Download Genetic tools for manipulating Acinetobacter baumannii genome: an

Journal of Medical Microbiology (2015), 64, 657– 669
Review
DOI 10.1099/jmm.0.000081
Genetic tools for manipulating Acinetobacter
baumannii genome: an overview
Indranil Biswas
Correspondence
Indranil Biswas
Department of Microbiology, Molecular Genetics and Immunology, University of Kansas
Medical Center, 3901 Rainbow Boulevard, Kansas City, KS 66160, USA
[email protected]
Acinetobacter baumannii is an emerging nosocomial pathogen involved in a variety of infections
ranging from minor soft-tissue infections to more severe infections such as ventilator-associated
pneumonia and bacteraemia. A. baumannii has become resistant to most of the commonly used
antibiotics and multidrug-resistant isolates are becoming a severe problem in the healthcare
setting. In the past few years, whole-genome sequences of .200 A. baumannii isolates have
been generated. Several methods and molecular tools have been used for genetic manipulation
of various Acinetobacter spp. Here, we review recent developments of various genetic tools
used for modification of the A. baumannii genome, including various ways to inactivate gene
function, chromosomal integration and transposon mutagenesis
Introduction
The genus Acinetobacter belongs to the family Moraxellaceae
within the order Gammaproteobacteria and is composed of
a taxonomically highly heterogeneous group of bacteria.
Bacteria belonging to Acinetobacter are ubiquitous in
the environment and are predominantly present in the
soil. At present, the genus comprises *35 validly named
species as listed at B@cterionet (http://www.bacterionet.
org/). This list is growing at an increasingly high rate;
within the last 2 years alone, six species were validated.
The genus also consists of a few unnamed genomic species
defined by nucleic acid hybridization and numerous isolates with uncertain taxonomic status (Espinal et al., 2012;
Visca et al., 2011). Acinetobacter baumannii is clinically the
most relevant species that has been frequently associated
with nosocomial infections and outbreaks. A. baumannii
can cause a variety of infections ranging from superficial
skin and soft-tissue infections to more severe diseases such
as bacteraemia and pneumonia (McConnell et al., 2013;
Peleg et al., 2008a; Roca et al., 2012). Hospital-acquired
pneumonia is the most common clinical infection caused
by A. baumannii where the mortality rate is high. Community-acquired pneumonia caused by A. baumannii is much
less frequent than nosocomial infections, but the mortality
rate is similar to hospital-acquired pneumonia. A. baumannii
is also responsible for bloodstream infections where the mortality rates are also high. Some other infections caused by A.
baumannii are burn and wound infections, urinary tract
infections, meningitis, osteomyelitis, and endocarditis
associated with prosthetic valves (Howard et al., 2012;
McConnell et al., 2013; Roca et al., 2012). The primary
Abbreviations: Flp, flippase; FRT, flippase recognition target; MDR,
multidrug-resistant; TraSH, transposon site hybridization.
000081 G 2015 The Authors
reason that A. baumannii is successful as a human pathogen
is due to its multidrug resistance, which is generally assimilated by acquisition of various genetic elements such as plasmids, insertion sequences and resistance islands (Dijkshoorn
et al., 2007; Gordon & Wareham, 2010; Imperi et al., 2011).
The other features that render this pathogen successful are
its ability to resist desiccation, which allows the pathogen
to persist on abiotic surfaces present in healthcare settings,
and to colonize asymptomatically within the human host,
enabling its rapid spread (Fournier et al., 2006; Jawad et al.,
1998; Marchaim et al., 2007; Wendt et al., 1997).
Biofilm formation on abiotic surfaces has been suggested to
play a role in A. baumannii persistence and is a debatable
issue (Antunes et al., 2011; Kaliterna & Goic-Barisic, 2013;
Longo et al., 2014; McQueary & Actis, 2011; RodriguezBano et al., 2008). A. baumannii expresses various proteins,
extracellular polysaccharide and type IV pili, all of which are
necessary for biofilm formation (Gayoso et al., 2014; Goh
et al., 2013; Shin et al., 2009). Although A. baumannii has
been described as non-motile, this organism displays
twitching motility and swarming-like motility that are also
mediated by type IV pili (Clemmer et al., 2011; Eijkelkamp
et al., 2011b; Harding et al., 2013; Skiebe et al., 2012). Furthermore, A. baumannii and other Acinetobacter spp. are
naturally competent, which enables them to take up genetic
elements encoding antibiotic resistance from the environment (Harding et al., 2013; Ramirez et al., 2010; Wilharm
et al., 2013). Type IV pili also play a significant role in
DNA uptake during competence. In addition to type IV
pili, a few other virulence factors have been studied to
some extent, but the role of the vast majority of the virulence factors in pathogenesis remains poorly understood
(Bahl et al., 2014; Eijkelkamp et al., 2014; Elhosseiny et al.,
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I. Biswas
2015; Harding et al., 2015; Marti et al., 2011; McConnell
et al., 2013; Roca et al., 2012; Rumbo et al., 2014).
