Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Pianetti, A. Articles by Papa, S. Search for Related Content PubMed PubMed Citation Articles by Pianetti, A. Articles by Papa, S. Agricola Articles by Pianetti, A. Articles by Papa, S.

 Previous Article  |  Next Article 

Applied and Environmental Microbiology, December 2005, p. 7948-7954, Vol. 71, No. 12
0099-2240/05/$08.00+0     doi:10.1128/AEM.71.12.7948-7954.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

Determination of the Viability of Aeromonas hydrophila in Different Types of Water by Flow Cytometry, and Comparison with Classical Methods

Anna Pianetti,^1* Tania Falcioni,² Francesca Bruscolini,¹ Luigia Sabatini,¹ Elivio Sisti,⁴ and Stefano Papa^2,3

Toxicological, Hygienic and Environmental Science Institute, Urbino University, Urbino,¹ Center of Cytometry and Cytomorphology, Urbino University, Urbino,² Institute of Morphological Science, Urbino University, Urbino,³ Center for Environmental Study, Rimini, Italy⁴

Received 15 April 2005/ Accepted 25 August 2005

Top Abstract Introduction Materials and Methods Results Discussion References

 
The presence of Aeromonas spp. in water can represent a risk for human health. Therefore, it is important to know the physiological status of these bacteria and their survival in the environment. We studied the behavior of a strain of Aeromonas hydrophila in river water, spring water, brackish water, mineral water, and chlorinated drinking water, which had different physical and chemical characteristics. The bacterial content was evaluated by spectrophotometric and plate count techniques. Flow cytometric determination of viability was carried out using a dual-staining technique that enabled us to distinguish viable bacteria from damaged and membrane-compromised bacteria. The traditional methods showed that the bacterial content was variable and dependent on the type of water. The results obtained from the plate count analysis correlated with the absorbance data. In contrast, the flow cytometric analysis results did not correlate with the results obtained by traditional methods; in fact, this technique showed that there were viable cells even when the optical density was low or no longer detectable and there was no plate count value. According to our results, flow cytometry is a suitable method for assessing the viability of bacteria in water samples. Furthermore, it permits fast detection of bacteria that are in a viable but nonculturable state, which are not detectable by conventional methods.

Top Abstract Introduction Materials and Methods Results Discussion References

 
During the last decade scientific interest has turned to members of the genus Aeromonas as human and animal pathogens. Many Aeromonas species have been associated with human diseases, including gastroenteritis, hemolytic-uremic syndrome, septicemia, etc. (21, 30, 48).

Aeromonas spp. are common aquatic microorganisms that occur in seawater, irrigation water, river water, brackish water, freshwater, groundwater, spring water, sewage-contaminated water, and activated sludge (4, 7, 9, 14, 15, 23, 24, 37, 43).

The wide distribution of aeromonads in different aquatic ecosystems underlines their capacity to adapt to environments with different trophic levels. Several studies have shown that the phenospecies Aeromonas hydrophila is prevalent in cleaner water, whereas Aeromonas caviae is prevalent in water with a high level of fecal pollution; the frequencies of the other phenospecies appear to be unrelated to environmental conditions (3, 15, 17, 23).

These bacteria have also been isolated from chlorinated and unchlorinated drinking water and from bottled mineral water, which shows that they are able to withstand long periods of nutrient limitation (6, 16, 19, 20, 29, 31, 46, 50). The survival of aeromonads in these ecosystems can be correlated with their capacity to produce and live within biofilms on the surfaces of pipes and bottles (26); on the other hand, it is known that Aeromonas spp. can cope with nutrient limitation conditions by entering into a starvation survival state (33).

The growing interest in these organisms addresses aquatic environments, because these environments represent an important reservoir of aeromonads. It is assumed that human infections arise from recreational activities, contaminated drinking water, or foods (23).

Therefore, due to the importance of water for survival and diffusion of the microorganisms, a study to evaluate the behavior of A. hydrophila in types of water with different physical and chemical characteristics was performed.

The bacterial content was evaluated by measuring absorbance and by using plate count agar; in addition, a flow cytometric analysis method was also used. Flow cytometry has become a method of choice for bacterial research. It has been used in basic (8, 13, 34, 39), clinical (2), and environmental (12, 47) studies. At present, the use of a flow cytometric technique in conjunction with a fluorescence technique provides a promising method for the detection and rapid enumeration of viable bacteria in environmental samples. Moreover, it provides information on the dynamics and physiological heterogeneity of bacterial populations (22).

