The role of assortment (spatial segregation) in evolutionary scenarios can be examined using simple microbial systems in the laboratory that allow controlled adjustment of spatial distribution. By modifying the founder cell density, various assortment levels can be visualized using fluorescently labelled bacterial strains in the colony biofilms of Bacillus subtilis.
Microbes provide an intriguing system to study social interaction among individuals within a population. The short generation times and relatively simple genetic modification procedures of microbes facilitate the development of the sociomicrobiology field. To assess the fitness of certain microbial species, selected strains or their genetically modified derivatives within one population, can be fluorescently labelled and tracked using microscopy adapted with appropriate fluorescence filters. Expanding colonies of diverse microbial species on agar media can be used to monitor the spatial distribution of cells producing distinctive fluorescent proteins.
Here, we present a detailed protocol for the use of green- and red-fluorescent protein producing bacterial strains to follow spatial arrangement during surface colonization, including flagellum-driven community movement (swarming), exopolysaccharide- and hydrophobin-dependent growth mediated spreading (sliding), and complex colony biofilm formation. Non-domesticated isolates of the Gram-positive bacterium, Bacillus subtilis can be utilized to scrutinize certain surface spreading traits and their effect on two-dimensional distribution on the agar-solidified medium. By altering the number of cells used to initiate colony biofilms, the assortment levels can be varied on a continuous scale. Time-lapse fluorescent microscopy can be used to witness the interaction between different phenotypes and genotypes at a certain assortment level and to determine the relative success of either.
In the last decades, microbes have been recognized as social communities associated with various ecosystems on earth1,2. In contrast to planktonic cultures used in general laboratory practice, microbes in the environment show a diverse range of spatial community structures depending on the ecological setting. Simple microbial systems can be utilized to understand the consequence of spatial structures on the evolution of social interactions3,4. Publications in the last 2-3 years using both eukaryotic and prokaryotic model systems highlighted the impact of spatial structures on the stability of cooperation within microbial populations5-8. Additionally, obligate interactions among microbes, e.g. metabolic cross-feeding, might also alter the spatial distribution of interacting partners9-11. The influence of spatial structure in these studies is mostly examined using surface attached sessile cells inhabiting the so-called biofilms or in colonies growing on the surface of an agar medium. Genetic drift resulting in high spatial assortment can be observed in microbial colonies where nutrient depletion at the edge of a cell division mediated expansion results in series of genetic bottlenecks that causes high local fixation probability for certain clonal linages12. Genetic drift can be therefore employed to examine the role of spatial segregation in microbial colonies.
In the environment, biofilms are multispecies communities surrounded by self-produced polymeric matrix13. Biofilm structure, function and stability depend on a complex network of social interactions where bacteria exchange signals, matrix components and resources, or compete for space and nutrients using toxins and antibiotics. Bacillus subtilis is a soil dwelling and root-colonizing bacterium that develops highly organized biofilm communities14. In analogy to social insects, B. subtilis cells employ a division of labor strategy, developing subpopulations of extracellular matrix producers and cannibals, motile cells, dormant spores and other cell types15,16. The differentiation process is dynamic and can be altered by environmental conditions17,18.
Strategies of surface colonization by bacteria can be easily manipulated under laboratory conditions by modifying the agar concentration in the growth media. At low agar levels (0.2-0.3%), bacteria harboring active flagella are able to swim, while semi-solid agar (0.7-1% agar) facilitates flagellum driven community spreading, called swarming19-21. In the absence of flagellum, certain bacterial strains are able to move over semi-solid medium via sliding, i.e. growth dependent population expansion facilitated by exopolysaccharide matrix and other secreted hydrophobin compounds22-24. Finally, bacteria which are capable of biofilm development form architecturally complex colonies on hard agar medium (1.2-2%)14,17,25. While these traits are examined in the laboratory by precisely adjusting the conditions, in natural habitats these surface-spreading strategies might transit gradually from one to another depending on the environmental conditions26. While single cell based motility is critical during initiation of biofilm development at the air-liquid interface in both Gram-positive and -negative bacteria27, complex colony biofilms of B. subtilis are not affected by deletion of flagellar motility28. However, spatial organization during the development of B. subtilis colony biofilms depends on the density of the bacterial inoculum used to initiate the biofilm8.
Here, we use B. subtilis to show that spatial segregation during surface colonization depends on the mechanism of population level motility (i.e. swarming or sliding), and colony biofilm development depends on the founder cell density. We present a fluorescent microscopy tool that can be applied to continuously monitor microbial biofilm growth, surface colonization and assortment at the macro scale. Further, a quantification method is presented to determine the relative strain abundance in the population.
