Article series: New technologies: methods and applications

Live from under the lens: exploring microbial motility with dynamic imaging and microfluidics

Journal name:
Nature Reviews Microbiology
Volume:
13,
Pages:
761–775
Year published:
DOI:
doi:10.1038/nrmicro3567
Published online

Abstract

Motility is one of the most dynamic features of the microbial world. The ability to swim or crawl frequently governs how microorganisms interact with their physical and chemical environments, and underpins a myriad of microbial processes. The ability to resolve temporal dynamics through time-lapse video microscopy and the precise control of the physicochemical microenvironment afforded by microfluidics offer powerful new opportunities to study the many motility adaptations of microorganisms and thereby further our understanding of their ecology. In this Review, we outline recent insights into the motility strategies of microorganisms brought about by these techniques, including the hydrodynamic signature of microorganisms, their locomotion mechanics, chemotaxis, their motility near and on surfaces, swimming in moving fluids and motility in dense microbial suspensions.

At a glance

Figures

  1. Microbial flow fields and motility mechanics.
    Figure 1: Microbial flow fields and motility mechanics.

    a | The flow field produced by a swimming Escherichia coli cell15. E. coli is a 'pusher', with the flagella at the back pushing the cell head, resulting in fluid moving away from the cell along the swimming direction and towards the cell from the sides (black streamlines). Colours represent magnitude of flow velocity. The arrows for the zoomed-in box show the forces exerted by the bacterium on the fluid. b | Planar projection of the 3D trajectory of an E. coli cell swimming in a 'run-and-tumble' pattern84. Circular markers are cell head positions at 79 ms intervals. cf | Many marine bacteria reorient by the 'flick' motility adaptation, an off-axis deformation of the flagellum that enables certain bacteria with a single flagellum to change their direction of swimming1, 2 (Supplementary information S1 (movie)). c | Schematic of the flick, which occurs at the onset of a forward run when the 'hook' is under compression. d | A trajectory of the bacterium Vibrio alginolyticus swimming in a 'run-reverse-flick' motility pattern. Circular markers represent cell head positions at 1 ms intervals. e | The image sequence, captured with high-intensity dark-field microscopy, shows the kinematics of the flagellum (magenta) during a flick2, for the trajectory shown in panel d. The flick occurs ~10 ms after the transition from backward to forward swimming. f | The flick results from a buckling instability of the hook, which for a pusher cell is compressed by the drag on the cell head and the propulsion force from the flagellum2. For a 'puller' cell the hook is under tension and cannot buckle. The image in part a is adapted from Ref. 15, National Academy of Sciences. The image in part b is adapted from Ref. 84, Nature Publishing Group. Figure parts d–f are adapted from Ref. 2, Nature Publishing Group.

  2. Microbial chemotaxis.
    Figure 2: Microbial chemotaxis.

    a | Snapshot of the phycosphere, the organic-matter-rich microzone surrounding individual phytoplankton cells (see also Supplementary information S2 (movie)). The maximum intensity projection image shows trajectories of natural marine bacteria (blue) strongly accumulating around a lysing Chaetoceros diatom by chemotaxis29. b | Schematic of the microfluidic 'microinjector' used to study microbial behavioural responses, particularly chemotaxis, to ephemeral resource hotspots41, 42, 43, 44, 45. The resource is ephemeral because it is initially in the form of a band of attractant that rapidly diffuses outwards. Bacterial locations and trajectories can be captured by time-lapse imaging. c | Bacterial pathogens can detect their coral hosts by chemotaxis. Motility is prevalent among putative coral pathogens, and microfluidic experiments showed that Vibrio coralliilyticus exhibits a strong chemotactic response towards coral mucus (yellow shading), which diffuses from the coral surface45. This response is likely to be a mechanism used by the pathogen to locate its coral host and is exacerbated under warming conditions45. d | Model for the coexistence of two closely related populations of marine bacteria based on trade-offs in their spatial behaviours48. Dynamic imaging in microfluidic devices showed that both populations of Vibrio cyclitrophicus use chemotaxis to migrate towards particles, but only one population attaches and forms biofilms on particles (red cells). The other population (blue cells) remains near the particle and retains the flexibility of rapidly migrating to new, more nutrient-rich particles. This represents the first microbial example of a competition–dispersal trade-off. PDMS, polydimethylsiloxane. The image in part c is adapted from Ref. 45, Nature Publishing Group.

