BAC Recombineering with the “Copeland” E.coli strains:
I. Preparing recombination-competent, electrocompetent BAC cells
for performing recombineering experiments
The strains DY380, EL250, El350, SW102, SW105, SW106 all have heatinducible expression of lambda prophage recombination proteins exo, bet and gam.
Therefore always culture these cells at 32˚ or lower except at the brief heat induction
step! They may die if cultured at 37˚ or higher; worse, they may survive but become
unable to catalyze recombination. Read the papers for more info about these strains.
Special reagents needed:
NO-Salt LB Media: 5 g yeast extract and 10 g tryptone per 1 L media. Do not add
sodium chloride.
Other reagents needed:
Sterile deionized water, 500 mls
Sterile 10% glycerol, 15 mls
LB plates + 6.25 µg/ml chloramphenicol + antibiotic used to select the
recombination event
Special equipment needed: Shaking water bath for small flasks; 32˚ bacterial culture
incubators; refrigerated centrifuge for 50 ml tubes and 1.5 ml tubes; electroporator;
IMPORTANT NOTES:
• “NO-Salt” LB media is used to grow the cells. This increases electroporation
efficiency.
• The cell rinsing steps require the use of extremely clean, sterile de-ionized water and
thorough rinsing of the cells to remove all traces of ions that could cause “arcing” during
electroporation.
1. The day before you want to do a recombination attempt: In afternoon, inoculate a
fresh colony of the BAC clone stock, in a recombinogenic strain, into a starter culture of
~2 ml no-salt LB + 12.5 µg/ml chloramphenicol; shake @ 32˚ overnight.
2. Next day, prepare a sterile flask containing 40 ml no-Salt LB = + 12.5 µg/ml
chloramphenical. Inoculate this with 1.5 ml of the overnight miniculture. One 40 ml
culture should generate enough cells for ~3 to 4 electroporations. Always prepare at
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Doug Mortlock updated Jan. 21, 2014
least 2 tubes per single recombination so you have an extra backup or control tube.
(Scale up the culture as needed to make additional electroporations.) Shake @ 32˚ for
about 2 hours.
3. Transfer cultures immediately to 42˚ shaker water bath, and shake for 15 minutes at
200 rpm to induce expression of the recombination proteins.
4. Incubate culture flasks on ice 20 min. For this and all the following steps, keep cells
on ice at all times. Prechill on ice the following items: two sterile 50 ml screw-cap
disposable tubes, the sterile deionized water , and the sterile 10% glycerol.
5. Transfer the culture evenly into two prechilled 50 ml screwcap tubes (20 mls per
tube). Spin down the culture at 3000 rpm, 15 minutes, 4˚ C.
6. Carefully remove and discard supernatant. Resuspend the cells in 20 mls of
prechilled sterile deionized water (0˚ C). Re-centrifuge the cells at 3000 rpm, 10
minutes, 4˚ C.
7. Carefully remove and discard supernatant. Resuspend the cells in 10 mls of
prechilled sterile deionized water (0˚ C). Re-centrifuge the cells at 3000 rpm, 10
minutes, 4˚ C.
8. Carefully remove and discard supernatant. Resuspend the cells in a total volume of
1.5 ml of prechilled sterile deionized water and transfer to a microcentrifuge tube.
9. Spin down the cells using a microcentrifuge, at 4˚ and full speed. Discard
supernatant.
10. Estimate the volume of the cell pellet. It should be around 50-80 µl. Add ~3x pellet
volumes of prechilled, sterile 10% glycerol and resuspend the cells carefully.
11. Transfer 40 µl aliquots of cells to 1.5 ml tubes. Use each for one electroporation
immediately, or freeze the tubes on crushed dry ice, and store at -80˚ until use.
Notes:
The above was originally derived from E-Chiang Lee et al. (Neal Copeland lab)
Genomics 73, 56–65 (2001) and modified according to recommendations from
members of the Mortlock lab; the use of media with reduced or no salt was suggested
by Joshua Hoffman, and Jean-Marc DeKeyser.