There is no doubt that A. baumannii is an important emerging pathogen. The importance is further strengthened by
the fact that w200 different isolates have already been
sequenced including several multidrug-resistant (MDR)
strains (Adams et al., 2008; Chen et al., 2011; Gao et al.,
2011; Iacono et al., 2008). The availability of a large
number of genome sequences of A. baumannii offers a
great advantage in understanding the role of various genetic
factors involved in the fitness of the organism and in virulence. However, genetic analysis of A. baumannii has thus
far been limited due to a lack of advanced genetic tools.
In general, the function of a gene is assessed by either inactivation or selective modification. In the case of A.
baumannii, the classical strategy for obtaining gene deletion
variants is based on either single-crossover insertion duplication mutagenesis (Aranda et al., 2010; Clemmer et al.,
2011; Heritier et al., 2005) or allelic exchange by doublecrossover recombination (Amin et al., 2013; Aranda
et al., 2010; Cerqueira et al., 2014; Harding et al., 2013).
In addition to these approaches, various transposon mutagenesis strategies are employed for obtaining gene deletion
variants (Clemmer et al., 2011; Dorsey et al., 2002; Jacobs
et al., 2010, 2014c; Smith et al., 2007; Umland et al.,
2012; Wang et al., 2014). Recently, Jacobs et al. (2014b)
published a short laboratory manual on the practical
aspects of genetic manipulation of A. baumannii. Whilst
their publication is very useful, it does not cover all of
the aspects of gene manipulation. In this review, we briefly
elucidate most of the methods currently available for the
manipulation of the A. baumannii genome. We also examine various transposon mutagenesis strategies for the
manipulation of the A. baumannii genome.
Methods for gene transfer in A. baumannii
Although naturally competent, the most commonly used
method for gene transfer is through electroporation.
Electroporation relies on the use of electric pulses to
transiently produce pores in the bacterial membrane
(Miller et al., 1988). These pores are large enough to allow
the passage of DNA, protein and DNA – protein complexes
from the outside milieu into the cells. The efficiency of
electroporation depends on the capacity of the membrane
to reassemble, which allows the cell to survive the electric
shock (Drury, 1996; Miller, 1994). The two main parameters that dictate the success of electroporation are the
strength and the duration of the electric pulse. For prokaryotic electroporation, the buffer containing the cells and
DNA should be free of electrolytes and salts so that the
solution can provide high resistance (Drury, 1996; Miller,
1994). Lowering the volume also increases the resistance.
Electroporation is highly efficient in A. baumannii and one
can obtain *106 transformants per microgram of replicative circular plasmid DNA. However, the efficiency
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depends on the size of the plasmids; smaller plasmids tend
to yield higher efficiency than larger plasmids (Wirth et al.,
1989). Linear DNA can also be used for efficient gene
transfer by electroporation in A. baumannii; although the
efficiency is somewhat low (v102 transformants mg21).
Linear DNA transformation is generally used for direct
gene replacement events (see below). As in Escherichia coli,
A. baumannii also encodes the RecBCD exonuclease that
generally degrades the unprotected linear DNA (de Vries &
Wackernagel, 2002; Kickstein et al., 2007). It appears that
electroporation transiently inactivates the exonucleolytic
function of RecBCD in E. coli, thus allowing gene replacement to occur (El Karoui et al., 1999). A similar
phenomenon might occur during electroporation in A.
baumannii. Electroporation is also the method of choice
for transposon mutagenesis where the DNA –transposase
complex is directly introduced in the cell by electroporation (see below).
A. baumannii also develops a high degree of competence for
natural transformation. Unlike other naturally competent
organisms, A. baumannii develops competence soon after
entry into the exponential growth phase and remains competent for a few hours after entry into the stationary phase,
after which competence gradually decreases (Palmen et al.,
1992). A recent study indicated that *36 % of clinical isolates are naturally competent (Wilharm et al., 2013).
Through this method, A. baumannii could take up any
plasmid DNA, linear DNA or genomic DNA from other
related bacteria. Plasmid DNA can yield i103 transformants
mg21. Transformation efficacy varies depending on the
isolate, and, in the case of linear DNA, on the locus of
homologous recombination. The transformation efficiency
varies from 1023 to 1028 transformants mg21 for linear DNA
substrates. It appears that some unknown environmental
signals often found in soil may boost natural transformation
in certain Acinetobacter spp. (Nielsen & van Elsas, 2001;
Nielsen et al., 1997). Whether A. baumannii needs additional
environmental cues for the development of natural competence has not been examined critically. Nevertheless,
natural transformation provides a fast and handy method to
transfer DNA into the organism.
Conjugation via triparental (or four-parental) mating is an
efficient method for transferring a non-conjugative but
mobilizable plasmid from a donor bacterium into A.
baumannii. In this method, two E. coli strains are used for
mobilizing a plasmid into a recipient A. baumannii (Fig. 1).
