- Research article
- Open Access
Identification of genes essential for pellicle formation in Acinetobacter baumannii
© Giles et al. 2015
- Received: 15 January 2015
- Accepted: 11 May 2015
- Published: 6 June 2015
Acinetobacter baumannii is an opportunistic pathogen, which has the ability to persist in the clinical environment, causing acute and chronic infections. A possible mechanism contributing to survival of A. baumannii is its ability to form a biofilm-like structure at the air/liquid interface, known as a pellicle. This study aimed to identify and characterise the molecular mechanisms required for pellicle formation in A. baumannii and to assess a broad range of clinical A. baumannii strains for their ability to form these multicellular structures.
Random transposon mutagenesis was undertaken on a previously identified hyper-motile variant of A. baumannii ATCC 17978 designated 17978hm. In total three genes critical for pellicle formation were identified; cpdA, a phosphodiesterase required for degradation of cyclic adenosine monophosphate (cAMP), and A1S_0112 and A1S_0115 which are involved in the production of a secondary metabolite. While motility of the A1S_0112::Tn and A1S_0115::Tn mutant strains was abolished, the cpdA::Tn mutant strain displayed a minor alteration in its motility pattern. Determination of cAMP levels in the cpdA::Tn strain revealed a ~24-fold increase in cellular cAMP, confirming the role CpdA plays in catabolising this secondary messenger molecule. Interestingly, transcriptional analysis of the cpdA::Tn strain showed significant down-regulation of the operon harboring the A1S_0112 and A1S_0115 genes, revealing a link between these three genes and pellicle formation. Examination of our collection of 54 clinical A. baumannii strains revealed that eight formed a measurable pellicle; all of these strains were motile.
This study shows that pellicle formation is a rare trait in A. baumannii and that a limited number of genes are essential for the expression of this phenotype. Additionally, an association between pellicle formation and motility was identified. The level of the signalling molecule cAMP was found to be controlled, in part, by the cpdA gene product, in addition to playing a critical role in pellicle formation, cellular hydrophobicity and motility. Furthermore, cAMP was identified as a novel regulator of the operon A1S_0112-0118.
- Cyclic AMP
Acinetobacter baumannii is an opportunistic Gram-negative human pathogen that causes severe nosocomial infections primarily in immune-compromised patients; disease states include pneumonia, meningitis and bacteraemia [1–5]. The bacterium can be found in intensive care units and is frequently isolated from medical devices [6–9]. The capacity to survive in these environments is possibly due to the ability of this organism to survive desiccation and resist a broad spectrum of antibiotics and disinfectants. As a result, A. baumannii has entrenched itself in the clinical environment [10–12].
Given the occurrence of A. baumannii in intensive care units and its high level of intrinsic and adapted resistance, there is increased pressure to identify novel drug targets. Therefore, understanding the molecular mechanisms contributing to virulence in A. baumannii is essential. Similar to numerous other organisms, A. baumannii has the ability to grow a multicellular structure known as a biofilm (reviewed by Longo et al. 2014)  of which the pellicle is a specialised form. The pellicle is localised to the interface between air and liquid, and is a structure of connected cells surrounded by a matrix of extracellular polymeric substance [14, 15]. The formation of a pellicle allows aerobic bacteria to have access in a stagnant environment to high levels of oxygen and nutrients . The pellicle can increase bacterial survival under antibiotic pressure as well as contribute to longer than usual persistence in a hostile environment, such as under drying conditions [17–19].
There are numerous molecular mechanisms potentially involved in pellicle formation by A. baumannii. One such mechanism is the csuA/BABCDE pilus chaperone-usher assembly system which is responsible for the production of pili and is essential in the formation of a biofilm on solid surfaces [2, 9, 20]. In a study on Bacillus subtilis, 288 regulatory and potential regulatory genes were associated with pellicle formation . Recent analyses of A. baumannii bacteria in the pellicle state have revealed alterations in the expression of at least 52 membrane proteins (32 up-regulated and 20 down-regulated) [22, 23]. These 52 proteins were found to be involved in; iron uptake systems, lipid and carbohydrate transport, cellular metabolism, starvation, in addition to porins and pili.
As bacteria encounter a wide range of environmental conditions, there is a requirement to respond to these changes, such as changing from a planktonic state to living within a biofilm and/or pellicle; this can involve deployment of a number of signalling molecules [24, 25]. Cyclic adenosine monophosphate (cAMP) is a ubiquitous signalling molecule used in conjunction with a variety of regulators for modulating gene expression in both prokaryotes and eukaryotes [26, 27]. In bacteria, it has been found to alter multiple virulence characteristics. For example, in Vibrio cholerae cAMP has been implicated in the regulation of cholera toxin, the toxin co-regulated pilus and other pathogenic factors . In Serratia marcesens the importance of the cAMP receptor protein (CRP complex) in the formation of biofilm has been established . In this case the type I fimbriae-dependant biofilm is tightly regulated and a deletion in the phosphodiesterase gene was found to increase cAMP within the bacteria cell and consequently decrease biofilm formation. Conversely, when increased copies of the phosphodiesterase gene are introduced into the bacterial cell, the bacteria enter into what is described as a “hyper biofilm” state . Similarly in Pseudomonas aeruginosa, cAMP has been identified as a key component in biofilm formation, where increased levels of cAMP can inhibit adherence capabilities . Furthermore, in this organism cAMP functions as a co-factor of the virulence factor regulator Vfr, where the cAMP-Vfr complex regulates type III secretion systems involved in exporting toxins and other compounds [30–32]. This complex also regulates virulence factors such as type IV pili, the las quorum sensing system and exotoxin A [32, 33]. Although a vfr-like gene is present in the A. baumannii genome, its role in regulation of virulence factors and interaction with cAMP has not been reported.
