The authors thank the FIRST microfabrication facility at ETH Zurich. This work was supported by an Australian Research Council Discovery Early Career Researcher Award DE180100911 (to D.R.B.), a Gordon and Betty Moore Marine Microbial Initiative Investigator Award GBMF3783 (to R.S.), and a Swiss National Science Foundation grant 1-002745-000 (to R.S.).

1. Fabrication of the Microfluidic Device for the Single Chemical-pulse Experiment
<ol>
	<li>Design the channel using computer-aided design (CAD) software and print it onto a transparency film to create the photo mask (Figure 1A).</li>
	<li>Fabricate the master by soft lithography (under clean-room conditions).
	<ol>
		<li>Clean a silicon wafer (4 inches) in quick succession with acetone, methanol and isopropanol, then dry using nitrogen. Bake the wafer in the oven at 130 &#176;C for 5 min.</li>
		<li>Place the wafer at the center of a spin-coater and pour SU-8 photoresist onto the wafer. Ramp the speed of the spin-coater up to 500 rpm over 5 s, and keep at 500 rpm for 10 s. Ramp up to the final speed over 10 s and maintain at this speed for 30 s. 
		NOTE: The exact value of the final speed depends on the targeted coating thickness and the SU-8 used.</li>
		<li>After the spin-coating process, bake the wafer at 65 &#176;C and then at 95 &#176;C. Let the wafer cool at room temperature (RT) for at least 5 min. 
		NOTE: The baking time depends on the targeted thickness and type of photoresist used. As a general rule, for every 100 &#181;m layer the wafer should be baked for 5 min at 65 &#176;C and 45 min at 95 &#176;C.</li>
		<li>Place the photo mask onto the wafer to ensure that just the region of interest is exposed and polymerized. Expose to UV light for the time recommended in the SU-8 manual. 
		NOTE: Here, with exposure energy of 200 mJ cm-2 at a wavelength of 350 nm, the wafer was exposed for 150 s.</li>
		<li>Bake the wafer at 65 &#176;C and 95 &#176;C for the time recommended in the SU-8 manual. 
		NOTE: As a general rule, for every 100 &#181;m layer the wafer should be baked for 5 min at 65 &#176;C and 45 min at 95 &#176;C.</li>
		<li>Immerse the wafer in a beaker filled with polymethyl methacrylate (PMMA) developer in order to obtain the master. Gently shake the beaker to ensure that the unpolymerized photoresist is washed out. Bake the master at 200 &#176;C to further cross-link the SU-8 layer.</li>
	</ol>
	</li>
	<li>Prepare a polydimethylsiloxane (PDMS) mixture by combining the elastomer with its curing agent (Table of Materials) at a 10:1 ratio in a beaker (here, 40 mL). Mix vigorously until the liquid is homogeneous. 
	NOTE: The PDMS mixture will look opaque because bubbles are generated during the mixing process. 
	CAUTION: In order to prevent skin coming into contact with potentially hazardous chemicals or biological material, always wear a lab coat and disposable plastic gloves throughout the protocol and follow any specific safety protocols according to the Material Safety Data Sheet (MSDS).</li>
	<li>Degas the PDMS mixture in a vacuum chamber for 45 min at RT. To expedite the process, periodically release the vacuum in order to burst the bubbles that form at the interface. 
	NOTE: The degassing process must be performed within 1 h to prevent the PDMS mixture from beginning to cure.</li>
	<li>Remove dust from the surface of the master with a pressurized cleaner, then pour the degassed PDMS mixture onto the master and bake in an oven at 80 &#176;C for 2 h. 
	NOTE: Alternatively, baking overnight (or for at least 12 h) at 60 &#176;C would achieve the same hardened PDMS.</li>
	<li>Cut the PDMS with a blade around the microstructures (at a distance of approximately 5 mm) and then carefully peel the PDMS from the master.</li>
	<li>Punch holes to serve as inlet and outlet of the microchannel (here, one inlet and one outlet, see Figure 1A). Ensure that nothing touches the bottom face of the PDMS where the features are located. 
	NOTE: The protocol can be paused here. The microchannel should be sealed with adhesive tape to prevent accumulation of dust and other particles during storage.</li>
	<li>Bond the PDMS microchannel to a glass slide.
	<ol>
		<li>Thoroughly clean a glass slide with soap, isopropanol and deionized water. Let the glass slide dry or use compressed air to speed up the process.</li>
		<li>Remove dust particles by dabbing with adhesive tape. Bond the PDMS microchannel on the clean glass slide, immediately after treating both surfaces with plasma (by using either a corona system or a plasma oxygen chamber) for 2 min.</li>
		<li>Place the microfluidic device to heat on a hot plate at 80 &#176;C for at least 1 h to strengthen the chemical bond with the glass.</li>
	</ol>
	</li>
</ol>
2. Fabrication of the 3D-printed Millifluidic Device for the Experiment with Multiple Pulses
<ol>
	<li>Design the 3D shape using 3D-design software and print the master for the PDMS mold with a high-resolution 3D printer (Figure 1B). 
	NOTE: See Table of Materials for the 3D printer and mold material used to create the master.</li>
	<li>Repeat steps 1.3&#8722;1.4, then clean the surface of the master by dabbing with adhesive tape. Put the master on a scale, then, while avoiding the central region of the master that will be occupied by the membrane, pour on the exact quantity of uncured PDMS mixture (here, 23.4 g) in order to obtain the desired height of PDMS by matching the height of the master (here, 0.5 mm). Remove any remaining bubbles with the help of compressed air.</li>
	<li>Place the PDMS cast on the master in an oven at 45 &#176;C to bake for at least 12 h.</li>
	<li>Bond the 3D PDMS mold to a Petri dish.
	<ol>
		<li>Gently peel off the hardened PDMS layer and punch inlet and outlet holes for the injection of the bacterial suspension (Figure 1C).</li>
		<li>Activate both the surface of a Petri dish (90 mm x 15 mm) and the PDMS mold with oxygen plasma for 2 min. Bond the PDMS mold on the Petri dish. Gently press the mold to the Petri dish, but do not press where the features are located as this can collapse the interrogation chamber.</li>