The first strain acts as a helper strain that carries a selftransmissible (conjugative) plasmid (helper plasmid)
encoding all the proteins necessary for transfer of itself and
other mobilizable plasmids into donor or recipient cells.
The donor strain carries a mobilizable plasmid (donor
plasmid) that encodes the origin of transfer (oriT). The
function of the helper plasmid is to provide single-strandspecific nicking activity at the oriT and target the nicked
DNA to the mating channel by relaxosome function,
allowing the donor plasmid to move from the donor cells
to A. baumannii recipient cells. The recognition of an oriT
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Genetic tools for A. baumannii
tra+, mob+
Helper
(E. coli )
tra+, mob–
Recipient
(A. baumannii )
Donor
(E. coli )
Transconjugant
(A. baumannii )
Fig. 1. Schematic representation of a typical triparental conjugation. A self-mobilizable plasmid (helper plasmid, e.g.
pRK2013), encoding both the transfer (‘tra’) and mobilization (‘mob’), is transferred into a donor strain harbouring the
recombinant plasmid. When the donor cell carries both the plasmids, the recombinant plasmid can be transferred to the
Acinetobacter recipient. If the recombinant plasmid contains a functional origin of replication, it can be established as the episome in the recipient. If the recombinant plasmid lacks a functional replication origin, but contains a homologous fragment, it
will be integrated into the Acinetobacter genome. When a transposon is present on the recombinant plasmid, it can be used
for transposition.
sequence of a mobilizable plasmid by the helper-plasmidencoded proteins is very specific (Hamilton et al., 2000;
Lessl et al., 1992). Helper plasmids belonging to the same
incompatibility group will recognize the cognate oriT of the
donor plasmid. Generally, the plasmids used for triparental
mating in A. baumannii belong to the IncP group; however,
other incompatibility group plasmids can be used (Jacobs
et al., 2014b). Some IncP group plasmids include derivatives of RK2, RP4 and R6. The most commonly used helper
plasmid is pRK2013, which carries the transfer and mobilization activities from RK2 on a ColE1 replicon (Ditta
et al., 1980, 1985; Figurski & Helinski, 1979; Singer et al.,
1986). Therefore, pRK2013 itself cannot replicate in A.
baumannii, but can mobilize a plasmid that carries oriT
from RK2 (Carruthers et al., 2013; Gohl et al., 2006;
Harding et al., 2013). The frequency of mobilization by
triparental mating can reach as high as 1021 per recipient
(Singer et al., 1986). Although not used frequently in A.
baumannii, triparental mating can also be used as a binary
cloning system. In this case, a cloning plasmid called
pRK290, also a derivative of the RK2 plasmid carrying the
RK2 replicon and oriT, but not self-transmissible, is used as
the cloning vector (Ditta et al., 1980). Genes of interest can
be cloned into this plasmid and the resultant plasmid can
be transferred into A. baumannii with the aid of a helper
plasmid such as pRK2013.
Methods for gene inactivation and gene
deletion in A. baumannii
The easiest and fastest method for inactivating a gene of
interest is by insertional inactivation using an integration vector. Integration vectors used for this type of gene
disruption must be conditional for replication to allow
selection for integration into the chromosome. This is
accomplished by using a plasmid that is able to replicate in
a permissive host or by using a conditional replication such
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as a temperature-sensitive replicon. The vector must also
carry a selectable marker, usually an antibiotic resistance
gene. Sometimes integration vectors also contain oriT so
that the construct can be transferred to other bacteria by
conjugation. This is especially helpful when the other
transfer methods, such as natural transformation or electroporation, are not very efficient. This is not an issue for
A. baumannii as all the modes of gene transfer work very
well. However, historically some of the vectors used for
integration contain the oriT locus and are often large in size
(Jacobs et al., 2014b). Integration at a particular locus on
the chromosome is achieved by including a homologous
sequence on the vector. If there is a homologous sequence
in the plasmid, then the plasmid will integrate into the
chromosome by the Campbell-type mechanism (Fig. 2).
This is a single-crossover event that was originally
proposed by Alan Campbell for the integration of bacteriophage into the E. coli genome (Campbell, 2007). For
integration to work successfully, the key recombination
protein RecA is required. The length of the homologous
sequence is also an important factor. A fragment length
of *100 bp is needed for homologous recombination
to occur and the frequency of integration is directly
proportional to the length of homology. To our benefit,
ColE1-derived E. coli plasmids do not replicate in
A. baumannii; therefore, they can serve as integration
vectors. In addition, some other E. coli plasmids, such as
those derived from R6K, also do not replicate in
A. baumannii. Some commonly and recently used integration plasmids for manipulation of the A. baumannii
genome are listed in Jacobs et al. (2014b).
Integration vectors can be used for other purposes. For
example, if there are two homologous sequences and the
sequences are relatively close on the chromosome, a double
crossover will result in integration of a resistance cassette
between the chromosomal targets (Niaudet et al., 1982).
