Pseudomonas aeruginosa is a globally distributed environmental bacterium, which is also a significant opportunistic pathogen of humans, animals and plants. It is considered that wide distribution of this bacterium is connected with its most significant constitutive property to form biofilms, and that this multicellular mode of growth, predominant in nature, serves as a protective mechanism against unfavourable environmental conditions. The work presented here examines the phenotypic diversity of Pseudomonas aeruginosa environmental isolates with respect to biofilm production capacity under different environmental conditions (temperature, pH, NaCl), production of virulence factors, and motility. The purpose of this work is to present the production of two quorum sensing-regulated virulence factors (rhamnolipids and pyocyanin), explore different motility tests (swimming, swarming and twitching) and discover potential relationship between assessed phenotypic features. Obtained results delineate environmental conditions coinciding with biofilm production and suggest a high correlation between rhamnolipid production levels and biofilm formation. Rhamnolipids affect motility competence, yet only the flagellum-mediated swimming motility has significant impact on the biofilm formation potential. Although it is challenging to demarcate a definitive, clear correlation between parameters tested, rhamnolipid content appears to serve as a link between the tested phenotypic factors.

INTRODUCTION

Pseudomonas aeruginosa is an environmentally significant, ubiquitous Gram-negative, rod shaped γ-proteobacterium. It is an important opportunistic human pathogen, also capable of causing infections in non-mammalian host species, such as insects, nematodes and plants. P. aeruginosa successfully colonizes different natural ecological habitats, including soil, water, rhizosphere, as well as wastewaters, wastewater treatment plants, wasted sludge leachate and, consequently, irrigation water or agricultural soil, thus increasing potential public health risks. Given that it is commonly present in public drinking water supply systems and enclosed spaces, such as hospitals and schools, many studies have investigated its major natural features (physical, physiological, biochemical, biological) that may be responsible for high survival and pathogenicity rates (Driscoll et al. 2007; Gellatly & Hancock 2013; Grosso-Becerra et al. 2014). It is thought that widespread distribution of P. aeruginosa is a consequence of its most significant constitutive property to form biofilms and that multicellular growth, predominantly encountered in nature, serves as a protective mechanism against unfavourable environmental conditions (antibiotics, chlorine and other disinfectants). Biofilm formation is linked to a form of inter-bacterial communication known as quorum sensing (QS), in which small diffusible signalling molecules (autoinducers) globally regulate gene expression. Using QS, bacterial populations can switch from acting as individual cells to operating in a concerted, multi-cellular fashion (Cady et al. 2012). Similarly, pathogenicity is dependent on the production and secretion of multiple virulence factors (pyocyanin, lectin, biosurfactant rhamnolipid, and others), also regulated at the transcriptional level by QS response. Common level of regulation indicates that biofilm production and virulence factor secretion may be linked, including clinical and environmental isolates equally. Indeed, a recent study argued that P. aeruginosa genomes of clinical and environmental isolates are highly conserved, and that all isolates are capable of producing virulence-associated traits, thus serving as potential pathogens (Grosso-Becerra et al. 2014).

P.aeruginosa's flagella provide swimming motility and are necessary for biofilm formation in liquid environments, while type IV pili enable twitching motility and are required for biofilm and microcolony formation on surfaces. It is speculated that swimming motility might enable bacteria to overcome repulsive forces at the water-surface interface, so that they reach the surface and that microcolonies are formed by twitching motility-driven cell aggregation. Additionally, P. aeruginosa is capable of swarming motility, a rapid and coordinated migration of bacterial population across a semi-solid surface. This multicellular phenomenon has recently gained increasing attention, as it is suspected to play a role in biofilm development of P. aeruginosa (Tremblay & Deziel 2008). Other reports propose that P. aeruginosa colonizes surfaces in vitro by either biofilm formation or swarming motility (Murray et al. 2010).