The availability of a fluorescent toolbox for bacteria facilitates not only the study of heterogeneous gene expression30,31 and protein localization32, but also the analysis of spatial distribution of strains within a population8. Fluorescent markers with sufficiently different excitation and emission wavelengths allow to distinctly localize two strains that otherwise are indistinguishable from each other when mixed. The described protocol can be employed for observing the population dynamics in mixed cultures, e.g. competition experiments or synergism between strains or species. The ability to determine the relative abundances of fluorescently labelled strains in a mixed population is not restricted to surface attached swarming, sliding, or biofilm colonies, but can also be used for other multicellular biofilm systems, including submerged, flow cell or air-medium interface biofilms27,33-35.
While the presented technique is a powerful tool to detect spatial distribution of strains and design competition experiments, it also allows following gene expression heterogeneity in expanding colonies. The culturing conditions described here apply for B. subtilis and the exact parameters for expansion on agar media might require optimization for other species or strains20. Placing the samples in an incubation chamber while imaging permits the experimenter to follow the population dynamics in time, although attention should be given to the humidity level within the chamber during the incubation.
The techniques described here also require the genetic modification of the examined bacterial strains so that the strains express fluorescent markers which can be distinguished from each other. Moreover, besides having distinct excitation and emission spectra, it is recommended that the two chosen fluorescent markers have similar quantum yields (i.e. ratio of absorbed photons that are emitted) and are expressed in a comparable level. In addition, relative intensity changes in time can be measured and normalized to an early time-point of an experiment. The relative increase or decrease can be then compared between different fluorophores with different quantum efficiencies. For the presented experimental system, different green- and red-fluorescent proteins were tested previously36,37 to select for the most optimal fluorescent pairs that can be detected in B. subtilis. The optimal exposure time should be determined for each fluorescent protein and sample. Certain cell densities or multiple layers of cells might be required to detect the signal efficiently within the population. Certain fluorescent proteins might have low intensities in the bacterial cells due to inefficient expression and/or translation of the protein and thus low quantum yield. Such inefficient fluorescent markers could reduce the sensitivity of the system and extend the time needed to detect the bacterial strains possibly resulting in cytotoxicity by the excitation light. The fluorescent intensities can be accordingly modified by altering the promoter used to express the fluorescent reporter coding gene. An expression level that is too high could result in unnecessary overproduction of the fluorescent protein leading to detrimental fitness costs for the bacterium. When performing competition experiments, one should consider the cost of particular fluorescent protein production in the cells. Control experiments, where the fluorescent markers are swapped between competed strains or where two isogenic strains differing only in their fluorescent markers are competed against each other, are always required to determine any bias toward one marker. The lifetimes of the fluorescent proteins within the cells might also affect the measured intensity. In addition, the autofluorescence of certain bacterial species might require the use of different fluorescent markers other than described here.
To precisely determine the spatial distributions and abundances of the distinct bacterial strains, the background signal originating from the first fluorescent protein while using the fluorescence filter for the second fluorescent marker and vice versa should be individually tested on monoculture samples (containing bacteria producing only one marker). This allows the subtraction of overlapping fluorescent signal intensities. Importantly, as the stereomicroscope records the fluorescence signal from above the expanding colony, the presented protocol is convenient to determine the spatial arrangement in two dimensions. The architecture of the expanding bacterial population could result in varying fluorescence levels (i.e. wrinkle-like structures might contain more cells displaying higher local fluorescence intensities). Therefore, the described analysis of the images determines the spatial distribution, but not the abundance of the strains within a certain location. Previous protocols described the sample preparation for swarming20 or fluorescence imaging of population dynamics in bacterial colonies38, but our protocol combines these techniques. Other microscopy techniques that permit the observation of three dimensional resolution of the population structure (e.g. confocal laser scanning microscopy39,40 or structured illumination microscopy41) can be applied for samples with increased structural complexities. These additional techniques also support single cell based detection of the strains31 that is not available using stereomicroscopes.
The authors have nothing to disclose.
This work was funded by grant KO4741/3-1 from the Deutsche Forschungsgemeinschaft (DFG). Further, the laboratory of Á.T.K. was supported by a Marie Skłodowska Curie career integration grant (PheHetBacBiofilm) and grant KO4741/2-1 from DFG. T.H., A.D., R.G.-M., and E.M. were supported by International Max Planck Research School, Alexander von Humboldt foundation, Consejo Nacional de Ciencia y Tecnologìa-German Academic Exchange Service (CONACyT-DAAD), and JSMC fellowships, respectively.