  3. Microbial interactions with surfaces.
    Figure 3: Microbial interactions with surfaces.

    a | Bacteria near a surface often swim in circles57. The rotation of the flagellar bundle near the surface induces a net reaction force on the flagellar bundle (Fflagella) from the surface; the cell head counter-rotates and experiences a force in the opposite direction (Fhead). Circular swimming results from the torque induced by these two forces57, 98. The inset shows observed trajectories of a smooth swimming (non-tumbling) mutant of Escherichia coli (HCB437) that also lacks most genes associated with chemotaxis57. b–d | Live imaging of cells on a surface revealed distinct modes of surface motility in Pseudomonas aeruginosa7, 59 (see also Supplementary information S3 (movie)). Cells can either stand up on the surface and 'walk' (part b) or lie on the surface and 'crawl' (part c). Walking results in jagged trajectories that are better for local exploration, whereas crawling has high directional persistence and enables bacterial cells to more effectively cover distance7. Simultaneous pulling of multiple type IV pili that results in steady crawling motility (part c) is interrupted by the rapid (100 ms), 'slingshot' release of a single pilus that causes an impulsive forward motion coupled with a change in direction6 (part d). The inset image in part a is republished with permission of Elsevier, from Swimming in circles: motion of bacteria near solid boundaries, Lauga, E., DiLuzio, W. R., Whitesides, G. M. & Stone, H. A. 90, 2, 2006; permission conveyed through Copyright Clearance Center, Inc. The images in parts b and c are adapted from Gibiansky, M. L. et al. Bacteria use type IV pili to walk upright and detach from surfaces. Science 330, 197 (2010). Reprinted with permission from AAAS. The image in part d is adapted with permission from Ref. 6, National Academy of Sciences.

  4. Microbial motility in moving fluids.
    Figure 4: Microbial motility in moving fluids.

    a | Elongated particles or microorganisms exposed to fluid velocity gradients ('shear') undergo periodic rotations, or 'Jeffery orbits'13. The angular velocity varies with orientation relative to the flow, being faster when the cell is oriented transverse to the flow and slower when it is aligned with the flow. Jeffery orbits can considerably affect the transport of microorganisms in the environment. b,c | Fluid flow biases the motility of swimming bacteria (see also Supplementary information S4 (movie)). Like many natural flows, the parabolic flow profile in a microfluidic channel has non-uniform shear. Bacteria are free to swim in all directions equally in the central, low-shear region, but preferentially align and become trapped in high-shear regions near the sides63 (part b). Therefore, motile bacteria are depleted in regions of low shear and accumulate in regions of high shear, resulting in strong spatial heterogeneity in the bacterial distribution63 (part c). This 'shear-trapping' increases surface attachment and hinders chemotaxis63. Part c is adapted from Ref. 63, Nature Publishing Group.

  5. Upstream motility and downstream bending in flowing fluids.
    Figure 5: Upstream motility and downstream bending in flowing fluids.

    a,b | Fluid flow causes upstream swimming of Escherichia coli64, 65 near a surface (part a) and upstream twitching of Pseudomonas aeruginosa66 on a surface (part b). Numbers (1 and 2) denote a sequence of events. The torque induced by the shear at the surface orients bacterial cells to point upstream in both cases. For twitching cells, the periodic extension and retraction of pili pulls the cell upstream66. Upstream motility can affect transport of bacteria in biomedical settings, including the urinary tract, catheters or blood vessels. c,d | Time-lapse imaging in a microchannel revealed the effect of the curved shape of Caulobacter crescentus in surface colonization under flow8. Curved wild-type cells (green) colonize the surface more successfully than straight cells (red) possessing a mutation in the gene encoding the cytoskeletal protein crescentin (creS) (part c), because fluid flow more effectively bends the dividing cell (part d, grey) towards the surface, conferring a higher surface-attachment probability to the daughter cell8 (part d, green). The image in part c is adapted from Ref. 8, Nature Publishing Group.