This is often the straightforward method for gene deletion;
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AbR
oriV
(ColE1)
AbR
Fig. 2. Gene inactivation by single-crossover integration. The integrative plasmid (suicide) carries a sequence that is homologous with a region in the bacterial chromosome. After a single-crossover event (mediated by RecA), the entire plasmid
sequence is integrated in the target region of the chromosome. If the homologous sequence is from an internal region within
an orf, the orf will be disrupted upon plasmid integration.
however, the major drawback with this method is the
possibility of a single-crossover event. If a single crossover
happens, it will simply integrate the entire plasmid without
creating a true deletion.
Integration plasmids can also be used for unmarked gene
deletion, i.e. deletion of a gene without inserting any resistance cassette. A schematic diagram for the overall
method is shown in Fig. 3. In this method, a homologous
fragment carrying either the mutated or the interrupted
target gene is cloned in an integration plasmid and the
integrants are selected for the single-crossover event with
the help of the antibiotic resistance cassette present in the
plasmid backbone. Then the integrated plasmid is allowed
to excise out by a second recombination event. If the
second recombination event occurs through the homology
that was not used for the original integration, it will result
in excision of the plasmid such that the native locus will
be replaced with the mutated (or the interrupted) gene
(Schweizer, 2008). The frequency of excision of the plasmid
depends on the length of homology and requires the activity of the RecBCD enzyme (Zaman & Boles, 1996). For
Gram-positive bacteria, it has been shown that rolling
circle plasmids with a thermosensitive replicon (such as
pGhost4, a pWV01 derivative; Biswas et al., 1993) are much
more efficient for the second recombination event due to
the production of single-stranded intermediates, which are
highly recombinogenic (Biswas et al., 1993). No such
conditional rolling circle plasmids have been tested for
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A. baumannii, although the pWV01 and its derivatives can
replicate in A. baumannii (Bryksin & Matsumura, 2010;
Murin et al., 2012) (unpublished). However, one can easily
screen for plasmid excision by using a counter-selection
marker to screen for the gene replacement event. Several
counter-selection markers are available for use in
A. baumannii. The most widely used is the sacB gene from
B. subtilis. The sacB gene product is toxic for Gram-negative bacteria when grown in the presence of 5 –10 %
sucrose. Therefore the transformants, which will have lost
the plasmid by excision (a second recombination event)
but carry the antibiotic resistance cassette, will survive on
plates containing sucrose and the antibiotics. Integration
plasmid pMo130-TelR (Amin et al., 2013; Hamad et al.,
2009) is one such vector that contains sacB for counterselection. This plasmid also carries a reporter gene xylE
that converts pyrocathechol to 2-hydroxymuconic semialdehyde, a yellow-coloured substance (Fig. 4). Some other
markers for counter-selection are pyrE that converts
5-fluoroorotic acid to 5-fluoro-UMP (Redder & Linder,
2012), gdmP (protease) that converts pregallidermin to
gallidermin (Prax et al., 2013) and tdk that confers
sensitivity to the base analogue azidothymidine (Metzgar
et al., 2004). If the disrupted gene construct contains a
selectable marker, then a direct selection for the marker
is possible (Fig. 5). One can then screen the transformants
for the loss of plasmid (suicide)-encoded marker to identify the double-crossover events (Fig. 5). Other vectors
with improved cloning strategies such as Gateway-based
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Genetic tools for A. baumannii
sacB
AbR
oriV
(ColE1)
A
B
A
B
Integration
A
B
sacB
AbR
A
B
Excision
A
A
B
WT
B
Gene replacement
Fig. 3. Gene replacement by a two-step strategy (integration and excision) using an integrative plasmid. The suicide plasmid
shown here is a generic construct that contains a narrow-host-range origin of replication (non-functional in the target host,
oriV), an antibiotic resistance gene (AbR), a counter-selection marker (sacB from Bacillus subtilis conferring sucrose sensitivity) and the target gene carrying appropriate mutations, such as in-frame insertion or deletion, or single-base mutation
(hatched box). Selection for the AbR phenotype produces integrants created by single-crossover recombination through
either region A or B. Selection for the counter-selection marker in the absence of antibiotic stimulates a second recombination event between the duplication created by the first recombination event, promoting plasmid excision. Excision through
region A restores the WT gene structure, whereas excision through region B creates gene replacement. However, if the initial
integration occurs through region B and excision through A, it will also generate gene replacement.
cloning or direct blunt-end cloning are also available both
for cloning and gene inactivation (Eijkelkamp et al., 2011a;
Oh et al., 2015).
Integration plasmids are also used for reporter gene
fusion. In this case, a reporter gene is placed under the
promoter of a target gene. After integration the reporter
gene will be expressed from the promoter of interest
in the chromosome. Several reporter genes, such as lacZ
(b-galactosidase), gusA (b-glucouronidase), xylE (catechol
2,3-dioxygenase), lux (luciferase) or gfp (green fluorescent
protein), can be used in A. baumannii (Davison, 2002).