The proteomic analyses by Marti et al. (2011) and Nait Chabane et al. (2014) suggested that pellicle formation in A. baumannii is not the result of a single gene or operon [22, 23]. Instead, it is likely that pellicle formation is multifactorial, requiring surface-exposed molecular structures and an adequate level of transcriptional regulation. The aim of this study is to provide insight into the genetic elements that contribute to the ability of A. baumannii to migrate to a surface and form a pellicle as well as to assess the prevalence of pellicle formation in a number of clinically-relevant A. baumannii strains.
Pellicle formation is a rare trait in clinical A. baumannii strains
Pellicle formation in glass and polypropylene, cell hydrophobicity, motility status and isolation site
A. baumannii isolate
Pellicle forming A. baumannii strains
0.65 ± 0.16
0.44 ± 0.08
32 ± 0.25
1.03 ± 0.01
0.94 ± 0.02
42 ± 0.19
0.13 ± 0.01
0.17 ± 0.04
98 ± 0.01
0.35 ± 0.08
0.28 ± 0.06
98 ± 0.03
1.01 ± 0.02
0.37 ± 0.08
94 ± 0.10
0.62 ± 0.00
0.79 ± 0.04
100 ± 0.00
0.65 ± 0.11
92 ± 0.07
0.47 ± 0.06
Surface film forming A. baumannii strains
9 ± 0.09
5 ± 0.07
Hydrophobicity across the clinical A. baumannii strains is not directly linked to pellicle formation ability
A1S_0112, A1S_0115 and cpdA are essential for pellicle formation in A. baumannii 17978hm
To further investigate the pellicle phenotype, we used our previously characterised A. baumannii 17978hm strain (harboring an insertion in hns) , which has the ability to produce increased amounts of pellicle material compared to the ATCC 17978 parent strain. This strain was chosen as it allows good discrimination between the pellicle and non-pellicle phenotype allowing ready identification in large scale screening procedures. To identify genes associated with this potential virulence factor a transposon bank of random mutants was constructed in the 17978hm strain using the plasmid pLOF mini::Tn10:gfp:kan . Approximately 4000 individual transposon insertion mutants were examined in microtitre trays for their ability to form a pellicle. Initial screening identified nine transposon insertion mutants that were incapable of pellicle formation. Subsequent evaluation of these strains in polypropylene and glass tubes confirmed the complete loss of the pellicle phenotype (data not shown). The exact insertion site of the individual transposon insertions were identified by cloning the kanamycin resistance gene associated with the transposon (mini::Tn10:gfp:kan) and subsequent sequencing. Insertions were found to be within three genes: A1S_0249 (cpdA), a cAMP phosphodiesterase; A1S_0112, acyl-CoA synthetase/adenosine monophosphate (AMP) acid ligase II; and A1S_0115, which is annotated to be involved in amino acid adenylation. The precise insertion site of the transposon is given as the nucleotide position relative to the A. baumannii ATCC 17978 genome sequence (GenBank CP000521) (Additional file 1). Seven transposon insertions were identified to be within the A1S_0115 gene. A single representative of each of the three distinct insertions, designated transposon mutants 1, 2 and 5, was taken for further investigation (Additional file 1).
A. baumannii 17978hm mutant derivatives constructed in this study
Tn10:gfp:kan insertion in cpdA, kanR
Tn10:gfp:kan insertion in cpdA containing pWH0249, kanR, ampR
Tn10:gfp:kan insertion in cpdA containing shuttle vector pWH1266, kanR, ampR, tetR
Tn10:gfp:kan insertion in A1S_0112, kanR
Tn10:gfp:kan insertion in A1S_0112 containing pWH0112, kanR, ampR
Tn10:gfp:kan insertion in A1S_0112 containing shuttle vector pWH1266, kanR, ampR, tetR
Tn10:gfp:kan insertion in A1S_0115, kanR
Tn10:gfp:kan insertion in A1S_0115 containing pWH0115, kanR, ampR
Tn10:gfp:kan insertion in A1S_0115 containing shuttle vector pWH1266, kanR, ampR, tetR
Fold change in gene expression of transposon mutant strains compared to the 17978hm parent strain
-2.8 ± 0.21
1.5 ± 0.44
-5.6 ± 0.032
-335.46 ± 1.42
-28.44 ± 1.33
-0.49 ± 1.39
-669.37 ± 1.24
-51.63 ± 1.70
-171.25 ± 1.68
-325.53 ± 1.22
-85.43 ± 1.46
-1.14 ± 1.59
-83.29 ± 1.20
-4.1 ± 1.09
-1.16 ± 1.63
-28.97 ± 1.54
-0.99 ± 1.32
-347.29 ± 1.52
-72.50 ± 1.14
Characterisation of pellicle-deficient mutant strains cpdA::Tn, A1S_0112::Tn and A1S_0115::Tn
Bacterial adherence is a critical virulence factor and aids in host and surface colonisation. The adherence of the A. baumannii mutant strains to human lung epithelial cells (A549) was assessed; all three transposon insertion strains had adherence capabilities that were not significantly different to that of the 17978hm parent strain (data not shown). This suggests that the mechanisms used by A. baumannii for pellicle formation are not essential for adherence to A549 human epithelial cells.
The levels of cAMP in the cpdA::Tn mutant strain are significantly increased compared to the 17978hm parent strain
Transcriptional profiling of the cpdA::Tn mutant strain under motility conditions
To assess the association between pellicle formation and motility qRT-PCR was undertaken on the A. baumannii 17978hm parent and the cpdA::Tn transposon mutant strain grown under previously identified motility conditions, i.e. on semi-solid LB agar . Analysis of the operon A1S_0112-0118 (Fig. 3) was undertaken by quantifying the transcription of the A1S_0112 gene, which is the first gene in the operon. In the cpdA::Tn strain, the transcription level of A1S_0112 was 2.8-fold down-regulated compared to the 17978hm parent (Table 3). This indicated that CpdA via cAMP plays a role, either directly or indirectly, in regulation of the A1S_0112-0118 operon. This potential regulatory role of CpdA and cAMP on the A1S_0112-0118 operon could explain the altered motility phenotype seen in the cpdA::Tn mutant strain, possibly due to alteration in the production of a biosurfactant or through another unknown mechanism(s).
