Thanks for sharing your protocol. It strikes me as quite difficult to do
correctly, whereas I have heard that making chemically competent cells via
Calcium Chloride is more straightforward. Is that your experience, or are
they just as challenging to prepare?
Has anyone got a straightforward protocol for chemically competent cells to
add? One which might be less sensitive to problems like autoclave
contamination or exacting water purity?
I ask because I will be trying to establish a lab soon and I may not
initially have access to a pristine water supply or perfect equipment.
Thanks,
Cathal
---
Twitter: @onetruecathal
Sent from my beloved Android phone.
On 24 Jul 2010 16:46, "DK" <d...@no.email.thankstospam.net> wrote:
In article <mailman.903.1279899487.25217.meth...@net.bio.net>, Christopher
Fu <christophe...@hotmail.com> wrote:
>
>Dear DK=2C
>
>Would you mind sharing your protocol for making generic home-made
>electrocompetent cells? I'm ...
>Quikchange XL ki and I'm obtaining either no or far too few colonies on
>my agar plates.
>
>Of course there are a whole list of possibilities why the experiment isn't
>work...
Indeed, it's probably not your competent cells. Below is a protocol that
our technician has written after few years of making the cells based on my
mish-mash protocol of various reports an trials. It is probably
too detailed and not generic enough but here it is. I added few notes
and minor edits.
*****************************************************************************************
Use MilliQ water directly from the MilliQ machine for everything
(DO NOT autoclave provided that the final water is filtered through
0.2 um filter). We use Top 10 E.coli (Invitrogen) for general cloning
purposes (culture w/out antibiotic) but DH5alpha work the same for us.
1.5 L of cell culture makes about 100 - 250 x 40ul single use aliquots.
You will need: 1 x 500mL flask
3 x 2L aeration flasks
6 x 250 mL autoclavable centrifuge bottles (PPCO)
3 x 1L glass bottles
1 x 250 mL glass bottle
~ 250 x 1.6 mL microcentrifuge tubes
2 x 50mL sterile disposable centrifuge tubes
DMSO (molecular biology grade; note: any other grade
was simply never tested)
Your E. coli strain, freshly plated
Preparation:
Inspect all glass and plastic ware to insure that it is clean and free from
cell debris; make sure that lids to all 250mL centrifuge bottles contain
red rubber gaskets and that they are seated properly (not twisted).
Vigorously rinse EVERYTHING twice with MilliQ water. Remember to
rinse all items that you will use to make LB.
Make and autoclave LB medium. Use MilliQ water. Do not use
pre-mixed LB reagents.
3 x 500mL in 2L flasks
1 x 200mL in 500mL flask
Dry autoclave microcentrifuge tubes, centrifuge bottles and glass
bottles. Cool to RT. After autoclaving, rince all bottles with MilliQ
right off the faucet, place on ice. (Note: "electrocompetent" E.coli
cells are very sensitive to water quality. Nearly all autoclaves
contaminate everything sterilized in them with all sort of stuff. Failure
to have pristine clean labware and water is the main reason for
troubles in making highly competent cells; tempting as it is, don't
cut corners here).
Fill sterile 1L bottles with MilliQ water.
Make 200mL 7% DMSO in the 250mL bottle
(14mL DMSO + 186mL MilliQ water).
Place all microfuge tubes, centrifuge bottles, MilliQ water bottles and
7% DMSO in the cold room O/N. Also chill the JS-14 rotor and swinging bucket
50mL tube holders. Place the 3-2L flasks of LB in the incubator at 37C O/N
(without shaking) to pre-warm.
Inoculate 200mL LB in 500mL flask with single, fresh colony.
Incubate @ 37C O/N w/shaking.
Protocol:
Inoculate each 2L flask of LB with 40-50mL overnight culture (this typically
gives starting O.D.600 = 0.2; note that OD600 is a function of the
spectrophotometer to be used! We have Beckman DU-640 and be aware
that other spec can give slightly or even widely different readings for the
same sample). Grow cells to O.D.600 = 0.55 - 0.60. Do not exceed 0.65
(this should take around 1 hr).
Place all centrifuge bottles, 50mL tubes and reagents in ice. Keep in the
cold room.
Make up fresh DYT, sterile filter into 15mL tube, bury in ice to chill:
25 mg tryptone
12.5 mg yeast extract
7 % DMSO to 10mL
When cells reach proper O.D., place all flasks on ice for 15+ minutes.
Turn on swinging bucket centrifuge and JS-14. Allow to chill to 2C.
Transfer cultures from flasks to 6- 250mL centrifuge bottles (in ice).
Spin in JS-14 at 6000 rpm at 2C for 20'. Decant supernatant, keep
bottles in ice.
