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Appl Environ Microbiol. Oct 2008; 74(19): 5925–5933.
Published online Aug 8, 2008. doi:  10.1128/AEM.00318-08
PMCID: PMC2565981

DNA Fragmentation in Microorganisms Assessed In Situ[down-pointing small open triangle]

Abstract

Chromosomal DNA fragmentation may be a direct or indirect outcome of cell death. Unlike DNA fragmentation in higher eukaryotic cells, DNA fragmentation in microorganisms is rarely studied. We report an adaptation of a diffusion-based assay, developed as a kit, which allows for simple and rapid discrimination of bacteria with fragmented DNA. Intact cells were embedded in an agarose microgel on a slide, incubated in a lysis buffer to partially remove the cell walls, membranes, and proteins, and then stained with a DNA fluorochrome, SYBR Gold. Identifying cells with fragmented DNA uses peripheral diffusion of DNA fragments. Cells without DNA fragmentation show only limited spreading of DNA fiber loops. These results have been seen in several gram-negative and gram-positive bacteria, as well as in yeasts. Detection of DNA fragmentation was confirmed by fluoroquinolone treatment and by DNA breakage detection-fluorescence in situ hybridization. Proteus mirabilis with spontaneously fragmented DNA during exponential and stationary growth or Escherichia coli with DNA damaged after exposure to hydrogen peroxide or antibiotics, such as ciprofloxacin or ampicillin, was clearly detected. Similarly, fragmented DNA was detected in Saccharomyces cerevisiae after amphotericin B treatment. Our assay may be useful for the simple and rapid evaluation of DNA damage and repair as well as cell death, either spontaneous or induced by exogenous stimuli, including antimicrobial agents or environmental conditions.

Chromosomal DNA fragmentation, resulting from massive DNA double-strand breaks, is a hallmark of cell death. In higher eukaryotic cells, this may be a consequence of active programmed cell death (PCD), i.e., apoptosis, where DNA is cleaved by an activated endonuclease (26). Otherwise, DNA fragmentation may occur passively through necrotic cell death. Passive DNA fragmentation is likely in microorganisms killed by various causes. Recent studies suggest, however, a possible PCD in unicellular bacteria and yeasts (38). In fact, bactericidal antibiotics may trigger a PCD response. It has been reported that bactericidal antibiotics can stimulate the production of hydroxyl radicals that contribute to cell death (20). Tolerant bacterial cells, which are resistant to the bactericidal action of some antibiotics, may be cells with a disabled PCD. Ecological pressure, differentiation processes, starvation, and certain DNA-damaging agents may activate PCD involving DNA fragmentation (1, 6, 18, 22, 35). DNA fragments released during autolysis may be absorbed by other bacteria, contributing to antigenic variation and the spread of antibiotic resistance (12). Bacterial autolytic mechanisms have been described primarily with reference to the cell wall level. Nevertheless, active DNA fragmentation has only partially been addressed.

The presence of DNA breakage is usually evaluated using biochemical or molecular procedures such as alkaline unwinding, DNA elution, gel electrophoresis, sucrose gradient sedimentation, melting curve analysis, viscoelastometry, or light scattering (2, 37). Unfortunately, evaluating DNA fragmentation within an individual cell is not possible, nor is it possible to identify important low-level intercellular variations. In situ procedures allow for cells with fragmented DNA to be discriminated from those without DNA fragmentation. However, in situ procedures have been developed primarily for use in higher eukaryotic cells. DNA breaks can be enzymatically labeled by attaching modified nucleotides that can be visualized by direct and indirect methods (10). Usually this process involves the Escherichia coli DNA polymerase I in the in situ nick translation assay or end labeling with the Klenow fragment of DNA polymerase I or the terminal deoxynucleotidyl transferase (TdT) in the TUNEL procedure (10). In each of these, a free and accessible 3′-OH group at the end of the break is necessary as a substrate for extension, but the ends of DNA breaks may be chemically modified by many DNA-damaging agents.

