Recombineering: genetic engineering in bacteria using homologous recombination.

Abstract

The bacterial chromosome and plasmids can be engineered in vivo by homologous recombination using PCR products and synthetic oligonucleotides as substrates. This is possible because bacteriophage-encoded recombination functions efficiently to recombine sequences with homologies as short as 35 to 40 bases. This recombineering allows DNA sequences to be inserted or deleted without regard to location of restriction sites. This unit first describes preparation of electrocompetent cells expressing the recombineering functions and their transformation with dsDNA or ssDNA. Support protocols describe a two-step method of making genetic alterations without leaving any unwanted changes, and a method for retrieving a genetic marker (cloning) from the E. coli chromosome or a co-electroporated DNA fragment and moving it onto a plasmid. A method is also given to screen for unselected mutations. Additional protocols describe removal of defective prophage, methods for recombineering.

Full text

Page 1

SECTION V
SPECIALIZED TECHNIQUES
UNIT 1.16
Recombineering: Genetic Engineering in
Bacteria Using Homologous Recombination
Over the past few years, in vivo technologies have emerged which, due to their efficiency
and simplicity, already complement and may one day replace standard genetic engineer-
ing techniques. The bacterial chromosome and episomes can be engineered in vivo by
homologous recombination using PCR products and synthetic oligonucleotides (oligos)
as substrates. This is possible because bacteriophage-encoded recombination functions
efficiently recombine sequences with homologies as short as 35 to 40 bases. This tech-
nology, termed recombineering, allows DNA sequences to be inserted or deleted without
regard to presence or location of restriction sites.
To perform recombineering, a bacterial strain expressing a bacteriophage recombination
system is required. The phage enzymes can be expressed from either their own promoter
or from a heterologous regulated promoter. Expressing the genes from their endogenous
phage promoter confers the advantage of tight regulation and coordinate expression,
which results in higher recombination frequencies. This is an important advantage, since
in many cases high recombination frequencies will be essential to obtaining a desired
recombinant. The authors of this unit routinely use a defective λ prophage located on the
E. coli chromosome, and have recently transferred the critical elements of this prophage
to a number of different plasmids (Thomason et al., 2005; also see Commentary). In
this prophage system, the phage recombination functions are under control of the bac-
teriophage λ temperature-sensitive cI857 repressor. At low temperatures (30◦to 34◦C),
the recombination genes are tightly repressed, but when the temperature of the bacterial
culture is shifted to 42◦C, they are expressed at high levels from the λ pLpromoter. In the
plasmid construct of Datsenko and Wanner (2000), the recombination genes are located
on a plasmid and expressed from the arabinose promoter. The Datsenko and Wanner
plasmid and some of the authors’ plasmid constructs have temperature-sensitive origins
of DNA replication. The plasmid-based systems have the advantage of mobility—they
can be transferred among different E. coli strains or to Salmonella typhimurium and
possibly other gram-negative bacteria. However, using the prophage system located on
the bacterial chromosome is more facile if the recombineering is targeted to a plas-
mid. After induction of the recombination functions, the modifying DNA, either a
double-stranded (ds) PCR product or a synthetic single-stranded (ss) oligonucleotide
(oligo), is introduced into the prophage-containing strain by electroporation. Recombi-
nants are obtained either by selection or screening of the population of cells surviving
electroporation. Once the desired construct is obtained, the prophage can be removed
by another recombination. Alternatively, engineered alleles on the chromosome can be
moved into a different host by P1 transduction. Plasmids with temperature-sensitive
replication origins can be lost from the recombinant strain by growth at the appropriate
temperature.
Preparation of electrocompetent cells that have expressed the recombineering functions
from the λ promoter and their transformation with double-stranded DNA (dsDNA) or
single-stranded DNA (ssDNA) is the first procedure described in this unit (see Basic
Protocol1).SupportProtocol1describesatwo-stepmethodofmakinggeneticalterations
without leaving any unwanted changes. In the latter protocol, the first of the two series
Contributed by Lynn Thomason, Donald L. Court, Mikail Bubunenko, Nina Costantino,
Helen Wilson, Simanti Datta, and Amos Oppenheim
Current Protocols in Molecular Biology (2007) 1.16.1-1.16.24
Copyright C ?2007 by John Wiley & Sons, Inc.
Escherichia coli,
Plasmids, and
Bacteriophages
1.16.1
Supplement 78

