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The aim of this study was to observe growth of isolated single bacterial cells in the absence of growth factors and intercellular contact. In order to exclude stochastic uncertainties induced by dilution series, a new micromanipulation method was developed to ensure explicit results under visual control. This was performed with particular care for production of single prokaryotic cells and subsequent investigation of their autonomous growth. Over 450 single isolated Listeria monocytogenes and Salmonella enterica subsp. enterica serovar Typhimurium cells in lag, log, and stationary growth phases were investigated by this method, which included thoroughly washing the cells. The proportion of living cells within the initial cultures was compared to the proportion of positive samples after enrichment of the separated single cells. This resulted in P values of ≥0.05 using the chi-square test for statistical analysis, indicating no significant difference, and clearly demonstrates reproduction of isolated single bacterial cells without the need for growth factors or intercellular contact. Ease of handling of the apparatus and good performance of the cleaning procedures were achieved, as was validation of the method, demonstrating its suitability for routine laboratory use.
The possibility of independent growth of isolated single prokaryotic cells has been discussed recently and remains controversial (15). Undoubtedly, cell-to-cell communication plays a key role in the establishment and development of bacterial communities. Both physical and chemical factors influence the organization of biofilms, sporulation, and resuscitation of bacterial populations. The analogy of the chemical factors to eukaryotic pheromones, as well as their role in bacterial cell division, has been postulated (34). These facts are evident and have been well investigated, and, in summary, the necessity of intercellular contacts, population effects, and intrinsic growth factors such as resuscitating promoting factor (rpf) is generally supposed (24, 36). Nevertheless, when it comes to cell division and growth of low inocula of prokaryotic cells, some questions appear to remain open. Considering evolutionary developments, asexual reproduction is a prerequisite for survival of single organisms in the environment and one of the necessities for the success of the prokaryotic kingdom. Taken the other way, asexual reproduction also suggests the possibility of independent growth of isolated single bacterial cells. The scientific community remains divided over this possibility, not least because of the possible heterogeneity of bacterial cultures in terms of the physiological status of every single cell and the effect of a Poisson distribution in highly diluted cultures, which influence experimental setups and results (17).
If one viable prokaryotic cell is to be shown to be capable of generating a population of daughter cells, a prerequisite is its isolation and physical manipulation as an individual cell. Simple serial dilution protocols, such as the most probable number (MPN) method, achieve this task, but they suffer from a lack of certainty that the diluted solution indeed contains only one viable cell, free from any adherent growth factors. In such cases the accuracy of the data obtained by multiple dilution procedures becomes uncertain regarding the Poisson distribution of highly diluted bacterial cell (≤10 CFU/ml) suspensions (11, 32).
Combined microinjection and micromanipulation methods are alternative applications. These are widespread routine techniques that have been developed recently for large eukaryotic cells (14, 38). However, application of these methods to prokaryotic cells leads to new physical conditions defined by the technical equipment secondary to the size of the handled cells (≤1 μm). These include limitations of fluorescence microscopy, general microscopic analysis, and photographic documentation. Over the past 30 years several attempts to improve the management of single prokaryotic cells using micromanipulation techniques have been made (10). The core issue of these studies has been isolation of single microbial cells from mixed populations under direct visual control (9, 13).
In contrast, the aim of this study was to develop a method capable of serial manipulation of single prokaryotic cells under visual control. This was accomplished to produce series of single prokaryotic cells and subsequently to investigate their autonomous growth. The growth of single Listeria monocytogenes and Salmonella enterica subspecies enterica serovar Typhimurium cells was investigated in highly diluted buffer systems without facultative growth factors being transferred from the primary enrichment. In this study over 450 single cells isolated from different growth phases (lag phase, log phase, and stationary phase) were manipulated using a novel single bacterial cell manipulation (SBCM) technique.