In situ detection of DNA breaks in higher eukaryotes can also be seen through the single-cell gel electrophoresis or comet assay (27). In this technique, the cells are trapped in an inert agarose microgel on a microscope slide, deproteinized by incubation with a lysing solution, and then electrophoresed. DNA staining with a fluorochrome reveals a comet with a head and a tail of chromatin in the direction of the positive pole of the electric field. Cells with DNA breaks have a higher tail and/or greater DNA concentration in the tail. In contrast to those used in enzymatic labeling, cells used in the single-cell gel electrophoresis assay are unfixed and fully accessible to lysis, and the DNA migration is not dependent on the chemical nature of the breaks. An image analysis system is habitually used for evaluation. Nevertheless, electrophoresis is not necessary to identify cells with fragmented DNA. After lysis in the agarose microgel, cells with fragmented DNA can be identified because they produce a peripheral halo of diffused DNA fragments in the agarose matrix. This is the underlying principle of the diffusion assay to detect cells with DNA fragmentation (33, 36).

Regarding microorganisms, the TUNEL assay has been used to determine DNA fragmentation in spheroplasts from yeasts, but only one paper has described its use in bacteria, specifically in E. coli and in the archaeon Haloferax volcanii (31). The procedure was time-consuming, requiring fixation, centrifugation, and permeabilization steps. Moreover, the evaluation used a flow cytometer, requiring a high number of cells. These technical factors make this procedure impractical for routine assessment of DNA fragmentation in the standard microbiology laboratory. For the comet assay, only one paper has described its use for E. coli (34). The procedure was also lengthy, requiring lysozyme digestion of the cell wall prior to incubation in the lysis solution and in conjunction with prolonged proteinase K digestion. The resulting images are difficult to interpret.

Despite its importance from both a basic research and a clinical point of view, DNA fragmentation has not been assessed in the microbiology laboratory. This may be due to the lack of a simple and rapid evaluation procedure. Most current techniques are complex, technically demanding, and not specifically adapted for different microorganisms. Here we present a diffusion-based assay to identify DNA fragmentation in bacteria and yeasts using fluorescence microscopy. This assay is assembled as a kit in order to implement a simple, fast, reproducible, and accurate method for studying DNA fragmentation in microorganisms.

MATERIALS AND METHODS

Microorganisms and cultures.

Chromosomal DNA fragmentation was assayed in gram-negative and gram-positive bacteria (Table (Table1).1). Gram-negative bacteria were grown in Luria-Bertani (LB) broth (1% Bacto tryptone, 0.5% yeast extract, 0.5% NaCl) or on LB agar at 37°C under aerobic conditions. Gram-positive bacteria were grown on Trypticase soy agar plates (Diagnostics Systems, Sparks, MD). Candida albicans and Saccharomyces cerevisiae yeasts were grown in yeast extract-peptone-dextrose broth and plates. Cell growth in liquid cultures was monitored with the spectrophotometer (Unicam 8625; Unicam, Cambridge, United Kingdom). In amphotericin B experiments, viability was determined by colony counting after sequential dilutions and plating.

TABLE 1.
Bacteria assessed with the Micro-Halomax kit

Experiments.

Five different experiments illustrated the use of the procedure to determine chromosomal DNA fragmentation. In the first experiment, aliquots of exponentially growing cultures of the E. coli strain TG1 were exposed to ciprofloxacin (5 μg/ml and 0.012 μg/ml) for 40 min in LB broth at 37°C. Fluoroquinolone is an inducer of DNA fragmentation; this experiment validated the assay. In the second experiment, Proteus mirabilis was incubated in LB medium at 37°C under aerobic conditions. The initial optical density in the spectrophotometer, measured at 600 nm, was 0.05. P. mirabilis isolates in the exponential growth phase and in the stationary phase were analyzed for the frequency of cells with fragmented DNA and for membrane permeability in aliquots that were removed periodically over 106 h. These were batch cultures, where growth uses an unsupplemented amount of nutrients so the nutrients will decrease in time, together with an increase in metabolites. In a third experiment, E. coli strain TG1, either exponentially growing or in the stationary phase, was incubated with 10 mM hydrogen peroxide for 10 min at room temperature in LB broth. Immediately, the cells were processed with the DNA fragmentation kit, as described below. The purpose of this experiment was twofold, as follows: (i) to assess the DNA damage induced by hydroxyl radicals and (ii) to explore the influence of the growth phase on the effect of hydrogen peroxide on DNA. In the fourth experiment, exponentially growing cultures of TG1 were exposed to ampicillin (300 μg/ml) for 40 min in LB broth at 37°C. This was used as a comparison of the effect on DNA of incubation with an antibiotic with a different mechanism of action than that of quinolones. Finally, exponentially growing cultures of S. cerevisiae were incubated with increasing doses of amphotericin B (0, 0.5, 1, 2, 4, 8, and 16 μg/ml) at 30°C for 3 and 24 h and in suspension in yeast extract-peptone-dextrose broth. After incubation, the cells were processed with the DNA fragmentation kit, as described below. A total of 1,000 to 5,000 microorganisms were scored per treatment.