The A1S_0112-0118 operon described above has previously been shown to be activated by quorum-sensing signals in the form of homoserine lactone (abaI, A1S_0109) . Furthermore, Saroj and Rather (2013) have shown that down-regulation of abaI leads to a reduction in motility in A. nosocomialis strain M2 [53, 54]. Therefore, to examine whether the insertional inactivation of cpdA in the cpdA::Tn mutant had a regulatory effect on the quorum sensing gene abaI when grown on motility media, we assessed the levels of transcription of abaI using qRT-PCR. The results revealed that, despite the altered motility phenotype of the derivative, abaI is expressed at a level comparable to the parent strain (~1.2-fold increase). This suggests that the A1S_0112-0118 operon is regulated independently by two different factors; quorum-sensing (AbaI) and cAMP. Levels of cAMP act as one of the major signalling molecules within the cell and have been show to regulate numerous genes in other bacterial species such as P. aeruginosa and Escherichia coli, possibly through activation of Vfr . In P. aeruginosa, vfr is auto-regulated and upon inactivation of cpdA, vfr expression increases by 12-fold [33, 45]. We investigated this possibility in our cpdA::Tn mutant strain by examining the transcript of the putative vfr gene (A1S_1182). The qRT-PCR results showed a 1.7-fold decrease in vfr expression in the cpdA::Tn mutant strain compared to the 17978hm parental strain, that, by a student t-test, was determined to be statistically insignificant. Thus, although a 1.7-fold reduction in vfr transcription was observed it is not known whether cAMP and Vfr work in concert in A. baumannii.
Whole transcriptome analysis of the CpdA mutant compared to the 17978hm A. baumannii parent strain
A number of the genes down-regulated in the cpdA::Tn mutant include genes encoding phage-related proteins (A1S_2021-2033), putative acyl carrier proteins, ferric enterobactin receptors, porins, signal peptides, membrane proteins and numerous hypothetical ORFs with no known function (Fig. 8). Variations in the expression of multiple potential transcriptional regulators were also observed; this may be due to the significant alterations that cAMP levels have on the general metabolism of the cell. Genes of particular interest in this study that are down-regulated include the CsuA/BABCDE system (A1S_2213-2218) (9 to 20-fold), the hms operon which consists of three genes designated A1S_2160 to A1S_2162) (10 to12-fold), and a chaperone usher system (A1S_1507-1510) (2-fold). Proteins encoded by the csu cluster and chaperone usher system have been identified in pellicle and biofilm formations [2, 20, 22, 23, 34, 55, 56]. Interestingly, a single polynucleotide polymorphism leading to truncation of the csuB gene potentially inactivating this operon has been identified within ATCC 17978 [2, 22, 56]. Within the hms operon two proteins (A1S_2160 and A1S_2161) share an average of 44 % identity with the PgaBC proteins (A1S_0938-0939), the third protein within Hms (A1S_2162) has been designated as a biofilm synthesis protein and shares no homology to the Pga proteins in A. baumannii ATCC 17978. The Pga system has previously been identified as being important in biofilm formation . Interestingly, a recent paper by Nait Chabane et al. (2014)  found that CsuA/BABCDE, PgaABC proteins and the chaperone usher system described above, are over represented within A. baumannii strains displaying a pellicle phenotype. This suggests that the Hms and chaperone usher pilus proteins could be key components in the formation of the pellicle matrix.
Numerous genes which are up-regulated encode proteins involved in oxidative phosphorylation (Fig. 8). In particular, these include those encoding ubiquinol oxidase subunit I and II (A1S_1434 and A1S_1433) which are 5.7- fold and 4.9-fold up-regulated, respectively. Other genes involved in oxidative phosphorylation include the NADH dehydrogenase complex (A1S_0762-0764) and the ATP synthetase complex (A1S_0148-0156). One of the most highly up-regulated genes (A1S_1387) is an oxidoreductase (18-fold); this class of enzymes is often involved in the FAD to FADH2 conversion, where FADH2 plays a major role in oxidative phosphorylation. Therefore, many of the up-regulated genes are involved in the energy, and by extension, the metabolic status of the cell which is not surprising given the central role cAMP plays in cell metabolism.
Other genes of interest which were significantly up-regulated are those involved in 50S and 30S ribosome assembly. The potential role of these up-regulated ribosomal proteins maybe in the assembly of 100S ribosomes, as these are required for cells to survive long term nutrient stress in situations where glucose is low and cAMP is high . Other up-regulated genes including those involved in fatty acid oxidation, trehalose synthase, membrane proteins, dehydrogenase proteins and a number of proteins which are annotated as hypothetical ORFs.
A small operon consisting of 4 genes (A1S_2088-2091) was identified as having 4-fold increase in expression. Proteins encoded by this region have recently been found in extracted pellicle material . Since, cpdA::Tn is not capable of pellicle production it would seem unlikely that the A1S_2088-2091 operon is involved in this phenotype, however, cpdA::Tn does show an increase in biofilm formation compared to the parent strain and as such this operon may contribute to liquid/surface biofilm production.
The previously described motility operon A1S_0112-0118  is up-regulated approximately 3.8-fold in the cpdA::Tn mutant transcriptome, which is in contrast to RT-PCR data (Table 3). These RT-PCR studies were conducted on motility agar to look at the influence of motility on the change in transcription of this operon. On motility agar both cAMP and quorum sensing may play a role in the control of this operon, however in liquid culture grown to mid-log phase used for the RNA-sequence analysis, cAMP appears to be solely responsible for the regulation of A1S_0112-0118.