This method has proven to routinely yield hundreds to thousands of recombinant
colonies per recombination, with the majority (50-100%) correctly targeted.
The electroporation efficiency depends critically on the time constant (TC) during
electroporation. The efficiency is optimal when the TC is 5.0-5.1. Typically, a no-DNA
control electroporation will give a time constant with 5.0-5.1. Adding DNA can often
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Doug Mortlock updated Jan. 21, 2014
decrease the TC to ~4.8 or lower depending on how clean it is. Below TC=4.5, the
transformation efficiency drops quickly. The primary causes of lower time constants
seem to be either having cells that are too dense (i.e. not resuspended in enough
volume), poor DNA quality, or poor washing.
Electroporation efficiency can be measured by doing a test with 0.1 ng of a control,
supercoiled plasmid (e.g. pBluescript). To measure efficiency, simply transform cells
with the plasmid, recover, plate on selective media and count. With fresh cells,
efficiencies of 1x10e8 colonies/µg or greater should be observed. Frozen cells tend to
lose an order of magnitude of efficiency through freezing, and by ~6-12 months may be
unsuable.
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II. General Electroporation protocol adapted for Recombineering
For each electroporation, use one aliquot of freshly prepared cells (or, cells freshly
thawed on ice, if they were previously stored frozen). heat-induced, electrocompetent
recombinogenic cells carrying the BAC to be targeted.
Special reagents needed:
0.1 cM gap electroporation cuvettes (e.g. Bio-Rad Life Science, catalog number 1652089Gene Pulser/MicroPulser Cuvettes, 0.1 cm gap)
1. Use 40-45 µl cells.On ice, add to the cells between 100 to 500 ng of the targeting
DNA fragment dissolved in H20 . The added DNA should itself be in no more than 5 ul
total volume. Mix briefly by gently flicking the tube or stirring with a pipette tip.
2. Transfer the cells/DNA mix to a prechilled cuvette on ice, avoiding bubbles. Tap the
cuvette lightly several times on a hard surface to help spread the cells into the bottom f
the cuvette evenly and to help remove any bubbles. Keep on ice.
3. Electroporate. (1800 kV for 0.1 cM gap cuvette).
e.g. for the BioRad Gene Pulser Xcell:
Turn on machine, and use the scroll buttons to select:
4 – Preset protocols
then
1 – Bacterial
then
1 – E.coli, 1 mM / 1.8 kV
then insert the cuvette and press the red button.
Make note of the time constant. It should be about 4.8-5.1. Lower values will
correlate with lower efficiency and can result from cells being too dense, not rinsed
enough or too much salt or other material in the DNA.
5. Immediately add 0.96 mls SOC or LB media to the cells and transfer to a sterile
falcon (snap-cap) tube.
6. Incubate at 32˚ for 90 minutes with shaking, 250 rpm.
7. Plate 100 µl cells on LB plate with 6.25 µg/ml chloramphenicol + other antibiotic as
needed to select for the recombination event. Incubate at 32˚ for 24 hours.
(see following page for notes on this protocol)
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Doug Mortlock updated Jan. 21, 2014
Technical Notes on BAC recombination
Using antibiotic selection, recombinant colonies usually become visible about 20-24
hours after plating at 32˚ and should be big enough to pick by 24 hours.
Using positive selection to identify recombinant clones is very efficient. We typically find
50%-100% of colonies have undergone the correct recombination event.
Negative controls: These may be useful when first learning the protocols.
(1) I usually do one test electroporation on each fresh batch of cells using no
DNA to check that the time constant is close to 5.0. It is convenient to use a pre-used,
rinsed cuvette for this to save the cost of a new cuvette. I usually do not save these
cells after the electroporation; however you may plate them out on selective media to
ensure that your cells are not contaminated with any resistant bacteria.