Lennox Broth (LB) | Carl Roth GmbH | X964 | |
Agar-agar, Kobe I | Carl Roth GmbH | 5210 | |
Petri dish (90 mm diameter) | any | NA | Use Petri dishes without ventillation cams |
Petri dish (35 mm diameter) | any | NA | Use Petri dishes without ventillation cams |
Difco Nutrient Broth | BD Europe | 234000 | |
KCl | any | NA | |
MgSO4 7H2O | any | NA | |
Ca(NO3)2 4H2O | any | NA | |
MnCl24H2O | any | NA | |
FeSO4 | any | NA | |
D-Glucose | any | NA | |
Fluorescence AxioZoom V16 time-lapse microscope | Carl Zeiss Microscopy GmbH | see bellow detailed description | |
AxioZoom V16 Microscope body | Carl Zeiss Microscopy GmbH | 435080 9030 000 | |
Phototube Z 100:0 for Axio Zoom V16 | Carl Zeiss Microscopy GmbH | 435180 9020 000 | without eyepieces |
Fluar Illuminator Z mot Fluorescence intermediate tube for Axio Zoom.V16 | Carl Zeiss Microscopy GmbH | 435180 9060 000 | |
Controller EMS 3 | Carl Zeiss Microscopy GmbH | 435610 9010 000 | |
System Control Panel SYCOP 3 | Carl Zeiss Microscopy GmbH | 435611 9010 000 | |
Reflector module Z | Carl Zeiss Microscopy GmbH | 435180 9160 000 | For Fluar Illuminator Z mot on Axio Zoom.V16 and SYCOP 3 |
Filter set 38 HE eGFP shift free (E) | Carl Zeiss Microscopy GmbH | 489038 9901 000 | EX BP 470/40, BS FT 495, EM BP 525/50 |
Filter set 63 HE mRFP shift free (E) | Carl Zeiss Microscopy GmbH | 489063 0000 000 | EX BP 572/25, BS FT 590, EM BP 629/62 |
Mount S | Carl Zeiss Microscopy GmbH | 435402 0000 000 | |
Objektive PlanApo Z 0,5x/0,125 FWD 114mm | Carl Zeiss Microscopy GmbH | 435280 9050 000 | 164mm parfocal length; M62x0.75 thread at front |
Coarse/fine drive with profile column | Carl Zeiss Microscopy GmbH | 435400 0000 000 | 490 mm, 10kg load capacity, compatible with stand bases 300/450 |
Stand base 450 | Carl Zeiss Microscopy GmbH | 435430 9902 000 | |
Cold-light source Zeiss CL 9000 LED CAN | Carl Zeiss Microscopy GmbH | 435700 9000 000 | |
CAN-bus cable | Carl Zeiss Microscopy GmbH | 457411 9011 000 | 2.5 m length |
Slit-ring illuminator | Carl Zeiss Microscopy GmbH | 417075 9010 000 | d=66 mm |
Flexible light guide 1500 | Carl Zeiss Microscopy GmbH | 417063 9901 000 | 8/1000 mm |
Illumination Adapter for light guide | Carl Zeiss Microscopy GmbH | 000000 1370 927 | |
Lightguide HXP with liquid fill | Carl Zeiss Microscopy GmbH | 000000 0482 760 | ø3mm x 2000mm |
Camera Adapter 60N-C | Carl Zeiss Microscopy GmbH | 426113 0000 000 | 2/3" 0.63x |
High Resolution Microscopy Camera AxioCam MRm Rev. 3 FireWire | Carl Zeiss Microscopy GmbH | 426509 9901 000 | |
AxioCam FireWire Trigger Cable Set | Carl Zeiss Microscopy GmbH | 426506 0002 000 | for direct shutter synchronization |
ZEN pro 2012 | Carl Zeiss Microscopy GmbH | 410135 1002 120 | Blue edition, requires min. Windows 7 64-bit |
ZEN Module Time Lapse | Carl Zeiss Microscopy GmbH | 410136 1031 110 | |
Standard Heating Stage Top Incubator | Tokai Hit | INUL-MS1-F1 | |
Zeiss Stereo Microscope Base Adapter | Tokai Hit | MS-V12 | |
Softwares | |||
Image J | National Institute of Health, Bethesda, MD, USA | v 1.49m | |
BioVoxxel plugin | BioVoxxel | http://www.biovoxxel.de/development/ |