		<li>Place the Petri dish bonded to the PDMS 3D mold in an oven at 45 &#176;C for at least 12 h to strengthen the chemical bond. 
		NOTE: The protocol can be paused here.</li>
	</ol>
	</li>
	<li>Membrane surface functionalization18
	<ol>
		<li>Activate the nanoporous polycarbonate (PCTE) membrane in an oxygen plasma chamber for 1 min at RT. 
		CAUTION: Avoid using a corona system for the plasma activation because it will damage the membrane.</li>
		<li>Under a chemical hood, dilute a commercial solution of 3-aminopropyltriethoxysilane (APTES) in deionized water to 1% by volume (here, 40 mL), by using a polypropylene tube. 
		CAUTION: The APTES solution is acutely toxic (MSDS health hazard score 3) and should be handled with extreme care exclusively under a chemical hood with gloves. Change gloves immediately after handling APTES solution. APTES fumes are destructive to the mucous membranes and the upper respiratory tract. The target organs of APTES are nerves, liver, and kidney. If a fume hood is not available, a face shield and full-face respirator must be implemented.</li>
		<li>Transfer the diluted APTES solution in a Petri dish and immerse the activated membrane in the APTES solution for 20 min.</li>
		<li>Remove the membrane from the APTES solution with tweezers and place it on a cleanroom wipe to dry.</li>
	</ol>
	</li>
	<li>For the fabrication of the PDMS microfluidic channels that will lie on the membrane and allow washing of the bacterial arena, repeat steps 1.1&#8722;1.7.</li>
	<li>Bond the PDMS washing channels to the functionalized membrane.
	<ol>
		<li>Activate both the PDMS washing channel and the PCTE membrane with an oxygen plasma chamber for 2 min. 
		NOTE: The membrane, which will be sandwiched between two PDMS layers, should be smaller than the PDMS washing channel.</li>
		<li>Immediately after the plasma treatment, bring the functionalized membrane and the PDMS washing channel into contact by gently pressing the PDMS washing channel onto the membrane. 
		NOTE: Do not apply excessive pressure at this stage, because it might cause (irreversible) attachment of the membrane to the channel&#39;s roof, blocking the channel.</li>
	</ol>
	</li>
	<li>Sandwich the membrane between two PDMS layers (Figure 1E).
	<ol>
		<li>Activate both the bonded laminate PDMS washing channel&#8722;PCTE membrane and the 3D PDMS mold previously bonded to the Petri dish with an oxygen plasma chamber for 2 min. Bring them into contact and press together.</li>
		<li>Place the Petri dish bonded to the sandwich structure in an oven at 45 &#176;C for at least 12 h to strengthen the chemical bond. 
		NOTE: The protocol can be paused here.</li>
	</ol>
	</li>
</ol>
3. Cell Culture
<ol>
	<li>Grow a population of V. ordalii (strain 12B09 or 12B09pGFP) overnight for 20 h in 2216 medium19 on an orbital shaker (600 rpm) at 30 &#176;C and harvest cells in late-exponential phase. For isolates harboring pGFP, add spectinomycin (50 &#181;g mL-1) to maintain the plasmid.</li>
	<li>Centrifuge a 1 mL aliquot of cells at 2500 x g for 3 min, remove the supernatant and re-suspend the cells in 1 mL of filtered artificial seawater. Repeat this step.</li>
	<li>Starve the population of V. ordalii on an orbital shaker (600 rpm) at 30 &#176;C for 3 h.</li>
	<li>Prepare a stock artificial seawater solution of 10 mM (here, 1 mL) of 4-methoxy-7-nitroindolinyl-caged-L-glutamate (MNI-caged-L-glutamate) and store at -20 &#176;C. 
	NOTE: Protect the MNI-caged-L-glutamate solution from ambient light to prevent photolysis.</li>
	<li>Dilute the cells 50x by re-suspending the starved cells in an artificial seawater solution of 1 mM MNI-caged-L-glutamate. 
	NOTE: This will ensure a final bacterial concentration lower than 5 x 107 mL-1 (the exact value depends on the initial concentration of cells in late exponential, which is typically ~109 mL-1). Bacterial concentration was estimated using a spectrophotometer at optical density (OD) of 600 nm.</li>
</ol>
4. Calibration of the Uncaging Protocol
<ol>
	<li>Place a droplet (20 &#956;L) of an artificial seawater solution of MNI-caged-L-glutamate20 at a concentration C0 = 10 mM on a glass slide. Encapsulate the droplet by covering it with a circular PDMS chamber (diameter d = 5 mm, height h = 250 &#956;m).</li>
	<li>Place the glass slide on a microscope stage and expose the entire chamber for 20 ms to an LED beam at a wavelength of 395 nm (power 295 mW) by using a 4x objective (numerical aperture [NA] = 0.13).</li>
	<li>Open the PDMS chamber and extract a droplet (1 &#956;L) for analysis with a UV-Vis spectrophotometer.</li>
	<li>Repeat steps 4.1&#8722;4.3 for different durations of LED illumination of 0.1 s, 0.5 s, 2.5 s, 25 s, 250 s (or until saturation of the chemical reaction) and 0 s (to obtain the background). Replicate the procedure steps 4.1&#8722;4.4 at least 3x (here, 4x).</li>
	<li>From the absorption spectrum, extract the value at 405 nm, which represent the peak in the absorption spectrum of the cage molecule21. To determine the rate k of the chemical uncaging reaction by the LED exposure14, use the following first-order kinetics equation for the numerical fit of the experimental data17 
	<img alt="Equation 1" src="/files/ftp_upload/60589/60589equ01.jpg" /> 
	where C(t) is the value of the absorption spectrum of the solution at 405 nm (after background removal) with an uncaging time of t seconds. For small uncaging time t such that kt 1, Eq. 1 can be simplified to the linear formulation 
	<img alt="Equation 2" src="/files/ftp_upload/60589/60589equ02.jpg" /></li>
</ol>
5. Single Chemical-pulse Experiment
<ol>
	<li>Maintain the microfluidic channel under vacuum for 20 min to reduce the gas concentration within the PDMS so that bubbles are less likely to form during the filling process.</li>
	<li>Extract the channel from the vacuum pump and immediately introduce the dilute bacterial suspension in 1 mM MNI-caged-L-glutamate in artificial seawater into the microchannel gently using a micropipette to avoid flagellar damage by mechanical shearing forces (here, the entire filling process took 5-10 s). After filling the channel with the bacterial suspension, suck the solution in excess with paper towel, and seal inlet and outlet with PDMS plugs by gently pressing them into the holes. 
