The Ecology of Bacterial Individuality

Individual-based microbial ecology (IBME) is a developing field of study in need of experimental tools to quantify the individual experience and performance of microorganisms in their natural habitats. We describe here the conception and application of a single-cell bioreporter approach with broad utility in IBME. It is based on the dilution of stable green fluorescent protein (GFP) in dividing bacteria. In the absence of de novo synthesis, GFP fluorescence of a daughter cell approximates half of that of its mother, from which follows that the fluorescence of a progeny cell is a quantitative measure for the reproductive success of its ancestor. To test this concept, we exposed GFP-filled bacteria to different degrees of environmental heterogeneity and assessed how this affected individual cells by the analysis of GFP content in their progeny. Reporter bacteria growing in rich medium in a shaking flask showed no variation in reproductive success, confirming that life in a broth is experienced much the same from one bacterium to the next. In contrast, when reporter bacteria were released onto plant leaf surfaces, representing a microscopically heterogeneous environment, clear intrapopulation differences in reproductive success were observed. Such variation suggests that individual cells in the founding population experienced different growthpermitting conditions, resulting in unequal contributions of individual bacteria to future offspring and population sizes. Being able to assess population changes bottom-up rather than top-down, the bioreporter offers opportunities to quantify single-cell competitive and facilitative interactions, assess the role of chance events in individual survivorship and reveal causes that underlie individualbased environmental heterogeneity. The Ecology of Bacterial Individuality 20 Introduction Environmental heterogeneity, defined as spatial and temporal variation in the physical, chemical and biological environment, is a fundamental property of ecosystems (Scheiner and Willig 2008). At the scale of individual organisms, it affects the ability to survive, reproduce, co-exist and interact with other organisms. For plants and animals, environmental impact can be assessed and quantified relatively simply at the level of individual organisms (Melbourne, Cornell et al. 2007). In microbial ecology, however, the effect of environmental variability on microbial activity and diversity is commonly assessed at a scale that is several orders of magnitude greater than the dimensions of the microorganisms under study (Hellweger and Bucci 2009). This ‘coarse-grained’ (Templeton and Rothman 1978) approach to environmental heterogeneity suffers from the averaging effect that is typical of many population-based approaches (Brehm-Stecher and Johnson 2004). It is increasingly being recognized that ‘fine-grained’ environmental heterogeneity, that is the one experienced by microscopic individuals at the micrometer-scale is a key factor in explaining microbial activity, diversity, distribution and evolution (Davey and Winson 2003; Green and Bohannan 2006; Prosser, Bohannan et al. 2007; Davidson and Surette 2008). However, due to the relative lack of tools to probe environments for micrometer-scale differences in physical, chemical or biological variables, little is known about the heterogeneity that individual micro-organisms are exposed to and, more importantly, how this affects their activity and reproductive success. Bioreporter technology (Leveau and Lindow 2001; Leveau and Lindow 2002; Harms, Wells et al. 2006; Leveau 2006; Tecon and van der Meer 2006) relies on microorganisms themselves to report on local environmental conditions. Many of these bioreporters involve the conditional expression of green fluorescent protein (GFP), a reporter that can be quantified with relative ease in individual cells by fluorescence image microscopy (Jaspers, Meier et al. 2001; Leveau and Lindow 2001) or flow cytometry (Axtell and Beattie 2002; Maksimow, Hakkila et al. 2002; Harms, Wells et al. 2006; Roostalu, Joers et al. 2008). When properly calibrated, the GFP signal becomes