put it in the
container containing the 'tea bags' of activated charcoal.
