Flow Cytometry 2015.pdf

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ScienceDirect Agriculture and Agricultural Science Procedia 6 (2015) 567 – 578

“ST26733”, International Conference "Agriculture for Life, Life for Agriculture"

Flow Cytometry Based Method for Evaluation of Biodegradative Potential of Pseudomonas fluorescens Robertina IONESCUa*, LuminiĠa MĂRUğESCUb, Ana-Maria TĂNASEa, Iulia CHICIUDEANa, Ortansa CSUTAKa, Diana PELINESCUa, Tatiana VASSUa, Ileana STOICAa* b

a Department of Genetics, University of Bucharest, 1-3 Portocalelor Street, Bucharest, 060101, Romania, Department of Botany and Microbiology, University of Bucharest, 1-3 Portocalelor Street, Bucharest, 060101, Romania

Abstract Flow cytometry is recognized as a useful tool in environmental microbiology studies, regarding oil contamination sites. This study was focused on cytometric analysis of cellular growth of P. fluorescens S7A strain, previously isolated from oil polluted soil, in order to assess bacterial response to different n-hexadecane and, respectively, standard oil concentration using two fluorescent tracers. In the same time, we measured the cell growth by spectrophotometric analysis and colorimetric method. Cytometry analysis, using propidium iodide and acridine orange helped us detect the increasing metabolic activity and cell density of the bacterial strain cultivated in MSM with various concentrations of n-hexadecane and standard-oil (1%, 2%, 4%, 6%, 15%, v/v) over 36 days. The resulted plots showed that the maximum cellular growth was on 1% n-hexadecane and 6% standard oil, underlined by the highest metabolic activity registered. Cytometry analysis indicated that cell viability and membrane integrity was not damaged by several hydrocarbons’ concentrations. Furthermore, our strain tolerated high concentration of pollutant, like 15% n-hexadecane and standard-oil. Spectrophotometric analysis and colorimetric assay using DCPIP (2,6dichlorophenol indophenol) confirmed cytometry results. All the applied methods showed that the bacterial cells needed a 4 days period of time in order to activate the transcriptional regulation of metabolic operons involved in the biodegradation of the pollutant chemical. Our results suggest the opportunity of using flow cytometry for metabolic activity research of pure strains isolated from polluted environments for xenobiotic pathways.

© Authors. Published Published by byElsevier ElsevierB.V. B.V. This is an open access article under the CC BY-NC-ND license © 2015 2015 The The Authors. (http://creativecommons.org/licenses/by-nc-nd/4.0/). Peer-review under responsibility of the University of Agronomic Sciences and Veterinary Medicine Bucharest. Peer-review under responsibility of the University of Agronomic Sciences and Veterinary Medicine Bucharest Keywords: flow cytometry, biodegradation, n-hexadecane, standard-oil, P. fluorescens

* Corresponding author. Tel.: +4021.311.80.77; Fax: +4021.311.80.77 E-mail address: [email protected] E-mail address: [email protected]

2210-7843 © 2015 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/). Peer-review under responsibility of the University of Agronomic Sciences and Veterinary Medicine Bucharest doi:10.1016/j.aaspro.2015.08.088