This report has shown that pellicle formation is a phenotypic trait seen in a limited number of A. baumannii strains. For the first time, the secondary signalling molecule cAMP has been shown to play a critical role in pellicle formation as well as motility in A. baumannii. Additionally, this study has shown that insertions in CpdA, A1S_0112 and A1S_0115 completely abolish pellicle formation. Furthermore, transcriptional analyses identified that cAMP effects the expression of the A1S_0112-0118 operon essential for motility, thereby altering the migration pattern seen in the cpdA::Tn mutant strain grown on semi-sold agar; insertions in the above operon in A1S_0112 and A1S_0115 resulted in a non-motile phenotype. We thus speculate, that pellicle and motility phenotypes of A. baumannii are linked via the expression of cAMP and the A1S_0112-0118 operon. Increased cell surface hydrophobicity was identified as an indicator of pellicle formation ability in the clinical strains tested herein. Whole transciptome analysis has identified an abundance of ORFs affected by the increase in cAMP within the cpdA::Tn mutant. A number of these correlate with genes previously known to be involved in pellicle formation including the hms locus and a chaperone usher system as well as the A1S_0112-0118 region identified in this study. Considering the importance of pellicle formation in the persistence and transmission of numerous bacterial species, this study has contributed to our understanding of the regulatory mechanisms behind these significant virulence traits in A. baumannii.
The 54 clinical A. baumannii bacterial strains used in this study and the hyper-motile derivative of A. baumannii ATCC 17978 designated 17978hm have been described previously [34, 59]. All A. baumannii strains were cultured on LB media containing 1 % agar overnight at 37 °C, and single colonies transferred to LB broth for subsequent use in assays, unless otherwise stated. For maintenance, conjugation, cloning and replicating plasmids E. coli DH5α, SM10 or JM109 cells were used [60–62].
Assessment of pellicle formation
The pellicle formation assay was adapted from previously described protocol . In brief, overnight bacterial cultures were diluted 1:100 in 5 mL of LB broth and grown stagnant in polypropylene or glass tubes for 72 h at 25 °C without shaking in the dark [63, 64]. Quantitative measurement of the pellicle material was assessed by the addition of 1 mL of ethanol into the tube underneath the pellicle material. The floating pellicle was then removed and resuspended in phosphate buffered saline (PBS) and the OD600 measured using a spectrophotometer (Beckman DU 640). Quantitative data were collected from at least three experiments on three different days.
Motility was assessed by migration on semi-solid surfaces using media containing 0.25 % agar as described previously . Briefly, an overnight colony was collected using a sterile loop and used to inoculate the centre of LB medium containing 0.25 % agar. The plate was then incubated overnight at 37 °C. Strains were considered motile if any movement of growth from the original inoculation site was observed.
Eukaryotic cell adherence assays
A. baumannii cell adherence to A549 (human type 2 pneumocytes) was investigated as previously described . In brief, cell lines were grown in Dulbecco’s Modified Eagle medium (Invitrogen, Australia) supplemented with 10 % foetal bovine serum (Bovogen, Australia). Washed A549 monolayers in 24-well tissue culture plates were infected with bacterial inoculums containing ~1x107 colony forming units. After incubation at 37 °C for 4 h the culture medium was removed and the monolayers washed with PBS. The cell monolayers were detached from the plate using a treatment of 100 μL of 0.25 % trypsin in PBS. Eukaryotic cells were subsequently lysed by the addition of 400 μL 0.025 % Triton X-100 and serial 10-fold dilutions were spread plated on LB agar to determine the number of colony forming units of adherent bacteria per well. The collated data for the adherence assay were obtained from at least three independent experiments and represent the data points from each experiment in quadruplicate wells.
Cell surface hydrophobicity tests
Cell surface hydrophobicity was examined as described previously . In brief, overnight cultures were diluted in LB broth and incubated at 37 °C for 2 h. Potassium urea magnesium buffer was used to wash the cells twice, before incubation at 30 °C for 20 min. The cell suspension was adjusted to OD600 ~ 0.25 in the same buffer in glass vials before the addition of xylene (3 mL). The sample was vortexed for 20 sec and the aqueous phase transferred to a 1.5 mL tube. The OD600 of the cell suspension was determined before (ODinitial) and after (ODfinal) the addition of xylene. The hydrophobicity index, expressed as a percentage, is calculated as (ODinital − ODfinal/ODinitial)*100. Quantitative data were collected from at least three experiments from three different days.
Transposon mutagenesis was undertaken using a modified mini Tn10 transposon (mini::Tn10:gfp:kan) carried on the plasmid pLof . The pLof mini::Tn10:gfp:kan plasmid was transformed into the donor E. coli SM10 strain and mated with the A. baumannii 17978hm recipient at a 1:10 ratio for 4 h at 37 °C on LB agar exconjugates were resuspended in 5 mL of LB broth and plated onto selective media containing 50 μg/mL kanamycin and 25 μg/mL chloramphenicol to select for transposition events and counter-select against the E. coli SM10 donor strain. Approximately, 4000 exconjugates were individually picked using sterile toothpicks to inoculate 200 μL of LB broth in 96 well microtitre plates. The cells were grown at 25 °C in the dark for 72 h and assessed visually for pellicle loss.
Identification of mini::Tn10:gfp:kan insertion in the chromosome
Chromosomal DNA isolated from A. baumannii 17978hm containing the mini::Tn10:gfp:kan was digested with restriction endonucleases XbaI and SacII for 4 h at 37 °C. Digested products were cloned into pBluescript SK II (Agilent technologies, CA, USA) and clones were confirmed by sequencing (Australian Genome Research Facility, Adelaide, South Australia). The insertion site of the transposons in the chromosome was determined by Sanger sequencing using an oligonucleotide designed to read out of the end of the transposon (see Additional file 2).