(2) When preparing recombination-competent BAC cells, prepare one batch that
is not heat-shocked, and electroporate this with your targeting fragment DNA. These
should not have recombination ability so should generate much fewer colonies than the
heat-shocked plate. This demonstrates that colony formation is dependent on induction
of recombination. If you get lots of colonies with this control, this usually means your
targeting fragment DNA is probably contaminated with intact circular plasmids
containing the antibiotic resistance cassette (usually results from poorly purified
targeting fragment.), which can propagate and cause a background of colony formation.
Homology arms: These should be >40 bp in size. 50 bp works fine. We have used
dozens of different 50 bp homology arms and they have all worked. However, longer
arms will work as well or better (higher efficiencies.)
How to verify BAC clone modifications: I do careful “fingerprint” digests (i.e. BamHI
or HindIII) of the final putative modified BACs and compare these to the original BAC to
check for unwanted rearrangements. Only the desired alterations in banding patterns
should be present. To do fingerprint digest, use a medium or long gel format, 0.8%
agarose, and run it slowly to get good band separation up to the 20 kb range.
I strongly advise performing PCR across the recombination sites and sequencing the
PCR products (or, direct sequencing of the BAC DNA if you can) to ensure the
recombination has gone correctly. If the BAC prep is destined for injection into mouse
or fish embryos or cell transfection, pulsed-field gel analysis is strongly advised to
ensure the BAC is still complete and intact.
Note that modified BACs do not have to be transferred out of the recombinogenic
strains before maxiprep scale DNA is purified. The recombinase proteins are strongly
repressed at 32˚ and so unwanted rearrangements are rare. However, fingerprinting is
still strongly advised.
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BAC Recombineering with the “Copeland” E.coli strains:
III. Inducing expression of Cre or FLPe with arabinose to induce sitespecific recombinations
Several of the Copeland lab recombinogenic E.coli strains also permit transient,
arabinose-inducible expression of Cre or FLPe recombinase, depending on the strain.
This is mainly used for 2 things: (1) efficient deletion of antibiotic cassettes preengineered with flanked lox or FRT sites, or (2) insertion of molecules into BACs via
site-specific recombination. Usually this is used for the former of these, and is typically
straightforward. For example, the pIBG-FTet plasmid has an FRT-flanked TetR
cassette that can easily be deleted this way after insertion into BACs via standard
homologous recombination. This is very efficient.
Note that almost all BAC vectors contain loxP and lox511 sites in the backbone,
complicating any use of Cre. Therefore the following text describes FLPe-mediated
deletion.
1. Inoculate several minicultures of the BAC containing the FRT-flanked cassette into
LB with chloramphenicol @ 12.5 µg/ml . Shake the independent clones at 32˚ overnight.
It is good to process more than one independent isolate, since occasional point
mutations may have altered an FRT sequence in individual transformants. This is
sometimes possible if the targeting fragment was generated by PCR, for example.
2. The next day, inoculate 0.5 ml each miniculture into 20 mls LB with chloramphenicol
@ 12.5 µg/ml. Shake at 32˚ until A600 ~ 0.4 (2-3 hours).
3. Add 0.2 mls of sterile 10% L-arabinose. Continue shaking at 32˚, 1 hour.
4. Discard all but 1 ml of the culture. Add 9 mls LB with chloramphenicol @ 12.5 µg/ml.
Continue shaking at 32˚, 1 hour.
5. Dilute culture and spread on plates with LB with chloramphenicol @ 12.5 µg/ml.
Incubate overnight at 32˚. (100 µl of a 10-4 or 10-5 dilution should result in ~200
colonies.)
6. Check about 10 colonies for loss of the FRT-flanked cassette. To do this, use two LB
plates each with chloramphenicol @ 12.5 µg/ml, but only one of the two should also
have the antibiotic for the resistance gene that you are trying to remove. Mark the
bottoms of the plates with numbers from 1 to 10. Using sterile toothpicks, pick a single
colony and make a small streak on the agar over the number on each of the two plates.