References

  1. Xie, L., Altindal, T., Chattopadhyay, S. & Wu, X.-L. Bacterial flagellum as a propeller and as a rudder for efficient chemotaxis. Proc. Natl Acad. Sci. USA 108, 22462251 (2011).
    This study reported the discovery of the 'flick', a new reorientation mechanism found among marine bacteria, which makes their motility drastically different from the run-and-tumble motility observed in E. coli.
  2. Son, K., Guasto, J. S. & Stocker, R. Bacteria can exploit a flagellar buckling instability to change direction. Nat. Phys. 9, 494498 (2013).
  3. Zhao, K. et al. Psl trails guide exploration and microcolony formation in Pseudomonas aeruginosa biofilms. Nature 497, 388391 (2013).
    This study mapped the chemical trails of individual bacteria on a surface, demonstrating that matrix-rich regions are self-reinforcing and form the skeleton of biofilms.
  4. Wang, P. et al. Robust growth of Escherichia coli. Curr. Biol. 20, 10991103 (2010).
  5. Zhang, Q. et al. Acceleration of emergence of bacterial antibiotic resistance in connected microenvironments. Science 333, 17641767 (2011).
  6. Jin, F., Conrad, J. C., Gibiansky, M. L. & Wong, G. C. L. Bacteria use type-IV pili to slingshot on surfaces. Proc. Natl Acad. Sci. USA 108, 1261712622 (2011).
    This study revealed that P. aeruginosa twitching on surfaces are capable of a rapid slingshot motion that can efficiently reorient cells.
  7. Gibiansky, M. L. et al. Bacteria use type IV pili to walk upright and detach from surfaces. Science 330, 197 (2010).
  8. Persat, A., Stone, H. A. & Gitai, Z. The curved shape of Caulobacter crescentus enhances surface colonization in flow. Nat. Commun. 5, 3824 (2014).
  9. Hol, F. J. H. & Dekker, C. Zooming in to see the bigger picture: microfluidic and nanofabrication tools to study bacteria. Science 346, 1251821 (2014).
  10. Sackmann, E. K., Fulton, A. L. & Beebe, D. J. The present and future role of microfluidics in biomedical research. Nature 507, 181189 (2014).
  11. Rusconi, R., Garren, M. & Stocker, R. Microfluidics expanding the frontiers of microbial ecology. Annu. Rev. Biophys. 43, 6591 (2014).
  12. Wessel, A. K., Hmelo, L., Parsek, M. R. & Whiteley, M. Going local: technologies for exploring bacterial microenvironments. Nat. Rev. Microbiol. 11, 337348 (2013).
  13. Guasto, J. S., Rusconi, R. & Stocker, R. Fluid mechanics of planktonic microorganisms. Annu. Rev. Fluid Mech. 44, 373400 (2012).
  14. Elgeti, J., Winkler, R. G. & Gompper, G. Physics of microswimmers-single particle motion and collective behavior: a review. Rep. Prog. Phys. 78, 056601 (2015).
  15. Drescher, K., Dunkel, J., Cisneros, L. H., Ganguly, S. & Goldstein, R. E. Fluid dynamics and noise in bacterial cell–cell and cell–surface scattering. Proc. Natl Acad. Sci. USA 108, 1094010945 (2011).
    This study reported the first experimental quantification of the flow field around a single swimming E. coli bacterium.
  16. Lighthill, M. J. Mathematical Biofluiddynamics (Society for Industrial and Applied Mathematics, 1975).
  17. Wu, X. L. & Liebchaber, A. Particle diffusion in a quasi-two-dimensional bacterial bath. Phys. Rev. Lett. 84, 30173020 (2010).