Complementation of transposon insertions
For genetic complementation of the non-pellicle forming A. baumannii mutants, the genes corresponding to A1S_0112, A1S_0115 and cpdA were PCR amplified, cloned into the shuttle vector pWH1266, transformed into DH5α E. coli cells and sequenced. Plasmids were extracted and subsequently transformed via electroporation using a MicroPulser (Bio-Rad, CA, USA) at 2.5 kV, 200 ohms and 25 microfarads into their respective A. baumannii transposon insertion strains . PCR was achieved with VELOCITY™ DNA polymerase (Bioline, Vic, Australia) and the oligonucleotides (Additional file 2) using the following PCR protocols. For amplification of cpdA, cycling conditions were 2 min at 94 °C, 30 cycles of 45 sec at 94 °C, 45 sec at 55 °C, and 90 sec at 72 °C, followed by a final extension of 4 min at 72 °C. Amplification of A1S_0112 used the same conditions with the exception of the annealing temperature which was increased to 60 °C. For amplification of A1S_0115, the cycling conditions were 2 min at 94 °C, 30 cycles of 90 sec at 94 °C, 90 sec at 60 °C, 4 min at 72 °C and a final extension of 6 min at 72 °C.
Measurement of intracellular cAMP levels
Assessment of cAMP levels within A. baumannii strains cpdA::Tn, cpdA::Tn(0249), cpdA::Tn(1266) and 17978hm was performed with cells were grown to an OD600 of 0.7. Cells were pelleted by centrifugation at 4500 g for 15 min, resuspended in PBS and disrupted at 30,000 lb per square inch using the One Shot Head from the TS series bench top disruptor (Constant Systems, United Kingdom). The resulting cellular material in the cell lysis solution was clarified by centrifugation at 4500 g for 30 min and the supernatant removed and stored at −80 °C until required. The concentration of cAMP was measured using the Cyclic Nucleotide XP Enzymatic Immunoassay kit (Cell Signalling Technology, MA, USA) according to the manufacturer’s instructions. In brief, 50 μL of horseradish peroxidase-linked cAMP solution and 50 μL of sample were added to the supplied 96-well tray, which was then covered and incubated at room temperature for 3 h on a horizontal orbital plate shaker. The contents of the plate were discarded and the plate was washed 4 times with 200 μL/well of wash buffer. Next, 100 μL of 3,3′,5,5′-tetramethylbenzidinesubstrate solution was added and incubated for 30 min at room temperature, after which 100 μL of stop solution was added and the absorbance measured at 450 nm. Using the inverse for the standards derived from a standard curve and adjusted for the amount of total protein added, the amount of cAMP was calculated. Total protein concentrations were quantified using the Lowry protein assay (Bio-Rad, CA, USA) following the manufacturer’s instructions.
Isolation of total cellular RNA
To measure transcriptional levels for either RT-PCR or in Hiseq RNA transcriptome analysis conducted by SA Pathology (Central Adelaide local Health Network via an Illumina platform), cells were isolated and RNA extracted as below. Bacterial cells were harvested from either semi-solid media (0.25 % LB agar) for RT-PCR using pre-chilled PBS or harvested from Mueller Hinton broth at OD600 of 0.6 for Hiseq. Bacterial cells were pelleted by centrifugation at 4500 g for 15 min and lysed in 1 mL TRIzol reagent (Life Technologies, Australia) and 200 μL of chloroform. Following phase separation, the aqueous layer was collected and RNA isolated using the Isolate RNA Mini Kit (Bioline, NSW, Australia) following the manufacturer’s recommendations. Samples for RT-PCR were subsequently treated with DNaseI (Promega, WI, USA) for 30 min at 37 °C and 1 μL of stop solution was added before a final incubation for 10 min at 65 °C. Triplicate samples were pooled and sent to SA Pathology for Ribosomal RNA reduction using Epicentre Ribo-Zero kit and RNA library bar-coding was undertaken on RNA for HiSeq. Samples were run on the Illumina HiSeq platform (1X50 base pair single reads). These data are accessible through GEO Series accession number GSE64935 (http://0-www.ncbi.nlm.nih.gov.brum.beds.ac.uk/geo/query/acc.cgi?acc= GSE64935).
The quality of the cDNA reads obtained from the HiSeq was checked using ‘Fastqc’ (http://www.bioinformatics.babraham.ac.uk/projects/fastqc/). Reads were subsequently mapped to the reference genome using ‘bowtie’ aligner (http://bowtie-bio.sourceforge.net/index.shtml). For the reads obtained, approximately 95 % were mapped to the ATCC 17978 genome of which 78 % mapped to the coding regions. In order to determine changes in gene expression the number of reads obtained for each ORF was normalised using reads per kilobase of transcript per million reads mapped.
Quantitative reverse transcription PCR
M-MLV reverse-transcriptase (Promega) and random hexamers (GeneWorks, SA, Australia) were used to synthesise complementary deoxyribonucleic acid (cDNA) following the manufacturer’s recommendations. qRT-PCR was performed on three biological replicates using DyNAmo SYBR green qRT-PCR kits (ThermoScientific, Vic, Australia) in conjunction with a Rotor-Gene RG-3000 (Corbett Life Science, Australia). A typical qRT-PCR run was as follows: 1 min at 95 °C, 40 cycles of 10 sec at 95 °C, 15 sec at 55 °C and 20 sec at 72 °C . Transcriptional differences were calculated using the ΔΔCT method . The 16S rRNA (A1S_r01) and gapDH (A1S_2501) transcription levels were used as a reference.
This project was financially supported by the National Health and Medical Research Council Australia Project Grant 535053 and a Flinders Medical Research Foundation Grant. SKG is the recipient of a Flinders University Research Scholarship (FURS).