Membrane permeability.

A mixture of SYBR Green II (21) and propidium iodide (PI) (Molecular Probes, Eugene, OR) was prepared in phosphate-buffered saline (PBS) at 333× and 0.17 μg/ml, respectively. A 4-μl aliquot of microorganisms growing in liquid medium was diluted in 16 μl of culture medium and incubated for 5 min with 4 μl of dye mixture in the dark. If the microorganisms were at a very low density, the dye mixture was added to a 20-μl aliquot of cell culture without dilution. Five microliters of the stained cell suspension was dropped onto a glass slide, covered with a coverslip, and examined by fluorescence microscopy. Permeable cells that did not exclude the PI appeared red, whereas those that were “alive” only fluoresced green. For S. cerevisiae, nonpermeable cells appeared unstained.

Determination of DNA fragmentation.

The commercial Sperm-Halomax kits (Halotech DNA SL, Madrid, Spain) used to determine DNA fragmentation in different mammalian spermatozoa were evaluated (13, 17, 30). Using the Sperm-Halomax kit as a reference, a prototype Micro-Halomax kit was developed for microorganisms (Halotech DNA SL, Madrid, Spain). While gram-negative bacteria could be directly processed, a previous brief cell wall digestion step was necessary for gram-positive bacteria. Streptococcus agalactiae was incubated with mutanolysin (0.1 mg/ml), Enterococcus faecalis with a mix of mutanolysin (0.1 mg/ml) and lysozyme (4 mg/ml), and Staphylococcus aureus with a mix of lysostaphin (0.05 mg/ml) and lysozyme (0.25 mg/ml). Streptococcus pyogenes was incubated with lysozyme (1 mg/ml). For this purpose, bacteria were scraped from the culture plate and resuspended in 0.25 ml of LB medium or PBS, pH 6.88, in 0.5-ml snap-cap microcentrifuge tubes. The enzyme was added at the desired final concentration and was incubated for 15 min at 37°C. C. albicans and S. cerevisiae were digested with lyticase, 2.5 U/ml for 15 min and 0.07 U/ml for 10 min, respectively, in 1 M sorbitol, 1 M EDTA, and 15 mM beta-mercaptoethanol, pH 7.5, at 30°C. All enzymes were purchased from Sigma (St. Louis, MO).