a measure for exposure to a particular environmental stimulus. For example, Leveau and Lindow (2001a) used a fructose-responsive promoter fused to the gene for GFP to probe the availability of this sugar to bacteria on plant leaf surfaces, also known as the phyllosphere (Leveau 2006). Temporal and spatial variation in single-cell green fluorescence indicated substantial heterogeneity in the availability of fructose to individual leaf colonizers (Leveau and Lindow 2001). Such heterogeneity has also been reported for other nutrients or stimuli that leaf bacteria are exposed to, including iron (Joyner and Lindow 2000), water (Axtell and Beattie Individual-based reproductive success of bacteria 21 2002), UV light (Gunasekera and Sundin 2006) and phenolic compounds (Sandhu, Halverson et al. 2007). While bacterial bioreporters, such as the ones described above, are useful in micrometer mapping of differences in the bacterial experience of single environmental variables, they cannot communicate how each of those variables, individually or jointly, impact the fate of bacteria in the environment under study. We therefore designed a bioreporter tool that describes micrometer-scale environmental heterogeneity in general terms, that is as a sum of all variables expressed into a single, quantifiable effect on the bacterium. The bioreporter we introduce here records environmental heterogeneity in terms of past reproductive success. In concept, it is based on the observation that upon cell division, GFP in a bacterial cell is distributed in a predictable manner between its two daughter cells (Rosenfeld, Perkins et al. 2006; Roostalu, Joers et al. 2008): one division leaves cells approximately half as green fluorescent as their parent, two divisions one-fourth as fluorescent, and so on. Thus, the GFP content of an individual offspring cell becomes a quantifiable measure of reproductive success. This approach resembles the method that was used (Mailloux and Fuller 2003) to estimate in situ doubling times for bacteria released into an aquifer after staining them with carboxy-fluorescein diacetate succinimidyl ester, a fluorescent protein stain that dilutes from the bacteria with every cell division. However, whereas these authors were interested solely in population averages of in situ growth, we tested our GFP-based bioreporter by exposure to microscopic conditions of low (that is, LB broth) and high (that is, the phyllosphere) environmental heterogeneity to reveal sub-population differences in the reproduction of single bacteria. The implications of our findings extend broadly to studies on other microbial habitats dealing with the question of how individual bacteria in founder populations differ in their contribution to future population sizes. Materials and methods Bacterial strains and culture conditions Erwinia herbicola 299R JBA28 (pCPP39) (Eh299R::JBA28 (pCPP39)) (Leveau and Lindow, 2001b) carries a chromosomal mini-Tn5-Km transposon insertion that expresses stable GFP from a LacIq-repressible PA1/O4/O3 promoter fusion to gfpmut3. The transposon confers resistance to kanamycin. The strain also harbors plasmid pCPP39, which confers tetracycline resistance and harbors a lacIq gene for control of PA1/O4/O3 activity, and thus GFP production by isopropyl-β-D-thiogalactopyranoside (IPTG). Bacteria were cultivated at 28 °C on LB agar or in LB broth at 300 r.p.m. Where appropriate, IPTG, kanamycin, or tetracycline were added to final concentrations of 1mM, 50 or 15 mg ml-1, respectively. Optical densities of bacterial cultures The Ecology of Bacterial Individuality 22 were measured at 600nm (OD600) in a Unico 1100 spectrophotometer (Unico, Dayton, NJ, USA). GFP-loading, release and recovery of bioreporter Eh299R::JBA28 (pCPP39) Exponentially growing cells of Eh299R::JBA28 (pCPP39) were diluted 300-fold into fresh LB broth containing 1mM IPTG and grown to mid-exponential phase. These GFP-loaded cells were used to inoculate plant leaves (see below) or LB broth. In the latter case, 25 ml of LB was inoculated with 200 ml of GFP-loaded bacteria and incubated at 28°C and 300 r.p.m. Samples were taken every 30 min to measure OD600 