Top Abstract Introduction Materials and Methods Results Discussion References

 
Bacteria.
A strain of A. hydrophila that was previously isolated from irrigation water and was identified as described elsewhere (38) was used in this study. This strain was stored at –80°C until it was used.

Water microcosms.
The water microcosms into which A. hydrophila was inoculated contained brackish water, river water, spring water, mineral water, and chlorinated drinking water. The samples of brackish water, river water, and spring water were collected with a "Mare-Lacus" sampler (International PBI, Milan, Italy); the drinking water was collected directly from the tap, and the mineral water was a commercial natural mineral water. Three liters of each type of water was filtered through a 0.22-µm membrane filter (Nalgene; International PBI) to remove autochthonous microbiota.

Inoculation of microcosms.
The A. hydrophila strain was regenerated in 5 ml of tryptone soya broth (Oxoid, Milan, Italy) with 0.6% (wt/vol) yeast extract and 3% (wt/vol) Casamino Acids with overnight incubation at 28°C. One loopful of the culture was streaked on m-Aeromonas selective agar base (Havelaar, Biolife-Italiana, Milan, Italy) and incubated for 18 to 24 h at 28°C. Two or three colonies were inoculated into 40 ml of nutrient broth (Oxoid) and incubated at 28°C for 18 to 24 h with agitation at 150 rpm. The broth cultures were centrifuged at 3,000 rpm for 30 min, and each pellet was resuspended in 10 to 15 ml of the same water used for the experiments. The tests were performed in 1-liter screw-cap Pyrex brown glass bottles. These bottles were cleaned with a 10% solution of K₂Cr₂O₇ in concentrated H₂SO₄ and rinsed in hot tap water, in a 10% HNO₃ solution, and finally in distilled water. Then they were autoclaved at 121°C for 15 min. The A. hydrophila strain was inoculated into 800 ml of each type of water to obtain a bacterial concentration of about 5 x 10⁶ CFU/ml, corresponding to an optical density at 600 nm of 0.07 (as determined with a UNICAM spectrophotometer) (45). The samples were incubated at room temperature (mainly at 22°C; range, 18°C to 25°C).

Control cultures were grown in nutrient broth and handled under the same conditions.

All the experiments were repeated three times, and the data were expressed as the mean for each sample.

Determination of the bacterial content.
The bacterial content was evaluated after inoculation into the water samples and at regular intervals (1 day) over a 30-day period using turbidimetric determination and the plate count method. Plate counting was performed by streaking 1 ml of inoculated water in triplicate from decimal dilutions onto Havelaar agar (Biolife). The plates were incubated at 28°C for at least 24 h, and viable counts were expressed as CFU/ml.

Flow cytometry.
The fluorescence intensity of stained bacteria was measured with a FACSCalibur flow cytometer (Becton Dickinson, Milan, Italy) equipped with an argon laser set at 15 mV and tuned to an excitation wavelength of 488 nm. All data were analyzed statistically with the Cell Quest software. The fluorescent filters and detectors were all standard filters and detectors; green fluorescence was collected in the FL1 channel (525 nm), orange fluorescence was collected in the FL2 channel (585 nm), and red fluorescence was collected in the FL3 channel (>670 nm). All parameters were collected as logarithmic signals. Fluorescent microspheres (Becton Dickinson liquid counting beads) were added to each sample as an internal reference. The solutions used were routinely filtered through Millipore membrane filters (pore size, 0.22 µm; Millipore, Bedford, Mass.) before passage through the flow cytometer. Debris was excluded from analysis by scatter gating. For the flow cytometric analysis, 5 x 10⁴ events were collected, and three independent experiments were carried out each time.

Viability staining with flow cytometry.
Cell membrane integrity was assessed by using the nucleic acid double-staining procedure of Barbesti et al. (5) based on the permanent SYBR Green I (Molecular Probes) and impermanent propidium iodide (PI) (Molecular Probes) fluorescent probes. Both of these probes stain RNA and DNA (18).

Five milliliters of each sample was centrifuged at 3,000 rpm for 10 min and resuspended in 5 ml of filtered phosphate-buffered saline. One milliliter of the bacterial suspension was stained with 1:10,000 (vol/vol) SYBR Green I and a 10-µg/ml commercial PI solution (5) and incubated for 15 min in the dark at room temperature. Each cell was characterized by two fluorescence parameters that measured green fluorescence emission (FL1) and orange-red fluorescence emission (FL3).