  18. Zhang, H.-P., Be'er, A., Florin, E.-L. & Swinney, H. L. Collective motion and density fluctuations in bacterial colonies. Proc. Natl Acad. Sci. USA 107, 1362613630 (2010).
  19. Dombrowski, C., Cisneros, L., Chatkaew, S., Goldstein, R. E. & Kessler, J. O. Self-concentration and large-scale coherence in bacterial dynamics. Phys. Rev. Lett. 93, 25 (2004).
  20. Berke, A. P., Turner, L., Berg, H. C. & Lauga, E. Hydrodynamic attraction of swimming microorganisms by surfaces. Phys. Rev. Lett. 101, 038102 (2008).
  21. Berg, H. C. E. coli in Motion (Springer, 2004).
  22. Turner, L., Ryu, W. S. & Berg, H. C. Real-time imaging of fluorescent flagellar filaments. J. Bacteriol. 182, 27932801 (2000).
  23. Stocker, R. Reverse and flick: hybrid locomotion in bacteria. Proc. Natl Acad. Sci. USA 108, 26352636 (2011).
  24. Leifson, E., Cosenza, B. J., Murchelano, R. & Cleverdon, R. C. Motile marine bacteria I. techniques, ecology, and general characteristics. J. Bacteriol. 87, 652666 (1964).
  25. Xie, L. & Wu, X. L. Bacterial motility patterns reveal importance of exploitation over exploration in marine microhabitats. part I: theory. Biophys. J. 107, 17121720 (2014).
  26. Taktikos, J., Stark, H. & Zaburdaev, V. How the motility pattern of bacteria affects their dispersal and chemotaxis. PLoS ONE 8, e81936 (2014).
  27. Wadhams, G. H. & Armitage, J. P. Making sense of it all: bacterial chemotaxis. Nat. Rev. Mol. Cell. Biol. 5, 10241037 (2004).
  28. Tu, Y. H. Quantitative modeling of bacterial chemotaxis: Signal amplification and accurate adaptation. Annu. Rev. Biophys. 42, 337359 (2013).
  29. Stocker, R. & Seymour, J. R. Ecology and physics of bacterial chemotaxis in the ocean. Microbiol. Mol. Biol. Rev. 76, 792812 (2012).
  30. Stocker, R. Marine microbes see a sea of gradients. Science 338, 628633 (2012).
  31. Ahmed, T., Shimizu, T. S. & Stocker, R. Microfluidics for bacterial chemotaxis. Integr. Biol. 2, 604629 (2010).
  32. Kalinin, Y. V., Jiang, L., Tu, Y. & Wu, M. Logarithmic sensing in Escherichia coli bacterial chemotaxis. Biophys. J. 96, 24392448 (2009).
  33. Fechner, G. T., Adler, H. E., Howes, D. H. & Boring, E. G. Elementary Psychophysics (Holt, 1966).
  34. Rieke, F. & Rudd, M. E. The challenges natural images pose for visual adaptation. Neuron 64, 605616 (2009).
  35. Lazova, M. D., Ahmed, T., Bellomo, D., Stocker, R. & Shimizu, T. S. Response rescaling in bacterial chemotaxis. Proc. Natl Acad. Sci. USA 108, 1387013875 (2011).
    This study revealed experimentally that E. coli is capable of rescaling its chemotactic response, a process termed fold-change detection, which ensures high chemotactic sensitivity across a broad range of environmental conditions.
  36. Zhu, X. et al. Frequency-dependent Escherichia coli chemotaxis behavior. Phys. Rev. Lett. 108, 128101 (2012).
  37. Kalinin, Y., Neumann, S., Sourjik, V. & Wu, M. Responses of Escherichia coli bacteria to two opposing chemoattractant gradients depend on the chemoreceptor ratio. J. Bacteriol. 192, 17961800 (2010).
    This study was the first to use microfluidics to examine the chemotactic decision-making process of E. coli cells that were exposed to two simultaneous chemical gradients.