	NOTE: In this way, fluid flow in the chamber is prevented.</li>
	<li>Place the microchannel onto a microscope stage and move the stage to set the field of view in mid-channel. Set up the microscope to perform imaging in phase contrast (4x objective) at a frame rate of 12 fps. 
	NOTE: This value can be varied but should not be less than 10 fps to allow reconstruction of the bacterial trajectories through image analysis (see protocol section 7).</li>
	<li>Set the pinhole of the LED beam at the minimum aperture. By setting the exposure time of the camera to 5 s, record a few frames to obtain precise measurements of the spatial location and size of the LED beam. 
	NOTE: The LED light source connects to the microscope&#39;s epi-fluorescence illuminator via liquid light guide connection. The LED beam does not affect video capturing, because video acquisition and LED stimulation are commanded independently via software.</li>
	<li>Perform continuous imaging for a total duration of 10 min (20 min for the largest chemical pulse), and simultaneously activate the focused LED beam at 395 nm for the desired duration (in this experiment, t = 20 ms, 100 ms, 500 ms) at a user-defined time point (here, 10 s after the start of the video acquisition) via microscope software. To obtain multiple replicates of the same process, move the stage to record the bacterial response over different positions of the microfluidic channel. Replicate this procedure for each pulse size, using a new microchannel. 
	NOTE: The uncaging process generates an axisymmetric cylindrical pulse that diffuses radially outwards in the imaging plane.</li>
	<li>Conduct separate experiments without chemical uncaging at higher magnification (20x objective, NA = 0.45) at a higher frame rate (50 fps) to record the unbiased swimming motion of the bacteria.</li>
</ol>
6. Multiple Chemical-pulse Experiment
<ol>
	<li>Maintain the millifluidic chamber under vacuum for 20 min.</li>
	<li>Place the Petri dish containing the millifluidic chamber onto a microscope stage. Fill the chamber below the membrane with the dilute bacterial suspension (here, the entire filling process took 10&#8722;15 s) of GFP-fluorescent V. ordalii in 1 mM MNI-caged-L-glutamate solution in artificial seawater. After filling the channel with the bacterial suspension, suck the solution in excess with paper towel, and seal inlet and outlet with PDMS plugs by gently pressing them into the holes. 
	NOTE: In this way, fluid flow in the chamber is prevented. To allow better visualization of bacteria in this setup where the imaging occurs through the nanoporous membrane, it is strongly recommended to use fluorescent strains instead of the wild type (as used in the single chemical-pulse experiment).</li>
	<li>Fill a syringe with an artificial seawater solution of 1 mM MNI-caged-L-glutamate and attach tubing to the inlet and outlet of the washing channel above the membrane.</li>
	<li>Connect the tubing to a waste reservoir and ensure that the tubing is entirely submerged in the fluid waste reservoir to avoid pressure oscillations.</li>
	<li>Set the appropriate flow rate on the syringe pump (here, 50 &#956;L min-1) to obtain the desired mean flow rate in the washing channel, which depends on the channel geometry.</li>
	<li>Start the syringe pump to establish the flow in the washing channel above the membrane.</li>
	<li>Run the software controlling the LED beam and microscope stage to generate user-defined sequences of pulses at different locations and with different intensities. 
	NOTE: The continuous flow of the artificial seawater solution of 1 mM MNI-caged-L-glutamate in the washing channel above the membrane replenishes the caged compound solution and washes the spent medium from the bacterial arena.</li>
	<li>Record video using a 4x objective at regular time intervals over a period of several hours and over multiple contiguous locations to cover a large surface (here, 1 cm x 1 cm, Figure 1F).</li>
</ol>
7. Image Analysis and Data Analysis
<ol>
	<li>Reconstruction of the bacterial trajectories
	<ol>
		<li>From the movies recorded with the 4x objective, reconstruct the bacterial trajectories using tracking software17. 
		NOTE: These trajectories represent the 2D projections of the 3D bacterial motion in the microchannel.</li>
		<li>From the reconstructed bacterial trajectories, determine the radial drift velocity of each swimming individual by projecting its swimming speed over the direction towards the center of the chemical pulse (Figure 2D). For the visualization of the spatio-temporal dynamics of the radial drift velocity, use a binning grid with spatial size 75 &#956;m x 75 &#956;m and temporal window of 5&#8722;10 s interval (Figure 2D,E). 
		NOTE: Because of the cylindrical symmetry of the process, data can be analyzed in polar coordinates and averaged over the angular dimension (Figure 3).</li>
		<li>From the reconstructed bacterial trajectories, determine the spatio-temporal dynamics of the distribution of bacteria as they respond to the chemical pulses (Figure 2E).</li>
		<li>From the higher resolution movies recorded with the 20x objective during the single-pulse experiments, extract the distribution of swimming speed and run time for the bacterial population as they respond to the chemical pulse (Figure 4).</li>
	</ol>
	</li>
	<li>Estimate the diffusion coefficient of bacteria by considering the dissipation dynamics of the bacterial population after chemotaxis is completed17 (here, for t &#62; 300 s; Figure 3).</li>
	<li>Estimate the diffusion coefficient of glutamate by applying the Stokes-Einstein equation 
	<img alt="Equation 3" src="/files/ftp_upload/60589/60589equ03.jpg" /> 
	where the amino acid molecules are assumed to be spherical particles with hydrodynamic radius22 rG, kB is the Boltzmann constant (1.38 x 10-23 m2 kg s-2 K-1), T is the temperature (296 K), and &#951; is the dynamic viscosity of the artificial seawater (salinity 36 g kg-1) at 23 &#176;C (10-3 Pa s)23</li>
</ol>

