Microfluidics expanding the frontiers of microbial ecology

Abstract

Microfluidics has significantly contributed to the expansion of the frontiers of microbial ecology over the past decade by allowing researchers to observe the behaviors of microbes in highly controlled microenvironments, across scales from a single cell to mixed communities. Spatially and temporally varying distributions of organisms and chemical cues that mimic natural microbial habitats can now be established by exploiting physics at the micrometer scale and by incorporating structures with specific geometries and materials. In this article, we review applications of microfluidics that have resulted in insightful discoveries on fundamental aspects of microbial life, ranging from growth and sensing to cell-cell interactions and population dynamics. We anticipate that this flexible multidisciplinary technology will continue to facilitate discoveries regarding the ecology of microorganisms and help uncover strategies to control microbial processes such as biofilm formation and antibiotic resistance.

Keywords: antibiotics; gradients; microenvironments; population dynamics; single-cell analysis; surface interactions.

Figure 1. Microfluidics provides a powerful platform for microbial ecology studies

Multiple features of natural microbial habitats can be included in microfluidic studies of the life of microbes: Multiple-inlet (1) and the incorporation of hydrogels (e.g., agarose; 2) to impose spatial gradients; constrictions and topological features to study the effect of flow in porous-like environments (3); funnel-shaped barriers to separate and concentrate swimming microbes (4); microfabricated topography to study bacterial adhesion (5); single-cell confinements to study growth and replication (6); population confinements to investigate competition and population dynamics (7).

Figure 2. Chemical gradients reveal microbial navigation strategies

a) Swimming trajectories of the dinoflagellate Oxyrrhis marina in response to a pulse of chemoattractant (grayscale background). Figure modified from Seymour et al. (2010b) with permission. b) Time evolution of the spatial distribution of a pathogen population (Vibrio coralliilyticus) exposed to a diffusing coral mucus (Pocillopora damicornis) gradient in a microfluidic channel. Figure modified from Garren et al. (2013) with permission. c) Experimental pulses of Escherichia coli traveling across a microfluidic channel at a constant propagation speed. Figure modified from Saragosti et al. (2010) with permission. Scale bars, 500 μm.

Figure 3. Fine control over microscale topography sheds light on microbe-surface interactions

a) Fluorescent Escherichia coli cells trapped and concentrated in arrow-shaped microfluidic ratchets. Figure modified from Hulme et al. (2008) with permission. b) Series of funnel arrays used to separate and concentrate motile from non-motile Escherichia coli cells. Figure modified from Galajda et al. (2007) with permission. c) Rectification of algal cells (Chlamydomonas reinhardtii) locomotion in microfluidic ratchets via secondary scattering. Figure modified from Kantsler et al. (2013) with permission. d) Fluorescent images (left-hand side) of bacterial cells, Pseudomonas aeruginosa, adhering to structured surfaces at decreasing (from left to right) spacing between posts, and a cross-sectional SEM image of the same process (right-hand side; cells are false-colored to highlight their orientation). Figure modified from Hochbaum & Aizenberg (2010) with permission. e) Pictorial visualization of the mechanism underlying the formation of Pseudomonas aeruginosa biofilm streamers (rendering from confocal images) in curved microchannels. Figure modified from Rusconi et al. (2011) with permission. f) Pseudomonas aeruginosa biofilm streamers highlighted by the subsequent injection of red-fluorescent cells. Figure modified from Drescher et al. (2013) with permission. g) Patterns of biofilm growth (Escherichia coli auto-fluorescence in green) and fluid flow (in red, slightly shifted to the right) in a microfluidic device modeling a porous soil environment. Figure modified from Durham et al. (2012) with permission. Scale bars, 2 μm (d), 20 μm (a,c), 200 μm (b,f,g).

Figure 4. Single-cell microconfinementa open new doors for understanding growth and persistence

a) Schematic representation, lineage tree, and snapshot of the microfluidic “mother machine”. Figure modified from Wang et al. (2010) with permission. b) Time-lapse images of a high-persistence mutant of Escherichia coli growing in a microfluidic chamber and exposed to ampicillin, showing the location of slowly-growing persisters (red arrows). Figure modified from Balaban et al. (2004) with permission. c) Morphogenesis of E. coli in ultra-thin microfluidic channels (dashed lines show approximate boundaries). Figure modified from Männik et al. (2009) with permission. Scale bars, 10 μm.

Figure 5. Multiple-cell microconfinements reveal new population-scale microbial processes

a) Bacterial growth and ordering in a quasi-2D microfluidic open channel. Figure modified from Volfson et al. (2008) with permission. b) Vibrio harveyi accumulation in a microfabricated maze inducing quorum sensing (dark-field image on the left; photon-counting image of the intrinsic quorum sensing luminescence on the right). Figure modified from Park et al. (2003b) with permission. c) Wild-type Escherichia coli collapsing into confining microfluidic chambers (experiments on the left; simulations on the right). Figure modified from Park et al. (2003a) with permission. d) Fluorescence images of competing Escherichia coli populations (GASP mutant in red; wild-type cells in green) in microhabitat patches. Figure modified from Keymer et al. (2008) with permission. e) Large array of connected microchambers showing the rapid emergence of antibiotic resistance within an antibiotic gradient for different inoculation densities and different times. Figure modified from Zhang et al. (2011) with permission. Scale bars, 20 μm (a,d), 200 μm (b), 500 μm (c), 5 mm (e).

Figure 6. Multi-layer microfluidic devices allow co-culturing

a) Schematic drawing of the gut-on-a-chip device showing the porous membrane lined by gut epithelial cells with or without mechanical strain exerted by suction. Figure modified from Kim et al. (2012b) with permission. b) Schematic drawing of the microfluidic device used to co-culture three species of soil bacteria by imposing spatial structure on three culture wells and providing a chemical communication channel. Figure modified from Kim et al. (2008) with permission.