An aliquot of each sample was diluted to a concentration of 5 × 106 to 10 × 106 microorganisms/ml in the broth culture medium specific for each microorganism. The yeast was centrifuged and resuspended in the lyticase buffer without the enzyme. Snap-cap microcentrifuge tubes (0.5 ml) containing gelled aliquots of 60 μl of low-melting-point agarose (Pronadisa, Laboratorios Conda, Madrid, Spain) in distilled water are provided with the Micro-Halomax kit. The tube was placed in a water bath at 90 to 100°C for 5 min to melt the agarose and then transferred to a water bath at 37°C (Memmert, Schwabach, Germany). After a 5-min incubation to allow for equilibration to 37°C, 25 μl of the diluted sample containing the microorganism was added to the tube and mixed with the melted agarose. Aliquots (20 μl) of the sample-agarose mixture were pipetted onto a precoated slide provided with the kit and were covered with a 22- by 22-mm coverslip. The coating of the slides consists of a dried agarose layer prepared with 1% standard agarose in water and drying in an oven at 80°C (Memmert, Schwabach, Germany). The slide was placed on a cold plate in the refrigerator (4°C) for 5 min to allow the agarose to solidify, producing a microgel with the intact cells trapped inside. The coverslip was gently removed, and the slide was immersed in 10 ml of lysis solution provided in the Micro-Halomax kit, previously tempered to 37°C, for 5 min in a closed tray at 37°C. This solution contains 2% sodium dodecyl sulfate, 0.05 M EDTA, and 0.1 M dithiothreitol, pH 11.5. The slide was always placed in horizontal position to prevent DNA stretching. After washing horizontally for 3 min in a tray with abundant distilled water, the slide was dehydrated by being incubated horizontally in cold (−20°C) ethanol baths, first 70%, and then 90%, and finally 100%, for 3 min each, followed by air-drying in an oven (Memmert, Schwabach, Germany).

DNA staining with the fluorochrome SYBR Gold (Molecular Probes, Eugene, OR) (7) could be performed immediately after drying. Before staining, the dried slide must be incubated in a microwave oven (Whirlpool, Norrköping, Sweden) at 750 W for 4 min to promote the attachment of DNA to the slide. The slide may also be placed in an oven at 80°C for 1 h to overnight. The slide was then stained with 25 μl of SYBR Gold diluted 1:100 in TBE buffer (0.09 M Tris-borate, 0.002 M EDTA, pH 7.5), covered with a plastic coverslip, and incubated for 5 min in the dark. The slide was briefly washed and mounted in TBE. Fluorescence microscopy must be performed immediately after staining to avoid drying. If needed, the slide may be stored at 4°C for hours in a self-made humid box in the dark to prevent drying. If dried, the coverslip may be removed by incubation in PBS and, after a brief wash, may be restained again. If immediate evaluation is not necessary, dried slides may be left overnight or a couple of days in a high-temperature oven (80°C) and then stored in a tightly closed box, in the dark at room temperature, for several months before staining.

Fluorescence microscopy allows for ×10 to ×100 magnification, but ×100 magnification is necessary for a precise visualization of the small spots from nucleoids with fragmented DNA. Three microgels can be placed on a same slide, including a control sample if required, so that all microgels are simultaneously processed under the same conditions.

DBD-FISH.

To confirm the presence of DNA breakage in cells with diffused DNA spots, the DNA breakage detection-fluorescence in situ hybridization (DBD-FISH) procedure (14, 15) was used for E. coli, P. mirabilis, Acinetobacter haemolyticus, Staphylococcus aureus, and S. cerevisiae nucleoids. The cells were immersed in agarose microgels and were lysed as described above. They were then washed in 0.9% NaCl and were incubated in an alkaline unwinding solution (0.03 M NaOH) for 2.5 min at 22°C. The gels were neutralized in 0.4 M Tris-HCl, pH 7.5, washed in distilled water, dehydrated in increasing ethanol baths (70%, 90%, and 100%) for 2 min each, and air-dried.

A DNA probe to label the total DNA from the microorganism was prepared. DNA from each microorganism was isolated using standard procedures and was labeled with biotin-16-dUTP using a nick translation kit according to the manufacturer's instructions (Roche Applied Science, San Cugat del Vallés, Spain). The DNA probe was mixed at 4.3 ng/μl in the hybridization buffer (50% formamide-2× SSC, 10% dextran sulfate, 100 mM calcium phosphate, pH 7.0) (1× SSC is 0.015 M sodium citrate plus 0.15 M NaCl, pH 7.0). The probe in hybridization buffer was denatured by incubation at 80°C for 8 min and then incubated on ice. This solution (30 μl) was pipetted onto the dried slide, covered with a glass coverslip (22 by 60 mm), and incubated overnight at room temperature, in the dark, in a humid chamber. The coverslip was removed, and the slides were washed twice in 50% formamide-2× SSC, pH 7.0, for 5 min and twice in 2× SSC, pH 7.0, for 3 min at room temperature. The slides were incubated with 80 μl of blocking solution (4× SSC, 0.1% Triton X-100, 5% bovine serum albumin) for 5 min, covered with a plastic coverslip, and stored in a humid chamber at 37°C. This solution was decanted, and the bound probe was detected by incubation with 80 μl of streptavidin-Cy3 (Sigma, St. Louis, MO) in 4× SSC, 0.1% Triton X-100, 1% bovine serum albumin (1:200), covered with a plastic coverslip, and stored in a humid chamber at 37°C. After washing in 4× SSC, 0.1% Triton X-100 three times for 2 min each, the slides were counterstained with 20 μl of DAPI (4′,6-diamidino-2-phenylindole) (2 μg/ml) in Vectashield (Vector, Burlingame, CA).