<ol>
	<li>Armitage, J. P., Lackie, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The biology of the chemotactic response. , (1991).</li><li>Azam, F., Malfatti, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microbial+structuring+of+marine+ecosystems.">Microbial structuring of marine ecosystems.</a> Nature Reviews Microbiology. 5 (10), 782-791 (2007).</li><li>Buchan, A., LeCleir, G. R., Gulvik, C. A., Gonz&#225;lez, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Master+recyclers:+features+and+functions+of+bacteria+associated+with+phytoplankton+blooms.">Master recyclers: features and functions of bacteria associated with phytoplankton blooms.</a> Nature Reviews Microbiology. 12 (10), 686-698 (2014).</li><li>Whitesides, G. M., Ostuni, E., Takayama, S., Jiang, X., Ingber, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Soft+lithography+in+biology+and+biochemistry.">Soft lithography in biology and biochemistry.</a> Annual Review of Biomedical Engineering. 3, 335-373 (2001).</li><li>Segall, J. E., Block, S. M., Berg, H. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporal+comparisons+in+bacterial+chemotaxis.">Temporal comparisons in bacterial chemotaxis.</a> Proceedings of the National Academy of Sciences of the United States of America. 83 (23), 8987-8991 (1986).</li><li>Waite, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-genetic+diversity+modulates+population+performance.">Non-genetic diversity modulates population performance.</a> Molecular Systems Biology. 12 (12), 895 (2016).</li><li>Stocker, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Marine+Microbes+See+a+sea+of+gradients.">Marine Microbes See a sea of gradients.</a> Science. 338 (6107), 628-633 (2012).</li><li>Ellis-Davies, G. C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Caged+compounds:+photorelease+technology+for+control+of+cellular+chemistry+and+physiology.">Caged compounds: photorelease technology for control of cellular chemistry and physiology.</a> Nature Methods. 4 (8), 619-628 (2007).</li><li>Kaplan, J. H., Somlyo, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flash+photolysis+of+caged+compounds:+New+tools+for+cellular+physiology.">Flash photolysis of caged compounds: New tools for cellular physiology.</a> Trends in Neurosciences. 12 (2), 54-59 (1989).</li><li>Ando, H., Furuta, T., Tsien, R. Y., Okamoto, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photo-mediated+gene+activation+using+caged+RNA/DNA+in+zebrafish+embryos.">Photo-mediated gene activation using caged RNA/DNA in zebrafish embryos.</a> Nature Genetics. 28 (4), 317-325 (2001).</li><li>England, P. M., Lester, H. A., Davidson, N., Dougherty, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Site-specific,+photochemical+proteolysis+applied+to+ion+channels+in+vivo.">Site-specific, photochemical proteolysis applied to ion channels in vivo.</a> Proceedings of the National Academy of Sciences of the United States of America. 94 (20), 11025-11030 (1997).</li><li>Fino, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RuBi-Glutamate:+Two-photon+and+visible-light+photoactivation+of+neurons+and+dendritic+spines.">RuBi-Glutamate: Two-photon and visible-light photoactivation of neurons and dendritic spines.</a> Frontiers in Neural Circuits. 3, (2009).</li><li>Khan, S., Spudich, J. L., McCray, J. A., Trentham, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemotactic+signal+integration+in+bacteria.">Chemotactic signal integration in bacteria.</a> Proceedings of the National Academy of Sciences of the United States of America. 92 (21), 9757-9761 (1995).</li><li>Sagawa, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-Cell+E.+coli+Response+to+an+Instantaneously+Applied+Chemotactic+Signal.">Single-Cell E. coli Response to an Instantaneously Applied Chemotactic Signal.</a> Biophysical Journal. 107 (3), 730-739 (2014).</li><li>Xie, L., Lu, C., Wu, X. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Marine+Bacterial+Chemoresponse+to+a+Stepwise+Chemoattractant+Stimulus.">Marine Bacterial Chemoresponse to a Stepwise Chemoattractant Stimulus.</a> Biophysical Journal. 108 (3), 766-774 (2015).</li><li>Jikeli, J. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sperm+navigation+along+helical+paths+in+3D+chemoattractant+landscapes.">Sperm navigation along helical paths in 3D chemoattractant landscapes.</a> Nature Communications. 6, 7985 (2015).</li><li>Brumley, D. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bacteria+push+the+limits+of+chemotactic+precision+to+navigate+dynamic+chemical+gradients.">Bacteria push the limits of chemotactic precision to navigate dynamic chemical gradients.</a> Proceedings of the National Academy of Sciences of the United States of America. 116 (22), 10792-10797 (2019).</li><li>Aran, K., Sasso, L. A., Kamdar, N., Zahn, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Irreversible,+direct+bonding+of+nanoporous+polymer+membranes+to+PDMS+or+glass+microdevices.">Irreversible, direct bonding of nanoporous polymer membranes to PDMS or glass microdevices.</a> Lab on a Chip. 10 (5), 548 (2010).</li><li>ZoBell, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+cultural+requirements+of+heterotrophic+aerobes.">The cultural requirements of heterotrophic aerobes.</a> Journal of Marine Research. 4, 42-75 (1941).</li><li>Canepari, M., Nelson, L., Papageorgiou, G., Corrie, J. E. T., Ogden, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photochemical+and+pharmacological+evaluation+of+7-nitroindolinyl-and+4-methoxy-7-nitroindolinyl-amino+acids+as+novel,+fast+caged+neurotransmitters.">Photochemical and pharmacological evaluation of 7-nitroindolinyl-and 4-methoxy-7-nitroindolinyl-amino acids as novel, fast caged neurotransmitters.</a> Journal of Neuroscience Methods. 112 (1), 29-42 (2001).</li><li>Trigo, F. F., Corrie, J. E. T., Ogden, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laser+photolysis+of+caged+compounds+at+405nm:+Photochemical+advantages,+localisation,+phototoxicity+and+methods+for+calibration.">Laser photolysis of caged compounds at 405nm: Photochemical advantages, localisation, phototoxicity and methods for calibration.</a> Journal of Neuroscience Methods. 180 (1), 9-21 (2009).</li><li>Ribeiro, A. C. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutual+diffusion+coefficients+of+L-glutamic+acid+and+monosodium+L-glutamate+in+aqueous+solutions+at+T=298.15K.">Mutual diffusion coefficients of L-glutamic acid and monosodium L-glutamate in aqueous solutions at T=298.15K.</a> The Journal of Chemical Thermodynamics. 74, 133-137 (2014).</li><li>Sharqawy, M. H., Lienhard, J. H., Zubair, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thermophysical+properties+of+seawater:+a+review+of+existing+correlations+and+data.">Thermophysical properties of seawater: a review of existing correlations and data.</a> Desalination and Water Treatment. 16 (1-3), 354-380 (2010).</li><li>. Nikon depth of field calculator Available from: <a target="_blank" href="https://www.microscopyu.com/tutorials/depthoffield">https://www.microscopyu.com/tutorials/depthoffield</a> (2019)</li><li>Ki&#248;rboe, T., Thygesen, U. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluid+motion+and+solute+distribution+around+sinking+aggregates:+II.+Implications+for+remote+detection+by+colonizing+zooplankters.">Fluid motion and solute distribution around sinking aggregates: II. Implications for remote detection by colonizing zooplankters.</a> Marine Ecology Progress Series. 211, 15-25 (2001).</li><li>Blackburn, N., Fenchel, T., Mitchell, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microscale+nutrient+patches+in+planktonic+habitats+shown+by+chemotactic+bacteria.">Microscale nutrient patches in planktonic habitats shown by chemotactic bacteria.</a> Science. 282 (5397), 2254-2256 (1998).</li><li>Stocker, R., Seymour, J. R., Samadani, A., Hunt, D. E., Polz, M. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+chemotactic+response+enables+marine+bacteria+to+exploit+ephemeral+microscale+nutrient+patches.">Rapid chemotactic response enables marine bacteria to exploit ephemeral microscale nutrient patches.</a> Proceedings of the National Academy of Sciences of the United States of America. 105 (11), 4209-4214 (2008).</li><li>Altindal, T., Chattopadhyay, S., Wu, X. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bacterial+chemotaxis+in+an+optical+trap.">Bacterial chemotaxis in an optical trap.</a> PLoS ONE. 6 (4), 18231 (2011).</li><li>Zhu, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Frequency-Dependent+Escherichia+coli+Chemotaxis+behavior.">Frequency-Dependent Escherichia coli Chemotaxis behavior.</a> Physical Review Letters. 108 (12), (2012).</li><li>Baraban, L., Harazim, S. M., Sanchez, S., Schmidt, O. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemotactic+behavior+of+catalytic+motors+in+microfluidic+channels.">Chemotactic behavior of catalytic motors in microfluidic channels.</a> Angewandte Chemie International Edition. 52 (21), 5552-5556 (2013).</li></ol>