Page 2

Genetic
Engineering in
Bacteria Using
Homologous
Recombination
1.16.2
Supplement 78Current Protocols in Molecular Biology
of steps is a recombineering reaction that replaces the sequence to be modified with an
antibiotic-resistance cassette and a counter-selectable marker (e.g., sacB, which is toxic
when cells are grown on medium containing sucrose; Gay et al., 1985). The second
series of steps is a subsequent recombination that replaces the antibiotic cassette and
the counter-selectable marker with the desired genetic alteration. Moreover, the same
selection and counter-selection can be reused to make further modifications. Support
Protocol 2 describes a method for retrieving a genetic marker (cloning) from the E.
coli chromosome or a co-electroporated DNA fragment and moving it onto a plasmid.
Whereastheaboveprotocolsgenerallyuseselectiontoidentifytherecombinants,Support
Protocols3and4describemethodstoscreenforunselectedmutations.SupportProtocol5
detailsmodificationsnecessarywhentargetingmulticopyplasmidswithrecombineering.
Basic Protocol 2 describes the removal of the defective prophage. Finally, Alternate
Protocols 1 and 2 present methods for recombineering with an intact prophage and for
introducing mutations onto bacteriophage λ, respectively.
STRATEGIC PLANNING
Before attempting to modify the E. coli chromosome or a plasmid, the DNA sequence of
thedesiredfinalconstructshouldbedetermined.ADNA-analysiscomputerprogramsuch
asGeneConstructionKit(GCK;TextcoSoftware;http://www.textco.com/)orVectorNTI
(Invitrogen) is invaluable for this task. Having the sequence of both the original genome
arrangement and the designed final construct as electronic files facilitates the design of
oligonucleotides to be used as primers for PCR or as ssDNA recombination substrates
themselves. The computer-determined sequences also allow rapid design of primers to
analyze and verify the potential recombinants. One must be aware of gene-regulation
issues when designing the constructs. Bringing in a promoter with an antibiotic cassette
can help in establishing drug resistance; however, transcription from this promoter can
extend beyond the drug marker and affect distal genes. The authors of this unit have
designedseveraldrugcassetteswiththeirpromoter,openreadingframe,andtranscription
terminator region, as described in Yu et al. (2000); primers for amplification are listed
in Table 1.16.2. One must also be careful of possible polarity effects and avoid creating
unwanted fusion proteins when generating recombinants.
The DNA substrate used to generate the recombinants depends on the desired change.
For a sizeable insertion, such as a drug cassette, a PCR product is generated that contains
40 to 50 base pairs of flanking homology to the chromosomal or plasmid target at each
end. This homology is provided at the 5?end of each synthetic primer. Following this
region of chromosomal homology, ∼20 bases of homology to the drug cassette provides
the primers to amplify the cassette sequence. Thus, two primers are designed that will
each be 60 to 70 nucleotides (nt) long: the 5?ends provide homology to the targeted
region and the 3?ends provide homology to the cassette (see Fig. 1.16.1). Careful primer
designiscrucial(seeabove).TheefficiencyofrecombineeringwithdsDNAcanapproach
0.1% (Yu et al., 2000). If deletions, small substitutions, or base changes are desired, a
synthetic single-stranded oligo of ∼70 to 100 nt can be used. The oligo should have
35 to 40 nt of complete homology flanking the alteration. Recombineering with ssDNA
in wild-type E. coli containing the defective λ prophage gives efficiencies approaching
1% (Ellis et al., 2001), and if host mismatch repair is inactivated, either by mutation or
by using an oligo that creates a C-C mismatch, a 20% to 25% recombination frequency
is achievable (Costantino and Court, 2003). This extremely high frequency means that,
for oligo recombination, it is possible to create recombinants without a selection and
find them by screening. Since an oligonucleotide corresponding to the lagging strand of
DNA replication is some 20-fold more efficient than its complement, it is worthwhile to
determine the direction of replication through one’s region of interest and use the oligo
that corresponds to the lagging strand (Costantino and Court, 2003).

Page 3

Escherichia coli,
Plasmids, and
Bacteriophages
1.16.3
Current Protocols in Molecular BiologySupplement 78
Figure 1.16.1
used to PCR amplify the antibiotic cassette. The PCR product is introduced by electroporation into
cells induced for the Red recombineering functions. The Red functions catalyze the insertion of
the cassette at the target site, which may be on the bacterial chromosome or on a plasmid.
Targeting of an antibiotic cassette. Two primers with 5?homology to the target are
It is also possible to rescue or clone gene(s) from the bacterial chromosome onto a
plasmid. To do this, create a linear PCR product of the plasmid using primers with
homologyflankingthetargetsequence,designedsothatthesequencewillbeincorporated
ontothecircularplasmidintheappropriateorientation.ThePCR-amplifiedlinearplasmid
will require an origin of DNA replication and a selective marker.
Multicopy plasmids can also be modified with recombineering, and changes occur at the
samefrequencyasmodificationsoftheE.colichromosome(Thomasonetal.,2007).This
means that point mutations can be made at a frequency of ∼5% to 30% in the absence of
mismatch repair. The frequency of insertion or removal of large DNA segments is much
lower,however,andisolationofrecombinantspecieswillbeproblematicintheabsenceof
a selection or screen. Designing the loss of a unique restriction site into the recombinant
plasmid permits enrichment of the recombinant class by allowing destruction of the
unmodified parental population by restriction digestion.
Some modification of Basic Protocol 1 is needed when targeting plasmids with recom-
bineering, due to the fact that they exist in multiple copies in the cell. Different plasmids
vary widely in their intracellular copy number, ranging from low copy vectors with only
a few molecules per cell, such as pSC101, to pUC-based plasmids that routinely have
∼500 copies per cell. This means that it is generally impossible to modify all copies
of the episome of interest during a recombineering reaction, and thus cells containing

Page 4

Genetic
Engineering in
Bacteria Using
Homologous
Recombination
1.16.4
Supplement 78Current Protocols in Molecular Biology
recombinant plasmids will also contain unmodified parental plasmid. The recombinant
class of molecules can be separated from the unmodified class by plasmid DNA isolation
and retransformation at a DNA concentration of less than one molecule per cell, which
creates pure clonal populations of the recombinant plasmid species. Parental plasmids
can also be eliminated by restriction digestion as mentioned above.
A complexity arising when plasmids are modified with recombineering is that circular
multimeric plasmid species are formed during the recombination reaction. In the absence
of linear substrate DNA, expression of the Red proteins does not cause plasmids to
multimerize under the conditions given here (Thomason et al., 2007), but when linear
single-strandedordouble-strandedsubstrateDNAisaddedandmodificationofthetarget
sequence occurs, circular multimeric species are formed. Multimer formation is thus a
hallmark of a recombinant plasmid molecule (and could theoretically be used to identify
recombinant species). It is important to begin recombineering with a monomer plasmid
species and to do the reaction in a recA mutant bacterial strain defective in homologous
recombination. Using a host defective in RecA-mediated recombination eliminates one
route to plasmid-by-plasmid recombination.
A decision should be made as to whether the plasmid will be introduced into the Red-
expressing cells by co-electroporation with the linear DNA after the recombination
functions are induced, or whether the plasmid should already be established in the
bacteria. Co-electroporation offers the advantage that it helps control plasmid copy
number at the time of recombineering and minimizes opportunities for plasmid multimer
formation. However, for very large plasmids of low copy number, co-electroporation
may not be feasible.
BASIC
PROTOCOL 1
MAKING ELECTROCOMPETENT CELLS AND TRANSFORMING WITH
LINEAR DNA
This basic protocol describes making electrocompetent cells that are preinduced for the
recombination functions and transforming them with the appropriate DNA to create
the desired genetic change. As noted in Strategic Planning, the phage recombination
functions are repressed by the phage λ temperature-sensitive cI857 repressor, so that
they are not expressed when the cells are grown at low temperature (30◦to 34◦C) but
are highly expressed when the culture is shifted to 42◦C. See Commentary for additional
considerations before executing the procedure.
Materials
Purified PCR product or oligonuclotide primers with ∼40 to 50 bases of flanking
homology on either side of desired change (also see UNIT 15.1)
Bacterial strain expressing the defective lambdoid prophage recombination system
λ Red (Table 1.16.1; strains are available from the Court Laboratory;
court@ncifcrf.gov)
LB medium and plates (UNIT 1.1), without antibiotic
Medium lacking carbon source: M9 medium (UNIT 1.1) or 1× TM buffer (APPENDIX 2)
Selective plates (UNIT 1.1)—minimal plates if selecting for prototrophy or rich plates
containing antibiotic (depending on drug cassette used):
30 µg/ml ampicillin
30 µg/ml kanamycin
10 µg/ml chloramphenicol
12.5 µg/ml tetracycline
50 µg/ml spectinomycin
30◦to 32◦C incubator
32◦and 42◦C shaking water baths