Fluorescence microscopy and digital image analysis.

Images were viewed with an epifluorescence microscope (Nikon E800) with a ×100 objective and appropriate fluorescence filters for fluorescein isothiocyanate-SYBR Gold (excitation 465 to 495 nm, emission 515 to 555 nm), PI-Cy3 (excitation, 540 to 525 nm; emission, 605 to 655 nm), and DAPI (excitation, 340 to 380 nm; emission, 435 to 485 nm). The images were captured with a high-sensitivity charge-coupled-device camera (KX32ME; Apogee Instruments, Roseville, CA). Groups of 16-bit digital images were obtained and stored as TIFF files. Image analysis used a macro in Visilog 5.1 software (Noesis, Gif sur Yvette, France). This allowed for thresholding, background subtraction, and measurement of the total fluorescence intensity (surface area in pixels times mean fluorescence intensity in gray level) of the signals. In the experiment concerning ciprofloxacin exposure, the surface area of diffusion of the DNA fragments from nucleoids, in number of pixels, was established for the control, for 0.012 μg/ml, and for 5 μg/ml. Since the data were not normally distributed, as ascertained by the Kolmogorov-Smirnov test, a nonparametric Mann-Whitney U test was performed to compare between doses.

RESULTS AND DISCUSSION

Technical implications.

Here we present an adapted single-cell diffusion assay, initially used to identify DNA fragmentation in mammalian sperm cells (13, 17, 30), for assessing chromosomal DNA integrity in microorganisms with relatively small genomes. Intact, unfixed microorganisms were immersed in an agarose microgel on a slide, lysed, and stained with a DNA fluorochrome. In higher eukaryotic cells, cells without DNA fragmentation release only DNA loops around a central core, but cells with fragmented DNA produce a large halo of diffusion of DNA spots or fragments.

Given the relatively small genome size of microorganisms, classical fluorochromes such as PI, DAPI, Hoechst, etc., are not suitable for staining. In order to visualize DNA fragments, it is necessary to use a highly sensitive fluorochrome, such as one from the SYBR family. SYBR Gold provides excellent sensitivity and photostability in comparison with other fluorochromes from the same family, giving an accurate visual assessment under the fluorescence microscope (7). Antifading solution was not used, since it diminishes the contrast between the small DNA dots and the background.

Validation of the assay by ciprofloxacin treatment and DBD-FISH.

Ciprofloxacin is a fluoroquinone that induces DNA double-strand breaks by trapping DNA gyrase and topoisomerase IV on DNA (19). DNA breaks have been shown by several methodologies, including viscosity measurements of cell lysates (24, 25). After processing with the Micro-Halomax kit and SYBR Gold staining, bacteria from untreated control cultures showed nucleoids with DNA loops spreading from a central core, which corresponds to the residual bacterium, with a compact, microgranular surface extended peripherally to many branches (Fig. 1a and b). Remarkably, a few bacteria, 0.4%, spontaneously had a very big halo of DNA spots radiating from the residual central core (Fig. (Fig.1a1a and see Fig. Fig.5a).5a). These images were similar to those visualized in higher eukaryotic cells with fragmented DNA with the diffusion-based assay (13, 17, 30, 36). After treatment with 5 μg/ml ciprofloxacin, all the nucleoids appeared similar to nucleoids with extensive diffusion of DNA spots as observed occasionally in the control cultures. Thus, results with ciprofloxacin demonstrate that (i) our procedure confidently detects DNA fragmentation and that (ii) images of nucleoids with a big halo of diffusion of DNA spots indicate extremely fragmented DNA. Quantitative analysis of digital images revealed that the mean surface area of the nucleoids was 11.5-fold larger than that of the untreated control bacteria (Fig. (Fig.1d;1d; Table Table22).