The authors have nothing to disclose.

This method allows researchers to study bacterial chemotaxis under controlled, dynamic gradients in micro- and millifluidic devices, enabling reproducible data acquisition. The near-instantaneous creation of chemical pulses at the microscale by photolysis aims to reproduce the types of nutrient pulses that bacteria encounter in the wild from a range of sources, for example, the diffusive spreading of plumes behind sinking marine particles25, or the nutrient spreading from lysed phytoplankton cells...

Microbes rely on chemotaxis, the process of detecting chemical gradients and modifying motility in response1, to navigate chemical landscapes, approach nutrient sources and hosts, and escape noxious substances. These microscale processes determine the macroscale kinetics of interactions between microbes and their environment2,3. Recent advances in microfluidics and microfabrication technologies, including soft lithography4, have revolutionized our ability to create controlled microenvironments in which to study the interactions of microbes. For example, past experiments have studied bacterial chemotaxis by generating highly controlled, stable gradients of intermediate to high nutrient concentrations5,6. However, in natural environments, microscale chemical gradients can be short-lived&#8213;dissipated by molecular diffusion&#8213;and background conditions are often highly dilute7. To directly measure the chemotactic response of microbial populations first exposed to unsteady chemical environments, we devised and here describe methods to combine microfluidic technology with photolysis, thereby mimicking gradients that wild bacteria encounter in nature.
Uncaging technology employs light sensitive probes that functionally encapsulate biomolecules in an inactive form. Irradiation releases the caged molecule, allowing the targeted perturbation of a biological process8. Due to the rapid and precise control of cellular chemistry that the uncaging affords9, photolysis of caged compounds has traditionally been employed by biologists, physiologists and neuroscientists to study the activation of genes10, ion channels11, and neurons12. More recently, scientists have leveraged the significant advantages of photolysis to study chemotaxis13, to determine the flagella switching dynamics of individual bacterial cells exposed to a stepwise chemoattractant stimulus14,15, and to investigate motility patterns of single sperm cells in three-dimensional (3D) gradients16.
In our approach, we implement photolysis of caged amino acids within microfluidic devices to study the behavioral response of a bacterial population to controlled chemical pulses, which become near-instantaneously available through photorelease. The use of a low-magnification (4x) objective (NA = 0.13, depth of focus approximately 40 &#956;m) allows both the observation of the population-level aggregative response of thousands of bacteria over a large field of view (3.2 mm x 3.2 mm), and the measurement of motion at the single-cell level. We present two applications of this method: 1) the release of a single chemical pulse to study bacterial accumulation&#8722;dissipation dynamics starting from uniform conditions, and 2i) the release of multiple pulses to characterize the bacterial accumulation dynamics under time-varying, spatially heterogeneous chemoattractant conditions. This method has been tested on the marine bacteria Vibrio ordalii performing chemotaxis toward the amino acid glutamate17, but the method is broadly applicable to different combinations of species and chemoattractants, as well as to biological processes beyond chemotaxis (e.g., nutrient uptake, antibiotic exposure, quorum sensing). This approach promises to help elucidate the ecology and behavior of microorganisms in realistic environments and to uncover the hidden trade-offs that individual bacteria face when navigating ephemeral dynamic gradients.