  38. Blackburn, N. Microscale nutrient patches in planktonic habitats shown by chemotactic bacteria. Science 282, 22542256 (1998).
  39. Bell, W. & Mitchell, R. Chemotactic and growth responses of marine bacteria to algal extracellular products. Biol. Bull. 143, 265277 (1972).
  40. Barbara, G. M. & Mitchell, J. G. Bacterial tracking of motile algae. FEMS Microbiol. Ecol. 44, 7987 (2003).
  41. Stocker, R., Seymour, J. R., Samadani, A., Hunt, D. E. & Polz, M. F. Rapid chemotactic response enables marine bacteria to exploit ephemeral microscale nutrient patches. Proc. Natl Acad. Sci. USA 105, 42094214 (2008).
  42. Seymour, J. R., Ahmed, T., Durham, W. M. & Stocker, R. Chemotactic response of marine bacteria to the extracellular products of Synechococcus and Prochlorococcus. Aquat. Microb. Ecol. 59, 161168 (2010).
  43. Seymour, J. R., Ahmed, T. & Stocker, R. Bacterial chemotaxis towards the extracellular products of the toxic phytoplankton Heterosigma akashiwo. J. Plank. Res. 31, 15571561 (2009).
  44. Seymour, J. R., Simó, R., Ahmed, T. & Stocker, R. Chemoattraction to dimethylsulfoniopropionate throughout the marine microbial food web. Science 329, 342345 (2010).
  45. Garren, M. et al. A bacterial pathogen uses dimethylsulfoniopropionate as a cue to target heat-stressed corals. ISME J. 8, 9991007 (2014).
  46. Seymour, J. R., Marcos & Stocker, R. Resource patch formation and exploitation throughout the marine microbial food web. Am. Nat. 173, E1529 (2009).
  47. Barbara, G. M. & Mitchell, J. G. Marine bacterial organisation around point-like sources of amino acids. FEMS Microbiol. Ecol. 43, 99109 (2003).
  48. Yawata, Y. et al. Competition–dispersal tradeoff ecologically differentiates recently speciated marine bacterioplankton populations. Proc. Natl Acad. Sci. USA 111, 56225627 (2014).
    This study revealed a competition–dispersal tradeoff among recently speciated sympatric marine bacteria, based on distinct behavioural interactions with particulate organic matter.
  49. Persat, A. et al. The mechanical world of bacteria. Cell 161, 988997 (2015).
  50. O'Toole, G., Kaplan, H. B. & Kolter, R. Biofilm formation as microbial development. Annu. Rev. Microbiol. 54, 4979 (2000).
  51. Karimi, A., Karig, D., Kumar, A. & Ardekani, A. M. Interplay of physical mechanisms and biofilm processes: review of microfluidic methods. Lab. Chip 15, 2342 (2015).
  52. Watnick, P. & Kolter, R. Biofilm, city of microbes. J. Bacteriol. 182, 26752679 (2000).
  53. Kearns, D. B. A field guide to bacterial swarming motility. Nat. Rev. Microbiol. 8, 634644 (2010).
  54. Teschler, J. K. et al. Living in the matrix: assembly and control of Vibrio cholerae biofilms. Nat. Rev. Microbiol. 13, 255268 (2015).
  55. Magariyama, Y. et al. Difference in bacterial motion between forward and backward swimming caused by the wall effect. Biophys. J. 88, 36483658 (2005).
  56. Molaei, M., Barry, M., Stocker, R. & Sheng, J. Failed escape: solid surfaces prevent tumbling of Escherichia coli. Phys. Rev. Lett. 113, 68103 (2014).
  57. Lauga, E., DiLuzio, W. R., Whitesides, G. M. & Stone, H. A. Swimming in circles: motion of bacteria near solid boundaries. Biophys. J. 90, 400412 (2006).