Another utility for the integration plasmids is to tag a
protein of interest at the C terminus. In this case the plasmid contains a homologous fragment carrying a tag (such
as His6, FLAG or Myc), which is inserted in-frame at the
end of the gene by removing the stop codon. After the
single-crossover integration, the target gene will be tagged at
the C terminus (Fig. 6). This strategy is particularly useful
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for tagging genes that are essential or whose expression level
cannot be altered. Moreover, with the help of commercially
available antibodies against the tags, it is possible to estimate the expression level of the tagged proteins in the cell.
One can also purify the tagged protein from the cell by
affinity chromatography (such as nickel-nitrilotriacetic acid
chromatography for His-tagged protein).
Recombineering systems for A. baumannii
Recombineering is an in vivo system originally developed in
E. coli for gene replacement purposes. Recombineering is
based on homologous recombination that allows precise
insertion, deletion or alteration of any sequence in the
genome (Sharan et al., 2009). Linear dsDNAs (such as
PCR fragments) or single-stranded oligonucleotides are
introduced in the cell to provide the homologous substrates
for genetic modification. Phage-encoded homologous
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MCS aphA
sacB
oriT
pMo130-TelR
xylE
kilA
telA
oriV
(ColE1)
telB
Fig. 4. Schematic diagram of an integration vector suitable for
manipulation of MDR A. baumannii. The vector encodes a tellurite-resistance gene cassette (kilA, telA and telB), a counterselection marker (sacB) and a reporter gene (xylE) that facilitates
screening. The vector also encodes a kanamycin-resistance gene
(aphA) suitable for selection in E. coli and a multicloning site
(MCS) for cloning of the target gene. Adapted from Amin et al.
(2013).
recombination proteins such as the lambda Red system or
the RecET system from Rac prophage catalyse the process.
The lambda Red system produces the phage-derived Gam,
Bet and Exo proteins (Murray, 2006). The function of the
Gam protein is to protect linear DNA from exonuclease
degradation by the RecBCD enzyme. The Exo protein
AbR
oriV
(ColE1)
Although the lambda Red system works very efficiently in
many Gram-negative organisms, including Agrobacterium
(Hu et al., 2014), it has been reported that the E. coli
lambda Red system does not work in A. baumannii and
expression of the Red system might be toxic to A. baumannii (Tucker et al., 2014). However, a RecET homologue has
been identified in one of the sequenced A. baumannii
strains (IS-123). This system, which was named RecETAb,
when overexpressed from a multicopy plasmid promotes
recombination (Fig. 7) (Tucker et al., 2014). The recombination efficiency was highest when the homology was 125 bp
and, depending on the target gene, the range varied
from 1029 to 1028 per viable cell (Tucker et al., 2014).
However if the homology is 75 bp, RecET-mediated recombineering is not productive at all in A. baumannii. Although the
recombineering system works in A. baumannii, due to its low
efficiency, the current system cannot be applied for
unmarked gene deletion or for the introduction of
mutation in the genome.
Methods for transposon
mutagenesis in A. baumannii
AbR*
AbR*
Fig. 5. Gene replacement through a direct double-crossover
event. If the integration plasmid contains the target gene disrupted by a selectable antibiotic resistance marker (AbR*), selecting directly for the marker and screening for the absence of
plasmid-encoded antibiotic resistance marker (AbR) leads to
gene replacement. Depending on the bacteria, direct gene replacement is relatively efficient and straightforward.
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displays a progressive 59 ! 39 exonuclease activity that creates an ssDNA overhang to which the Bet protein binds and
promotes annealing of complementary DNA strands. The
synergic activity of Bet and Exo leads to insertion of linear
DNA at the desired target site producing the recombinant.
The RecET system is analogous to the Red system, where
RecE is a 59 ! 39 dsDNA exonuclease that uses a processive
mode of digestion and therefore is very similar to lambda
Exo (Shiraishi et al., 2002). However, RecT is an ssDNA
annealing protein that is similar in both structure and function to the lambda Bet protein.
Transposon mutagenesis is a very powerful laboratory tool
that facilitates genome-scale studies of gene function. It
offers a rapid and economical means of generating large
numbers of independent insertions in the target genome
with little experimental manipulation. Transposons that
are developed for other Gram-negative bacteria such as
E. coli, Pseudomonas aeruginosa and Vibrio cholerae work
very well in A. baumannii (Cameron et al., 2008; Goryshin
et al., 2000; Liberati et al., 2006). The two most commonly
used transposon mutagenesis systems are derived from Tn5
and himar1 (a mariner transposon). Other transposons,
such as Tn3171 or Tn10, have also been used in various
Acinetobacter spp. with some degree of success (Ely, 1985;
Leahy et al., 1993). The EZ : : TN Transposome system,
particularly EZ : : TN vR6Kcori/KAN-2w that is based
on Tn5, which has a transposition frequency 1000-fold
higher than the WT Tn5 (Goryshin et al., 2000), is widely
used in A. baumannii and other Acinetobacter spp.