The system permitted us to distinguish cells with intact membranes (live cells) (green), cells with damaged membranes (green plus orange-red), and cells with compromised membranes (orange-red; dead). Thus, the system was able to place the bacterial cells in three different cytometric clusters. The probes were tested using exponential-phase bacterial cells as a positive control and bacteria that were heat killed at 70°C as a negative control.

Statistical analysis.
Analysis of variance applied to the regression model was used for statistical analysis for the correlation between the absorbance and CFU/ml. Cytometric data were analyzed using the Cell Quest software (Becton Dickinson); percentages of cells were determined by gating processing on acquired data files.

Data were acquired in triplicate for each test sample, and means, calculated with the Cell Quest software, were used for analysis of statistical significance by a Student paired t test.

Physical and chemical parameters.
The physical and chemical parameters of the water samples that were determined before bacterial inoculation were conductivity at 20°C, total hardness, total alkalinity, pH, and the concentrations of chlorine, sulfates, phosphates, nitrates, ammonia, calcium, iron, silver, copper, magnesium, and total organic carbon (TOC). The concentrations of nitrates, ammonia, phosphates, sulfates, and TOC and the pH were also monitored at the end of the experiments in order to evaluate possible changes following bacterial growth. The tests were performed by APAT IRSA-CNR methods (1).

Top Abstract Introduction Materials and Methods Results Discussion References

 
As shown in Fig. 1, the optical densities of the brackish water increased slightly (from 0.07 to 0.095) in the first 24 h and then gradually decreased to 0.022 in the last 4 days of the trial. A similar curve, but without an initial increase, was observed for the mineral water; in this case the optical density fell to 0.005 on day 30.

View larger version (18K): [in this window] [in a new window]

FIG. 1. Survival curves for A. hydrophila inoculated into different types of water as detected by measurement of absorbance.

In the other types of water marked decreases in the optical density were observed in the first 6 to 10 days (from 0.07 to 0.008 to 0.009), and the optical density was 0 on day 27 for the river water and on day 29 for the spring water and the drinking water.

The survival curves, as determined by the standard plate count method, exhibited a similar behavior for the brackish water and mineral water until day 23; on day 30 the brackish water contained log 5.84 CFU/ml, and the mineral water contained log 0.69 CFU/ml. The lowest values were obtained for the drinking water; for this water no growth was found on day 23 or thereafter (Fig. 2).

View larger version (10K): [in this window] [in a new window]

FIG. 2. Survival curves for A. hydrophila inoculated into different types of water as detected by plate counting.

There was a correlation between the absorbance values and the plate count values (P < 0.001; R² = 0.096 for river water; R² = 0.097 for brackish water; R² = 0.099 for the other types of water; R² = 0.098 for the control culture).

Cytofluorimetric analysis showed that the highest percentages of viable bacteria were generally found in drinking water, and the values ranged from 47 to 74% throughout the experiment. In contrast, the lowest values were observed in the mineral water, in which the percentages of viable cells were 10% from day 12 onward. In the other types of water the values were still around 50% on day 30 (Fig. 3).

View larger version (35K): [in this window] [in a new window]

FIG. 3. Viable A. hydrophila evaluated by flow cytometry, as revealed in the FL1 channel. N. broth, nutrient broth.

The percentages of damaged cells did not change much during the experiment; damaged cells were prevalent in the mineral water, and the percentages ranged from 9 to 30% during the experiment (Fig. 4 and 5).

View larger version (24K): [in this window] [in a new window]

FIG. 4. Damaged A. hydrophila evaluated by flow cytometry, as revealed in the FL1 and FL3 channels. N. broth, nutrient broth.

View larger version (47K): [in this window] [in a new window]

FIG. 5. Flow cytometric analysis of fractions of damaged and membrane-compromised cells in nutrient broth (a) and fractions of viable, damaged, and membrane-compromised cells in mineral water (b), drinking water (c), river water (d), spring water, (e) and brackish water (f).

In the control culture in nutrient broth, after increases in the optical density and CFU/ml on day 3 (1.72 and log 9.4 CFU/ml, respectively), there were gradual decreases, and the optical density and number of CFU/ml were 0.042 and log 8.4 CFU/ml at the end of the trial (Fig. 6). In contrast, cytofluorimetric analysis showed that there were low percentages of viable cells and high proportions of damaged cells beginning on day 3 (Fig. 3, 4, and 5).