Add a little of ice-cold MilliQ water to each cell pellet. Swirl while
keeping
cold until cells are completely resuspended. (This can take a long time;
do all wash steps in cold room). Add MilliQ water to each 250mL bottle to
at least 200mL (about 1" below top); invert several times. Spin in JS-14
at 6000 rpm at 2C for 20'
Aspirate off supernatant (use the aspirator connected to a sterile pipette.
The pellet will be very loose; try to remove as much liquid as possible
without disturbing the pellet.
Repeat MilliQ water wash. Pellet. Aspirate.
Add ~10mL 7% DMSO to each pellet and resuspend by swirling. Combine
contents of the three 250mL bottles into one 50mL centrifuge tube. Top off
50mL tubes with 7% DMSO, mix thoroughly. It is extremely important to
keep everything ice-cold for the remainder of the protocol.
Spin at at the maximum speed your rotor and adapters allow to spin
50 ml Falcon tubes. We use 3500 rpm for 20' at 2C in JS-42 swinging
rotor. Use pre-chilled tube adapters. Carefully aspirate the supernatant,
keeping tubes in ice.
Add fresh DYT medium to 2.5mL mark in each tube. Resuspend pellets
by pipetting up and down. Pool cells together in one tube; keep deeply
in ice.
Test for arcing: remove 50ul cells to a 1mm electroporation cuvette.
Zap at 1.9 kV (200 Ohm/25 uF). If arcing occurs, wash cells once more
and re-test. If arcing occurs again, the cells are no good.
Have eppendorf tubes on ice firmly. We use a shop-made 48-positions
aluminum tube rack to make this step easier and cleaner. Aliquot cells at
40 ul per tube into ice-cold tubes. You should have about 5mL of cells at
this point. If you did a good job, this will be more concentrated than you
need. I make 25 x 40ul aliquots of "concentrated" cells for super-demanding
applications and then add 4 mL more of DYT to the remaining cells, mix
and finish aliquoting. Each time you fill the rack, transfer the tubes to a
freezer storage box in the -80. Repeat until done. Store "concentrated"
cells separately. (Note: most protocols call for snap freezing in liquid
nitrogen; we've never tested this back to back with cells slow frozen
in DMSO; as "real" cells are routinely frozen slowly in 7-10% DMSO,
one would think that E.coli would survive such freezing better, too).
Test Competency:
It is ideal to do a series: 1.3, 1.4, 1.5, 1.6 and 1.7 kV at 200 Ohm, 25 uF.
Electroporate each aliquot with 1 ng of a reliable plasmid in 1 ul of 1X T4
DNA ligase buffer. Recover in 1 mL SOC. Dilute 100ul of recovered cells
with 900 ul of SOC, plate 50 ul. Grow overnight and count colonies. At least
one condition should give 500-5000 colonies, a transformation efficiency of
>10^8-10^9 colonies per ug plasmid. Anything less should be considered
a poor prep.
Electroporation Protocol (E. coli):
Place 1 mm electroporation cuvettes on ice, place frozen cell aliquots
on ice. let thaw. Add 1 ul of ligation reaction or DpnI-treated plasmid to
each tube of cells, pipette up and down briefly to mix. Transfer to cold
cuvette, tap cuvette on bench to remove air bubbles. Insert cuvette into
the holder, zap and IMMEDIATELY add 500uL - 1mL of SOC to the cuvette
and transfer to a culture tube. Recover for 1 hour at 37C with shaking.
Plate appropriate amounts on LB+antibiotic.
If you have extremely competent cells, or you are using your cells for
a less demanding application, dilute one aliquot of your cells 2 - 4x with
ice-cold MQ water and split it for two to four transformations.
The same frozen cells can be used for a low efficiency chemical
transformation using Inoue buffer (useful to save on electroporation
cuvettes, yet avoiding making separate cells whenever transforming
with high plasmids): make 2X sterile Inoue buffer. Add 40 ul of it
to an aliquote of frozen cells, thaw on ice, add DNA, incubate for
5-30 min, heat shock for 45-60 sec at 42C or 90 min at 37C, cool
on ice for 5 min, add LB or SOC.
Final note: most protocols use 10% glycerol, which is much cheaper
than DMSO. There are hints that doing all washes in 10% glycerol
instead of water is better. We never tested any of this in a reproducible
manner. But then, DMSO is not that expensive, easy to pipet, is
ALWAYS STERILE and used here only in the final step, so we never
bothered.
***********************************
As far as Quick Change mutagenesis goes, we never use any kit
and we routinely do mutagenesis and whole gene cloning using
this approach and plasmids up to 9 kbp. Literally never failed
in my hands.
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