    This study rationalized why many species of bacteria swim in circular trajectories when near a surface.
  58. Utada, A. S. et al. Vibrio cholerae use pili and flagella synergistically to effect motility switching and conditional surface attachment. Nat. Commun. 5, 2913 (2014).
  59. Conrad, J. C. et al. Flagella and pili-mediated near-surface single-cell motility mechanisms in P. aeruginosa. Biophys. J. 100, 16081616 (2011).
  60. Rusconi, R. & Stocker, R. Microbes in flow. Curr. Opin. Microbiol. 25, 18 (2015).
  61. Durham, W. M., Kessler, J. O. & Stocker, R. Disruption of vertical motility by shear triggers formation of thin phytoplankton layers. Science 323, 10671070 (2009).
  62. Durham, W. M. et al. Turbulence drives microscale patches of motile phytoplankton. Nat. Commun. 4, 2148 (2013).
  63. Rusconi, R., Guasto, J. S. & Stocker, R. Bacterial transport suppressed by fluid shear. Nat. Phys. 10, 212217 (2014).
    This study revealed that the coupling of motility and flow can result in high levels of bacterial accumulation in certain regions of the flow, hampering chemotaxis and favouring surface attachment.
  64. Hill, J., Kalkanci, O., McMurry, J. L. & Koser, H. Hydrodynamic surface interactions enable Escherichia coli to seek efficient routes to swim upstream. Phys. Rev. Lett. 98, 68101 (2007).
  65. Kaya, T. & Koser, H. Direct upstream motility in Escherichia coli. Biophys. J. 102, 15141523 (2012).
  66. Shen, Y., Siryaporn, A., Lecuyer, S., Gitai, Z. & Stone, H. A. Flow directs surface-attached bacteria to twitch upstream. Biophys. J. 103, 146151 (2012).
  67. Marcos, Fu, H. C., Powers, T. R. & Stocker, R. Bacterial rheotaxis. Proc. Natl Acad. Sci. USA 109, 47804785 (2012).
  68. Meng, Y. Z. et al. Upstream migration of Xylella fastidiosa via pilus-driven twitching motility. J. Bacteriol. 187, 55605567 (2005).
    This study revealed that bacteria twitching on surfaces migrate upstream in the presence of fluid flow, owing to a hydrodynamic torque that orients them against the flow.
  69. Bures, J. et al. Small intestinal bacterial overgrowth syndrome. World J. Gastroenterol. 16, 29782990 (2010).
  70. Dunkel, J. et al. Fluid dynamics of bacterial turbulence. Phys. Rev. Lett. 110, 228102 (2013).
  71. Sokolov, A., Goldstein, R., Feldchtein, F. & Aranson, I. Enhanced mixing and spatial instability in concentrated bacterial suspensions. Phys. Rev. E Stat. Nonlin. Soft Matter Phys. 80, 18 (2009).
  72. Saintillan, D. & Shelley, M. J. Active suspensions and their nonlinear models. Comptes Rendus Phys. 14, 497517 (2013).
  73. Kaiser, A. et al. Transport powered by bacterial turbulence. Phys. Rev. Lett. 112, 158101 (2014).
  74. Butler, M. T., Wang, Q. & Harshey, R. M. Cell density and mobility protect swarming bacteria against antibiotics. Proc. Natl Acad. Sci. USA 107, 37763781 (2010).
  75. Sitti, M. Miniature devices: voyage of the microrobots. Nature 458, 11211122 (2009).
  76. Mesquita, A. R. & Hespanha, J. P. Jump control of probability densities with applications to autonomous vehicle motion. IEEE T. Automat. Contr. 57, 25882598 (2012).
  77. Rubenstein, M., Cornejo, A. & Nagpal, R. Programmable self-assembly in a thousand-robot swarm. Science 345, 795799 (2014).