Swarming motility requires the production of two biosurfactants – rhamnolipids and 3-hydroxyalkanoic acids. Since rhamnolipids are also implicated in many aspects of biofilm development, it has been proposed that these two multicellular behaviours are linked. The aim of this paper is to examine pyocyanin and rhamnolipid production in biofilm formation and compare it with bacteria's capacity for motility. Investigated P. aeruginosa isolates were collected from Lake Palić in northern Serbia, which is mainly filled with fresh water by neighbouring wastewater treatment plant effluent, but, unofficially, also with unknown quantities of numerous other point and diffuse water resources of unknown quality. In summer months it is also filled with water coming from the Tisa river. Together with Krvavo Lake, Great Park and the Zoo, it is part of the Palić Nature Park (category III protected area). Lake Palić is an interesting ecosystem with intensive eutrophication process, bioremediation and recultivation possibilities, as well as water and sediment quality from the ecological and sanitary points of view. However, investigations of individual bacterial strains, such as P. aeruginosa, in the region are missing, despite their significant impact on the health status of animals and plants in the entire protected area, as well as drinking water resources and human environmental conditions.

MATERIAL AND METHODS

Sample collection

Investigated P. aeruginosa isolates were collected monthly during 2014 from Lake Palić water and sediment. A total of 150 water and sediment samples were collected, from which four isolates were isolated: isolate P_V_2 (location: ‘Muški štrand’), isolate P_V_3 (location: Sector 2–Sector 3 crosspoint), isolate KTP_V_9 (location: channel between the Tisa and Lake Palić) and isolate KPL_V_4 (location: channel between Palić and Ludaš Lakes).

Isolation, cultivation and identification procedure

Water samples were incubated for 24 h at 37 °C/120 rpm, before 200 μL of each sample was inoculated into cetrimide agar (HiMedia Laboratories, India). Inoculated plates were incubated 24–48 h at 42 °C. Typical colonies were reisolated and identified with API 20 NE (BioMérieux, France). Molecular identification based on an internal fragment of a housekeeping gene, acetyl-coenzyme A synthetase (acsA), confirmed that all four isolates are indeed P. aeruginosa (Vujović et al. in preparation). P. aeruginosa PAO 1 strain was used as a reference strain in all the assays. For each performed test, overnight bacterial culture, grown in lysogeny broth (LB) medium (tryptone 10 g/L, yeast extract 5 g/L, NaCl 10 g/L) at 37 °C in a rotary shaker (120 rpm), was used. All measures were completed in triplicate.

Pyocyanin and rhamnolipid production

Pyocyanin assay was performed according to Grosso-Becerra et al. (2014). Briefly, bacterial cultures were centrifuged for 10 min at 4 °C/10,000 rpm, 7.5 mL of supernatant was extracted with 4.5 mL of chloroform and reextracted into 1.5 mL of 0.2 N HCl (resulting in pink to deep red colour shift). Absorbance of the solution was measured at 520 nm and the resulting optical density reading (OD520) multiplied by 17.072 (molar attenuation coefficient) to yield final pyocyanin concentration (μg pyocyanin/mL culture supernatant).

Rhamnolipids concentration was estimated by the orcinol method (Fang et al. 2005). 500 μL of filtered culture supernatant was extracted twice with 1 mL of chloroform: ethanol (2:1, v/v). The organic phase was evaporated to dryness and dissolved in 0.2 mL of deionized water. 900 μL of a solution containing 0.19% orcinol (in 53% sulfuric acid) was added to 100 μL of each sample. Samples were heated at 80 °C in a water bath for 30 min and cooled for 15 min at room temperature, followed by A421 measuring. Concentrations of rhamnolipids were determined by comparing the data with those obtained with l-rhamnose standards between 0 and 50 μg/mL, multiplied by 3.4 and expressed as rhamnose equivalents.