<table><tbody><tr><td>(3-Aminopropyl) triethoxysilane (APTES)</td><td>Sigma-Aldrich</td><td>A3648</td><td>&gt;98% purity, highly toxic</td></tr><tr><td>CELLSTAR tube</td><td>Greiner Bio-One</td><td>210261</td><td>50 ml</td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5424R</td><td>to eliminate spent media from the bacterial culture</td></tr><tr><td>Digital Incubators Incu-Line</td><td>VWR-CH</td><td>390-0384</td><td>to bake 3D master</td></tr><tr><td>Duster</td><td>VWR-CH</td><td>16650-22</td><td>to clean the wafer and microchannels</td></tr><tr><td>Hot plate</td><td>VWR-CH</td><td>444-0601</td><td>to bond the microchannels</td></tr><tr><td>Isopropanol</td><td>Sigma-Aldrich</td><td>W292907</td><td></td></tr><tr><td>LightSafe micro centrifuge tubes</td><td>Sigma-Aldrich</td><td>Z688312</td><td>1.5 ml</td></tr><tr><td>MATLAB</td><td>Mathworks</td><td></td><td>for image analysis and bacterial tracking</td></tr><tr><td>Microcentrifuge tube</td><td>Eppendorf</td><td>30120086</td><td>1.5 ml</td></tr><tr><td>Microscope glass slide</td><td>VWR-CH</td><td>631-1552</td><td></td></tr><tr><td>Microscope Nikon Eclipse TiE</td><td>Nikon Instruments</td><td>MEA53100</td><td>with motorized stage</td></tr><tr><td>MNI-Glutamate</td><td>Tocris Bioscience</td><td>1490</td><td>&gt;98 % purity, photosensitive</td></tr><tr><td>Mold printing equipment</td><td>Stratasys</td><td></td><td>Objet30 3D printer</td></tr><tr><td>Mold printing service</td><td>3D Printing Studios</td><td>Custom</td><td>&lt;a href=&quot;https://www.3dprintingstudios.com/&quot; target=&quot;_blank&quot;&gt;https://www.3dprintingstudios.com/&lt;/a&gt;</td></tr><tr><td>Nanodrop One UV-Vis Spectrophotometer</td><td>Thermo Fisher Scientific</td><td>ND-ONE-W</td><td>to calibrate the uncaging</td></tr><tr><td>NIS Elements</td><td>Nikon Instruments</td><td></td><td>Microscope Imaging Software</td></tr><tr><td>Oven Venti-Line</td><td>VWR-CH</td><td>466-3516</td><td>to bake PDMS (with forced convection)</td></tr><tr><td>Photoresist SU-8-3050</td><td>MicroChem Corp.</td><td>SU8-3050</td><td></td></tr><tr><td>Plasma chamber Zepto</td><td>Diener Electronic</td><td>ZEPTO-1</td><td>to functionalize the surfaces before bonding</td></tr><tr><td>Polycarbonate membrane</td><td>Sterlitech</td><td>PCT0447100</td><td>0.4 &amp;micro;m pore size, 19 % open area, 24 &amp;micro;m thickness</td></tr><tr><td>Polyethylene microtubing</td><td>Scientific Commodities</td><td>BB31695-PE/2</td><td>I.D. x O.D.: 0.015&quot; x 0.043&quot; / 0.38mm x 1.09mm</td></tr><tr><td>Polystyrene Petri dish</td><td>VWR-CH</td><td>25373-100</td><td>bottom surface (90 mm x 15 mm) to bond the millifluidic device</td></tr><tr><td>Scale</td><td>VWR-CH</td><td>611-2605</td><td>to weight PDMS mixture</td></tr><tr><td>sCMOS camera Andor Zyla</td><td>Oxford Instruments</td><td></td><td>for phase contrast and fluorescence microscopy (max 100 fps)</td></tr><tr><td>Sea salt</td><td>Instant Ocean</td><td>Product No. SS1-160p</td><td></td></tr><tr><td>SolidWorks 2015</td><td>Dassault Systemes SolidWorks</td><td></td><td>Used to design the mold</td></tr><tr><td>Spectra X light engine</td><td>Lumencolor</td><td></td><td>for LED 395 nm</td></tr><tr><td>Sylgard 184</td><td>Dow Corning</td><td>110-41-155</td><td>PDMS Si Elastomer Kit; curing agent</td></tr><tr><td>Syringe (Luer-Lok)</td><td>B Braun Omnifix</td><td>4616308F</td><td></td></tr><tr><td>Syringe Needle</td><td>Agani</td><td>A228</td><td>from 10 to 30 ml</td></tr><tr><td>Syringe Pump 11 Pico Plus Elite</td><td>Harvard Apparatus</td><td>70-4506</td><td>Terumo Agani 23 gauge 5/8 inch (16mm)</td></tr><tr><td>VeroGrey</td><td>Stratasys</td><td></td><td>Dual Syringe Pump</td></tr><tr><td>Vortex-Genie</td><td>Scientific Industries</td><td>SI-0236</td><td>Mold Material</td></tr></tbody></table>

generating controlled, dynamic chemical landscapes to study microbial behavior

We demonstrate a method for the generation of controlled, dynamic chemical pulses&#8213;where localized chemoattractant becomes suddenly available at the microscale&#8213;to create micro-environments for microbial chemotaxis experiments. To create chemical pulses, we developed a system to introduce amino acid sources near-instantaneously by photolysis of caged amino acids within a polydimethylsiloxane (PDMS) microfluidic chamber containing a bacterial suspension. We applied this method to the chemotactic bacterium, Vibrio ordalii, which can actively climb these dynamic chemical gradients while being tracked by video microscopy. Amino acids, rendered biologically inert (&#39;caged&#39;) by chemical modification with a photoremovable protecting group, are uniformly present in the suspension but not available for consumption until their sudden release, which occurs at user-defined points in time and space by means of a near-UV-A focused LED beam. The number of molecules released in the pulse can be determined by a calibration relationship between exposure time and uncaging fraction, where the absorption spectrum after photolysis is characterized by using UV-Vis spectroscopy. A nanoporous polycarbonate (PCTE) membrane can be integrated into the microfluidic device to allow the continuous removal by flow of the uncaged compounds and the spent media. A strong, irreversible bond between the PCTE membrane and the PDMS microfluidic structure is achieved by coating the membrane with a solution of 3-aminopropyltriethoxysilane (APTES) followed by plasma activation of the surfaces to be bonded. A computer-controlled system can generate user-defined sequences of pulses at different locations and with different intensities, so as to create resource landscapes with prescribed spatial and temporal variability. In each chemical landscape, the dynamics of bacterial movement at the individual scale and their accumulation at the population level can be obtained, thereby allowing the quantification of chemotactic performance and its effects on bacterial aggregations in ecologically relevant environments.

We used the microfluidic and millifluidic devices (Figure 1) to study bacterial accumulation profiles under dynamic nutrient conditions. Bacterial trajectories were extracted from recorded videos acquired by phase contrast microscopy of the accumulation-dissipation dynamics of a bacterial population following a chemical pulse released by photolysis (Figure 2 and Figure 3). By averaging millions of trajectories, the spatiotemporal dy...

Watch this Scientific Journal Video about Generating Controlled, Dynamic Chemical Landscapes to Study Microbial Behavior at JoVE.com

Generating Controlled, Dynamic Chemical Landscapes to Study Microbial Behavior

A protocol for the generation of dynamic chemical landscapes by photolysis within microfluidic and millifluidic setups is presented. This methodology is suitable to study diverse biological processes, including the motile behavior, nutrient uptake, or adaptation to chemicals of microorganisms, both at the single cell and population level.

We demonstrate a method for the generation of controlled, dynamic chemical pulses&#8213;where localized chemoattractant becomes suddenly available at the ...

generating-controlled-dynamic-chemical-landscapes-to-study-microbial

Research

JoVE Journal

Bioengineering

6.0K Views.  Institute of Environmental Engineering, Department of Civil, Environmental and Geomatic Engineering. Deze methode maakt het mogelijk om de microschaal ecologie en het gedrag van micro-organismen navigeren dynamische chemische gradiënten te verduidelijken en om hun verborgen trade-offs, nutriënten kinetiek, en de dynamiek van de bevolking in ecologisch relevante micro-omgevingen te ontdekken. Door microfluïde technologie te combineren met fotolyse, genereren we gecontroleerde chemische pulsen, waarbij gelokaliseerde chemoattractant plotseling beschikbaar komt op microschaal, om de chemotac...