L. monocytogenes EGDe (1/2a, internal number 2964) and S. Typhimurium (NCTC 12023) were maintained at −80°C using MicroBank technology (Pro-Lab Diagnostics, Richmont Hill, Canada) and were part of the collection of bacterial strains at the Institute of Milk Hygiene, Milk Technology and Food Science, Department of Veterinary Public Health and Food Science, University of Veterinary Medicine, Vienna, Austria. All bacterial strains were grown overnight in tryptone soy broth with 0.6% (wt/vol) yeast extract (TSB-Y; Oxoid, Hampshire, United Kingdom) at their respective optimal growth temperatures (37°C, L. monocytogenes; 42°C, S. Typhimurium). To estimate the required individual inoculation time to reach the different growth phases (lag phase, log phase, and stationary phase), growth curves for L. monocytogenes EGDe and S. Typhimurium were determined. After inoculation of 1 ml of an overnight bacterial culture into 50 ml TSB-Y, the bacteria were incubated at their optimal growth temperatures. Aliquots were taken in duplicate every hour for 10 h and additionally after 24 and 36 h for measurement and plating. Cell counting was performed by plating and measurement of optical density at 600 nm (OD600) as described below.
To evaluate the suitability of the staining method for our application, the growth of pure cultures (unstressed and heat injured) was compared with that of stained ones. Staining of Gram-positive and Gram-negative bacteria was performed with SYBR Safe DNA gel stain (SYTO9; Invitrogen, Molecular Probes, Eugene, OR). Four different types of overnight cultures were prepared for L. monocytogenes and S. Typhimurium. The control samples contained 1.1 × 102 CFU (relative standard deviation [RSD], ±13.6%) S. Typhimurium cells or 4.8 × 102 CFU (RSD, ±6.8%) L. monocytogenes cells transferred into 9 ml of fresh medium (TSB-Y). The stained samples additionally contained 0.25 μl/ml SYTO9. The samples were incubated for 20 h at the respective optimal growth temperatures (37°C, L. monocytogenes; 42°C, S. Typhimurium). One hundred microliters was plated on tryptone soy agar plates supplemented with 0.6% (wt/vol) yeast extract (TSA-Y; Oxoid, Hampshire, United Kingdom), and the plate count method was used for quantification after incubation at the respective optimal growth temperatures for 24 h. The heat injury settings for L. monocytogenes (55°C, 20 min) and S. Typhimurium (48°C, 30 min) were determined as published previously (1, 22). The heat-injured cells were exposed to the same procedure (control and stained samples) with SYTO9 as described above.
A prerequisite for easy and fast manipulation of the bacterial cells is the removal of debris from the enrichment broth; therefore, four wash steps were established. One milliliter of the initial culture in either lag phase, log phase, or stationary phase was pelleted at 7,000 × g for 3 min and washed twice in 1 ml of double-distilled water (ddH2O; Mayerhofer Pharmazeutika, Leonding, Austria) at 7,000 × g. An aliquot of stain equivalent to 0.25 μl/ml SYTO9 was added to the resuspended bacterial pellet.
To test the influence of several ingredients of the transport buffer with respect to osmotic stress and reduction of surface tension, the bacterial cells were washed twice in 1 ml of ddH2O at 7,000 × g and resuspended in 1 ml of transfer buffer A (1× phosphate-buffered saline [PBS]), B (ddH2O), C (1% Tween 80 [Merck, Darmstadt, Germany] in ddH2O), or D (1% Tween 80 in Ringer's solution [Mayerhofer Pharmazeutika, Leonding, Austria]). The manipulation protocol was performed as described below, and the rates of recovery of the isolated single bacterial cells in comparison with the viability staining data of the respective initial cultures were determined.
Viability staining was performed by adding 1 μl of component A and 1 μl of component B of the BacLight Live/Dead bacterial viability kit (Molecular Probes, Willow Creek, OR) to 1 ml of an appropriate dilution of the bacterial cultures in sterile filtrated Ringer's solution (Merck, Darmstadt, Germany). The samples were incubated for 15 min protected from light, and 400 μl was filtered onto 0.22-μm-pore-size, 13-mm black polycarbonate filters (Millipore, Billerica, MA) using a 5-ml syringe and a Swinnex filter holder (Millipore). Filter discs (diameter, 12.7 mm) to test antibiotics (Schleicher & Schuell GmbH, Dassel, Germany) were placed beneath the polycarbonate filters in the filter holder for support. Ten fields per filter were analyzed, and two filters were prepared for each sample. The following formula was used to calculate the number of stained cells (N) per ml sample: N = mean number of cells per field × (effective filtration area/area of the field) × (1/dilution factor) × (1/filtrated volume in ml). A DM-IRB fluorescence microscope (Leica, Wetzlar, Germany) with a 470-nm filter was used for microscopic analysis at 1,000-fold magnification.