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1. Introduction During the last years scientists focused on developing and optimization of new techniques to characterize metabolic changes in bacterial cells exposed to various environmental conditions, including various pollutants (Nielsen et al., 2009). One major pollutant is represented by the petroleum hydrocarbons that cause disturbances in the ecological balance/environmental damage. Use of microorganisms is one important procedure able to remove pollutants from the environment (Das and Chandran, 2011). Several studies have reported various bacterial species with metabolic capabilities for pollutant degradation, including Pseudomonas sp. (Kaczorek, 2011; Zhong, 2014). Characterization of the microorganisms with biodegradative capacities is a mandatory step in designing optimised methods for decontamination. There are current attempts to use flow cytometry in studying the structural and functional features of microorganisms in different environmental samples (Bressan, 2015; De Roy, 2012). Flow cytometry techniques are able to characterize near-real time number, size and biochemical characteristics of cells. Several studies highlighted the usefulness of this method to monitor the dynamics of microbial subpopulations at different physiological states (Matos and daSilva, 2013). This is a reliable method, mainly used for determination of viable cells number and also for morphological and physiological characterization of cells (cell size, internal complexity of the cells as a measure of metabolic activity, cell membrane integrity as a label for viable cells) (Diaz, 2010; Pianetti et al, 2005). In the present study, we investigate the effects of various concentrations (1%, 2%, 4%, 6%, 15%, v/v) of nhexadecane and, respectively, standard-oil, on metabolic activity and cell viability for Pseudomonas fluorescens S7A using a flow-cytometry-based technique. We also estimated the cellular multiplication rate using a colorimetric assay based on 2,6-dichlorophenol indophenol (DCPIP) that becomes colourless in reduced state resulted from the metabolic use of substrates, in our case hydrocarbons. 2. Research Methods Microbial strains Bacterial strain analysed in this study was Pseudomonas fluorescens S7A previously isolated from oil polluted soil (site Berca – Buzau); taxonomic identification, as well as phenotypic and biochemical characterization was performed as described by Ionescu et al. (2013). Culture experiments P. fluorescens S7A was grown in liquid Luria-Bertani media (peptone 10g, yeast extract 5g, NaCl 10g, agar 20g, pH 7,5) for 18h, 120rpm, at 28°C. Cells were washed twice with Bushnell-Haas media (BH) (MgSO4 0,2g, CaCl2 0,02g, KH2PO4 1,0g, K2HPO4 1,0g, NH4NO3 1,0g, FeCl3 0,05g, per litre, pH 7,0). An inoculum of OD600= 1 was added to 100ml BH media supplemented with n-hexadecane (Promega) and, respectively, standard-oil (Fluka) in various concentrations (1%, 2%, 4%, 6%, 15%, (v/v) as sole carbon sources. Flasks (500ml) were incubated at 28°C, 120rpm, for 36 days. All experiments were performed in duplicates. Two control flasks were used, one without cells and the other without carbon source. Cell multiplication was spectrophotometrically monitored (Ultrospec 3000), periodically (days 1, 2, 4, 8, 14, 23, 30 and 36) using OD600. Optical density readings were correlated with the estimation of viable cells -colony- forming units (CFU/ml). Flow cytometry Flow cytometry analysis was performed with a Facs Calibur (Becton Dickinson) equipped with 488nm excitation from an argon-ion laser at 15mW. Cells from 1ml cultures were harvested by centrifugation for 5min at 10.000rpm and washed twice with 500μl distilled water. Bacterial cells were resuspended in 0,5ml distilled water with 10μM acridine orange, and respectively, propidium iodide (BD Bioscience). Samples were vigorously vortexed and incubated at room temperature for 30 min in the dark. To estimate the metabolic activity we analyzed cell size (FSC), granularity (SSC) and ratio FL1/FL3. Data were processed with the software WinMDI version 2.8. Cell growth and degradation capacity of strain P. fluorescens S7A was checked by a colorimetric and rapid test using 2,6-dichlorophenol indophenol (DCPIP). According to Varjani, the level of discoloration of DCPIP is directly proportional with the hydrocarbon metabolism (Varjani et al., 2013). Bacterial inoculum was prepared as previously described and was added in 1% concentration into tubes (30ml) containing liquid BH medium (7,5ml), DCPIP 20mg/ml and n-hexadecane/ standard-oil 1%, 2%, 4%, 6%, 15% (v/v). Duplicates samples were processed and analyzed in similar conditions (28°C, 120rpm, in the dark). Controls without cells or carbon source were also performed. We observed the change of colour after 20 days.

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3. Results and Discussion In order to evaluate the cellular multiplication and viability at various concentrations (1%, 2%, 4%, 6% and 15%, v/v) of n-hexadecane, respectively, oil, we used three different techniques – OD600 (cell number expressed as optical density at 600nm), CFU (viable cell number expressed as Colony Forming Units on solid media) and flow cytometry. 3.1. Our results obtained by estimating viable cells (CFU/ml) showed that strain P. fluorescens S7A undergoes a much longer lag time on n-hexadecane (8 days) than on standard oil (4days) (Figure 1). On the other hand, P. fluorescens S7A reached the maximum growth rate on the same day for both hydrocarbon systems, e.g. the 23rd day. Our data also showed that after a longer adaptation time on n-hexadecane, this bacterial strain multiplied much more efficient on this linear hydrocarbon (14 x 109 cells/ml) than on standard oil (6 x 109cells/ml). In both cases, cell growth was completely inhibited at high hydrocarbon concentrations (15%), probably due to toxic effects on the general physiological state of the cells. These results were expected, as the bacterial strain P. fluorescens S7A was previously isolated from oil-polluted soil samples, so it was not already specialized to rapidly use a single linear hydrocarbon as carbon source and it needed a longer adaptation time for the metabolic switch. After this lag time, cell metabolism proved to be more efficient on n-hexadecane as sole carbon source supporting a double multiplication rate. Moreover, based on our data we could conclude that this strain preferentially uses linear hydrocarbons, as it needed 6% standard oil concentration (standard oil is a mixture that includes linear hydrocarbons) to reach a maximum growth rate, compared to the 1% pure n-hexadecane that was sufficient.