  78. Rusconi, R., Lecuyer, S., Guglielmini, L. & Stone, H. A. Laminar flow around corners triggers the formation of biofilm streamers. J. R. Soc. Interface 7, 12931299 (2010).
  79. Drescher, K., Shen, Y., Bassler, B. L. & Stone, H. A. Biofilm streamers cause catastrophic disruption of flow with consequences for environmental and medical systems. Proc. Natl Acad. Sci. USA 110, 43454350 (2013).
  80. Malfatti, F. & Azam, F. Atomic force microscopy reveals microscale networks and possible symbioses among pelagic marine bacteria. Aquat. Microb. Ecol. 58, 114 (2009).
  81. Kiviet, D. J. et al. Stochasticity of metabolism and growth at the single-cell level. Nature 514, 376379 (2014).
  82. Rojas, E., Theriot, J. A. & Huang, K. C. Response of Escherichia coli growth rate to osmotic shock. Proc. Natl Acad. Sci. USA 111, 78077812 (2014).
  83. Brumley, D. R., Wan, K. Y., Polin, M. & Goldstein, R. E. Flagellar synchronization through direct hydrodynamic interactions. eLife 3, e02750 (2014).
  84. Berg, H. C. & Brown, D. A. Chemotaxis in Escherichia coli analyzed by three-dimensional tracking. Nature 239, 500504 (1972).
  85. Liu, B. et al. Helical motion of the cell body enhances Caulobacter crescentus motility. Proc. Natl Acad. Sci. USA 111, 1125211256 (2014).
  86. Wu, M. M., Roberts, J. W., Kim, S., Koch, D. L. & DeLisa, M. P. Collective bacterial dynamics revealed using a three-dimensional population-scale defocused particle tracking technique. Appl. Environ. Microb. 72, 49874994 (2006).
  87. Taute, K. M., Gude, S., Tans, S. J. & Shimizu, T. S. High-throughput 3D tracking of bacteria on a standard phase contrast microscope. Nat. Commun. 6, 8776 (2015).
  88. Brumley, D. R., Polin, M., Pedley, T. J. & Goldstein, R. E. Metachronal waves in the flagellar beating of Volvox and their hydrodynamic origin. J. R. Soc. Interface. 12, 20141358 (2015).
  89. Weibel, D. B., DiLuzio, W. R. & Whitesides, G. M. Microfabrication meets microbiology. Nat. Rev. Microbiol. 5, 209218 (2007).
  90. Xia, Y. N. & Whitesides, G. M. Soft lithography. Annu. Rev. Mater. Sci. 28, 153184 (1998).
  91. Tabeling, P. Introduction to Microfluidics (Oxford Univ. Press, 2005).
  92. Kantsler, V., Dunkel, J., Blayney, M., Goldstein, R. E. & Hyman, A. A. Rheotaxis facilitates upstream navigation of mammalian sperm cells. eLife 3, e02403 (2014).
  93. Kim, S., Kim, H. J. & Jeon, N. L. Biological applications of microfluidic gradient devices. Integr. Biol. 2, 584603 (2010).
  94. Adler, M., Erickstad, M., Gutierrez, E. & Groisman, A. Studies of bacterial aerotaxis in a microfluidic device. Lab. Chip 12, 48354847 (2012).
  95. Wong, I. & Ho, C. M. Surface molecular property modifications for poly(dimethylsiloxane) (PDMS) based microfluidic devices. Microfluid Nanofluid 7, 291306 (2009).
  96. Cheng, S. Y. et al. A hydrogel-based microfluidic device for the studies of directed cell migration. Lab. Chip 7, 763769 (2007).
  97. Weibel, D. B. et al. Bacterial printing press that regenerates its ink: Contact-printing bacteria using hydrogel stamps. Langmuir 21, 64366442 (2005).
  98. Lauga, E. & Powers, T. R. The hydrodynamics of swimming microorganisms. Rep. Prog. Phys. 72, 096601 (2009).