(Clemmer et al., 2011; Jacobs et al., 2010; Smith et al., 2007;
Tomaras et al., 2003). The EZ : : TN Transposome is a
nucleoprotein complex in which the transposase is bound
to the inverted repeat sequences present at the end. The
main advantage of the EZ : : TN system is that the complex
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Genetic tools for A. baumannii
(a)
AbR
Reporter
AbR
(b)
AbR
Tag
AbR
Fig. 6. Strategies for reporter gene fusion and gene tagging using integration vectors. (a) In gene fusion, a fragment carrying
a promoter region of the target gene is either transcriptionally or translationally fused with a reporter gene (hatched arrow).
Upon integration, the reporter gene will be expressed from the native promoter. (b) During gene tagging, the 39 end of the
gene is translationally fused with a suitable tag (solid circle) and upon integration the tagged protein will be expressed instead
of the native protein.
is directly introduced into A. baumannii by electroporation
and there is no need for delivery by suicide vector or
conjugation. The frequency achieved through this system
can be very high. In one report, as many as 2 % of the
colonies recovered without antibiotic selection after electroporation were mutants (Dorsey et al., 2002). Several Tn5
delivery plasmids are available that can be used instead of
the EZ : : TN system (Fig. 8). To obtain a 95 % saturation
mutagenesis, one would require between 12 000 and 42 000
mutants depending on the estimation method used
(Phogat et al., 2001). With the high frequency of Tn5mediated transposition, saturation mutagenesis in A. baumannii is easily achievable. Mapping of the insertion site is
also very straightforward with the EZ : : TN system. In the
case of the EZ : : TN v R6Kcori/KAN-2w system, a direct
cloning approach termed ‘rescue’ cloning can be used. This
transposon contains an E. coli origin of replicon (R6Kcori)
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that requires the p-protein. When genomic DNA from the
mutant is appropriately restricted and self-ligated and
transformed into an E. coli strain expressing the p-protein
(encoded by pir), ‘rescued’ clones containing plasmids are
produced. These plasmids are used for sequencing using
transposon specific primers. Two other methods have been
used to identify the insertion site: direct PCR using genomic DNA and random amplification of terminal ends PCR.
Both are single-primer PCRs and, as they do not require
cloning, they are therefore relatively fast.
A himar1-based transposon mutagenesis system, MAR2xT7,
which was developed originally for P. aeruginosa, has also
been adapted for use in A. baumannii (Liberati et al., 2006;
Peleg et al., 2008b). The transposon is delivered to this
organism through conjugation using a helper plasmid such
as pRK2013. The frequency of transposon mutagenesis is
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I. Biswas
(a)
ME-I
MCS ME-O
aphA
AbR*
oriV
(R6Kγ)
pBAM1
PCR
oriT
tnpA
AbR*
(b)
RecET(AB)
ME-I
aphA
FRT
bla
MCS
T0λ P
trc
FRT
ME-O
lacIq
oriV
(R6Kγ)
pBEL
oriT
Ab
R*
tnpA
Fig. 7. Strategy for recombineering using a selectable marker.
The targeting construct is generated by PCR amplification to
introduce the region of homology (double lines) and an antibiotic
resistance gene. The construct is electroporated into the A. baumannii cells that express phage recombination RecET and is
selected for antibiotic (AB)-resistant transformants. The current
method for A. baumannii requires that the homologous region
should be at least 100 bp long.
also very high with MAR2xT7. One of the advantages of using
MAR2xT7 is to allow genetic footprinting of the mutants
with transposon-site hybridization (TraSH) analysis – a
method for tracking mutants in the library under various
conditions. Two T7 promoters, placed at both ends of
MAR2xT7 (Fig. 9), can be used for identification of the
insertion site by linker-ligation-mediated PCR amplification
using the genomic DNA. The PCR products obtained can be
used for transcription using T7 polymerase and the RNA
products can be used for making probes for hybridization to
microarrays to identify the amplified genomic fragments,
and thus the individual mutants present in the library. The
MAR2xT7 transposon has been used in A. baumannii to
identify genes necessary for interaction with Candida albicans. A derivative of the himar1 transposon was recently used
in conjunction with insertion sequencing to identify
A. baumannii genes required for persistence in the lung
(Wang et al., 2014).
In addition to their use in genome-wide mutagenesis, transposons can be used as molecular tools for facilitating complementation, expression analysis and molecular tagging.
A Tn7-based system has been developed for single-copy gene
integration in many Gram-negative bacteria, including A.
baumannii (Choi et al., 2005; Kumar et al., 2010). Unlike other
transposons, Tn7 integrates in both a site and orientationspecific manner at neutral attachment sites (attTn7, 24 bp)
generally located downstream of glmS (glucosamine
664
bla
Fig. 8. Schematic diagram of Tn5 delivery plasmids. (a) A multipurpose mini-Tn5 plasmid pBAM1. This plasmid can be delivered
to the recipient by conjugation or by electroporation. This mini-Tn5
can generate a transposition frequency as high as 1023.Adapted
from Martı́nez-Garcı́a et al. (2011). (b) Plasmid pBAM1 can be
modified to generate a cassette for regulated expression of heterologous genes in Gram-negative bacteria. The expression module
in plasmid pBEL contains the lacI q element and the Ptrc promoter.