and to collect bacteria for fixation (see below). For plant inoculations, GFP-loaded bacteria were diluted in Milli-Q water to a final concentration of 5 x 104 colony-forming units ml-1. Leaves of 12–14-day old Phaseolus vulgaris plants (green snap bean, variety Blue Lake Bush 274) were inoculated by brief submersion into this bacterial suspension, shaken to dispose of excessive liquid and transferred to a closed translucent box for high-humidity incubation at 21°C. At different time intervals, two leaves were transferred to a 50-ml Falcon tube with 20 ml 1 x PBS buffer, vortexed briefly and sonicated for 7 min. Part of the bacterial cells in the leaf washing was plated on agar for counting colony forming units, whereas the rest was collected on 0.2-μm Durapore filters (Millipore, Amsterdam, The Netherlands), recovered by vortexing for 15s in 1ml 1 x PBS, and fixed (see below). Fluorescence in situ hybridization, fluorescence microscopy and image cytometry Bacterial cells collected from LB broth or plant leaves were fixed as described previously (Leveau and Lindow 2001) and stored at -20 °C in 50% 1 x PBS/50% ethanol for no longer than 2 weeks. To distinguish cells of Eh299R::JBA28 (pCPP39) from indigenous bacteria on the bean leaves, fixed leaf washings were subjected to fluorescence in situ hybridization using an Eh299R-specific, TAMRA-labeled probe (Brandl, Quinones et al. 2001) at a final concentration of 5.5 ng μl-1. LB-or leaf-exposed cells were examined with an Axio Imager.M1 (Zeiss, Oberkochen, Germany) using 470/20 nm excitation for the visualization of GFP and 546/6 nm for TAMRA. Digital images were captured at 1000-fold magnification with an AxioCam MRm camera (Zeiss) in phase contrast and through a 525/25nm (GFP) or 575–640 nm (TAMRA) filter set. Using AxioVision 2.6 Software (Zeiss), single-cell GFP fluorescence was quantified as the mean-pixel intensity (Leveau and Lindow 2001), and expressed in units of Sfere (Standardized fluorescence reference), where 1 milliSfere equals one-thousandth of the average mean-pixel intensity of 1-mm Tetraspeck Fluorescent Microsphere Standards (Molecular probes, Eugene, OR, USA). Data analyses and simulations were performed in Microsoft Excel 2003 (Microsoft CorIndividual-based reproductive success of bacteria 23 poration, Redmond, WA, USA). Computer simulations For computer simulations presented in Figure 3a, reproductive success was calculated for 100 cells with a green fluorescence (GF) equal to X/2t, in which X equals the Excel formula ‘=norminv(rand(),1000,250)’. Figure 3b shows the temporal changes in reproductive success of individual bacteria as a function of t from a population of 90 cells with GF = X and 10 * 2t cells with GF = X/2t. Figure 3c shows the reproductive success of 20 bacteria with GF = X, 20 * 20.125 * t bacteria with GF = X/20.125* t, 20 * 20.25 * t bacteria with GF = X/20.25 * t, 20 * 20.5 * t bacteria with GF = X/20.5 * t and 20 * 2t bacteria with GF = X/2t. Figure 3d shows the reproductive success in a population of 20 bacteria with GF = X, 20 * 2t bacteria with GF = X/2t (t ≤ 1) or 20 * 21 bacteria with GF = X/21 (t>1), 20 * 2t bacteria with GF = X/2t (t>2) or 20 * 22 bacteria with GF = X/22 (t>2), 20 * 2t bacteria with GF = X/2t (t>3) or 20 * 23 bacteria with GF = X/23 (t>3) and 20 2t bacteria with GF = X/2t. Results GFP dilution is a quantitative measure for reproductive success A previously formulated mathematical model of GFP expression in bacteria (Leveau and Lindow 2001) predicts that in the absence of de novo synthesis, GFP dilutes from dividing cells at a rate equal to growth rate μ. We verified this prediction here using Eh299R::JBA28 (pCPP39), which accumulates stable GFP in the presence of the synthetic inducer IPTG. Upon transfer of IPTG-induced GFP-loaded cells to broth that lacked IPTG, GFP fluorescence declined exponentially and at a rate that was not significantly different from μ (Figure 1). No such decrease in fluorescence was observed when GFP-loaded cells were transferred to sterile Milli-Q water (data 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 0.00 0.05 0.10 0.15 0.20 0.25 0.30 0.35 0 200 400 600 80