View larger version (11K): [in this window] [in a new window]

FIG. 6. Survival curves for A. hydrophila inoculated into nutrient broth as detected by plate counting and measurement of absorbance. OD, optical density.

In drinking water on day 23 of incubation, there were prominent cytometric clusters of live cells (74%), even though there was no plate growth during the same period (Fig. 2).

The initial analysis of physical and chemical parameters (Table 1) showed that the highest values for electrical conductivity, total hardness, and the concentrations of chlorine, sulfates, calcium, magnesium, and TOC were the values for the brackish water. At the end of the experiments (Table 2) the nitrate concentrations were lower in all types of water, while the concentrations of ammonia, phosphates, sulfates, and TOC were higher.

View this table: [in this window] [in a new window]

TABLE 1. Physical and chemical parameters before the bacterial inoculum was added

View this table: [in this window] [in a new window]

TABLE 2. Physical and chemical parameters before the bacterial inoculum was added and at the end of the experiments

Top Abstract Introduction Materials and Methods Results Discussion References

 
The aim of this study was to evaluate the capacity of a strain of A. hydrophila inoculated into different types of water to survive by using conventional methods of detection, such as absorbance and plate counting, and to compare these methods with the flow cytometric technique.

The data from the spectrophotometric analysis showed that the bacterial content was variable and depended on the type of water. In fact, when the bacterial strain was inoculated into brackish water, the optical densities were higher throughout the trial and were still 0.022 on day 30. In mineral water the bacteria behaved similarly; in contrast, in the river water, spring water, and drinking water the optical densities decreased after 8 to 10 days of incubation, and values could not be determined on the last days of the experiment. In some cases, including mineral water and drinking water, cytofluorimetric data were different from spectrophotometric data. In fact, from 12 day of incubation, the percentages of live bacteria in drinking water (20%) were less than the values for the other types of water, while the optical densities were higher than those obtained for the other types of water (Fig. 1 and 3).

Estimating microbial growth by measurement of absorbance has the advantage of being rapid, nondestructive, inexpensive, and relatively easy to automate. However, absorbance is an indirect measure of bacterial behavior; it depends on the total number of cells present in the sample and does not allow workers to distinguish between live and dead cells. Furthermore, the sensitivity of this method appears to be correlated with microbiological culture density (10, 11).

When bacteria are monitored, it is very important to determine the viability of cells in addition to their abundance, since this is a critical issue in assessing water quality and preventing sanitation problems (41).

The plate count method is a widely used technique for determining the bacterial charge, but it supplies information related only to viability and growth capacity. In this study the results obtained from the plate count analysis statistically correlated with the absorbance data. However, the turbidimetric method for determining bacterial concentrations appeared to be less sensitive than the plate count method. In fact, in the river water and spring water a number of viable cells were detected when the optical density was no longer detectable. These findings agree with the results of other studies (11) in which the workers reported that the usefulness of turbidimetry was restricted to conditions under which high cell densities were reached. In such a case a more sensitive method, such as plate counting, might yield higher estimates. In any case, standard plate counting also has major disadvantages because it requires long incubation times and for many bacterial species there are not efficient growth media.

Recent investigations (35, 40) have demonstrated that many nondifferentiating bacteria, including Aeromonas spp., can respond to a number of environmental stresses by entrance into a viable but nonculturable state. Such forms cannot be detected by standard plating techniques, and their detection requires special resuscitation procedures (27, 28, 49). As a consequence, the routine plating method could underestimate the real bacterial presence, which could be a public health concern, especially if the cells maintain their virulence in the viable but nonculturable state (36).

New approaches have been developed to assess bacterial viability. Flow cytometry is a powerful tool for analyzing a cell population at the single-cell level, since it can be used both to identify and enumerate bacterial populations from environmental samples and to characterize functional properties of an individual cell (18, 42).

The flow cytometric nucleic acid double-staining protocol is based on simultaneous use of permeable fluorescent probes (SYBR Green dyes) (25) and an impermeable fluorescent probe (PI) and can distinguish viable, membrane-damaged, and membrane-compromised cells.