Page 5

Escherichia coli,
Plasmids, and
Bacteriophages
1.16.5
Current Protocols in Molecular Biology Supplement 78
Table 1.16.1
Bacterial Strains Commonly Used for Recombineering
Strain Genotype
DY329
DY330
DY331
W3110 ?lacU169 nadA::Tn10 gal490 pgl?8 λcI857?(cro bioA) (TetR)
W3110 ?lacU169gal490 pgl?8 λcI857 ?(cro-bioA)
W3110 ?lacU169 srlA::Tn10 ?recA gal490 pgl?8 λcI857
?(cro-bioA) (TetR)
W3110 λcI857 ?(cro-bioA)
mcrA ?(mrr-hsdRMS-mcrBC) φ80dlacZ?M15 lacX74 deoR recA1
endA1 araD139 ?(ara, leu)7697 galU gal490 pgl?8 rpsL nupG
λ(cI857ind1) ?{(cro-bioA)<>tetRA} (TetR)
W3110 gal490 pgl?8 λcI857 ?(cro-bioA) int<>cat-sacB
W3110 ?lacU169 λcI857 ?(cro-bioA)
W3110 gal490 pgl?8 λcI857 ?(cro-bioA)
W3110 ?lacU169 λcI857 ?(cro-bioA) galKam mutS<>amp
W3110 ?lacU169 λcI857 ?(cro-bioA) galKam uvrD<>kan
aDH10B derivative (Invitrogen).
bGives less background on low concentrations of chloramphenicol than DY378.
DY378
DY380a
DY441
HME5
HME45b
HME63
HME64
125- and 250-ml Erlenmeyer flasks, preferably baffled
Refrigerated low-speed centrifuge with Sorvall SA-600 rotor (or equivalent)
35- to 50-ml plastic centrifuge tubes
0.1-cm electroporation cuvettes (Bio-Rad), chilled
Electroporator (e.g., Bio-Rad E. coli Pulser)
Additional reagents and equipment for PCR (UNIT 15.1), agarose gel electrophoresis
of DNA (UNITS 2.5A & 2.6), purification of DNA by ethanol precipitation (UNIT 2.1A;
optional; commercially available PCR cleanup kit may be substituted),
electroporation (UNIT 1.8), isolation of bacterial colonies by streaking (UNIT 1.3),
restriction enzyme digestion (UNIT 3.1), and DNA sequencing (Chapter 7)
Prepare DNA for transformation
1. Design and procure the oligos to use for PCR-mediated generation of a dsDNA
product, or for use in single-stranded oligo engineering.
The sequences of the primers used to amplify the common drug cassettes are listed in
Table 1.16.2. Remember to add the homologous targeting sequence to the 5?ends of the
oligos.
UNIT 15.1 describes general considerations for primer design.
2. MakethePCRproduct(UNIT15.1),examineitbyagarosegelelectrophoresis(UNIT2.5A),
and gel purify by isolating the desired band (UNIT 2.6) if unwanted products are
obtained.
If the DNA is gel purified, avoid exposing it to ultraviolet light, which will damage it and
result in lower recombination frequencies.
3. CleanupthePCRproductbyethanolprecipitation(UNIT2.1A)orusingacommercially
available kit to remove salt.
Ifaplasmid template isused toconstruct a PCR-amplified drug cassette, any intact circu-
lar plasmid remaining will transform the cells efficiently and give unwanted background.
This background can be minimized by using a linear plasmid template for the PCR and
by digesting the completed PCR reaction with DpnI before using it for electroporation.
Always include a control reaction of uninduced cells transformed with the PCR product,
to give a measure of any unwanted intact plasmid background.