One milliliter of an overnight culture was transferred to 9 ml of fresh medium and incubated until the respective growth phase was reached, according to the growth curve obtained as described above. Subsequently, 100 μl of the appropriate dilutions in TSB-Y was added to the samples. One milliliter of the initial culture in either the lag phase, log phase, or stationary phase was pelleted at 7,000 × g for 3 min and washed twice in 1 ml ddH2O at 7,000 × g. An aliquot of stain equivalent to 0.25 μl/ml SYTO9 was added to the resuspended bacterial pellet. The bacterial cells were washed twice in 1 ml of ddH2O at 7,000 × g and resuspended in 1 ml of transfer buffer D (Ringer's solution and 1% Tween 80). An approximately 1:1,000 dilution with transfer buffer, depending on the concentration of the initial culture, was introduced to ensure a low cell concentration prior to sample loading. For positive controls, each 10 μl of the respective culture before and after the washing procedure was transferred to TSB-Y and incubated overnight.
A conventional manual hydraulic micromanipulation-microinjection system (Narishige International Limited, London, United Kingdom) was further developed as follows. The originally air-driven microinjection system (Narishige; IM-5B) was filled with red mineral oil to reduce the response time of the sample transport and to improve detection of air bubbles within the system. The original 50-ml syringe was replaced by a 500-μl 1750 AD syringe (Hamilton, Bonaduz AG, Bonaduz, Switzerland) to obtain a hydraulic ratio better suited for the manipulation of bacterial cells. A second microinjection system was constructed on a lathe, assembled with 10- to 50-μl 1701/1705 RN Hamilton syringes to provide a second hydraulic ratio for fine manipulation of the cells, and connected to the system with a medical three-way tap (Discofix; B. Braun AG, Melsungen, Germany). Another medical three-way tap was connected to provide separate access to the sample from behind the capillary (Fig. (Fig.1)1) and to separate the sample from the hydraulic transport part of the system. The sample was fed to the system via a 10-ml syringe through the three-way tap. A hydraulic micromanipulator (MMO-202; Narishige) was used to control the capillary (Femtotip; Eppendorf, Hamburg, Germany), which was fixed in a Narishige IM-H injection holder. To clean the system, the sample part was disconnected from the hydraulic transport part by closing the three-way tap and the fraction of mineral oil, which was transported to the sample part of the system, was discarded. Subsequently, the sample part was flushed several times with liposoluble cleaners, Microzid (Schülke und Mayr, Vienna, Austria), 70% ethanol, and ddH2O via 20-ml syringes using the sample path through the system. The whole system was mounted in a fume hood and sterilized periodically by UV radiation. For documentation a DFC-300FX charge-coupled device (CCD) camera (Leica, Solms, Germany) was used.
Manipulation was carried out on a sterile microscopic coverslip (24 mm by 50 mm, no. 0 [0.08 to 0.12 mm thick]; Menzel, Braunschweig, Germany) by building a double glass layer (overall thickness: ~0.16 mm) with four superior sterile round coverslips (diameter, 5 mm, no. 00 [0.055 to 0.08 mm thick]; Menzel, Braunschweig, Germany) as the carrier. The sample part of the manipulator was filled with buffer solution containing the bacterial cells by injection into the apparatus with a 10-ml syringe. The sample part was then connected to the hydraulic transport part of the system by switching the respective three-way tap (Fig. (Fig.1,1, no. 2, and and2C).2C). Cell transport through the capillary onto the sterile round coverslips was supported hydraulically by the microinjectors. The microinjector fitted with the 500-μl syringe was used to transport the cells to the tip of the capillary. The second microinjector, fitted with the 10- to 50-μl syringes, was used to pull the cells enclosed in a drop of buffer solution out of the capillary onto the glass slip. The glass slips were subsequently put into the reaction tubes with sterile tweezers. Three out of four round coverslips were designated for manipulation of single cells; the fourth one represented the negative control. After the manipulation, each 5-mm carrier glass slide was carefully transferred to a reaction tube containing 9 ml TSB-Y and incubated overnight at the respective optimal growth temperature.
Cell count and live/dead ratio for the respective growth phases of the initial cultures were evaluated before the washing procedure by viable cell counting using BacLight Live/Dead staining as described above.