The efficiency of combined staining is magnified by the energy transfer from SYBR Green I to PI when both probes are bound to nucleic acids, as described by Barbesti et al. (5)

In this study flow cytometric analysis revealed viable A. hydrophila cells even when the optical densities were low or not detectable and there was no plate growth (e.g., drinking water) (Fig. 2).

For the control culture in nutrient broth, the optical densities were higher than those found in the water samples tested. At the same time cytofluorimetric analysis revealed the lowest viability and the highest percentage of damaged cells. In the control culture the live cell clusters tended to disappear, while the damaged clusters persisted. The percentages of live cells were high in all types of water, ranging from 50% to 80%, and the live cell clusters were well defined during all of the trials with the exception of the mineral water and control culture trials (Fig. 5). This behavior could have been due to excess nutrients that have inhibitory effects on microbial growth, especially when the microorganisms are oligotrophic (44).

The apparent discordance between the plate count data and the cytofluorimetric data requires further investigation. The analysis of subpopulations (sorting) performed by flow cytometry allows us to quantify and characterize cells with different physiological parameters, such as enzyme activity, respiration, membrane potential, intracellular pH, and membrane integrity; this could provide an explanation for the behavior of the bacteria when they are inoculated into media.

No significant correlation was found between survival and any single physical or chemical parameter for the brackish water, spring water, and drinking water; in any case, high percentages of viable cells usually corresponded to high values for at least one of the parameters tested. The mineral water was characterized by the absence of iron and phosphates, which are essential for bacterial growth (32). This fact, together with the low organic carbon concentration, could have been responsible for the reduced viability percentages that were found from day 12 onward. An interesting result is the increases in the concentrations of ammonia, phosphates, sulfates, and TOC observed at the end of the experiments. The increase in the ammonia content could have been due to anaerobic respiration, in which nitrates were utilized as an electron acceptor. On the other hand, a suffering state of the bacteria with dead cells and the resulting release of cellular components into the surrounding environment could explain the increases in the other parameters. Thus, the bacterial presence in bottled water could contribute to the changes in chemical parameters during storage.

Our data are preliminary, and further studies should be performed to better define the factors that can affect bacterial survival, such as storage temperature and interference by autochthonous flora.

According to our results, cytofluorimetric analysis had several advantages. SYBR Green I coupled with propidium iodide allowed us to assess the bacterial viability in water; the system provided rapid direct counting of high numbers of cells; and the system allowed us to directly analyze water samples regardless of cultivation, and therefore sample handling and possible contamination due to sample handling were reduced. Thus, the original characteristics of samples were likely to be analyzed.

Furthermore, the independence from cultivation makes flow cytometry an appealing technique for fast detection of the viability of pathogenic bacteria in the viable but nonculturable state, which can be present in the aquatic environment but not detectable by conventional methods.

In conclusion, flow cytometric analysis appears to be a valid tool for assessment of bacterial viability and for better microbiological characterization of water in order to control and prevent health risks.

 
This work was supported by "CIPE" research project 36-2002 of the Marche Region of Italy.

 
* Corresponding author. Mailing address: Toxicological, Hygienic and Environmental Science Institute, Via S. Chiara, 27, University "Carlo BO," 61029 Urbino, Italy. Phone: 39 722 303544. Fax: 39 722 303541. E-mail: pianetti{at}uniurb.it.

Top Abstract Introduction Materials and Methods Results Discussion References

 