Page 6

Genetic
Engineering in
Bacteria Using
Homologous
Recombination
1.16.6
Supplement 78 Current Protocols in Molecular Biology
Table 1.16.2
(from Yu et al., 2000)
PCR Primers and Suggested Source of Template for Amplifying Drug Cassettes
Gene Source Primer sequence
Ampicillin pBluescript SK(+)
(Stratagene)
Tn10
5?CATTCAAATATGTATCCGCTC
5?AGAGTTGGTAGCTCTTGATC
5?CAAGAGGGTCATTATATTTCG
5?ACTCGACATCTTGGTTACCG
5?TGTGACGGAAGATCACTTCG
5?ACCAGCAATAGACATAAGCG
5?TATGGACAGCAAGCGAACCG
5?TCAGAAGAACTCGTCAAGAAG
5?ACCGTGGAAACGGATGAAGGC
5?AGGGCTTATTATGCACGCTTAA
Tetracycline
Chloramphenicol pPCR-Script Cam
(Stratagene)
Tn5
Kanamycin
Spectinomycin DH5αPRO (Clontech)
Prepare bacterial cultures
4. Inoculate the suitable bacterial strain (Table 1.16.1) from frozen glycerol stock or a
single colony into 3 to 5 ml LB medium. Shake at 30◦to 32◦C overnight.
Most of the authors’ strains containing the defective lambdoid prophage are W3110
derivatives; however, the prophage can be moved into other backgrounds. See
Commentary for details. Plasmids expressing the recombination functions can be put
into any strain of choice.
Either 30◦or 32◦C is acceptable for the low temperature throughout the procedure, since
either temperature allows good repression by the cI857 repressor. The cultures will grow
more rapidly at 32◦C.
5. Add ∼0.5 ml of the overnight culture to 35 ml of LB medium in a 250-ml (baffled)
Erlenmeyer flask.
This is a 70-fold dilution. One must make sure that one’s dilution is at least 50-fold.
Higher dilutions will also work, but the cells will take longer to grow to the appropriate
density. If targeting the recombineering to a plasmid, or expressing from a plasmid, add
antibiotic as appropriate to maintain selection during growth. If an alternate method of
inducing the recombination functions is used (i.e., addition of arabinose), the inducer
should be added to the medium. In this case remember to include an additional flask
containing an uninduced culture as a negative control. Expression of the λ Red functions
fromtheplasmidsofDatsenkoandWanner(2000)isenhancedbyuseof10mMarabinose
(for Ara+strains).
6. Place the flask in the 32◦C shaking water bath and grow cells at 32◦C with shaking
for ∼2 hr.
The time will vary with different strains and dilutions. The cells are ready when the A600
is between 0.4 and 0.6. It is important not to over-grow the cells, since stationary phase
cells do not express the recombination functions well.
Induce recombination functions
7. Transfer half the culture to a 125-ml (baffled) Erlenmeyer flask and place that flask
in the 42◦C water bath. Shake 15 min at 220 rpm to induce. Leave the remainder
of the culture at 32◦C; this will be used as the uninduced control that lacks recom-
bination activity. While the cells are inducing, fill an ice bucket with an ice-water
slurry.

Page 7

Escherichia coli,
Plasmids, and
Bacteriophages
1.16.7
Current Protocols in Molecular BiologySupplement 78
8. Immediately after inducing for 15 min at 42◦C, rapidly cool the flask in the ice-
water slurry with gentle swirling. Leave on ice for ≥5 min. Follow the same cooling
protocol with the uninduced 32◦C culture. While the cells are on ice, precool the
centrifuge to 4◦C and chill the necessary number of 35- to 50-ml plastic centrifuge
tubes, labeled for induced and uninduced cells.
The temperature shift is unnecessary when a chemical inducer like arabinose is used.
Make electrocompetent cells
9. Transferboththeinducedanduninducedculturestotheappropriatelylabeledchilled
35- to 50-ml centrifuge tubes. Centrifuge 7 min at 4600 × g (6700 rpm in a Sorvall
SA-600 rotor), 4◦C. Aspirate or pour off supernatant.
10. Add 1 ml ice-cold distilled water to the cell pellet in the bottom of each tube and
gently resuspend cells with a large pipet tip (do not vortex). Add another 30 ml
ice-cold distilled water to each tube, seal, and gently invert to mix, again without
vortexing. Centrifuge tubes again as in step 9.
All subsequent resuspensions of cells through step 16 should be done gently and without
vortexing. Preparation of the cells for electroporation washes out any added chemical
inducing agent.
11. Decant the 30-ml supernatant very carefully from the soft pellet in each tube and
resuspend each cell pellet in 1 ml ice-cold distilled water.
Remove tubes from the centrifuge promptly. The pellet is very soft and care should be
taken not to dislodge it, especially when processing multiple tubes.
12. Transfer resuspended cells to microcentrifuge tubes. Microcentrifuge 30 to 60 sec at
maximumspeed,4◦C.Carefullyaspiratesupernatant.Ineachofthetubes,resuspend
the cell pellet in 200 µl cold distilled water, which will provide enough material for
four or five electroporations.
Forroutineprocedureswhenoptimalrecombinationfrequencyisnotnecessary,e.g.,when
selection is used to find recombinants, electrocompetent cells can be stored at −80◦C
after resuspending the cell pellet in 15% (v/v) glycerol. For highest efficiency, use freshly
processed cells.
Introduce DNA by electroporation
13. Chill the desired number of 0.1-cm electroporation cuvettes on ice. Turn on the
electroporator and set to 1.80 kV.
Brands of cuvettes and electroporator other than Bio-Rad may work, but have not been
tested in the authors’ laboratory. The larger 0.2-cm cuvettes may require different elec-
troporation conditions (consult electroporator instruction manual) and standardization
to obtain optimal recombination frequencies.
14. In microcentrifuge tubes on ice, mix 100 to 150 ng of salt-free PCR fragment (from
step 3) or 10 to 100 ng of single-stranded oligonucleotide with 50 to 100 µl of the
suspension of induced or uninduced cells (from step 12). Do the mixing and subse-
quent electroporation rapidly; do not leave the DNA-cell mixes on ice for extended
periods. Be sure to include the following electroporation reactions and controls:
a. Induced cells plus DNA.
This is the culture that should yield the designed recombinants.
b. Induced cells without DNA.
This is a control to identify contamination, determine the reversion frequency, and
obtain some idea of the efficiency of the selection.