A

B Figure 1. Cell growth of Pseudomonas fluorescens S7A on n-hexadecane (A) and, respectively, standard oil (B), detected by plate counting (CFU/ml)

3.2. Cell growth was also measured by optical density at 600nm (Figure 2). Results were similar to CFU, curves followed the same profile, but absolute values were higher than CFU as they quantify both living and dead bacterial cells, the latter still having integral cell walls.

A

B

Figure 2. Cell growth of Pseudomonas fluorescens S7A on n-hexadecane (A) and, respectively, standard oil (B), detected by OD600

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Compared to other reports in which the highest tolerated concentrations were 10% for n-hexadecane and, respectively, 2% for diesel oil (Partovinia, 2010; Zhang, 2011), in our experiments P. fluorescens S7A tolerated 15% n-hexadecane and standard oil. 3.3. Flow cytometry analysis During the last decade flow-cytometry has been adapted for microbial biodegradation studies (Satyanarayana et al., 2012). In our experiments we used propidium iodide and acridine orange to investigate the dynamics of the metabolic state of the P. fluorescens S7A cells exposed to various concentrations of n-hexadecane and, respectively, standard oil. Our results show that on 1%, 2%, 4% and, respectively, 6% n-hexadecane as sole carbon source for 30 days, the ratio of metabolically active cells increases in the microbial population, e.g. both relevant parameters - cell size (determined as FSC - Forward Angle Light Scatter) and cell complexity (determined as SSC - Right Angle Light Scatter) – (Figure 3). 15 days

30 days

n-hexadecane 6%

n-hexadecane 4%

n-hexadecane 2%

n-hexadecane 1%

2 days

Figure 3. Cell size (FSC) and internal cell complexity (SSC) of P. fluorescens S7A grown on various concentrations of n-hexadecane as sole carbon source for 30 days, determined by flow cytometry assay

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Our data shows that the concentration of 1% n-hexadecane is the most suitable for the tested bacterial strain. Thus, the highest proportion of metabolically active cells of P. fluorescens S7A (measured as internal cell complexity – SSC) was recorded on 1% n-hexadecane 30 days of cultivation (Figure 4).

1%

2%

4%

6%

Figure 4. Proportion of metabolically active cells measured as internal cell complexity (SSC) by flow cytometry in P. fluorescens S7A populations grown on n-hexadecane 1%, 2%, 4%, 6% for 30 days.

In contrast, the P. fluorescens S7A cells showed an opposite dynamics on 15% n-hexadecane, thus proving the toxic effect of such high hydrocarbon concentration (Figure 5). Both cytometric analysis and UFC recordings indicated similar results, e.g. 1% n-hexadecane yielded highest growth rate and ratio of metabolically active cells of P. fluorescens S7A. Cytometric analysis of P. fluorescens S7A cells grown on standard oil showed that best results were on 6% concentration. Thus, the highest multiplication rate, as well as the the highest ratio of metabolically active cells was sustained by a 6% concentration of standard oil (Figure 6 and 7).

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15 days

30 days

n-hexadecane 15%

2 days

Figure 5. Cell size (FSC) and internal cell complexity (SSC) of P. fluorescens S7A grown on 15% n-hexadecane as sole carbon source for 30 days, determined by flow cytometry assay

15 days

30 days

Standard oil 6%

Standard oil 4%

Standard oil 2%

Standard oil 1%

2 days

Figure 6. Cell size (FSC) and internal cell complexity (SSC) of P. fluorescens S7A grown on various concentrations of standard oil as sole carbon source for 30 days, determined by flow cytometry assay

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1%

2%

4%

6%

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Figure 7. Proportion of metabolically active cells measured as internal cell complexity (SSC) by flow cytometry in P. fluorescens S7A populations grown on standard oil 1%, 2%, 4%, 6% for 30 days

Flow cytometry experiments also allowed to monitor the dynamics of the population cell viability during cultivation on hydrocarbons as sole carbon source. In this respect we used propidium iodide (PI) that enters only altered-membrane cells, and acridine orange (AO) that enters both live and dead cells. Thus a combination of these two dyes provides a rapid and reliable method for discriminating live and dead bacteria. Using these two compounds, our flow cytometry data showed that cell population increased in whole number, while dead cells maintained at low values, when P. fluorescens S7A was grown on 1% n-hexadecane, while 15% alkane was toxic for the bacterial cells that depicted a high proportion of dead cells. The metabolic activity increased from the 2nd to the 23th day of cultivation (when it reached a maximum) and after 30 days fluorescence intensity detected in the two channels (FL1 and FL3) decreased (Figure 8). Therefore, these results are similar to those based on CFU and, respectively, OD600. In the same manner, flow cytometry experiments proved that 6% standard oil was the most suitable concentration of this hydrocarbon mixture for P. fluorescens S7A. On this concentration of standard oil the microbial population increased in number, while dead cells were fewest (Figure 9).