1
1. Agenzia per la Protezione dell'Ambiente e per i Servizi Tecnici (APAT), Istituto di Ricerca sulle Acque del Consiglio Nazionale delle Ricerche (IRSA-CNR). 2003. Manuali e linee guida 29/2003. Agenzia per la Protezione dell'Ambiente e per i Servizi Tecnici (APAT), Istituto di Ricerca sulle Acque del Consiglio Nazionale delle Ricerche (IRSA-CNR), Rome, Italy.
2
2. Alvarez-Barrientos, A. M., J. Arroyo, R. Cantón, C. Nombela, and M. Sánchez-Pérez. 2000. Applications of flow cytometry to clinical microbiology. Clin. Microbiol. Rev. 13:167-195.[Abstract/Free Full Text]
3
3. Araujo, R. M., R. M. Arribas, F. Lucena, and R. Pares. 1989. Relation between Aeromonas and faecal coliforms in fresh waters. J. Appl. Bacteriol. 76:213-217.
4
4. Ashbolt, N. J., A. Ball, M. Dorsch, C. Turner, P. Cox, A. Chapman, and S. M. Kirov. 1995. The identification and human health significance of environmental aeromonads. Water Sci. Technol. 31:263-269.
5
5. Barbesti, S., S. Citterio, M. Labra, M. D. Baroni, M. G. Neri, and S. Sgorbati. 2000. Two- and three-color fluorescence flow cytometric analysis of immunoidentified viable bacteria. Cytometry 40:214-218.[CrossRef][Medline]
6
6. Biscardi, D., A. Castaldo, O. Gualillo, and R. de Fusco. 2002. The occurrence of cytotoxic Aeromonas hydrophila strains in Italian mineral and thermal waters. Sci. Total Environ. 292:255-263.[CrossRef][Medline]
7
7. Borrel, N., M. J. Figueras, and J. Guarro. 1998. Phenotypic identification of Aeromonas genospecies from clinical and environmental sources. Can. J. Microbiol. 44:103-108.[CrossRef][Medline]
8
8. Bouix, M., and J.-Y. Leveau. 2001. The applications of flow cytometry in microbiology. Bull. Soc. Fr. Microbiol. 16:210-218.
9
9. Boussaid, A., B. Baleux, L. Hassani, and J. Lesne. 1991. Aeromonas species in stabilization ponds in the arid region of Marrakesh, Morocco, and relation to fecal pollution and climatic factors. Microb. Ecol. 21:11-20.[CrossRef]
10
10. Dalgaard, P., T. Ross, L. Kamperman, K. Neumeyer, and T. A. McMeekin. 1994. Estimation of bacterial growth rates from turbidimetric and viable count data. Int. J. Food Microbiol. 23:391-404.[CrossRef][Medline]
11
11. Dalgaard, P., and K. Koutsoumanis. 2001. Comparison of maximum specific growth rates and lag times estimated from absorbance and viable count data by different mathematical models. J. Microbiol. Methods 43:183-196.[CrossRef][Medline]
12
12. Davey, H. M., and D. B. Kell. 1996. Flow cytometry and cell sorting of heterogeneous microbial populations: the importance of single-cell analysis. Microbiol. Rev. 60:641-696.[Abstract/Free Full Text]
13
13. Davey, H. M., D. H. Welchart, D. B. Douglas, and A. S. Kaprelyants. 1999. Estimation of microbial viability using flow cytometry, p. 11.3.11-11.3.20. In Current protocols in cytometry. Wiley, New York, N.Y.
14
14. Farmer, J. J., III, M. J. Arduino, and F. W. Hickman-Brenner. 1992. The genera Aeromonas and Plesiomonas p. 3012-3028. In A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (ed.), The procaryotes, 2nd ed. Springer-Verlag, New York, N.Y.
15
15. Fiorentini, C., E. Barbieri, L. Falzano, P. Matarrese, W. Baffone, A. Pianetti, M. Katouli, I. Kühn, R. Möllby, F. Bruscolini, A. Casiere, and G. Donelli. 1998. Occurrence, diversity and pathogenicity of mesophilic Aeromonas in estuarine waters of the Italian coast of the Adriatic sea. J. Appl. Microbiol. 85:501-511.[CrossRef][Medline]
16
16. Graviel, A. A., J. P. Landre, and A. J. Lamb. 1998. Incidence of mesophilic Aeromonas within a public drinking water supply in north-east Scotland. J. Appl. Microbiol. 84:383-392.[CrossRef][Medline]
17
17. Hanninen, M. L., and A. Siitonen. 1995. Distribution of Aeromonas phenospecies among strains isolated from water, foods or from human clinical samples. Epidemiol. Infect. 115:39-50.[Medline]
18