Repeat for all 10 colonies and incubate plates overnight at 32˚. You should see that
most or all clones grow only on the chloramphenicol-only plate. Save one or more of
these clones and analyze them by restriction digest to confirm loss of the cassette as
expected. Typically all 10 colonies have undergone successful deletion.
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BAC Recombineering:
IV. Using KanaRpsL selection cassette for precise mutagenesis via
positive/negative selection. - Steve Pregizer and Doug Mortlock
This method can be used to introduce precise mutations such as SNPs into BACs
without leaving traces of exogenous DNA behind. The cassette was described by
Wang et al. Mol Biotechnol 2009 vol. 42 (1) pp. 110-6. Details of this process are
described in their paper and so are not reproduced here. This method is superior to
previous methods such as galK selection for various reasons, mostly because the
reagents are cheaper and the colony growth is not very slow.
Positive selection to introduce the cassette at the desired mutagenesis site is easily
achieved with kanamycin selection. Negative selection (to select clones that have
recombined out the RpsLKana cassette) is achieved using regular LB+chloramphenicol
plates supplemented with streptomycin.
Annealing primers for RpsLKana cassette, which should generate a
1991-bp amplicon from pSK-RpsLKana
Forward: 5’-GTTTTCCCAGTCACGACGTT-3’
Reverse: 5’-TGTGGAATTGTGAGCGGATA-3’
• We have the following modification to the Wang et al protocol: We increased the
streptomycin concentration in LB plates to 5 mg/ml (that is not a typo, it is very high.)
The reason is that streptomycin does not kill cells that retain the cassette but impairs
their growth. Cells that have lost the cassette form larger colonies among a
background of small ones. The streptomycin dose must be stringent to allow the size
separation to be clearly seen.
Issues and troubleshooting:
• As with all negative selection strategies, some undesired clones will grow at this step
due to random mutations disrupting the cassette. Thus, the desired recombination must
be done efficiently. This is needed in order to screen out the truly recombined clones
at a reasonable frequency above the background of untargeted, undesired mutations
(typically, these are deletions that take out the region of the BAC containing the
cassette).
• Screening of the clones by restriction digest is recommended. For subtle
modifications (e.g. SNPs), the pattern should revert to the “wild type” after replacement
of the inserted RpsLKana cassette with the desired mutation. These clones must be
sequenced to verify the mutation.
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• The negative selection step can often permit a small percentage of viable Kana+ cells
to contaminate the colonies on the Kan+Strep plates. Therefore, you should restreak
this clone on a fresh plate and follow up and isolated clone, and confirm it is truly 100%
Kan-sensitive.
• Example of negative selection on an LB/chloramphenicol/streptomycin plate:
A BAC clone containing the KanaRpsL cassette in SW105 cells was used for
recombination to replace the cassette with a single nucleotide. Recombinationcompetent cells were prepared and transformed with ~100 ng of double-stranded oligo
with the homology arms and the SNP base pair. After electroporation and recovery,
100 µl of a 1:100 dilution of cells were plated on LB + choloramphenicol (12.5 µg/ml) +
streptomycin (5 mg/ml). The appearance of the plate after 24 hours inc. at 32˚C is
shown. ~20 large colonies are visible among thousands of small colonies. Of 16 large
colonies picked for screening, 4 had the desired SNP replacement. The rest had
unwanted deletions or mutations that inactivated the RpsL-Kana cassette and failed to
incorporate the SNP mutation, restore the normal restriction digest pattern, or both.
DPM, 2012.
Arrows = some large colonies that have replaced/inactivated the RpsL gene.