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30 days

n-hexadecane 15%

n-hexadecane 1%

2 days

(PI) 30 days

n-hexadecane 15%

n-hexadecane 1%

2 days

(AO) Figure 8. Density plots of P. fluorescens S7A grown on n-hexadecane. Propidium iodide (PI); acridine orange (AO)

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30 days

oil 15%

oil 1%

2 days

(PI) 30 days

oil 15%

oil 1%

2 days

(AO) Figure 9. Density plots of P. fluorescens S7A grown in standard-oil. Propidium iodide (PI); acridine orange (AO)

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Both propidium iodide and acridine orange bind nucleic acids in a situs-nonspecific manner, acting as intercalating agents on DNA molecules. When bound to nucleic acids, PI has an emission maximum at 620nm, whereas AO at 525nm. Live cells, both prokaryotic and eukaryotic, have intact membranes and are impermeable to PI that enters only cells with damaged membranes. In contrast, AO enters all cells, live and dead. Thus using both dyes, flow cytometry can make the difference between live and dead cells in a bacterial population (Diaz, 2010; Pieretti, 2012). During last decade, flow cytometry techniques were adapted from eukaryotic cells to prokaryotes, e.g. food microbiology and, respectively, environmental microbiology (Backman, 2004; Czechowska, 2008; Diaz, 2010; Matos, 2013; Percival, 2011; Pieretti, 2012 Bressan, 2015). Moreover, flow cytometry techniques can be used in investigating non-culturable bacteria (Khan et al., 2010), as well as microbial communities in aquatic and soil ecosystems (De Roy, 2012; Bressan, 2015). There are also studies that compared flow cytometry technique with classical microbiology methods, e.g. CFU and OD at 600nm (Pianetti et al, 2005). In our study we addressed a complex analysis in which flow cytometry experiments have brought new and significant data regarding cell population dynamics during bacterial biodegradation of hydrocarbons. Thus, besides fast operating with huge cell numbers, this technique enabled us to monitor metabolically active cells during the biodegradation process and provided valuable insights on optimum hydrocarbon concentrations. 3.4. DCPIP Colorimetric Assay The principle of this technique lies in discoloration redox indicator DCPIP, which in the reduced state (when incorporating electrons during bacterial metabolism of hydrocarbons) changes colour from blue to colourless. The ability of P. fluorescens S7A to grow and biodegrade on various concentrations of n-hexadecane, respectively standard-oil (1%, 2%, 4%, 6%, 15%, v/v) was confirmed by decolourization of the DCPIP indicator. Discoloration of DCPIP was first noticed for samples with standard-oil in concentration 2% and 6% after 4 days and for the other concentration after 20 days. The samples with hexadecane were discoloured after 6 days and completely after 20 days (Figure 10), indicating slow response compared with samples with standard-oil. Our results are similar to those obtained by Kubota et al. (2008) and Thenmozhi et al. (2013), using DCPIP for testing the biodegradation of different pollutants such as alkanes (n-decane, n-hexadecane, n-eicosane, dodecylcyclohexane), aromatic (benzene, toluene, phenol, xylene) and polycyclic hydrocarbons (anthracene, naphtalene). The bioremediation potential of Pseudomonas sp. demonstrated by DCPIP test was confirmed by many studies (Pirôllo, 2008; de Oliveira, 2014; Varjani, 2013). Also, Buþková et al. (2013) used DCPIP for selecting bacterial strains with biodegradative capacity.

A.

C.

B.

D. Figure 10. DCPIP assay of P. fluorescens S7A on n-hexadecane (1%, 2%, 4%, 6%, 15%) (A, B) and, respectively, standard oil (1%, 2%, 4%, 6%, 15%) (C, D), after 0 day (A, C) and 20 days (B, D)