Page 8

Genetic
Engineering in
Bacteria Using
Homologous
Recombination
1.16.8
Supplement 78 Current Protocols in Molecular Biology
c. Uninduced cells plus DNA.
This control tells whether there is some contaminating factor in the DNA that is
contributing to the selected colonies (for example, intact plasmid template from the
PCR reaction will give rise to drug-resistant colonies here).
15. Introduce the DNA into the cells by electroporation (UNIT 1.8).
The time constant should be greater than 5 msec for optimal results. Low time constants
indicate problems with the cells, the DNA, or even the equipment.
16. Immediately after electroporation, add 1 ml LB medium to the cuvette using a
micropipettor with a 1000-µl pipet tip. If transforming with a drug cassette, transfer
the electroporation mix to sterile culture tubes and incubate the tubes with shaking
at 30◦to 34◦C for 1 to 2 hr to allow expression of the antibiotic resistance gene.
Even if not selecting for drug resistance, it is still recommended that the cells outgrow to
recover from the shock of electroporation. It has been observed in the authors’ laboratory
that omitting the outgrowth reduces the cell viability ∼10-fold.
An alternative, and in the author’s experience more reliable, method for outgrowth is to
spread appropriate dilutions of cells (see step 18 below) on a sterile 82-mm-diameter
nitrocellulose filter atop a rich (LB) plate. Incubate this plate for 3 hr at 30◦to 34◦C.
After the incubation, transfer the filter to the appropriate selective drug plate using
sterile forceps. This method is preferable because the cells are less dense and more
efficient outgrowth is achieved.
Determine cell titers
17. Makeserial1:10dilutionsoftheelectroporationmixthrough10−6usingM9medium
or 1× TM buffer, dispensing 0.9 ml M9 or TM and 0.1 ml of the cell suspension per
tube.
Thedilutionscanbemadeinrichmediumifaselectionforantibioticresistanceisapplied.
18. To determine total viable cell count, spread 100 µl of 10−5and 10−6dilutions on LB
plates (rich plates without drug). Incubate the plates at 30◦to 34◦C for 1 to 2 days,
depending on the growth requirements of the recipient strain.
19. To determine recombinant cell count, plate cells on selective plates as follows de-
pending on the anticipated recombinant frequency.
a. If efficient recombination is expected, spread both 10 and 100 µl of the 10−1and
10−2dilutions.
b. If low numbers of recombinants are expected, spread 100 µl each of a 1:5 and
1:10 dilution.
Theauthors routinely obtain ten-fold or higher recombinant yields withthe prophage
system than with the Datsenko and Wanner plasmids.
For the no-DNA and uninduced controls, plate 200 µl directly on selective plates.
Since targeting to the chromosome results in a lower copy number of the drug cassette
than is present with a multicopy plasmid, antibiotic concentrations must be adjusted
accordingly. The authors routinely use the drug concentrations recommended above for
chromosomal constructs. Minimal plates are used for selection based on prototrophy.
20. Incubate plates at the appropriate temperature (30◦to 34◦C).
At 30◦C, colonies may take two days to come up on LB plates and 3 to 4 days on minimal
plates. Candidates should be purified by streaking for single colonies and retested for the
appropriate phenotype.

Page 9

Escherichia coli,
Plasmids, and
Bacteriophages
1.16.9
Current Protocols in Molecular BiologySupplement 78
Analyze recombinants
21. Once recombinant clones are identified, confirm the presence of the desired mu-
tation(s) by PCR analysis (UNIT 15.1) followed by DNA sequencing (Chapter 7) or
restriction digestion analysis (UNIT 3.1), if appropriate.
ThedesignofthePCRprimersdependsonthechangesmade.Foranantibioticcassetteor
other insertion, the primer pair used to amplify the insertion can also be used to confirm
its presence. These primers will not determine whether the cassette has integrated at
the desired location, however. The recombinant junctions can be confirmed with the
help of two additional primers (all four should have compatible annealing temperatures)
pointingoutwardsfromthecassette;useoneprimerflankingthecassetteandoneinternal
cassette primer to amplify the unique junctions created by the recombination reaction. A
primerpairthathybridizestotheexternal flankingsequences oneachsideratherthanthe
insertion itself can also be used to demonstrate loss of the target sequences and presence
of the insertion. Unwanted mutations can be introduced by heterologies (variations) in
the synthetic primer population (Oppenheim et al., 2004); therefore, it is important to
confirm the final construct by sequence analysis, especially the regions derived from the
original primers.
SUPPORT
PROTOCOL 1
MANIPULATING cat-sacB FOR COUNTER-SELECTION AND GENE
REPLACEMENT
This protocol describes a two-step method to create precise genetic changes without
otherwise altering the DNA sequence. First, cat-sacB (or another counter-selectable
cassette) is placed on the DNA; this is then replaced with the desired alteration in a
second recombineering event. The final construct will not have a drug marker.
Additional Materials (also see Basic Protocol1)
Template for amplification of cat-sacB: bacterial strain DY441 (DY329 with a
cat-sacB insertion on the E. coli chromosome) or the plasmid pEL04 (Lee et al.,
2001). pEL04 has previously been called both pK04 and pcat-sacB. Both
DY441 and pEL04 are available from the Court Laboratory; court@ncifcrf.gov.
SpeI and DpnI restriction endonucleases (UNIT 3.1)
Primer L sacB: 5?-homology sequence-ATC AAA GGG AAA ACT GTC CAT
AT-3?
Primer R cat: 5?-homology sequence-TGT GAC GGA AGA TCA CTT CG-3?
Invitrogen Platinum High Fidelity enzyme
LB-Cm plates: LB plates (UNIT 1.1) containing 10 µg/ml chloramphenicol
M63 minimal glycerol plates with sucrose (see recipe) or LB plates (UNIT 1.1)
lacking NaCl but containing 6% (w/v) sucrose
Medium lacking carbon source: e.g., M9 medium (UNIT 1.1)
Thermal cycler (MJ Research PTC-100)
Amplify the cat-sacB element for recombineering
1. If using pEL04, completely cleave pcat-sacB with SpeI.
This step is unnecessary if the chromosomal insertion of cat-sacB is used as template for
PCR. Failure to digest when the plasmid is used as template will give a high background
of intact plasmid transformants.
2. Amplifycat-sacB,eitherfromthecleavedpEL04orfromDY441withtheInvitrogen
Platinum High Fidelity enzyme and an MJ Research thermal cycler, using 50 pmol
of each primer and the following cycling program:

Page 10

Genetic
Engineering in
Bacteria Using
Homologous
Recombination
1.16.10
Supplement 78Current Protocols in Molecular Biology
1 cycle:
9 cycles:
2 min
15 sec
30 sec
3.5 min
15 sec
30 sec
3.5 min
(adding 5 sec/cycle)
94◦C
94◦C
55◦C
68◦C
94◦C
55◦C
(denaturation)
(denaturation)
(annealing)
(extension)
(denaturation)
(annealing)
19 cycles:
68◦C
68◦C
4◦C
(extension)
(extension)
(hold)
1 cycle:
1 cycle:
7 min
indefinite
The PCR primers used to amplify the cat-sacB element are at the 3?end of chimeric
primers that include 5?segments of bacterial target homology ∼40 to 50 nt in length.
Hence each primer is 60 to 70 nt long.
Since the PCR product is greater than 3 kb, it can be difficult to amplify. The above
conditions have been used successfully in the authors’ laboratory.
3. If pEL04 was used as template, digest the completed PCR reaction with DpnI to
specifically eliminate the methylated plasmid template. Purify the PCR product to
remove salt (UNIT 2.1A).
Perform electroporation and recombination to insert cat-sacB at the desired location
4. Insert the cat-sacB cassette into the chromosome as described above (see Basic
Protocol1, steps 4 to 21; also see Background Information for important tips) using
the following techniques specific for the cat-sacB cassette.
a. Select chloramphenicol-resistant (CmR) colonies and purify on LB-Cm plates to
isolate single colonies.
b. Test several CmRisolates for sensitivity to sucrose, either on minimal glycerol
plates containing 5% sucrose or on LB plates lacking NaCl but containing 6% su-
crose (Blomfield et al., 1991). Use the parental transformation strain as a sucrose-
resistant control. Determine that the insertion is correct before proceeding with
the next step of the procedure.
Thesucrose-resistantparentservesasacontrolforgrowthinthepresenceofsucrose.
Sucrose sensitivity needs to be tested because the PCR process generates mutations
that inactivate sacB in a fraction of the clones.
Perform electroporation and recombination to replace cat-sacB with chosen allele
5. UseaconfirmedCmR/sucrose-sensitivecandidatefromstep4asthestartingbacterial
strain for a second round of recombineering by carrying out Basic Protocol1, but
suspend the electroporated cells at step 16 in a final volume of 10 ml instead of 1 ml
of LB medium, and incubate with aeration (applied via shaking in a shaking water
bath) at 30◦to 32◦C for 4 hr to overnight for outgrowth following electroporation.
The higher dilution promotes better cell recovery and allows complete segregation of
recombinant chromosomes that no longer carry the cat-sacB cassette from nonrecom-
binant sister chromosomes that still contain it. If outgrowth is inadequate and sister
chromosomes are not fully segregated, the presence of the cassette on one chromosome
will confer sucrose sensitivity to the entire cell, thus preventing recovery of recombinants.
This is generally true in counter-selection experiments and illustrates a common problem
encountered with them.
6. Centrifuge cells 7 min at 4600 × g, 4◦C. Remove supernatant, then wash the cells
twice, each time by resuspending in l ml minimal medium lacking a carbon source,
such as M9, centrifuging again as before, and removing the supernatant. Resuspend
and dilute the cells for plating, and spread appropriate dilutions of the cells on either

Page 11

Escherichia coli,
Plasmids, and
Bacteriophages
1.16.11
Current Protocols in Molecular Biology Supplement 78
LB sucrose plates or minimal glycerol plates (if the defective prophage was moved
to another strain, the metabolic requirements of that strain must be considered). In
parallel, be sure to include a control of electroporated cells to which no PCR product
was added, to determine the frequency of spontaneous sucrose-resistant mutants;
these spontaneous mutants will retain the CmR cassette.
The frequency of spontaneous sucrose-resistant cells is normally ∼1 in 104. Thus, in
the recombination experiment, the sucrose-resistant colonies that arise are of two types,
spontaneous mutants like those found in the control, and deletions caused by replacing
cat-sacB by recombination with the PCR product. The frequency of the latter is optimally
10- to 100-fold greater than that of spontaneous mutation.
7. Purify∼10sucrose-resistantcoloniesbystreakingtoisolatesinglecolonies(UNIT 1.3)
and then test for chloramphenicol sensitivity (CmS)—spontaneous mutants will be
CmRwhiletherecombinantswillbeCmS.ScreentheCmS/sucrose-resistantcolonies
by PCR (see Basic Protocol1, step 21), then sequence to confirm the presence of the
desired change.
Screen a minimum of ten colonies. If high-efficiency recombination is not achieved, more
colonieswillneedtobescreened.Notethatthefrequencyofspontaneousmutantsremains
relatively constant and provides an internal control for determining the efficiency of the
recombination.
SUPPORT
PROTOCOL 2
RETRIEVAL OF ALLELES ONTO A PLASMID BY GAP REPAIR
Often it is desirable to retrieve a DNA sequence from the bacterial chromosome, either
to clone and amplify a gene, to express a gene under a given promoter, or to create a
gene or operon fusion. To do this with recombineering, a PCR product with homology to
the target at the ends is made from a linearized plasmid DNA and introduced into cells
expressing the Red system. This homology will allow recombination with the sequence
to be retrieved, yielding a circular plasmid containing the sequence. It is important to
linearize the plasmid DNA used as template. This retrieval method works with ColEI
and p15A (pACYC) replicons, but not with pSC101 (Lee et al., 2001).
If the desired gene is not present on the chromosome, it can be provided as a PCR
product and introduced into the cells along with the PCR-amplified vector DNA by co-
electroporation. Always remember to provide flanking homology so that the plasmid can
recombine with the additional PCR product (it is easier to provide the homology on the
short product to be cloned rather than on the vector).
Additional Materials (also see Basic Protocol1)
Plasmid onto which sequence of choice is to be rescued
Restriction enzyme(s) (UNIT 3.1) that do not cut within plasmid region to be
amplified
Synthetic chimeric primers providing homology to sequence flanking gene of
choice and to the plasmid sequence to be amplified
Amplify linear plasmid PCR product with homology to the target
1. Design primers with homology that flanks the desired target.
Drawing a sketch of the plasmid as a gapped circle interacting with the target sequence
will help one visualize the recombination reaction (see Fig. 1.16.2), since the plasmid
template has the linear ends pointing toward each other. Both of the primers will have
∼50 nt of bacterial sequence homology at the 5?ends linked to 3?plasmid sequence.
It is not necessary to amplify the entire plasmid. Any portion can be amplified as long
as the minimal requirements of a selectable marker and an origin of DNA replication
are met. If the DNA to be retrieved is adjacent to an antibiotic resistance gene, only a
plasmid replication origin need be amplified; the origin can be used to retrieve both the
desired sequence and the nearby drug marker. Avoid having other regions of the plasmid