- Bergogne-Bérézin E. Treatment of Acinetobacter infections. Expert Opin Investig Drugs. 1997;6(2):119–27.PubMedView ArticleGoogle Scholar
- McQueary CN, Actis LA. Acinetobacter baumannii biofilms: Variations among strains and correlations with other cell properties. J Microbiol. 2011;49(2):243–50.PubMedView ArticleGoogle Scholar
- Dijkshoorn L, Nemec A, Seifert H. An increasing threat in hospitals: multidrug-resistant Acinetobacter baumannii. Nat Rev Microbiol. 2007;5(12):939–51.PubMedView ArticleGoogle Scholar
- Shin JA, Chang YS, Kim HJ, Kim SK, Chang J, Ahn CM, et al. Clinical outcomes of tigecycline in the treatment of multidrug-resistant Acinetobacter baumannii infection. Yonsei Med J. 2012;53(5):974–84.PubMed CentralPubMedView ArticleGoogle Scholar
- Roca I, Espinal P, Vila-Farrés X, Vila J. The Acinetobacter baumannii oxymoron: commensal hospital dweller turned pan-drug-resistant menace. Front Microbiol. 2012;23(3):148.Google Scholar
- Peleg AY, Seifert H, Paterson DL. Acinetobacter baumannii: emergence of a successful pathogen. J Clin Microbiol Rev. 2008;21(3):538–82.View ArticleGoogle Scholar
- Ramirez MS, Don M, Merkier AK, Bistué AJ, Zorreguieta A, Centrón D, et al. Naturally competent Acinetobacter baumannii clinical isolate as a convenient model for genetic studies. J Clin Microbiol. 2010;48(4):1488–90.PubMed CentralPubMedView ArticleGoogle Scholar
- Neely AN, Maley MP, Warden GD. Computer keyboards as reservoirs for Acinetobacter baumannii in a burn hospital. Clin Infect Dis. 1999;29(5):1358–60.PubMedView ArticleGoogle Scholar
- Donlan RM, Costerton JW. Biofilms: survival mechanisms of clinically relevant microorganisms. Clin Microbiol Rev. 2002;15(2):167–93.PubMed CentralPubMedView ArticleGoogle Scholar
- Karaiskos I, Galani L, Baziaka F, Giamarellou H. Intraventricular and intrathecal colistin as the last therapeutic resort for the treatment of multidrug-resistant and extensively drug-resistant Acinetobacter baumannii ventriculitis and meningitis: a literature review. Int J Antimicrob Agents. 2013;41(6):499–508.PubMedView ArticleGoogle Scholar
- Strassle P, Thom KA, Johnsonm JK, Leekha S, Lissauer M, Zhu J, et al. The effect of terminal cleaning on environmental contamination rates of multidrug-resistant Acinetobacter baumannii. Am J Infect Control. 2012;40(10):1005–7.PubMedView ArticleGoogle Scholar
- Karageorgopoulos DE, Falagas ME. Current control and treatment of multidrug-resistant Acinetobacter baumannii infections. Lancet Infect Dis. 2008;8(12):751–62.PubMedView ArticleGoogle Scholar
- Longo F, Vuotto C, Donelli G. Biofilm formation in Acinetobacter baumannii. New Microbiol. 2014;37(2):119–27.PubMedGoogle Scholar
- Yamamoto K, Arai H, Ishii M, Igarashi Y. Involvement of flagella-driven motility and pili in Pseudomonas aeruginosa colonization at the air-liquid interface. Microbes Environ. 2012;27(3):320–3.PubMed CentralPubMedView ArticleGoogle Scholar
- Moonmangmee S, Kawabata K, Tanaka S, Toyama H, Adachi O, Matsushita K. A novel polysaccharide involved in the pellicle formation of Acetobacter aceti. J Biosci Bioeng. 2002;93(2):192–200.PubMedView ArticleGoogle Scholar
- Yuan J, Chen Y, Zhou G, Chen H, Gao H. Investigation of roles of divalent cations in Shewanella oneidensis pellicle formation reveals unique impacts of insoluble iron. Biochim Biophys Acta. 2013;1830(11):5248–57.PubMedView ArticleGoogle Scholar
- Espinal P, Martí S, Vila J. Effect of biofilm formation on the survival of Acinetobacter baumannii on dry surfaces. J Hosp Infect. 2012;80(1):56–60.PubMedView ArticleGoogle Scholar
- Kanchanarach W, Theeragool G, Inoue T, Yakushi T, Adachi O, Matsushita K. Acetic acid fermentation of Acetobacter pasteurianus: relationship between acetic acid resistance and pellicle polysaccharide formation. J Biosci Bioeng. 2010;74(8):1591–7.Google Scholar
- Sambandan D, Dao DN, Weinrick BC, Vilchéze C, Gurcha SS, Ojha A, et al. Keto-mycolic acid-dependent pellicle formation confers tolerance to drug-sensitive Mycobacterium tuberculosis. MBio. 2013;4(3):e00222–00213.PubMed CentralPubMedView ArticleGoogle Scholar
- Tomaras AP, Dorsey CW, Edelmann RE, Actis LA. Attachment to and biofilm formation on abiotic surfaces by Acinetobacter baumannii: involvement of a novel chaperone-usher pili assembly system. Microbiol. 2003;149(12):3473–84.View ArticleGoogle Scholar
- Kobayashi K. Bacillus subtilis pellicle formation proceeds through genetically defined morphological changes. J Bacteriol. 2007;189(13):4920–31.PubMed CentralPubMedView ArticleGoogle Scholar
- Martí S, Nait Chabane Y, Alexandre S, Coquet L, Vila J, Jouenne T, et al. Growth of Acinetobacter baumannii in pellicle enhanced the expression of potential virulence factors. PLoS One. 2011;6(10):e26030.PubMed CentralPubMedView ArticleGoogle Scholar
- Nait Chabane Y, Martí S, Rihouey C, Alexandre S, Hardouin J, Lesouhaitier O, et al. Characterisation of pellicles formed by Acinetobacter baumannii at the air-liquid interface. PLoS One. 2014;9(10):e111660.PubMed CentralPubMedView ArticleGoogle Scholar