The gene of interest can be cloned in the multicloning site (MCS)
just downstream of the promoter and the expression can be regulated by IPTG. Adapted from Nikel & de Lorenzo (2013).
6-phosphate synthetase). Most bacterial genomes contain a
single glmS gene and therefore a single insertion site for Tn7.
Interestingly, although A. baumannii encodes two glmS genes,
it encodes only one attTn7 and thus allows for a single integration (Kumar et al., 2010). The commonly used Tn7 vector
(mini-Tn7) does not encode transposase and therefore the
activity must be supplied by another helper plasmid. However, mini-Tn7 contains two flippase recognition target (FRT)
sites for flippase (Flp)-mediated site-specific recombination
that allows for excision of the antibiotic resistance marker
(aacC1) to create the unmarked insertion. A schematic diagram for the insertion of genes through Tn7-based systems in
A. baumannii is shown in Fig. 10.
Transposons are also useful for expression analysis. Several
mini-Tn5-based systems have been developed for Gramnegative bacteria. These constructs encode a variety of
promoterless reporter genes ( phoA, lacZ, gfp and lux) to
generate transcriptional or translational fusions to the
inactivated genes. In addition, transposons may also contain
outward-facing inducible promoters to avoid polar mutations and thus can be used for the identification of
essential genes under non-inducing conditions (Judson &
Mekalanos, 2000). Furthermore, like mini-Tn7, mini-Tn5
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Journal of Medical Microbiology 64
Genetic tools for A. baumannii
bla (ApR)
(a)
Mariner Tnp
pMAR2xT7
T7
T7
IR (29 bp)
aacC1
IR (29 bp)
oriT
(b)
bla (ApR)
oriV (R6Kγ)
tra
pSC189
Mariner Tnp
oriV (R6Kγ)
3×FLAG
aphA
IR FRT
FRT IR
Fig. 9. Schematic diagram of two useful mariner-based transposons. (a) Plasmid pMAR2xT7 encodes the himar1 transposase
that is outside the MAR2xT7 transposon cassette. MAR2xT7 contains the aacC1 gene encoding gentamicin resistance and short
inverted repeats (IR, 29 bp) and outward-facing T7 promoters at
both ends. The T7 promoters allow transcription of the nearby
sequences at the transposition sites. The resultant transcripts can
be used for hybridization to a microarray – a process known as
TraSH analysis. Adapted from Liberati et al. (2006). (b) A plasmid
that facilitates epitope tagging of the inserted genes. Plasmid
pSC189 encodes a transposon module with FLAG epitopes, a
conditional origin of replication and FRT sites that flank the kanamycin resistance gene (aphA). The plasmid also encodes the
Himar1 transposase that is outside the transposon module and
the plasmid can be delivered via conjugation. This transposon
module can generate tagged truncated proteins. However, the
aphA gene can be excised out using Flp/FRT-mediated recombination to generate a tagged full-length functional protein, if the
final construct is in-frame. Adapted from Chiang & Rubin (2002).
derivatives often contain sequences for site-specific
recombination within the inverted repeats that allow the use
of epitope or His6 tags for localization or native protein
purification (Jacobs et al., 2003; Ross-Macdonald et al.,
1999). Although mini-Tn5 is an attractive and powerful
system, it has not been used in Acinetobacter. Similarly,
various mariner-based transposons have also been developed for tagging Gram-negative bacteria (Cameron et al.,
2008; Chiang & Rubin, 2002). These systems have the potential to be used as tools for manipulation of Acinetobacter
spp., including A. baumannii.