For the single cell growth experiments, an inoculum of 1 milliliter containing approximately 3.5 × 107 CFU/ml (RSD, ±1.4%) L. monocytogenes to obtain an adequate concentration of bacteria for subsequent SBCM was transferred to 9 ml TSB-Y and incubated at 37°C for 2 h (lag phase), 5 h (log phase), and 10 h (stationary phase). Incubation time was calculated according to a calibration curve based on the determination of the growth rate of the respective bacteria. For S. Typhimurium cultures 1 ml bacterial suspension, containing approximately 1.5 × 107 CFU/ml (RSD, ±1.4%) bacterial cells, was transferred to 9 ml TSB-Y and incubated at 37°C for 1.5 h (lag phase), 3 h (log phase), and 8 h (stationary phase). L. monocytogenes and S. Typhimurium initial cultures in the respective growth phases were washed and subjected to SBCM as described above. Subsequently, growth in TSB-Y at 37°C for Listeria and 42°C for Salmonella within 24 h was evaluated for single isolated cells. Due to the high number of cultures obtained, the cell count of the recultivated single cells was evaluated by measurement of optical density and comparison of the obtained values with a calibration function. Measurement of the optical density was performed at 600 nm (OD600) in duplicate with an HP 8452 spectrophotometer (Hewlett Packard, Palo Alto, CA). All offspring populations were plated on unselective agar after micromanipulation and checked for L. monocytogenes and S. Typhimurium by plating on selective agar and real-time PCR. For Listeria Palcam (Biokar Diagnostics, Beauvais, France) and Oxoid chromogenic listeria agar base (OCLA; Oxoid, Basingstoke, Great Britain) agar plates were used in parallel. For Salmonella XLD4 (Merck, Darmstadt, Germany) agar plates were used. Selective plates for Listeria were incubated at 37°C for 48 h and for Salmonella at 42°C for 24 h.
For confirmation of suspected colonies on TSB-Y agar plates, the colonies were suspended in 1 ml 0.01 M Tris-HCl (pH 7.0), pelleted at 5,000 × g for 5 min, and subjected to Chelex-based DNA isolation. The pellet was resuspended in 100 μl 0.01 M Tris-HCl and 400 μl lysis solution (0.25 mM Tris-HCl [pH 7.0], 2.5% [wt/vol] Chelex 100 resin [Bio-Rad, Hercules, CA]) and incubated for 10 min at 100°C. The debris was pelleted at 14,000 × g for 5 s. The remaining supernatant was stored at 4°C until real-time PCR was performed.
Real-time PCR was performed as published previously by targeting a 274-bp fragment of the prfA gene of L. monocytogenes (4, 30). S. Typhimurium was detected using the SureFood kit (R-Biofarm, Darmstadt, Germany), according to the instruction manual. Real-time PCR was performed in a Mx3000p real-time PCR thermocycler (Stratagene, La Jolla, CA). The 25-μl volume contained 5 μl of DNA template. All real-time PCR sampling was performed in duplicate.
Data resulting from bacterial growth (CFU/ml) are represented as means ± RSDs. Two-group comparisons were analyzed by chi-square test for the single cell growth tests and by two-tailed Student's t test for the evaluation of the staining method. P values were calculated, and values ≤0.05 were considered significant.
A total of 450 L. monocytogenes and S. Typhimurium cells have been handled with the newly developed method, 165 cells during establishment of the protocol and 285 cells for the purpose of single cell growth evaluation. The respective negative controls were all unspoiled, indicating sterile handling during the procedure and reliable performance of the cleaning protocol. The method allowed an output of up to 25 single cells per hour. Visual control of the stained bacterial cells was adequate (Fig. (Fig.2),2), and, together with the two leverages provided by the different syringe volumes, routine application of the method was achievable.
To determine the influence of the staining dye used for visualization of the bacterial cells, the maximum cell densities of L. monocytogenes and S. Typhimurium in enrichment media (TSB-Y) containing 0.25 μl/ml SYTO9 were monitored. Unstressed and heat-injured cultures were investigated for both species. Within the control group, grown in TSB-Y, heat-injured Salmonella multiplied to a mean of 1.48 × 108 CFU/ml (RSD, ±27.6%) and the unstressed to a mean of 2.12 × 108 CFU/ml (RSD, ±31.3%). Unstressed Salmonella cultures incubated with TSB-Y and 0.25 μl/ml of SYTO9 multiplied to a mean of 2.30 × 108 CFU/ml (RSD, ±22.4%) and heat-injured to a mean of 1.89 × 108 CFU/ml (RSD, ±19.5%) within 20 h.