Motility assay

Plates containing LB medium solidified with 0.3% (swimming motility), 0.5% (swarming motility) and 1.0% (twitching motility) agar were used. For swarming motility assay, plates with modified M8 medium (200 mL of 5x M9 salt solution (64 g Na2HPO4 × 7H2O, 15 g KH2PO4, 2.5 g NaCl and dH2O up to 1,000 mL), supplemented with 0.4% glucose, 0.05% tryptone and 2 mL 1 M MgSO4), solidified with 0.5% agar, were also used. After pouring the media, plates were allowed to dry for 60 min (first 20 min under UV light) in the laminar flow cabinet (Tremblay & Deziel 2008). 5 μL of bacteria suspended in LB adjusted to OD600 = 1.0 were spotted on each plate and incubated at 22 and 37 °C for 24 h. Motility zone was expressed in mm motility and motility plates captured with a digital camera.

Biofilm formation

Biofilm formation was analysed by microtiter plate assay (O'Toole 2011). A microtiter dish was filled with 200 μL bacteria suspended in LB, adjusted to OD600 = 0.3–0.4 (≈109 CFU/mL). Following 24 h incubation at 22 and 37 °C, planktonic bacteria were removed and the plate washed twice with 200 μL phosphate-buffered saline (PBS) buffer (8.1 g/L NaCl, 0.2 g/L KCl, 0.61 g/L Na2HPO4, 0.2 g/L KH2PO4, pH = 7). The plates were air dried and biofilms fixed with methanol for 15 min and stained with 0.4% crystal violet dye (20 min). Dye was washed out with tap water and the colour resuspended with 33% glacial acetic acid. Colour intensity was measured by microtiter plate reader at λ = 630 nm. Possible effects of different environmental conditions on biofilm formation were examined in the same way, using LB medium adjusted to pH = 6 and LB medium with doubled NaCl concentration (10 g/L tryptone, 5 g/L yeast extract, 20 g/L NaCl). Bacterial suspension was prepared from bacterial colonies grown overnight on nutritious agar and then suspended in modified LB medium, adjusted to OD600 = 0.3–0.4 (≈109 CFU/ml). P. aeruginosa PAO 1 strain was used as a positive control.

Statistical analyses

Descriptive statistics and relationship between observed results were obtained using Microsoft Excel (2013). Mean values of data were compared by Spearman's rank correlation coefficient at significance level ɑ = 0.01.

RESULTS AND DISCUSSION

Results obtained in this study show that all investigated isolates were capable of biofilm formation (Figure 1(a)). Testing the potential to produce biofilms at different temperatures, given that temperature is expected to impact bacterial growth and biofilm formation (due to differential nutrient intake, enzyme reaction rates and physical properties of the compounds within and surrounding the cells), indicated a more successful biofilm formation at 22 °C than at 37 °C (Figure 1(a)). These findings are congruent with previous reports proposing decrease (up to 98%) in biofilm production at 37 °C (Hostacka et al. 2010). However, opposing findings have also been published. For instance, Kannan & Gautam (2015) suggested stronger biofilm formation and its higher mechanical stability at 37 °C, compared to 28, 33 and 42 °C. Results from this study, on the other hand, speak in favour of the idea that the initial interaction between bacteria and substrate may increase with lowering the temperature, increasing the likelihood of adhesion (Garrett et al. 2008). More uniform properties of polysaccharides at lower temperatures may increase the possibility of biofilm adhesion, since many microbial polysaccharides undergo transition from an ordered state at lower temperatures and in the presence of ions, to a disordered state at elevated temperatures in low ionic environments (Garrett et al. 2008).
Figure 1

Biofilm quantity measured in LB medium (a), LB medium under pH = 6 (b) and LB medium with increased NaCl concentration (c). Histograms show median values, vertical lines link minimal and maximal values reached and circles are at the standard deviation.

Figure 1

Biofilm quantity measured in LB medium (a), LB medium under pH = 6 (b) and LB medium with increased NaCl concentration (c). Histograms show median values, vertical lines link minimal and maximal values reached and circles are at the standard deviation.