Conclusion and future perspective
As a result of its clinical significance, the vast majority of
studies on A. baumannii are focused mainly on the
http://jmm.sgmjournals.org
emergence and nature of multidrug resistance. As research
on the pathogenesis of A. baumannii advances and more
complete genome sequences become available, the need
for sophisticated and powerful genetic tools for functional
genomic studies becomes necessary. Until recently, genetic
tool development was primarily centred towards a few
important Gram-negative pathogens such as Agrobacterium,
Pseudomonas and Vibrio. Some of the tools developed
for these organisms can be applied directly to the study
of A. baumannii (Davison, 2002). However, because of
the inherent genetic differences, the tools developed for
other Gram-negative bacteria are often not adaptable for
A. baumannii. Therefore, there is a need for the development of Acinetobacter-specific tools to study the clinical
isolates. The choice of available antibiotic resistance markers is also a limiting factor, as it depends largely on the
susceptibility of the isolate. Only a handful of antibiotic
resistance markers are available to use in A. baumannii
and recycling of these few valuable markers is necessary
when multiple genes are targeted for modification in
the same cell. Site-specific recombination is an efficient
method that allows removal of antibiotic resistance
markers from the genome. The two most commonly used
methods are the Flp/FRT system from yeast (Schweizer,
2003) and the Cre/loxP system from bacteriophage (Marx
& Lidstrom, 2002). Whilst the former system has been
applied to several Acinetobacter spp., including A. baumannii, this system is not very efficient when multiple gene
deletions are desired. When multiple gene deletions are
required, the Cre/loxP system is ideal as the Cre recombinase can utilize two mutated loxP sites to generate an inactive loxP site that is no longer recognized by Cre. Thus, the
system can be used repeatedly to delete multiple genes in
a given strain (Banerjee & Biswas, 2008). Although the
Cre/loxP system has not yet been adapted for A. baumannii,
a thermosensitive derivative of the pWV01 plasmid carrying Cre recombinase (pCrePA; Pomerantsev et al., 2006)
is available and can be used directly in A. baumannii
without the need for further modification.
The MDR phenotype poses another difficulty in manipulating the MDR strains and necessitates the use of nonantibiotic resistance markers. Two such markers, tellurite
and heavy metal resistance, have already been used in
A. baumannii (Amin et al., 2013). Other non-antibiotic
resistance markers, such as triclosan resistance, bailaphos
(or phosphinothricin) resistance and glyphosate resistance,
have been used in various Gram-negative bacteria such as
Burkholderia and Pseudomonas, but these markers have
never been used in A. baumannii (Davison, 2002). Both
integrating plasmids and transposons containing these
non-antibiotic markers are available, and can be adapted
to A. baumannii either directly or with very little
modification.
It is noteworthy to mention that the RecET-mediated
recombineering system is functional in A. baumannii
(Tucker et al., 2014). However, the system requires much
improvement to accommodate shorter flanking fragments,
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665
I. Biswas
(a)
ααcC1
FRT
tnsB
pUC18T-Tn7T-Gm
tnsA
Pm
oriV
(R6Kγ)
oriV
(ColE1)
FRT
tnsC
Tn7R
(c)
MCS
Tn7L
T 0 T1
pTNS2
oriT
oriT
tnsD
bla
bla
(b)
MCS
T0T1
glmS1
ααcC1
glmS2
Tn7R FRT
FRT
Tn7L
Fig. 10. Tn7-mediated gene insertion in Gram-negative bacteria. (a) Schematic diagram of a mobilizable mini-Tn7 cassette
on a pUC18-based suicide plasmid. The plasmid encodes right and left ends of Tn7 (Tn7 R and Tn7 L), FRT sites that flank
the gentamicin resistance gene (aacC1), a multicloning site (MCS) for the cloning of the gene of interest, and the transcriptional terminators T0 and T1 from l phage and rrnB operon of E. coli, respectively. (b) Integration of mini-Tn7 at the attTn7
site on the A. baumannii chromosome. There is only one functional att site adjacent to the glmS2 gene on the A. baumannii
genome. (c) Depiction of the transposon helper plasmid pTNS2 that encodes the TnsABCD transposase subunits. The miniTn7 is delivered to the recipient A. baumannii cells using a four-parental mating involving three E. coli strains each with miniTn7, pTNS2 and helper plasmid for mobilization, and the recipient A. baumannii strain. Adapted from Choi & Schweizer
(2006), Choi et al. (2005) and Kumar et al. (2010).
to allow recombination without any selection and to
permit insertion of single amino acid changes in the
genome. Though the lambda Red system does not function
in A. baumannii, it is worth modifying the system so that it
can be successfully adapted to A. baumannii. Alternatively,
a different RecET system, perhaps from another Acinetobacter sp., would be helpful to develop a more efficient
and powerful recombineering tool.
The transposon mutagenesis systems based on Tn5 or mariner transposons are highly efficient in A. baumannii. Several mutant libraries have been constructed in highly
virulent clinical and laboratory isolates (Harding et al.,
2013; Jacobs et al., 2014a, c; Umland et al., 2012; Wang
et al., 2014). Some of these libraries are not comprehensive,
whilst others are. In the case of a widely used A. baumannii
isolate, ATCC 17987, at least two mutant libraries have
been generated and one of them contains *150 000
unique insertions, thereby surpassing 95 % coverage
(Smith et al., 2007; Wang et al., 2014). This particular
library has been used for insertion sequencing (INSeq, a
deep-sequencing method to comprehensively identify the
insertion sites) to reveal novel virulence factors required
for A. baumannii persistence (Goodman et al., 2009). Similar high-density libraries in other MDR clinical isolates
would be necessary to systematically understand the pathogenesis of this organism.
666
Acknowledgements
This work was made possible by a Fulbright – Nehru Senior Research
Fellowship award to I. B. The author would like to thank Dr V. Balaji
(Christian Medical College, Vellore, India) for helpful discussion.
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