Pure cultures of unstressed and heat-injured Listeria multiplied to means of 1.27 × 109 CFU/ml (RSD, ±16.9%) and 9.46 × 108 CFU/ml (RSD, ±23.3%), respectively, in TSB-Y at 37°C within 20 h. Stained cultures (0.25 μl/ml SYTO9 in TSB-Y) of unstressed and heat-injured Listeria multiplied to means of 1.29 × 109 CFU/ml (RSD, ±22.1%) and 1.11 × 109 CFU/ml (RSD, ±15.0%), respectively. Thus, the deviations of the growth of both species were within the overall standard deviation, indicating no significant influence of the staining protocol on the growth of L. monocytogenes and S. Typhimurium (P ≥ 0.05).
The effect of the transport buffer composition was critical, as the ingredients could have affected the process and impaired the viability of the manipulated bacterial cells. One hundred sixty-five Salmonella cells in four different buffer compositions were compared, and growth was evaluated after SBCM in TSB-Y within 24 h. Table Table11 presents the results of the comparison of the tested buffer systems. The use of 1× PBS (buffer A) led to immediate crystallization of the buffer in and around the capillary notch, due to the high salt concentration, the rise in temperature caused by light, and the very small work volume (<0.1 μl). Microscopic observation became impossible, as was further processing, due to the obstructed capillary. Buffer B (ddH2O) performed without impairing handling, and microscopic observation and a recultivation rate of 80.9% could be obtained. The addition of 1% Tween 80 (buffer C: ddH2O with 1% Tween 80) resulted in improved manipulation performance and a recultivation rate of 92.4%. The last-tested composition, Ringer's solution with 1% Tween 80 (buffer D), yielded a recultivation rate of 94.1%. The significant (P ≤ 0.05) improvement of 13.2% combined with facilitation of the work flow favored this buffer composition, and thus it was used for the manipulation of Listeria and Salmonella.
To evaluate single cell growth, the percentages of viable cells in the initial Listeria and Salmonella cultures were initially determined by BacLight Live/Dead staining. These numbers were then compared with the percentages of viable cells after SBCM by means of growing offspring based on the reproduction of the single cells after manipulation and incubation.
The results are outlined in Table Table2.2. The corresponding percentages of viable L. monocytogenes cells in the initial culture were 79.6% for the lag phase, 75.8% for the log phase, and 53.6% for the stationary phase. The corresponding percentages of viable S. Typhimurium cells in the initial culture were 86.4% for the lag phase, 77.9% for the log phase, and 60.8% for the stationary phase.
L. monocytogenes and S. Typhimurium initial cultures of the respective growth phases were washed and subjected to SBCM as described in Materials and Methods. Offspring cultures of single L. monocytogenes cells grew to a mean OD600 of 1.27 for all growth phases, corresponding to a mean cell count of 1.21 × 109 CFU/ml (RSD, ±9.0%). Offspring cultures of single S. Typhimurium cells grew to a mean OD600 of 1.66 for all growth phases, corresponding to a mean cell count of 7.45 × 108 CFU/ml (RSD, ±6.0%).
The rates of recultivated single cells after SBCM were 84.5%, 70.0%, and 60.0% for Listeria in lag, log, and stationary growth phases, respectively, and 80.0%, 73.3%, and 66.7% for Salmonella in lag, log, and stationary growth phases, respectively. These rates of recultivated single cells after SBCM resulted in overall concordance with the live/dead ratio of the initial cultures providing the source for SBCM. The values for recultivation corresponded to the respective mean values of the live/dead ratios of the initial cultures resulting in P values of ≥0.05 (chi-square test) (Table (Table2).2). This indicates no significant difference in the mean percentage of viable cells compared to the mean percentage of recultivated cells after SBCM. All samples containing the offspring populations were checked for contamination, and monocultures of L. monocytogenes and S. Typhimurium were verified (see also Materials and Methods).