Bacteria have the potential to change in response to internal and external pH conditions by adjusting the activity and synthesis of proteins associated with many different cellular processes (Olsen 1993). This suggests that bacteria contain mechanisms in place, allowing their populations to adapt to small environmental changes in pH values. Therefore, the potential impact of pH on biofilm formation was also examined. Obtained data indicate that lower pH conditions do not inhibit biofilm development (Figure 1(b)), as biomass quantities were either equivalent to or higher than the values at neutral pH. The exception was isolate KTP_V_9, which exhibited reduced biofilm formation at pH 6 and 22 °C (Figure 1(b)). On the other hand, NaCl increased biofilm abundance in all tested isolates, particularly at 22 °C (Figure 1(c)), indicating that this condition favours biofilm formation. Martinez (2011), on the contrary, reported the inhibitory effect of high NaCl concentrations (2, 4, and 7%) on P. aeruginosa biofilms. Additionally, saline concentrations in tears, for example, have been proven an effective antimicrobial agent against organisms that could potentially infect the eye (Kwong et al. 2007), including P. aeruginosa. Yet, results of this work do not support the idea that hypertonic saline aerosol treatment may be useful to prevent P. aeruginosa growth and biofilm development.

Factors which may influence the ability of P. aeruginosa to form biofilms were examined next, by assessing Pseudomonas virulence factors, given that they are also regulated by QS response at the molecular level. Pyocyanin is a water-soluble, blue-green extracellular pigment with antimicrobial properties. Rhamnolipids are glycolipid biosurfactants, cytotoxic to eukaryotic cells, which have a role in nutrient distribution in biofilm structure. Obtained results regarding pyocyanin and rhamnolipid production in tested isolates are presented in Table 1. Pyocyanin concentrations varied between 1.0 and 3.40 μg/mL and were significantly lower in comparison to 5.29 μg/mL obtained for the clinical isolate PAO1 (Grosso-Becerra et al. 2014), thus indicating limited pathogenicity of environmental isolates analysed in this study.

Table 1

Pyocyanin and rhamnolipids content

 Pyocyanin [μg/ml]Rhamnose equivalent [μg/ml]
P_V_2 2.32 13.0 
P_V_3 2.87 11.43 
KPL_V_4 1.0 11.67 
KTP_V_9 3.40 14.17 
PAO 1 5.29 178.2 
 Pyocyanin [μg/ml]Rhamnose equivalent [μg/ml]
P_V_2 2.32 13.0 
P_V_3 2.87 11.43 
KPL_V_4 1.0 11.67 
KTP_V_9 3.40 14.17 
PAO 1 5.29 178.2 

Results obtained in this study suggest moderate negative correlation between pyocyanin and biofilm production (data not shown) and moderate correlation between rhamnolipid production and the formation of biofilms, particularly at 22 °C (rs = 0.400), although the correlation is not statistically significant at ɑ = 0.01 (rs < rc, n = 5). Rhamnolipid production is controlled by the rhamnosyltransferase (rhl) mechanism, part of the QS system, whose activation is implicated in population level increase. However, rhamnolipids concentrations were similar in all investigated isolates in this study, thus it is challenging to argue a relationship between biofilm production, motility and rhamnolipid production.

Results of performed motility tests are depicted in Figures 2 and 3. Flagellum-mediated swimming in aqueous environment is shown at low concentration (<0.3%) of agar, type IV pilus-mediated twitching on solid surface (1% agar), while swarming motility is presented on semisolid (0.5%) agar, with normal (LB) or limited nitrogen sources (M8).
Figure 2

Qualitative motility assay for environmental P. aeruginosa isolates and PAO 1 reference strain: swimming motility (rows 1 and 2), swarming motility (rows 3, 4, 7 and 8) and twitching motility (rows 5 and 6).

Figure 2

Qualitative motility assay for environmental P. aeruginosa isolates and PAO 1 reference strain: swimming motility (rows 1 and 2), swarming motility (rows 3, 4, 7 and 8) and twitching motility (rows 5 and 6).

Figure 3

Quantitative motility assay for four environmental P. aeruginosa isolates and PAO 1 reference strain.