18. Haugland, R. P. 1998. Handbook of fluorescent probes and research chemicals, p. 680. Molecular Probes, Inc., Eugene, Oreg.
19
19. Havelaar, A. H., F. M. Schets, A. van Silfhout, W. H. Jansen, G. Wieten, and D. van der Kooij. 1992. Typing of Aeromonas strains from patients with diarrhoea and from drinking-water. J. Appl. Bacteriol. 72:435-444.[Medline]
20
20. Ivanova, E. P., N. V. Zhukova, N. M. Gorshkova, and E. L. Chaikina. 2001. Characterization of Aeromonas and Vibrio species isolated from a drinking water reservoir. J. Appl. Microbiol. 90:919-927.[CrossRef][Medline]
21
21. Janda, J. M., and S. L. Abbott. 1996. Human pathogens, p. 151-173. In B. Austin et al. (ed.), The genus Aeromonas. Wiley, London, United Kingdom.
22
22. Kaouther, B. A., P. Breeuwer, P. Verbaarschot, F. M. Rombouts, A. D. L. Akkermans, W. M. De Vos, and T. Abee. 2002. Multiparametric flow cytometry and cell sorting for the assessment of viable, injured, and dead Bifidobacterium cells during bile salt stress. Appl. Environ. Microbiol. 68:5209-5216.[Abstract/Free Full Text]
23
23. Kirov, S. M. 1997. Aeromonas and Plesiomonas species, p. 265-287. In M. P. Doyle, L. R. Beuchat, and T. J. Montville (ed.), Food microbiology. Fundamentals and frontiers. AMS Press, Washington, D.C
24
24. Krovacek, K., V. Pasquale, S. B. Baloda, V. Soprano, M. Conte, and S. Dumontet. 1994. Comparison of putative virulence factors in Aeromonas hydrophila strains isolated from the marine environment and human diarrheal cases in southern Italy. Appl. Environ. Microbiol. 60:1379-1382.[Abstract/Free Full Text]
25
25. Lebaron, P., N. Parthuisot, and P. Catala. 1998. Comparison of blue nucleic acid dyes for flow cytometric enumeration of bacteria in aquatic systems. Appl. Environ. Microbiol. 64:1725-1730.[Abstract/Free Full Text]
26
26. Leclerc, H., and A. Moreau. 2002. Microbial safety of natural mineral water. FEMS Microbiol. Rev. 26:207-222.[CrossRef][Medline]
27
27. Maalej, S., M. Denis, and S. Dukan. 2004. Temperature and growth-phase effects on Aeromonas hydrophila survival in natural seawater microcosms: role of protein synthesis and nucleic acid content on viable but temporarily nonculturable response. Microbiology 150:181-187.[Abstract/Free Full Text]
28
28. Mary, P., N. E. Chihib, O. Charafeddine, C. Defives, and J. P. Hornez. 2002. Starvation survival and viable but nonculturable states in Aeromonas hydrophila. Microb. Ecol. 43:250-258.[CrossRef][Medline]
29
29. Massa, S., C. Altieri, and A. D'Angela. 2001. The occurrence of Aeromonas spp. in mineral water and well water. Int. J. Food Microbiol. 63:169-173.[CrossRef][Medline]
30
30. Merino, S., X. Rubiers, S. Knøchel, and J. M. Tomàs. 1995. Emerging pathogens: Aeromonas spp. Int. J. Food Microbiol. 28:157-168.[CrossRef][Medline]
31
31. Messi, P., E. Guerrieri, and M. Bondi. 2002. Survival of an Aeromonas hydrophila in an artificial mineral water microcosm. Water Res. 36:3410-3415.[Medline]
32
32. Miettinen, I. T., T. Vartiainen, and P. J. Martikainen. 1997. Phosphorus and bacterial growth in drinking water. Appl. Environ. Microbiol. 63:3242-3245.[Abstract]
33
33. Morita, R. Y. 1993. Bioavailability of energy and the starvation state, p. 1-23. In S. Kjellerberg (ed.), Starvation in bacteria. Plenum Press, New York, N.Y.
34
34. Nebe-von-Caron, G., P. J. Stephens, C. J. Hewitt, J. R. Powell, and R. A. Badley. 2000. Analysis of bacterial function by multi-colour fluorescence flow cytometry and single cell sorting. J. Microbiol. Methods 42:97-114.[CrossRef][Medline]
35
35. Oliver, J. D. 1993. Formation of viable but nonculturable cells, p. 239-272. In S. Kjellerberg (ed.), Starvation in bacteria. Plenum Press, New York, N.Y.
36
36. Oliver, J. D. 2000. The public health significance of viable but nonculturable bacteria, p. 277-300. In R. R. Colwell and D. J. Grimes (ed.), Nonculturable microrganisms in the environment. ASM Press, Washington, DC.
37