Video: Generating Controlled, Dynamic Chemical Landscapes to Study Microbial Behavior

Monitoring Spatial Segregation in Surface Colonizing Microbial Populations

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.

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 ...

Genetics

Chemotactic Response of Marine Micro-Organisms to Micro-Scale Nutrient Layers

The degree to which planktonic microbes can exploit microscale resource patches will have considerable implications for oceanic trophodynamics and biogeochemical flux. However, to take advantage of nutrient patches in the ocean, swimming microbes must overcome the influences of physical forces including molecular diffusion and turbulent shear, which will limit the availability of patches and the ability of bacteria to locate them. Until recently, methodological limitations have precluded direct examinations of microbial behaviour within patchy habitats and realistic small-scale flow conditions. Hence, much of our current knowledge regarding microbial behaviour in the ocean has been procured from theoretical predictions. To obtain new information on microbial foraging behaviour in the ocean we have applied soft lithographic fabrication techniques to develop 2 microfluidic devices, which we have used to create (i) microscale nutrient patches with dimensions and diffusive characteristics relevant to oceanic processes and (ii) microscale vortices, with shear rates corresponding to those expected in the ocean. These microfluidic devices have permitted a first direct examination of microbial swimming and chemotactic behaviour within a heterogeneous and dynamic seascape. The combined use of epifluorescence and phase contrast microscopy allow direct examinations of the physical dimensions and diffusive characteristics of nutrient patches, while observing the population-level aggregative response, in addition to the swimming behaviour of individual microbes. These experiments have revealed that some species of phytoplankton, heterotrophic bacteria and phagotrophic protists are adept at locating and exploiting diffusing microscale resource patches within very short time frames. We have also shown that up to moderate shear rates, marine bacteria are able to  fight the flow  and swim through their environment at their own accord. However, beyond a threshold high shear level, bacteria are aligned in the shear flow and are less capable of swimming without disturbance from the flow. Microfluidics represents a novel and inexpensive approach for studying aquatic microbial ecology, and due to its suitability for accurately creating realistic flow fields and substrate gradients at the microscale, is ideally applicable to examinations of microbial behaviour at the smallest scales of interaction.  We therefore suggest that microfluidics represents a valuable tool for obtaining a better understanding of the ecology of microorganisms in the ocean.

The fabrication of microfluidic channels and their implementation in experiments for studying the chemotactic foraging behaviour of marine microbes within a patchy nutrient seascape and the swimming behaviour of bacteria within shear flow are described.

The degree to which planktonic microbes can exploit microscale resource patches will have considerable implications for oceanic trophodynamics and ...

Biology

Soil Lysimeter Excavation for Coupled Hydrological, Geochemical, and Microbiological Investigations

Studying co-evolution of hydrological and biogeochemical processes in the subsurface of natural landscapes can enhance the understanding of coupled Earth-system processes. Such knowledge is imperative in improving predictions of hydro-biogeochemical cycles, especially under climate change scenarios. We present an experimental method, designed to capture sub-surface heterogeneity of an initially homogeneous soil system. This method is based on destructive sampling of a soil lysimeter designed to simulate a small-scale hillslope. A weighing lysimeter of one cubic meter capacity was divided into sections (voxels) and was excavated layer-by-layer, with sub samples being collected from each voxel. The excavation procedure was aimed at detecting the incipient heterogeneity of the system by focusing on the spatial assessment of hydrological, geochemical, and microbiological properties of the soil. Representative results of a few physicochemical variables tested show the development of heterogeneity. Additional work to test interactions between hydrological, geochemical, and microbiological signatures is planned to interpret the observed patterns. Our study also demonstrates the possibility of carrying out similar excavations in order to observe and quantify different aspects of soil-development under varying environmental conditions and scale.

This study presents an excavation method for investigating subsurface hydrological, geochemical, and microbiological heterogeneity of a soil lysimeter. The lysimeter simulates an artificial hillslope which was initially under homogeneous condition and had been subjected to approximately 5,000 mm of water over eight cycles of irrigation in an 18-month period.

Studying co-evolution of hydrological and biogeochemical processes in the subsurface of natural landscapes can enhance the understanding of coupled ...

Environment

Microfluidic Picoliter Bioreactor for Microbial Single-cell Analysis: Fabrication, System Setup, and Operation

In this protocol the fabrication, experimental setup and basic operation of the recently introduced microfluidic picoliter bioreactor (PLBR) is described in detail. The PLBR can be utilized for the analysis of single bacteria and microcolonies to investigate biotechnological and microbiological related questions concerning, e.g.&#160;cell growth, morphology, stress response, and metabolite or protein production on single-cell level. The device features continuous media flow enabling constant environmental conditions for perturbation studies, but in addition allows fast medium changes as well as oscillating conditions to mimic any desired environmental situation. To fabricate the single use devices, a silicon wafer containing sub micrometer sized SU-8 structures served as the replication mold for rapid polydimethylsiloxane casting. Chips were cut, assembled, connected, and set up onto a high resolution and fully automated microscope suited for time-lapse imaging, a powerful tool for spatio-temporal cell analysis. Here, the biotechnological platform organism Corynebacterium glutamicum&#160;was seeded into the PLBR and cell growth and intracellular fluorescence were followed over several hours unraveling time dependent population heterogeneity on single-cell level, not possible with conventional analysis methods such as flow cytometry. Besides insights into device fabrication, furthermore, the preparation of the preculture, loading, trapping of bacteria, and the PLBR cultivation of single cells and colonies is demonstrated. These devices will add a new dimension in microbiological research to analyze time dependent phenomena of single bacteria under tight environmental control. Due to the simple and relatively short fabrication process the technology can be easily adapted at any microfluidics lab and simply tailored towards specific needs.

In this protocol the fabrication, setup and basic operation of a microfluidic picoliter bioreactor (PLBR) for single-cell analysis of prokaryotic microorganisms is introduced. Industrially relevant microorganisms were analyzed as proof of principle allowing insights into growth rate, morphology, and phenotypic heterogeneity over certain time periods, hardly possible with conventional methods.

In this protocol the fabrication, experimental setup and basic operation of the recently introduced microfluidic picoliter bioreactor (PLBR) is described ...