Figure 3

Quantitative motility assay for four environmental P. aeruginosa isolates and PAO 1 reference strain.

All tested isolates were capable of all three motility types (Figure 2). The largest motility zones were observed in flagellum-dependent movement, as well as in swarming under nitrogen deficiency, while swarming (LB) and twitching diameters were the smallest (Figures 2 and 3). Opposite the results for the biofilm forming ability, motility strength and tendril sharpness were more pronounced at 37 °C. This increase in swarming colony expansion with the increase in temperature is likely due to higher metabolic activity at higher temperatures (Murray et al. 2010). The results suggest moderate (rs = 0.300, ɑ ≤ 0.01) and strong (rs = 0.600, ɑ ≤ 0.01) correlation at 22 and 37 °C, respectively, between flagellum-mediated motility and swarming on M8 plates, as well as moderate correlation (rs = 0.400, ɑ ≤ 0.01) between flagellum-mediated motility and swarming at 37 °C. This confirms the importance of functional flagella for swarming motility, as indicated by Murray et al. (2010).

Swarming motility involves many conditions, including rhamnolipid production. Weak correlation was found between rhamnolipid production and swimming (rs = 0.200, ɑ ≤ 0.01) and twitching motility (rs = 0.300, ɑ ≤ 0.01). Very strong and statistically significant correlation was detected between rhamnolipid production and swarming on LB medium (rs = 0.900, rs = rc, ɑ ≤ 0.01), while strong negative correlation was attained between rhamnolipid production and swarming on oligo-nutrient medium (M8) (rs = −0.600, ɑ ≤ 0.01). These results are not surprising, given that the influence of QS upon swarming motility has been shown to be conditional and that changes to the growth medium (e.g. chemical composition) can significantly impact both surface motility and the importance of cell–cell signalling as bacteria attach to surfaces (Du et al. 2011). Interestingly, isolate KTP_V_9 produced the highest level of rhamnolipids (Table 1) and showed the most intensive propensity for flagellum-mediated motility. This isolate was the only one with clearly defined tendrils under 0.3% agar content at 37 °C (Figure 3). This finding favours the proposal that rhamnolipid production is a major factor regulating the expression of swarming motility (Croda-Garcia et al. 2011).

Finally, moderate correlation (rs = 0.500, ɑ ≤ 0.01) between swarming motility on M8 agar plates and biofilm producing potential was achieved, which is somewhat in agreement with findings of others (Caiazza et al. 2005; Murray et al. 2010), suggesting the importance of swarming motility under oligo-nutrient conditions in biofilm formation. Further, moderate correlation between swimming motility and biofilm forming potential at 37 °C (rs = 0.300, ɑ ≤ 0.01) was obtained, very weak correlation (rs = 0.100, ɑ ≤ 0.01) between swarming motility and biofilm quantity at 37 °C, as well as strong correlation, although not statistically significant, between twitching motility and biofilm production at 22 °C (rs = 0.775, rs < rc, ɑ ≤ 0.01). These findings support the importance of bacterial motile competence (flagellum and pili IV) in the biofilm forming process – while flagellum allows constant colonization, pili IV mediate twitching motility and virulence potential (O'Toole & Kolter 1998; Murray et al. 2010).

CONCLUSIONS

The presented study demonstrates high phenotypic diversity in P. aeruginosa isolates and points out the correlation between certain phenotypic characteristics. Higher potential for biofilm production at lower temperature and increased NaCl concentration (2%), together with decreased pyocyanin production, compared to the referent PAO1 clinical isolate, indicate a bacterial population adapted to environmental conditions. Rhamnolipid production correlates to generation of biofilm mass in tested isolates, indicating them as a useful tool for testing biofilm formation potential. Further, obtained data have shown that the flagellum-mediated motility is of great importance in the biofilm forming process, as well as that the flagellum-mediated transport is in correlation with swarming ability under low nutrient conditions. As rhamnolipids content is in moderate to high correlation with all tested motility types (except swarming on M8), this factor might be the most significant link between all phenotypic attributes.