Page 12

Genetic
Engineering in
Bacteria Using
Homologous
Recombination
1.16.12
Supplement 78 Current Protocols in Molecular Biology
Figure 1.16.2
to the target at the ends (indicated by dark arrows) is generated by PCR. The plasmid is introduced
by electroporation into cells expressing the Red functions, which catalyze recombination of the
vector with the target site, resulting in incorporation of the gene onto the plasmid.
Cloning genes by gap repair of a plasmid. A linear plasmid with flanking homology
that are homologous to the bacterial chromosome (e.g., lac); these can lead to unwanted
rearrangements.
2. To minimize background, digest the plasmid with one or more restriction enzyme(s)
that do not cut within the region to be amplified. Amplify the linear plasmid by PCR
using the primers (reaction conditions will need to be established empirically).
The amplified product will be a linear gapped plasmid with flanking homology to either
side of the allele to be rescued from the chromosome.
UsetheleastamountofplasmidDNApossibleforthePCRtemplate,tominimizetheback-
groundoffalsepositives.DigestionofthecompletedPCRreactionmixwithDpnIwillhelp
remove the template plasmid. Purify the PCR product to remove salt before proceeding.
Transform induced cells with the linear plasmid and select recombinants
3. Introduce the linear plasmid PCR product into the strain (see Basic Protocol1, steps
4 to 21). If necessary, also add the PCR product to be co-electroporated. Select
for the marker on the plasmid, and transform the uninduced cells with the linear
plasmid PCR product, to determine the background of intact plasmid present. Purify
candidate colonies and screen them with PCR. Isolate the recombinant plasmids and
use them to retransform a standard cloning strain such as DH5α or XL2 Blue, to
generate pure clones.
Transformation into a recA mutant host ensures that the newly engineered plasmid does
not undergo additional rearrangement.
IftheDNAofinterestislinkedtoadrugmarkeranditisretrievedwithareplicationorigin
lacking a drug marker, dilute the electroporation mix into 10 ml LB medium and grow the
culture overnight nonselectively. The next day, isolate plasmid DNA and transform into
a high-efficiency cloning strain, selecting for the drug resistance of the rescued marker.
The transformation should be done with a low concentration of DNA, to minimize uptake
of multiple plasmids into the same cell. The advantage of using only the plasmid origin
for retrieval is that any possible background of religated vector is eliminated.
Gap repair is less efficient than targeting to the chromosome. The maximal yield achieved
in the authors’ laboratory is ∼1000 recombinants/108cells. A possible side reaction
is joining of the plasmid ends without incorporation of the chromosomal marker. This
is caused by small (>5 base) repeats near the linear ends (Zhang et al., 2000); it is
likely that the Red functions facilitate the short repeat recombination that generates this
background. The short repeat recombination can be reduced or eliminated by designing
primers that are free of such repeated sequences.

Page 13

Escherichia coli,
Plasmids, and
Bacteriophages
1.16.13
Current Protocols in Molecular BiologySupplement 78
SUPPORT
PROTOCOL 3
SCREENING FOR UNSELECTED RECOMBINANTS
When recombinant frequencies approach 1/1000, direct screening can often be used to
find recombinant colonies from total viable cells plated out nonselectively on LB. The
authors have successfully used the nonradioactive Roche DIG (digoxigenin) system for
colony hybridization to detect the recombinant bacterial colonies (Thomason, unpub.
observ.). For this method to be feasible, the sequence inserted by recombineering must
be unique to the recombinant and absent in the starting strain. The authors have used a
21-nt long DIG-labeled oligonucleotide probe to detect insertion of the same sequence.
For larger insertions or fusion proteins such as GFP derivatives, a labeled PCR product
orgel-purifiedfragmentcouldalsobeusedasaprobe.Botholigoprobes andlarger DNA
fragments could be radioactively labeled with32P.
The authors have also successfully isolated unselected recombinants when the genetic
change confers a slow growth phenotype by simply looking for colonies that grow more
slowly than the majority class. Another useful method is the mismatch amplification
mutationassay-PCR(MAMA-PCR;Chaetal.,1992;Swaminathanetal.,2001).MAMA-
PCR is capable of identifying single base changes by screening colonies.
Additional Materials (also see Basic Protocol1)
Flanking primers for PCR analysis of mutation of interest (also see UNIT 15.1)
Additional reagents and equipment for colony hybridization (e.g., UNIT 6.6)
1. Introduce the genetic change of interest (see Basic Protocol1, steps 1 to 18).
Possible changes include small in-frame deletions or a protein tag. For subtle changes,
it is helpful to engineer a restriction-site change that can be detected in a PCR product.
For detection by hybridization, confirm beforehand that the sequence does not exist in
the parental bacterial strain—e.g., using a BLAST search (UNIT 19.3; Altschul et al.,
1990).
2. Assuming an expected viable cell count in the transformation mix of ∼108, plate the
cells nonselectively on rich plates so that the expected number of colonies per plate
is ∼500.
Six plates should be adequate if the recombination is efficient. More crowded plates will
mean that fewer plates must be screened, but if the plates are too crowded it will be more
difficult to locate positive clones.
3. Screenforrecombinantsbyperformingcolonyhybridizationusingestablishedproce-
dures(e.g., UNIT 6.6).Ifusinganonradioactivelabelingmethod,followtheconditions
suggested by the manufacturer.
The required length of oligonucleotide probes will depend on the sensitivity of the system.
The authors have detected positives colonies at frequencies as low as 5 × 10−2to 1 ×
10−3.
4. Streak positive candidates to obtain pure clones and retest.
Since the recombination only alters one of several copies of the chromosome existing
in a single cell, the colony from that cell will be heterozygous for the allele. Thus, on
re-streaking, some fraction of the colonies will not give a positive signal. The signal can
again be detected by colony hybridization. If a new restriction site has been designed into
the construct, perform PCR and cut the product with the appropriate restriction enzyme
to detect the recombinant (also see UNIT 3.1).