The topic of single bacterial cell growth is a basic question in both fundamental and applied microbiological research. Bacterial growth factors and ecosystems such as biofilms on the one hand and the lag phase of individual bacterial cells under various conditions on the other hand are the two major microbiological research areas where notable efforts have been made in illuminating bacterial single cell growth. In addition, in food-related topics the investigation of the individual lag phase of single cells plays a key role, as very low bacterial contamination levels are expected in this area (8). Approaches applied so far can be classified as follows: MPN-derived methods, mathematical modeling, direct investigation of single cells in a population, and drawing conclusions from the investigation of bacterial populations derived from one supposed starter cell. Common to these approaches is that either a single cell is observed, or the entire population is monitored; the two qualities are never simultaneously included. Elfwing et al. (6) designed a flow chamber capable of monitoring growth of a great number of single cells during the first stages of cell division under a microscope. With this method, single cells could be observed but no conclusions could be made about the entire population. Based on this flow chamber the kinetics of the lag phase of single cells was investigated by several authors (18, 21, 28). In these studies the characteristics of single cell growth were extensively investigated but the growth of single prokaryotic cells without growth factors and intercellular communication was not investigated.
A statistical approach was established by Francois et al. (7). Here an MPN-derived method of obtaining single cells was used by producing series of 2-fold dilutions of bacterial suspensions on microtiter plates starting with a mean of 100 cells. Comparison with a simulation model showed good concordance of this statistical approach. The possibility of attached cells occurring, as is frequently observed in log-phase cultures, has not, however, been taken into account, nor has the possibility of insufficient dilution of potential growth factors. Swinnen et al. (33) and Dens et al. (5) have commented on the theoretical foundations of single cell growth with reference to cell division theory and individual cell-based modeling of the microbial lag phase and comparison of existing stochastic population models such as that of Baranyi (2). Both authors used mathematical tools based on recently published experimental data. The Bioscreen automated microtiter plate incubator was used by Robinson et al. (29) in their study. The results indicated an impact of salt concentration and inoculum size on the growth of L. monocytogenes and reproducible growth of the control cells, which were incubated in TSB-Y at low inoculum quantities. Nevertheless, no details about the setting of these highly diluted inocula were given, and the Bioscreen method has been compared with microscopy in a study by Wu et al. (37), which concludes that there are remarkable variations among microscopic methods. Investigation of the lag phase of microbial growth does not specifically prove the potential for autonomous growth of single prokaryotic cells. Nevertheless, some data such as MPN-derived results suggest this indirectly. In conclusion, these studies aimed at observing growing cells, not their rate of growth within a batch of isolated cells or the influence of growth factors.
Growth factors and cell communication as prerequisites for bacterial growth are linked to the topic of viable but nonculturable (VBNC) bacteria (26). Based on data derived from the investigation of dormant cells and the use of poor media, the roles of intercellular contacts and growth factors such as rpf have been investigated for Rhodococcus rhodochrous, Micrococcus luteus, Mycobacterium spp., other Actinobacteria, and S. Typhimurium (15, 16, 25, 27, 31, 35, 36). The influence of growth factors provided by the actual hosts of the bacteria, such as interleukin-1 (IL-1), IL-6, and granulocyte colony-stimulating factor (G-CSF), has also been demonstrated (15). The relevance of growth factors and intercellular contact for growth regulation in populations is undeniable but not applicable to single isolated bacterial cells, as suggested by evolutionary necessities and the capacity for asexual reproduction. This is all the more relevant as the above-mentioned studies were based on observations of bacterial populations, not single cells.
Thus, it was necessary to develop techniques to isolate single cells, and this has recently been acknowledged (33). Some such efforts have been made by Li et al., Fröhlich and König, Kvist et al., and Ishoy et al. (9, 13, 19, 20). However these techniques focused on the isolation of single cells from heterogeneous mixed populations. Due to the methods used, the rates of growing cells after single cell manipulation varied between 30% and 90% in these studies and the proportions of living and dead cells before the manipulation process were not determined. This mainly resulted from the study designs, which were developed for observation of the composition of mixed populations and not single cell growth.
The specific experimental needs for investigating single bacterial cells led to the development of a new isolation method in this study (Fig. (Fig.1).1). A single cell was to be observed, and the entire population arising from this ancestor cell was to be monitored. The microscopic approach was chosen to ensure the actual presence of one cell. Sterile glass slides with a diameter of 5 mm were established as the carrier to avoid the influence of electrostatic attachment of target cells during pipetting and to ensure the transport of the cells into the enrichment media after manipulation under visual control (Fig. (Fig.3).3). To prevent cross contamination with cells flowing out of optical focus onto the glass carrier, the diameter of the tip of the capillary was chosen to be as small as possible (~0.5 μm). This small-opening capillary diameter led, however, to significant obstruction. Thus, removal of debris from the enrichment broth and particles of the dye solution was very important and necessitated a washing protocol. However, a consequence of this extensive washing procedure was occasionally dilution of cell adhesion and growth-promoting molecules.