FIG. 1.
Images after application of the Micro-Halomax kit to E. coli cultures. Cells were embedded in an agarose microgel, lysed, and stained with SYBR Gold. (a) Nucleoids from control untreated cells, showing the spread of DNA loops from a central core. One ...
FIG. 5.
Exponentially growing cultures from E. coli control cells (a) and from those exposed to 10 mM hydrogen peroxide for 10 min (b), evaluated with the diffusion-based assay using the Micro-Halomax kit. (a) Bacterial nucleoids from control cultures only show ...
TABLE 2.
Digital image analysis of nucleoids from E. colia

Interestingly, the effect of ciprofloxacin on DNA integrity was increased with respect to untreated control cultures at the MIC, i.e., 0.012 μg/ml (Fig. (Fig.1c).1c). After this low dose, the DNA damage was less and was constant among the bacteria. In fact, the nucleoids appeared more spread, with their average surface area being 3.9-fold larger than that of the untreated control cells (Table (Table2)2) and having larger peripheral DNA fragments than those detected after the high dose. As postulated by Drlica et al. (11), ciprofloxacin at low doses like the MIC and short incubation times may block growth without killing the cells, suggesting the formation of reversible complexes. At higher doses, like 5 μg/ml, the DNA is extremely fragmented, as observed here, perhaps causing rapid death. The experiment demonstrates the sensitivity and potential value of our procedure for determining the activity of quinolones, both in basic and in clinical research. This approach is currently under investigation in our laboratory.

To further confirm the presence of DNA breaks in nucleoids with diffused DNA fragments in control E. coli cultures, the DBD-FISH technique was employed (14, 15). This procedure uses the same microgel as the diffusion assay, allowing simultaneous or sequential visualization of the nucleoids with or without fragmented DNA and labeling of DNA breaks. DBD-FISH is a powerful procedure that involves microgel embedding, lysis, and incubation with a limited alkaline DNA unwinding step (3, 32). This final step transforms DNA breaks into limited single-stranded DNA (ssDNA) segments generated from the ends of the breaks, which hybridize to fluorescent DNA probes. As DNA breaks increase, more ssDNA is produced, increasing probe hybridization and fluorescence intensity. Fluorescence may be quantified using image analysis software. When hybridizing with a whole-genome probe, DNA breaks in the entire genome are assessed. Damage within specific DNA sequence areas may be evaluated by hybridizing specific DNA probes.

The DBD-FISH procedure was applied to E. coli (Fig. (Fig.2)2) and other microorganisms lysed in the microgel. Nucleoids with diffused spots were strongly labeled, further confirming massive DNA breaks.

FIG. 2.
DBD-FISH-detected DNA breaks in nucleoids from control cultures of E. coli. Cells in an agarose microgel were lysed and treated with an alkaline unwinding treatment to transform DNA breaks into restricted ssDNA motifs detected by hybridization with a ...

Three kinds of experiments demonstrated the potential of the procedure to determine chromosomal DNA fragmentation, as follows: (i) analysis of cells with spontaneous fragmented DNA in culture, (ii) reactive oxygen species-induced DNA damage, and (iii) analysis of antibiotic and antifungal agent effects.

Batch cultures.

P. mirabilis was incubated in liquid LB medium for 106 h, while the turbidity was monitored and aliquots were removed periodically to determine membrane permeability and DNA fragmentation. Bacteria with fragmented DNA were easily separated from nucleoids without fragmented DNA (Fig. (Fig.3a).3a). The frequency of bacteria with DNA fragmentation was established using the Micro-Halomax kit. This evaluated nucleoids with diffused DNA fragments in 1,000 to 5,000 cells per experimental point. Membrane permeability was determined by SYBR Green II and PI staining. All cells were permeable to SYBR Green II, but PI is a vital dye, so only cells with membrane permeability, which do not exclude the dye, appear red under the PI filter set of the microscope. The frequency of PI-permeable cells was established in 1,000 to 5,000 microorganisms per experimental point (Fig. (Fig.44).