Page 14

Genetic
Engineering in
Bacteria Using
Homologous
Recombination
1.16.14
Supplement 78Current Protocols in Molecular Biology
SUPPORT
PROTOCOL 4
SCREENING FOR UNSELECTED PLASMID RECOMBINANTS
With the extremely high oligonucleotide recombination frequencies obtainable in the
absence of mismatch repair (20% to 25% of total viable cells), direct sequencing of
unselected plasmid clones can be used to find recombinants. If the oligonucleotide
carrying the mutation to be introduced creates a C-C mismatch when paired at the target
site, the recombination can be done in wild-type cells containing the defective prophage,
since a C-C mismatch is not repaired. Otherwise, a bacterial strain with prophage and
mutant for the mismatch repair system can be used. It is helpful if the mutation to
be inserted creates a restriction site change that can be monitored by digestion of a
PCR-amplified fragment covering the region of interest.
Additional Materials (also see Basic Protocol1)
Cells expressing Red but mutant for host mismatch repair system or
oligonucleotide that creates C-C mismatch when annealed to target DNA strand:
e.g., HME63 and/or HME64 (see Table 1.16.1)
High-efficiency cloning strain (lacking the Red system)
1. Perform Basic Protocol1, introducing both the plasmid and the oligonucleotide by
co-electroporation into cells mutant for the host mismatch repair system, or using an
oligo that creates a C-C mismatch when annealed to the target DNA strand.
2. After the outgrowth, dilute the electroporation mix into 10 ml broth containing the
appropriate antibiotic for plasmid selection and grow the culture overnight.
3. Isolate plasmid DNA from this culture.
4. Using a low DNA concentration, transform the engineered plasmid into a high
efficiency cloning strain (lacking the Red system), selecting for drug resistance.
Purify colonies.
Use a low DNA concentration to minimize uptake of more than one molecule/cell.
5. Screen to find the mutation, either by direct sequencing or, if there is a restriction site
change, by amplifying a fragment with PCR and digesting it with the appropriate
enzyme(s).
Theauthorsrecommendlookingat25to50colonies.Thisprocedurehasbeensuccessfully
used for plasmid mutagenesis (Thomason, unpub. observ.).
SUPPORT
PROTOCOL 5
MODIFYING MULTICOPY PLASMIDS WITH RECOMBINEERING
Multicopy plasmids can be modified by recombineering with efficiencies similar to those
obtained when targeting the E. coli chromosome. This protocol optimizes the basic
procedure to deal with additional complexities arising when plasmids are targeted (see
Commentary). Addition of both the plasmid and the linear substrate DNA, either double-
or single-stranded, by co-electroporation allows better control over the initial ratio of
plasmid molecules to number of cells and minimizes opportunity for plasmid multimer
formation. After recombineering, the recombinant species will be contaminated with
unmodified parental plasmid and the two species must be separated.
Additional Materials (also see Basic Protocol 1)
Plasmid to be modified: monomer species, freshly isolated from recA mutant host
such as DH5α; determine plasmid DNA concentration by A260
recA mutant bacterial strain expressing Red functions (e.g., DY331 or DY380; see
Table 1.16.1)

Page 15

Escherichia coli,
Plasmids, and
Bacteriophages
1.16.15
Current Protocols in Molecular Biology Supplement 78
Appropriate selective plates, if needed (when selecting for drug resistance, use
drug concentrations appropriate for the multicopy plasmid used)
Additional reagents and equipment for plasmid miniprep (UNIT 1.6) and agarose gel
electrophoresis (UNIT 2.5A)
1. Follow Basic Protocol 1, steps 1 to 13, using a recA mutant strain containing the Red
functions.
2. In step 14 of Basic Protocol 1, mix the plasmid DNA and the linear DNA, either
single-stranded or double-stranded, with the cells. The DNAs will be introduced
into the cells in one electroporation reaction. Include the following electroporation
reactions and controls:
a. Induced cells plus plasmid and modifying DNA.
This is the culture that should yield the recombinant plasmids.
b. Induced cells without plasmid or linear DNA.
This is a control to identify contamination in the bacterial culture.
c. Induced cells plus plasmid only.
This control will provide a measure of the plasmid transformation efficiency and serve as
a negative control for recombinant selection or screening.
d. Uninduced cells plus plasmid and linear DNA.
This control will confirm that the “recombinants” are dependent on expression of the Red
system.
If the plasmid is introduced by co-electroporation with the linear substrate, enough
molecules of plasmid must be added to obtain a high transformation frequency, since
recombineering will occur only in cells receiving the plasmid. In most cases, ∼10 ng
of plasmid DNA per electroporation is sufficient, but it may be necessary to determine
empirically the appropriate amount of plasmid DNA to add, since the transformation
efficiency of the plasmid being used may differ. Ideally, each cell should receive about
one plasmid molecule.
If the plasmid is already resident in the cell, add only the modifying linear DNA at this
step.
3. Perform electroporation as in step 15 of Basic Protocol 1. After electroporation,
follow step 16 for outgrowth, using the longer 2 hr time.
4. Add9mlLBmediumandtheappropriateamountofantibioticforplasmidselection;
let the culture grow overnight at 30◦to 32◦C.
5. The following day, isolate plasmid DNA from the culture with a standard miniprep
procedure (UNIT 1.6).
If the recombinant has been designed to remove a unique restriction site, digest several
microliters of the DNA with this enzyme. This will help to eliminate the inevitable back-
ground of unmodified parental plasmid, thus enriching for the recombinant population.
6. Transform the plasmid miniprep DNA or the clean restriction digest into a standard
recA cloning strain such as DH5α at a low DNA concentration (less than one DNA
molecule/cell).
Electroporation is highly recommended, since it is much more efficient than chemical
transformation.
7. After a 2-hour outgrowth, plate for single colonies on the appropriate media. Plate
selectively to select recombinant colonies, or nonselectively on LB plates (plus