Three-way taps were introduced to maintain steady flow of bacteria from the side to minimize stress to the target cells and to enable easy handling. The apparatus described provides the novelty of loading the sample from behind the capillary instead of aspiration (9, 13). Due to possible unfavorable effects during the micromanipulation procedure caused by a small capillary notch and extended residence time, the composition of the transport buffer was of major importance. Previous studies suggested that osmotic stress mainly accounts for low recovery of single cells during micromanipulation procedures (10, 13). Therefore, an optimal osmotic-stress-reducing transport buffer system was established. Commonly used Ringer's solution is well known to protect bacterial spheroblasts against osmotic lysis, and its use, in combination with the positive effects of the surfactant Tween 80, resulted in an overall recultivation rate of 85.8% for L. monocytogenes and S. Typhimurium (3, 12).
For manipulation of individual bacterial cells, visualization is necessary. In this study one of the main modifications to already-described micromanipulation systems was the use of bacterial stain instead of fluorescent labeling or an inverse phase-contrast microscope (9, 13). Initial experiments were performed with green fluorescent protein-labeled L. monocytogenes strains, but these resulted in weak fluorescence emission (data not shown). Thus, the use of SYTO9 as the bacterial stain was tested instead, and the results indicated its suitability and potential applicability for broad-range use with different bacterial species.
L. monocytogenes and S. Typhimurium were chosen as model organisms for Gram-positive and Gram-negative bacteria, respectively. The combination of the washing procedure and the SBCM technique eliminated the influence of deviating effects caused by dilution series and the possibility of coexisting growth-stimulating factors and cellular contacts. In this way the tested S. Typhimurium and Listeria monocytogenes cells were forced to grow in a solitary and unaffected manner.
Comparison of the distributions of living and dead cells within the population before the isolation of single cells and the growth of these separated cells after manipulation resulted in surprisingly good concordance. Single L. monocytogenes and S. Typhimurium cells from all investigated physiological states grew according to their live/dead distributions within the original culture. The concentrations of the cultures after 24 h of incubation, as well as the deviations of the single values, indicate unaffected optimal growth.
Optimal growth also enhanced the frequency of appearance of cells cohering in duplicate at the end of cell division prior to their isolation by micromanipulation. These cells have to be considered when the ratio of living to dead cells is estimated and are not in the pool of single cells to be isolated when SBCM is applied. Duplets or longer chains of bacterial cells resulting from cell division bias the results of MPN-based methods and other culture-dependent quantitative microbiological methods.
Mukamolova et al. (23) reported sufficient removal of rpf from M. luteus cultures in lactate minimal medium following 5-fold washing, and this resulted in a dilution of 3.2 × 10−6. The dilution factor obtained during the cell washing procedure was 1 × 10−10 in this study. With regard to these data, sufficient removal of any growth factor within the transport buffer of the system could be assumed. The data presented give no indication of additional growth factors for the propagation of the isolated L. monocytogenes and S. Typhimurium cells.
The question as to whether “a critical cell concentration is necessary in an inoculum, below which cells cannot grow,” as postulated by Kaprelyants and Kell (15), has thus been answered for the first time. The data presented include unstressed cells in rich media, which constitute the basis for further investigations. These include the roles of various factors which necessarily influence bacterial cell growth in different environments.
The method presented in this study is sufficient to prove the possibility of independent growth of single isolated bacterial cells, in the absence of growth factors or intercellular contacts. In comparison with recent approaches, clear evidence is provided without stochastic effects or deficiencies of experimental design. Over 450 single cells have been manipulated in this way without any contamination, attesting to easy handling of the apparatus and efficiency of the cleaning procedures. Compared with sample numbers investigated in earlier studies, application of this modified SBCM technique is apparently less time-consuming and more suitable for routine laboratory use. The study also solidly validates the method, recommending it for further basic research as well as for routine applications, including validation of enrichment media and precise performance with low-level contamination.
We gratefully acknowledge the financial support of the Christian Doppler Society for facilitating this work.
We thank Sonja Klinger providing the engineering drawing of the instrument.
Published ahead of print on 19 February 2010.