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Appendix
A. Copeland lab bacterial strains and plasmids for recombineering:
Summary chart of key strains, links and references
For more information see also:
http://web.ncifcrf.gov/research/brb/recombineeringInformation.aspx
E. coli STRAINS FOR RECOMBINEERING:
Strain
Heatinducible
recombination
system
Tetracyclineresistant
Arabinoseinducible FLP
Arabinoseinducible
CRE
Gal
phenotype
DY380
EL250
EL350
SW102
SW105
SW106
YES
YES
YES
YES
YES
YES
YES
NO
NO
YES
NO
NO
NO
YES
NO
NO
YES
NO
NO
NO
YES
NO
NO
YES
Gal+
Gal+
Gal+
Gal-
Gal-
Gal-
Reference for DY380, EL250, EL350 strains:
E.C. Lee et al.
A highly efficient Escherichia coli-based chromosome engineering system adapted for
recombinogenic targeting and subcloning of BAC DNA.
Genomics. 2001 Apr 1;73(1):56-65.
Reference for SW102, SW105, SW106 strains:
S. Warming et al.
Simple and highly efficient BAC recombineering using galK selection.
Nucleic Acids Research, 2005, Vol. 33, No. 4
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Appendix
B. Annealing oligos
Reagents needed:
Complementary oligonucleotides
10X Annealing Buffer: 1 M NaCl / 10 mM EDTA pH 8.0 / 100 mM Tris pH 7.5
ADB (Adapter dilution buffer): 50 mM NaCl / 10 mM Tris pH 7.5
1. Resuspend oligos in water to approximately 1 µg/µl concentration. (The actual total
µg’s of oligo is usually supplied by the oligo manufacturer on the spec sheet or on the
tube).
2. Make up annealing reaction with 20 µg each oligo in total of 400 µl 1X annealing
buffer, using 10X annealing buffer stock and water.
3. Heat several hundred ml of water in a beaker till just boiling. Remove from heat.
Float the annealing reaction in the beaker. Let cool to room temperature on the
benchtop.
Alternative to step 3: Transfer the 400 µl oligo mix to 4 PCR tubes. Cycle in
thermal cycler with a program with initial step of 100˚ for 5 minutes, then ramp the
temperature down by 1 degree per minute for 75 minutes, so the final temp should be
about 25 degrees (room temp).
4. Centrifuge briefly. The final concentration of double-stranded, annealed oligo should
be close to 100 ng / µl. It can be assumed that most of the oligos have annealed.
Dilute in ADB so that an appropriate amount can be removed for ligations (usually
around 1 ng, see notes below)
5. (optional): To precipitate, add 40 µl 3M sodium acetate, pH 5.2 and 1.0 ml ethanol.
Inc. at -20˚ for > 30 min. Centrifuge 15 min., remove sup., add 100 µl 70% ethanol,
respin 5 min., remove sup. and let air-dry 5 min. Resuspend annealed oligos in ADB.
Determine concentration by UV spec.
Note: Short annealing sequences may denature in low salt buffers or water. Therefore
storage or dilution of double-stranded oligos in 50 mM NaCl (such as ADB) is advised.
Tips for ligating adapters to cut plasmids:
• Excessive molar ratios of adapters will inhibit recovery of desired ligation products.
Remember to adjust adapter concentration in downstream ligations to have a good
adapter/plasmid molar ratio; i.e. 10:1 is fine.
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• Most adapters will be around 20-50 base pairs long. A typical ligation with a 5 kb
plasmid and 20 bp adapter should work well with 25 ng/µl plasmid and 1.0 ng/µl
adapter. Relatively few ng of adapter is required because they are so small.
• Remember that synthesized oligos do not have phosphorylated 5' ends, so they will
not ligate to dephosphorylated plasmids (i.e. linearized plasmids should NOT be
pretreated with CIAP, SAP, Antarctic phophatase or other phosphatases).
• If you plan to cut your plasmid with two enzymes (e.g. to insert an adapter oligo via
directional cloning), gel-purification of the linearized plasmid is useful for getting rid of
the small internal “stuffer” fragment. This fragment could otherwise inhibit adapter
ligation by re-ligating back to the vector. We have found this really helps for good
efficiency,
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C. Basic Alkaline Lysis Plasmid Miniprep Protocol
For crude purification of plasmid and BAC DNA from E. coli. The DNA recovered from
this protocol is suitable for routine restriction digests and subcloning, but usually not for
sequencing.