- Lazar V. Quorum sensing in biofilms - How to destroy the bacterial citadels or their cohesion/power? Anaerobe. 2011;17(6):280–5.PubMedView ArticleGoogle Scholar
- de Kievit TR. Quorum sensing in Pseudomonas aeruginosa biofilms. Environ Microbiol. 2009;11(2):279–88.PubMedView ArticleGoogle Scholar
- Kalivoda EJ, Brothers KM, Stella NA, Schmitt MJ, Shanks RM. Bacterial cyclic AMP-phosphodiesterase activity coordinates biofilm formation. PLoS One. 2013;8(7):e71267.PubMed CentralPubMedView ArticleGoogle Scholar
- Shemarova IV. cAMP-dependent signal pathways in unicellular eukaryotes. Crit Rev Microbiol. 2009;35(1):23–42.PubMedView ArticleGoogle Scholar
- Skorupski K, Taylor RK. Cyclic AMP and its receptor protein negatively regulate the coordinate expression of cholera toxin and toxin-coregulated pilus in Vibrio cholerae. Proc Natl Acad Sci U S A. 1997;94(1):265–70.PubMed CentralPubMedView ArticleGoogle Scholar
- Ono K, Oka R, Toyofuku M, Sakaguchi A, Hamada M, Yoshida S, et al. cAMP signaling affects irreversible attachment during biofilm formation by Pseudomonas aeruginosa PAO1. Microbes Environ. 2014;29(1):104–6.PubMed CentralPubMedView ArticleGoogle Scholar
- Wolfgang MC, Lee VT, Gilmore ME, Lory S. Coordinate regulation of bacterial virulence genes by a novel adenylate cyclase-dependent signaling pathway. Dev Cell. 2003;4(2):253–63.PubMedView ArticleGoogle Scholar
- Yahr TL, Vallis AJ, Hancock MK, Barbieri JT, Frank DW. ExoY, an adenylate cyclase secreted by the Pseudomonas aeruginosa type III system. Proc Natl Acad Sci U S A. 1998;95(23):13899–904.PubMed CentralPubMedView ArticleGoogle Scholar
- Jimenez PN, Koch G, Thompson JA, Xavier KB, Cool RH, Quax WJ. The multiple signaling systems regulating virulence in Pseudomonas aeruginosa. Microbiol Mol Biol Rev. 2012;76(1):46–65.PubMedView ArticleGoogle Scholar
- Fuchs EL, Brutinel ED, Jones AK, Fulcher NB, Urbanowski ML, Yahr TL, et al. The Pseudomonas aeruginosa Vfr regulator controls global virulence factor expression through cyclic AMP-dependent and -independent mechanisms. J Bacteriol. 2010;192(14):3553–64.PubMed CentralPubMedView ArticleGoogle Scholar
- Eijkelkamp BA, Stroeher UH, Hassan KA, Papadimitrious MS, Paulsen IT, Brown MH. Adherence and motility characteristics of clinical Acinetobacter baumannii isolates. FEMS Microbiol Lett. 2011;323(1):44–51.PubMedView ArticleGoogle Scholar
- Martí S, Rodríguez-Bano J, Catel-Ferreira M, Jouenne T, Vila J, Seifert H, et al. Biofilm formation at the solid-liquid and air-liquid interfaces by Acinetobacter species. BMC Res Notes. 2011;4(5):1–4.Google Scholar
- Pour NK, Dusane DH, Dhakephalkar PK, Zamin FR, Zinjarde SS, Chopade BA. Biofilm formation by Acinetobacter baumannii strains isolated from urinary tract infection and urinary catheters. FEMS Immunol Med Microbiol. 2011;62(3):328–38.PubMedView ArticleGoogle Scholar
- Costa GF, Tognim MC, Cardoso CL, Carrara-Marrone FE, Garcia LB. Preliminary evaluation of adherence on abiotic and cellular surfaces of Acinetobacter baumannii strains isolated from catheter tips. Braz J Infect Dis. 2006;10(5):346–51.PubMedView ArticleGoogle Scholar
- Furuhata K, Kato Y, Goto K, Hara M, Fukuyama M. Diversity of heterotrophic bacteria isolated from biofilm samples and cell surface hydrophobicity. J Gen Appl Microbiol. 2009;55(1):69–74.PubMedView ArticleGoogle Scholar
- Kouidhi B, Zmantar T, Hentati H, Bakhrouf A. Cell surface hydrophobicity, biofilm formation, adhesives properties and molecular detection of adhesins genes in Staphylococcus aureus associated to dental caries. Microb Pathog. 2010;49(1-2):14–22.PubMedView ArticleGoogle Scholar
- Chiku K, Tsunemi K, Yamamoto M, Ohnishi-Kameyama M, Yoshida M, Ishii T, et al. Defects in D-rhamnosyl residue biosynthetic genes affect lipopolysaccharide structure, motility, and cell-surface hydrophobicity in Pseudomonas syringae pathovar glycinea race 4. Biosci Biotechnol Biochem. 2013;77(3):505–10.PubMedView ArticleGoogle Scholar
- Raut J, Rathod V, Karuppayil SM. Cell surface hydrophobicity and adhesion: a study on fifty clinical isolates of Candida albicans. Nippon Ishinkin Gakkai Zasshi. 2010;51(3):131–6.View ArticleGoogle Scholar
- Delcour AH. Outer membrane permeability and antibiotic resistance. Biochim Biophys Acta. 2009;1794(5):808–16.PubMed CentralPubMedView ArticleGoogle Scholar
- Eijkelkamp BA, Stroeher UH, Hassan KA, Elbourne LD, Paulsen IT, Brown MH. H-NS plays a role in expression of Acinetobacter baumannii virulence features. Infect Immun. 2013;81(7):2574–83.PubMed CentralPubMedView ArticleGoogle Scholar
- Stretton S, Techkarnjanaruk S, McLennan AM, Goodman AE. Use of green fluorescent protein to tag and investigate gene expression in marine bacteria. Appl Environ Microbiol. 1998;64(7):2554–9.PubMed CentralPubMedGoogle Scholar
- Fuchs EL, Brutinel ED, Klem ER, Fehr AR, Yahr TL, Wolfgang MC. In vitro and in vivo characterization of the Pseudomonas aeruginosa cyclic AMP (cAMP) phosphodiesterase CpdA, required for cAMP homeostasis and virulence factor regulation. J Bacteriol. 2010;192(11):2779–90.PubMed CentralPubMedView ArticleGoogle Scholar