Given that clinical and environmental isolates constitute a single population, this investigation provides initial knowledge regarding P. aeruginosa phenotype diversity in Lake Palić area and represents the basis for future studies of P. aeruginosa populations in Serbia, which are an absolute necessity for adequate environmental and public health protection.

ACKNOWLEDGEMENTS

This research was partially supported by the Ministry of Education, Science and Technological Development of the Republic of Serbia (grant numbers TR 31080 and TR34019) and the EU Commission project AREA, contract number 316004. The authors would like to thank Dr Branka Vasiljević from the Institute of Molecular Genetics and Genetic Engineering, University of Belgrade for kindly providing P. aeruginosa PAO1 strain.

REFERENCES

REFERENCES
Cady
N. C.
McKean
K. A.
Behnke
J.
Kubec
R.
Mosier
A. P.
Kasper
S. H.
Burz
D. S.
Musah
R. A.
2012
Inhibition of biofilm formation, quorum sensing and infection in Pseudomonas aeruginosa by natural products-inspired organosulfur compounds
.
PLoS ONE
7
(
6
),
e38492
.
Caiazza
N. C.
Shanks
R. M. Q.
O'Toole
G. A.
2005
Rhamnolipids modulate swarming motility patterns of Pseudomonas aeruginosa
.
Journal of Bacteriology
187
(
21
),
7351
7361
.
Du
H.
Xu
Z.
Shrout
J. D.
Alber
M.
2011
Multiscale modelling of Pseudomonas aeruginosa swarming
.
Math. Models Methods Appl. Sci.
21
(
1
),
939
954
.
Fang
X.
Wang
Q.
Shuler
P.
Bai
B.
2005
Bio engineering high performance microbial strains for MEOR by directed protein evolution technology
.
Final Report for US DOE Award No. DE-FC26-06NT15525. California Institute of Technology
,
Covina, CA, USA
.
Garrett
T. R.
Bhakoko
M.
Zhang
Z.
2008
Bacterial adhesion and biofilms on surfaces
.
Progress in Natural Science
18
(
9
),
1049
1056
.
Gellatly
S. L.
Hancock
R. E. W.
2013
Pseudomonas aeruginosa: new insights into pathogenesis and host defences
.
Pathogens and Disease
67
(
3
),
159
173
.
Grosso-Becerra
M. V.
Santos-Medellin
C.
Gonzales-Valdez
A.
Delgado
G.
Morales Espinosa
R.
Servin-Gonzalez
L.
Alcaraz
L. D.
Soberon-Chavez
G.
2014
Pseudomonas aeruginosa clinical and environmental isolates constitute a single population with high phenotypic diversity
.
BMC Genomics
15
(
318
).
Hostacka
A.
Ciznar
I.
Stefkovicova
M.
2010
Temperature and pH affect the production of bacterial biofilm
.
Folia Microbiologica
55
(
1
),
75
78
.
Kannan
A.
Gautam
P.
2015
A quantitative study on the formation of Pseudomonas aeruginosa biofilm
.
Springer Plus
4
(
379
).
Kwong
M. S. F.
Evans
D. J.
Ni
M.
Cowell
B. A.
Fleiszig
S. M. J.
2007
Human tear fluid protects against Pseudomonas aeruginosa keratitis in a murine experimental model
.
Infection and Immunity
75
(
5
),
2325
2332
.
Martinez
R. F.
2011
Effect of iron and sodium chloride on biofilm development of Stenotrophomonas maltophilia. College of Liberal Arts and Social Sciences Theses and Dissertations. Paper 102. http://via.library.depaul.edu/etd/102
.
Olsen
E. R.
1993
Influence of pH on bacterial gene expression
.
Molecular Microbiology
8
,
5
14
.
O'Toole
G. A.
2011
Microtiter dish biofilm formation assay
.
Journal of Visualized Experiments
47
,
2437
.