1. Place 1.5 ml of LB media (with appropriate antibiotic if needed; for ampicillin resistant
bugs, use 40-100 µg/ml ampicillin) in a sterile tube such as 14 ml snap-cap tubes.
Incoulate from a single bacterial colony, and shake overnight at 37˚.
2. The next day, decant the culture into 1.5 ml microfuge tubes. Spin down in
microcentrifuge, at max. speed (~12,000 rpm), 5 min.
3. Decant or aspitrate the cleared media from bacterial pellet . Invert the tube and tap
briefly on paper towels to “drip-dry”.
3. Add 200 µl cold (4˚) Solution I. Vortex to resuspend cells completely; make sure no
clumps of cells remain.
4. Add 200 µl fresh Solution II. Do not vortex. Invert tubes about 6 times to lyse the
cells. The lysate should become relatively clear.
5. Add 200 µl cold (4˚) Solution III. Do not vortex. Invert tubes about 6 times to
thoroughly mix. A dense fluffy white precipitate should form.
6. Spin down in microcentrifuge, at max. speed (~12,000 rpm), 5 min.
7. Transfer 500 µl of the supernatant to a fresh tube, carefully avoiding solid clumps.
Discard the tube with the dregs of liquid and pelleted material.
8. Add 1.0 ml of cold (-20˚) 100% ethanol. Mix well by inverting and brief vortexing.
(Optional: incubate at –20˚ for 30 minutes which can increase final yield). Spin down in
microcentrifuge, at max. speed (~12,000 rpm), 15 min.
9. A pellet should be readily visible. Decant supernatant. Invert the tube and tap briefly
on paper towels. Add 100 µl of cold (-20˚) 70% ethanol. Spin down in microcentrifuge,
at max. speed (~12,000 rpm), 2 min.
10. Remove all of 70% ethanol supernanant carefully from pellet by aspirating. Leave
tubes open on benchtop for ~5 min to dry briefly. Add 25 µl TE pH 7.5. Pellets should
readily dissolve.
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For plasmids, 1 µl of resuspended DNA is usually plenty for a restriction digest.
For BACs, use at least 5-10 µl per digest if using standard 6-cutter enzymes.
Since BACs are low copy vectors you may need to prep more than 1.5 ml of culture to
get enough DNA. Simply scale up the protocol accordingly. For PFGE analysis of rarecutter enzyme digests, I suggest ~3 µl per digest. Do not repeatedly freeze-thaw BAC
DNAs – they will shear and not be suitable for PFGE. On the other hand the crude
miniprep DNA can degrade if stored at 4˚.
Solutions for alkaline lysis:
Solution I (a.k.a. GTE) + RNAse A
100 ml
500 ml
final conc:
50 mM Glucose
0.9 g
4.5 g
25 mM Tris pH 8.0
2.5 ml 1 M
12.5 ml 1 M
10 mM EDTA
2.0 ml 0.5 M
10.0 ml 0.5 M
RNAse A 50 µg/ml
0.5 ml 10mg/ml
2.5 ml 10mg/ml
ddH20:
to 100 ml
to 500 ml
Filter-sterilize and store at 4˚ C.
Solution II
100 ml
Make fresh or store in plastic tubes (not glass bottles).
final conc:
0.2 M NaOH
2 ml 10 M
1 % SDS
10 ml 10%
88 ml ddH20
Solution III
1 liter
Dissolve 294.5 g Potassium Acetate in approx. 500 ml ddH20. Add glacial acetic acid to
lower pH to 5.5 (will require 150-200 mls). Bring final volume up to 1 liter with ddH20.
Store at 4˚ C.
TE pH 7.5 is:
10 mM Tris, diluted from 1M pH 7.5 stock
1 mM EDTA, diluted from 0.5M pH 8.0 stock
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Doug Mortlock updated Jan. 21, 2014