FIG. 3.
Images after the application of Micro-Halomax to cultures of P. mirabilis (a) and C. albicans (b), processed as indicated in Materials and Methods. Nucleoids without DNA fragmentation released DNA loops around a central core from the residual cell. Otherwise, ...
FIG. 4.
Kinetics of the frequency of P. mirabilis cells with fragmented DNA and with a PI-permeable membrane. Identification of bacteria with fragmented DNA was performed using the Micro-Halomax kit, as indicated in Materials and Methods. OD600, optical density ...

The culture changed from the exponential growth phase to the stationary phase after 9 h. The percentage of cells permeable to PI increased at 48 h from 0.5% to 5% and then progressively rose through the end of the experiment to 88%. The proportion of bacteria with fragmented DNA significantly increased after 81 h from 0.5 to 1% up to 9.5%. It remained constant at 35% from 99 to 103 h and then rose to 52% 3 h later.

These results suggest that membrane permeability does not indicate the presence of fragmented DNA, being that these are independent parameters related to different initial targets and not correlated in time. Moreover, the stationary phase does not seem to be steady in the frequency of bacteria with fragmented DNA. In the initial period of the stationary phase, the proportion of bacteria with DNA fragmentation did not increase over that in the exponential phase. The percentage increased later, probably reflecting a progressive change in the turnover rate between dead and dividing cells. Perhaps with the accumulation of metabolites and the depletion of nutrients, the fraction of cells with fragmented DNA should increase further.

Hydrogen peroxide treatment.

The effect of reactive oxygen species on DNA integrity was evaluated. Hydrogen peroxide decomposes into hydroxyl radicals (OH) through catalysis by low-valence-transition metal ions in a Fenton-Haber-Weiss reaction. These oxidizing agents strongly reacted with macromolecules. OH attack on DNA results in a variety of base damages and DNA breaks (8, 9). In the only report using the TUNEL assay for bacteria (31), labeling of DNA breaks was detected in exponentially growing cultures of E. coli after exposure to extremely high doses of H2O2 (86 mM for 30 min). Surprisingly, in stationary-phase cultures, even doubling the H2O2 dose (172 mM) did not result in DNA breakage. This suggested that H2O2 does not directly cause DNA breaks, which could be transient intermediates in DNA repair produced by the DNA repair enzymes (31).

We tested this hypothesis by using our DNA fragmentation assay. The percentage of E. coli cells with fragmented DNA was 0.4% and 37.6% in untreated control cells growing exponentially and in the stationary phase, respectively (Fig. 5a and c). Using a lower dose for a shorter length of time than in the previous report (10 mM and 10 min), 100% of nucleoids showed extensively fragmented DNA, either in the exponential or stationary growth phase (Fig. 5b and d). This result suggests a higher sensitivity in the microgel-based assay than in the TUNEL assay for bacteria and illustrates a difficulty with enzymatic procedures for labeling DNA breaks. In the case of TdT, a free 3′-OH group at the terminus of the DNA break is essential as a substrate in order to polymerize the nucleotides (10). Attack by agents like H2O2 does not produce “clean” DNA termini but rather chemically modified ends, such that direct DNA breaks could be undetectable to enzymes (9). To allow for DNA repair, exonuclease III removes blocking groups at the 3′ terminus (9). Labeling by TdT should therefore be possible. The absence of enzymatic labeling in stationary-phase cultures could be explained if end processing is impaired at this stage. Nevertheless, H2O2-induced DNA breaks are visible with our assay, since it is independent of the chemical nature of the DNA break. Overall, our diffusion assay identifies DNA damage by OH, in both the exponential and stationary growth phases.

Ampicillin incubation.