37. Pianetti, A., W. Baffone, F. Bruscolini, E. Barbieri, M. R. Biffi, L. Salvaggio, and A. Albano. 1998. Presence of several pathogenic bacteria in the Metauro and Foglia rivers (Pesaro-Urbino, Italy). Water Res. 32:1515-1521.[CrossRef]
38
38. Pianetti, A., L. Sabatini, F. Bruscolini, F. Chiaverini, and G. Cecchetti. 2004. Faecal contamination indicators, Salmonella, Vibrio and Aeromonas in water used for the irrigation of agricultural products. Epidemiol. Infect. 132:231-238.[CrossRef][Medline]
39
39. Porter, J., D. Deere, R. Pickup, and C. Edwards. 1996. Fluorescent probes and flow cytometry: new insights into environmental bacteriology. Cytometry 23:91-96.[CrossRef][Medline]
40
40. Roszak, D. B., and R. R. Colwell. 1987. Survival strategies of bacteria in the natural environment. Microbiol. Rev. 51:365-379.[Free Full Text]
41
41. Sgorbati, S., S. Barbesti, M. G. Neri, C. Davegna, and S. Citterio. 1997. Rapid flow cytometric immunodetection of bacteria to monitor aquatic environments. Eur. J. Histochem. 41:187-188.
42
42. Shapiro, H. M. 2000. Microbial analysis at the single-cell level: tasks and techniques. J. Microbiol. Methods 42:3-16.[CrossRef][Medline]
43
43. Soler, L., M. J. Figueras, M. R. Chacon, J. Vila, F. Marco, A. J. Martinez-Murcia, and J. Guarro. 2002. Potential virulence and antimicrobial susceptibility of Aeromonas popoffii recovered from freshwater and seawater. FEMS Immunol. Med. Microbiol. 32:243-247.[CrossRef][Medline]
44
44. Thorne, S. H., and H. D. Williams. 1999. Cell density-dependent starvation survival of Rhizobium leguminosarum bv. phaseoli: identification of the role of an N-acyl homoserine lactone in adaptation to stationary-phase survival. J. Bacteriol. 181:981-990.[Abstract/Free Full Text]
45
45. Thornley, J. P., J. G. Shaw, I. A. Gryllos, and A. Eley. 1996. Adherence of Aeromonas caviae to human cell lines Hep-2 and Caco-2. J. Med. Microbiol. 45:445-451.[Abstract/Free Full Text]
46
46. Tsai, G. J., and S. C. Yu. 1997. Microbiological evaluation of bottled uncarbonated mineral water in Taiwan. Int. J. Food Microbiol. 37:137-143.[CrossRef][Medline]
47
47. Vives-Riego, J., P. Lebaron, and G. Nebe-von-Caron. 2000. Current and future applications of flow cytometry in aquatic microbiology. FEMS Microbiol. Rev. 24:429-448.[CrossRef][Medline]
48
48. Wadstrom, T., and A. Ljungh. 1991. Aeromonas and Plesiomonas as food and waterborne pathogens. Int. J. Food Microbiol. 12:303-312.[CrossRef][Medline]
49
49. Wai, S. N., Y. Mizunoe, and A. Takade. 2000. A comparison of solid and liquid media for resuscitation of starvation- and low-temperature-induced nonculturable cells of Aeromonas hydrophila. Arch. Microbiol. 173:307-310.[CrossRef][Medline]
50
50. Warburton, D. W., J. K. McCormick, and B. Bowen. 1994. Survival and recovery of Aeromonas hydrophila in water: development of methodology for testing bottled water in Canada. Can. J. Microbiol. 40:145-148.[Medline]

---

Applied and Environmental Microbiology, December 2005, p. 7948-7954, Vol. 71, No. 12
0099-2240/05/$08.00+0     doi:10.1128/AEM.71.12.7948-7954.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

This article has been cited by other articles:

• Falcioni, T., Papa, S., Gasol, J. M. (2008). Evaluating the Flow-Cytometric Nucleic Acid Double-Staining Protocol in Realistic Situations of Planktonic Bacterial Death. Appl. Environ. Microbiol. 74: 1767-1779 [Abstract] [Full Text]  
• Quiros, C., Herrero, M., Garcia, L. A., Diaz, M. (2007). Application of Flow Cytometry to Segregated Kinetic Modeling Based on the Physiological States of Microorganisms. Appl. Environ. Microbiol. 73: 3993-4000 [Abstract] [Full Text]  

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Pianetti, A. Articles by Papa, S. Search for Related Content PubMed PubMed Citation Articles by Pianetti, A. Articles by Papa, S. Agricola Articles by Pianetti, A. Articles by Papa, S.