- Hunger M, Schmucker R, Kishan V, Hillen W. Analysis and nucleotide sequence of an origin of DNA replication in Acinetobacter calcoaceticus and its use for Escherichia coli shuttle plasmids. Gene. 1990;87(1):45–51.PubMedView ArticleGoogle Scholar
- Clemmer KM, Bonomo RA, Rather PN. Genetic analysis of surface motility in Acinetobacter baumannii. Microbiol. 2011;157(9):2534–44.View ArticleGoogle Scholar
- Stewart CR, Rossier O, Cianciotto NP. Surface translocation by Legionella pneumophila: a form of sliding motility that is dependent upon type II protein secretion. J Bacteriol. 2009;191(5):1537–46.PubMed CentralPubMedView ArticleGoogle Scholar
- Rumbo-Feal S, Gómez MJ, Gayoso C, Álvarez-Fraga L, Cabral MP, Aransay AM, et al. Whole transcriptome analysis of Acinetobacter baumannii assessed by RNA-sequencing reveals different mRNA expression profiles in biofilm compared to planktonic cells. PLoS One. 2013;8(8):e72968.PubMed CentralPubMedView ArticleGoogle Scholar
- Henrichsen J. The influence of changes in the environment on twitching motility. Acta Pathol Microbiol Scand B. 1975;83(3):179–86.PubMedGoogle Scholar
- Skiebe E, de Berardinis V, Morczinek P, Kerrinnes T, Faber F, Lepka D, et al. Surface-associated motility, a common trait of clinical isolates of Acinetobacter baumannii, depends on 1,3-diaminopropane. Int J Med Microbiol. 2012;302(3):117–28.PubMedView ArticleGoogle Scholar
- McDonough KA, Rodriguez A. The myriad roles of cyclic AMP in microbial pathogens: from signal to sword. Nat Rev Microbiol. 2012;10(1):27–38.Google Scholar
- Saroj SD, Rather PN. Streptomycin inhibits quorum sensing in Acinetobacter baumannii. Antimicrob Agents Chemother. 2013;57(4):1926–9.PubMed CentralPubMedView ArticleGoogle Scholar
- Carruthers MD, Harding CM, Baker BD, Bonomo RA, Hujer KM, Rather PN, et al. Draft genome sequence of the clinical isolate Acinetobacter nosocomialis strain M2. Genome Announc. 2013;1(6):e00906–13.PubMed CentralPubMedView ArticleGoogle Scholar
- Vallet I, Olson JW, Lory S, Lazdunski A, Filloux A. The chaperone/usher pathways of Pseudomonas aeruginosa: identification of fimbrial gene clusters (cup) and their involvement in biofilm formation. Proc Natl Acad Sci U S A. 2001;98(12):6911–6.PubMed CentralPubMedView ArticleGoogle Scholar
- Eijkelkamp BA, Stroeher UH, Hassan KA, Paulsen IT, Brown MH. Comparative analysis of surface-exposed virulence factors of Acinetobacter baumannii. BMC Genomics. 2014;15:1020.PubMed CentralPubMedView ArticleGoogle Scholar
- Choi AH, Slamti L, Avci FY, Pier GB, Maira-Litrán T. The pgaABCD locus of Acinetobacter baumannii encodes the production of poly-b-1-6-N-acetylglucosamine, which is critical for biofilm formation. J Bacteriol. 2009;191(19):5953–63.PubMed CentralPubMedView ArticleGoogle Scholar
- Wada A, Mikkola R, Kurland CG, Ishihama A. Growth phase-coupled changes of the ribosome profile in natural isolates and laboratory strains of Escherichia coli. J Bacteriol. 2000;182(10):2893–9.PubMed CentralPubMedView ArticleGoogle Scholar
- Eijkelkamp BA, Hassan KA, Paulsen IT, Brown MH. Investigation of the human pathogen Acinetobacter baumannii under iron limiting conditions. BMC Genomics. 2011;12(1):126.PubMed CentralPubMedView ArticleGoogle Scholar
- Hanahan D. Studies on transformation of Escherichia coli with plasmids. J Mol Biol. 1983;166(4):557–80.PubMedView ArticleGoogle Scholar
- Simon R, Priefer U, Puhler A. A broad host range mobilization system for invivo genetic-engineering - transposon mutagenesis in Gram-negative bacteria. BioTechnol. 1983;1(9):784–91.View ArticleGoogle Scholar
- Yanisch-Perron C, Vieira J, Messing J. Improved M13 phage cloning vectors and host strains: nucleotide sequences of the M13mp18 and pUC19 vectors. Gene. 1985;33(1):103–19.PubMedView ArticleGoogle Scholar
- Mussi MA, Gaddy JA, Cabruja M, Arivett BA, Viale AM, Rasia R, et al. The opportunistic human pathogen Acinetobacter baumannii senses and responds to light. J Bacteriol. 2010;192(24):6336–45.PubMed CentralPubMedView ArticleGoogle Scholar
- Golic A, Vaneechoutte M, Nemec A, Viale AM, Actis LA, Mussi MA. Staring at the cold sun: blue light regulation is distributed within the genus Acinetobacter. PLoS One. 2013;8(1):e55059.PubMed CentralPubMedView ArticleGoogle Scholar
- Rosenberg E, Gottlieb A, Rosenberg M. Inhibition of bacterial adherence to hydrocarbons and epithelial cells by emulsan. Infect Immun. 1983;39(3):1024–8.PubMed CentralPubMedGoogle Scholar
- Matthysse AG, Stretton S, Dandie C, McClure NC, Goodman AE. Construction of GFP vectors for use in gram-negative bacteria other than Escherichia coli. FEMS Microbiol Lett. 1996;145(1):87–94.PubMedView ArticleGoogle Scholar
- Brazma A, Vilo J. Gene expression data analysis. Microbes Infect. 2001;3(10):823–9.PubMedView ArticleGoogle Scholar
- Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) method. Methods. 2001;25(4):402–8.PubMedView ArticleGoogle Scholar
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