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Affiliations

  1. Department of Mechanical Engineering, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139, USA.

    • Kwangmin Son
  2. Ralph M. Parsons Laboratory, Department of Civil and Environmental Engineering, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139, USA.

    • Kwangmin Son,
    • Douglas R. Brumley &
    • Roman Stocker
  3. Department of Civil, Environmental and Geomatic Engineering, ETH Zurich, 8093 Zurich, Switzerland.

    • Douglas R. Brumley &
    • Roman Stocker

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The authors declare no competing interests.

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Author details

  • Kwangmin Son

    Kwangmin Son is a postdoctoral associate at Massachusetts Institute of Technology (MIT), Cambridge, USA. He received his Ph.D. in mechanical engineering from MIT in 2015, studying the physical ecology of marine microorganisms, and in particular microbial motility and chemotaxis. His current research is aimed at understanding bacteria–virus encounters and adsorption dynamics using single-virus dynamic imaging.

  • Douglas R. Brumley

    Douglas R. Brumley is a postdoctoral fellow at Massachusetts Institute of Technology (MIT), Cambridge, USA, and a cross-disciplinary fellow of the Human Frontier Science Program. He received his Ph.D. from the Department of Applied Mathematics and Theoretical Physics at the University of Cambridge, UK, in 2013. His current research focuses on the microscale fluid dynamics of microbial transport at coral surfaces.

  • Roman Stocker

    Roman Stocker is a professor in civil, environmental and geomatic engineering at ETH Zürich, Switzerland. His research centres on the physical ecology and biophysics of microorganisms, with a focus on marine microorganisms and how they shape ocean health and biogeochemistry.

Supplementary information

Movies

  1. Supplementary information S1 (movie) (8.4 MB)

    Motility mechanics. Many marine bacteria reorient by a 'flick', an off-axis deformation of the flagellum that enables bacteria with a single flagellum to change their direction of swimming. This video shows the flick process of Vibrio alginolyticus (see also Fig. 1c–f), recorded using high-speed, high-intensity dark-field microscopy (40X objective lens, 420 frames s−1). On the left is the raw video, on the right a processed version showing the (single, polar) flagellum in magenta. Note the buckling of the flagellum (see also Fig. 1e, 50–70 ms) shortly after the reversal in swimming direction ( Fig. 1e, 20 ms). This movie is reproduced from Ref. 2, Nature Publishing Group.

  2. Supplementary information S2 (movie) (1.90 MB)

    Chemotaxis. Using chemotaxis, natural marine bacteria can cluster around photosynthetic diatoms, here Chaetoceros affinis, in response to the gradients in dissolved organic matter originating from the diatom (see also Fig. 2a). Courtesy of Steven Smriga and Vicente Fernandez, Department of Civil, Environmental and Geomatic Engineering, ETH Zurich, 8093 Zurich, Switzerland.

  3. Supplementary information S3 (movie) (5.97 MB)

    Surface motility. Two-point tracking of a single Pseudomonas aeruginosa bacterium as it crawls along a surface (see also Fig. 3d). Markers 1 and 2 represent the leading and trailing poles, respectively. The video corresponds to 700 s in real time, with playback sped up by a factor of 40. This movie is reproduced with permission from Ref. 6, National Academy of Sciences.

  4. Supplementary information S4 (movie) (452 KB)

    Motility in flow. Trajectory of a smooth-swimming Bacillus subtilis bacterium in a microfluidic channel (see also Fig. 4b). The raw video of the motile cell is shown first, followed by a replay in which the tracked cell trajectory (green) and position and orientation (red) are included. The flow in the channel is from left to right, and the video is recorded in the reference frame comoving with the mean speed of the flow (mean speed = 500 μm s−1, mean absolute shear rate = 2.5 s−1). The looped trajectory results from the velocity gradient generating a hydrodynamic torque that continually reorients the cell while it swims. The video was captured at 70.6 frames s−1 using dark-field microscopy, and is replayed 1.7 times slower than real time. This movie is reproduced from Ref. 63, Nature Publishing Group.

Additional data