To evaluate the influence of ampicillin treatment on chromosomal DNA, exponentially growing cultures of TG1 were exposed to 300 μg/ml ampicillin for 40 min or 24 h. This dose was much higher than the MIC of 3 μg/ml. In contrast to the target of ciprofloxacin, which affects DNA, the cell wall is the primary target for ampicillin; it inhibits peptidoglycan synthesis after binding to penicillin binding proteins and activating autolysins (5, 16).

Unlike with ciprofloxacin, a 40-min treatment with ampicillin barely increased the frequency of cells with fragmented DNA or with the appearance of DNA damage. The nucleoids were similar to those from untreated control cells. When incubated for 24 h, the density of bacteria and corresponding nucleoids was reduced but had a uniform background of DNA spots that were probably from spontaneously lysed cells that released the DNA fragments to the medium (Fig. (Fig.6).6). This suggests that cell death initially appears to be independent of DNA damage but may evolve later in DNA degradation.

FIG. 6.
E. coli cultures processed with the Micro-Halomax kit after ampicillin treatment, 300 μg/ml, for 24 h. Nucleoids from residual cells appear more relaxed, accompanied by a dense background of DNA fragments. Bar: 5 μm.

Amphotericin B incubation in yeast.

A presumed PCD has been described in S. cerevisiae, following acidic, oxidative, or osmotic stress and after UV exposure (23). This has also been reported for Candida albicans after acetic acid, H2O2, or amphotericin B treatment (28, 29). Apoptotic cells were very significantly increased after a 200-min incubation with 4 μg/ml amphotericin B (28). Apoptotic cells were considered those not growing and not permeable to PI. DNA fragmentation was not assessed. After a dose of 16 μg/ml, practically no Candida cells grew, with the cells appearing 10% PI impermeable assumed to be apoptotic and the PI-permeable cells presumed to be necrotic.

An image of C. albicans showing DNA fragmentation is presented in Fig. Fig.3b.3b. Nevertheless, we assessed the possible induction of DNA fragmentation by amphotericin B in S. cerevisiae. In untreated control cultures, no cells with fragmented DNA were detected in 6,000 yeasts. There was no evidence of fragmented DNA with any dose of the antifungal agent when incubated for 3 h. After a 24-h incubation with amphotericin B, yeast cells with fragmented DNA were recorded in a dose-dependent manner (Fig. (Fig.7).7). The frequency of cells with fragmented DNA was lower than that with the PI-permeable membrane. In fact, with the highest dose assayed, 5% of the cells contained fragmented DNA, whereas 85% were PI permeable. This decouples membrane permeability from DNA fragmentation as being indicative of death in these microorganisms, at least after treatment with antifungal agents, like amphotericin B, that target yeast cell membranes (4). A substantial and proportional decrease in viability with the dose, assayed 48 h after treatment, was demonstrated (Fig. (Fig.7),7), suggesting that DNA fragmentation after amphotericin B treatment is either a rare phenomenon or a late response.

FIG. 7.
S. cerevisiae cells with fragmented DNA (right scale), a PI-permeable membrane, and viability (left scale) after incubation for 24 h with increasing doses of amphotericin B.

Conclusion.

The experiments presented here illustrate the ability of the technique, assembled as a kit, to determine the presence of fragmented DNA in microorganisms. Its simplicity, short assay time (50 min), and efficacy make this technique useful for the routine determination of DNA fragmentation and intercellular variation. Applications may be extensive for both basic and clinical research. Only a fluorescence microscope is required. Though direct visual identification is quite sharp, the scoring process may be partially automated by adapted image analysis software. This automation could be more complete by integrating a microscope with a motorized plate and focus, a charge-coupled-device camera for image capture, and image analysis software. This could be useful when scoring many thousands of microorganisms, resembling the conditions in flow cytometer facilities.

Acknowledgments

J. L. Fernández, V. Goyanes, and J. Gosálvez collaborate as scientific advisers of Halotech DNA SL.

This work was supported by public grants from the Xunta de Galicia (07CSA050916PR and INCITE07PXI916201ES) and by FIS PI061368.

We are grateful to Christopher de Jonge, from the University of Minnesota, for the critical reading of the manuscript.

Footnotes

[down-pointing small open triangle]Published ahead of print on 8 August 2008.

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