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Our results are consistent with those obtained using previous methods. Furthermore, DCPIP method is a fast, inexpensive and reliable assessment for screening the ability of microorganisms to use hydrocarbons as sole carbon sources. 4. Conclusions Our experiment analysed the biodegradation capacity of P. fluorescens S7A using 3 different approaches, (i) cell multiplication on hydrocarbon as sole carbon source, measured as colony forming units and optical density at 600nm, (ii) dynamics of the ratio of metabolically active cells analysed by flow cytometry, and (iii) biochemical visualization of hydrocarbon biodegradation by DCPIP decolorization assay. All our results confirmed that this bacterial strain has high biodegradation abilities, best proved on 1% n-hexadecane and, respectively, 6% standard oil. Moreover, flow cytometry provides a most effective method to examine the metabolic activity of the pure strains isolated from oil-polluted soils. Acknowledgements This work was supported by the strategic grant POSDRU/159/1.5/S/133391, Project “Doctoral and Post-doctoral programs of excellence for highly qualified human resources training for research in the field of Life sciences, Environment and Earth Science” cofinanced by the European Social Fund within the Sectorial Operational Program Human Resources Development 2007 – 2013. References Backman, A., Maraha, N., Jansson, J.K., 2004. Impact of temperature on the physiological status of a potential bioremediation inoculant, Arthrobacter chlorophenolicus A6. Applied and Environmental Microbiology, 70(5), 2952–2958. Bressan, M., Gattin I. T., Desaire S., Castel L., Gangneu, C., Laval K., 2015. A rapid flow cytometry method to assess bacterial abundance in agricultural soil. Applied Soil Ecology, 88, 60-68. Buþková M., Puškarová A., Chovanová K., Kraková L., Ferianc P., Pangallo D., 2013. A simple strategy for investigating the diversity and hydrocarbon degradation abilities of cultivable bacteria from contaminated soil. World Journal of Microbiology and Biotechnology, 29(6),1085-1098. Czechowska K., Johnson D.R., van der Meer J.R., 2008. Use of flow cytometric methods for single-cell analysis in environmental microbiology. Current Opinion in Microbiology, 11, 205-212. Das N., Chandran P., 2011. Microbial degradation of petroleum hydrocarbon contaminants: an overview. Biotechnology Research International, 2011: 1-13 (http://dx.doi.org/10.4061/2011/94181). de Oliveira N.C., Rodrigues A.A., Alves M.I., Antoniosi Filho N.R., Sadoyama G., Vieira J.D.G., 2014. Endophytic bacteria with potential for bioremediation of petroleum hydrocarbons and derivatives.African Journal of Biotechnology, 11(12), 2977-2984. De Roy K., Clement L., Thas O., Wang Y., Boon N., 2012. Flow cytometry for fast microbial community fingerprinting. Water Research, 46(3), 907-919. Diaz M., Herrero M., Garcia L.A., Quiros C., 2010. Application of flow cytometry to industrial microbial bioprocesses. Biochemical Engineering Journal 48, 385–407. Ionescu R., Tanase A.-M., Vassu T., Pelinescu D., Chiciudean I., Csutak I., Stoica I., 2013, Characterization of Pseudomonas strains with hydrocarbons-degrading potential, Romanian Biotechnological Letters, 18(3), 8372-8380. Kaczorek E., Olszanowski A., 2011. Uptake of Hydrocarbon by Pseudomonas fluorescens (P1) and Pseudomonas putida (K1) strains in the presence of surfactants: a cell surface modification. Water, Air and Soil Pollution, 214(1-4), 451–459. Khan M.M.T., Pyle B.H., Camper A.K., 2010. Specific and rapid enumeration of viable but nonculturable and viable-culturable gram-negative bacteria by using flow cytometry. Applied and Environmental Microbiology, 76(15), 5088-5096. Kubota K., Koma D., Matsumiya Y., Chung S.Y., Kubo M., 2008. Phylogenetic analysis of long-chain hydrocarbon-degrading bacteria and evaluation of their hydrocarbon-degradation by the 2, 6-DCPIP assay. Biodegradation, 19(5), 749-757. Matos C.T., da Silva T.L., 2013. Using multi-parameter flow cytometry as a novel approach for physiological characterization of bacteria in microbial fuel cells.Process Biochemistry, 48(1), 49-57. Nielsen T.H., Sjoholm O.R., Sorensen J., 2009. Multiple physiological states of a Pseudomonas fluorescens DR54 biocontrol inoculant monitored by a new flow cytometry protocol. FEMS Microbiology Ecology, 67(3), 479-490. Partovinia A., Naeimpoor F., Hejazi P., 2010. Carbon content reduction in a model reluctant clayey soil: slurry phase n-hexadecane bioremediation. Journal of Hazardous Materials, 181(1), 133-139. Percival S.L., Slone W., Linton S., Okel T., Corum L., Thomas J.G., 2011. Use of flow cytometry to compare the antimicrobial efficacy of silvercontaining wound dressings against planktonic Staphylococcus aureus and Pseudomonas aeruginosa. Wound Repair and Regeneration, 19, 436–441.

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