In Situ Chemotaxis Assay to Examine Microbial Behavior in Aquatic Ecosystems

Microbial behaviors, such as motility and chemotaxis (the ability of a cell to alter its movement in response to a chemical gradient), are widespread across the bacterial and archaeal domains. Chemotaxis can result in substantial resource acquisition advantages in heterogeneous environments. It also plays a crucial role in symbiotic interactions, disease, and global processes, such as biogeochemical cycling. However, current techniques restrict chemotaxis research to the laboratory and are not easily applicable in the field. Presented here is a step-by-step protocol for the deployment of the in situ chemotaxis assay (ISCA), a device that enables robust interrogation of microbial chemotaxis directly in the natural environment. The ISCA is a microfluidic device consisting of a 20 well array, in which chemicals of interest can be loaded. Once deployed in aqueous environments, chemicals diffuse out of the wells, creating concentration gradients that microbes sense and respond to by swimming into the wells via chemotaxis. The well contents can then be sampled and used to (1) quantify strength of the chemotactic responses to specific compounds through flow cytometry, (2) isolate and culture responsive microorganisms, and (3) characterize the identity and genomic potential of the responding populations through molecular techniques. The ISCA is a flexible platform that can be deployed in any system with an aqueous phase, including marine, freshwater, and soil environments.

Presented here is the protocol for an in situ chemotaxis assay, a recently developed microfluidic device that enables studies of microbial behavior directly in the environment.

Microbial behaviors, such as motility and chemotaxis (the ability of a cell to alter its movement in response to a chemical gradient), are widespread ...

Modeling Methanotroph Niches – The Gradient Syringe Model Ecosystem

Agarose-Based Model Ecosystem for Cultivating Methanotrophs in a Methane-Oxygen Counter Gradient

Aerobic methane-oxidizing bacteria, known as methanotrophs, serve important roles in biogeochemical cycling. Methanotrophs occupy a specific environmental niche within methane-oxygen counter gradients found in soils and sediments, which influences their behavior on an individual and community level. However, conventional methods to study the physiology of these greenhouse gas-mitigating microorganisms often use homogeneous planktonic cultures, which do not accurately represent the spatial and chemical gradients found in the environment. This hinders scientists' understanding of how these bacteria behave in situ. Here, a simple, inexpensive model ecosystem called the gradient syringe is described, which uses semi-solid agarose to recreate the steep methane-oxygen counter gradients characteristic of methanotrophs' natural habitats. The gradient syringe allows for the cultivation of methanotrophic strains and the enrichment of mixed methane-oxidizing consortia from environmental samples, revealing phenotypes only visible in this spatially resolved context. This protocol also reports various biochemical assays that have been modified to be compatible with the semi-solid agarose matrix, which may be valuable to researchers culturing microorganisms within other agarose-based systems.

Author Spotlight: Designing Simple and Inexpensive Techniques to Grow Methane-Oxidizing Bacteria in the Laboratory

A protocol is described for preparing a simple model ecosystem that recreates the methane-oxygen counter gradient found in the natural habitat of aerobic methane-oxidizing bacteria, enabling the study of their physiology in a spatially resolved context. Modifications to common biochemical assays for use with the agarose-based model ecosystem are also described.

Aerobic methane-oxidizing bacteria, known as methanotrophs, serve important roles in biogeochemical cycling. Methanotrophs occupy a specific environmental ...

Window on a Microworld: Simple Microfluidic Systems for Studying Microbial Transport in Porous Media

Microbial growth and transport in porous media have important implications for the quality of groundwater and surface water, the recycling of nutrients in the environment, as well as directly for the transmission of pathogens to drinking water supplies. Natural porous media is composed of an intricate physical topology, varied surface chemistries, dynamic gradients of nutrients and electron acceptors, and a patchy distribution of microbes. These features vary substantially over a length scale of microns, making the results of macro-scale investigations of microbial transport difficult to interpret, and the validation of mechanistic models challenging. Here we demonstrate how simple microfluidic devices can be used to visualize microbial interactions with micro-structured habitats, to identify key processes influencing the observed phenomena, and to systematically validate predictive models. Simple, easy-to-use flow cells were constructed out of the transparent, biocompatible and oxygen-permeable material poly(dimethyl siloxane). Standard methods of photolithography were used to make micro-structured masters, and replica molding was used to cast micro-structured flow cells from the masters. The physical design of the flow cell chamber is adaptable to the experimental requirements: microchannels can vary from simple linear connections to complex topologies with feature sizes as small as 2 &mu;m. Our modular EcoChip flow cell array features dozens of identical chambers and flow control by a gravity-driven flow module. We demonstrate that through use of EcoChip devices, physical structures and pressure heads can be held constant or varied systematically while the influence of surface chemistry, fluid properties, or the characteristics of the microbial population is investigated. Through transport experiments using a non-pathogenic, green fluorescent protein-expressing Vibrio bacterial strain, we illustrate the importance of habitat structure, flow conditions, and inoculums size on fundamental transport phenomena, and with real-time particle-scale observations, demonstrate that microfluidics offer a compelling view of a hidden world.

Microfluidic devices can be used to visualize complex natural processes in real time and at the appropriate physical scales. We have developed a simple microfluidic device that mimics key features of natural porous media for studying growth and transport of bacteria in the subsurface.

Microbial growth and transport in porous media have important implications for the quality of groundwater and surface water, the recycling of nutrients in ...

Generation of Dynamical Environmental Conditions using a High-Throughput Microfluidic Device

Mimicking in vivo environmental conditions is crucial for in vitro studies on complex life machinery. However, current techniques targeting live cells and organs are either highly expensive, like robotics, or lack nanoliter volume and millisecond time accuracy in liquid manipulation. We herein present the design and fabrication of a microfluidic system, which consists of 1,500 culture units, an array of enhanced peristaltic pumps and an on-site mixing modulus. To demonstrate the capacities of the microfluidic device, neural stem cell (NSC) spheres are maintained in the proposed system. We observed that when the NSC sphere is exposed to CXCL in day 1 and EGF in day 2, the round-shaped conformation is well maintained. Variation in the input order of 6 drugs causes morphological changes to the NSC sphere and the expression level representative marker for NSC stemness (i.e., Hes5 and Dcx). These results indicate that dynamic and complex environmental conditions have great effects on NSC differentiation and self-renewal, and the proposed microfluidic device is a suitable platform for high throughput studies on the complex life machinery.

We present a microfluidic system for high throughput studies on complex life machinery, which consists of 1500 culture units, an array of enhanced peristaltic pumps and an on-site mixing modulus. The microfluidic chip allows for the analysis of the highly complex and dynamic micro-environmental conditions in vivo.

Mimicking in vivo environmental conditions is crucial for in vitro studies on complex life machinery. However, current techniques targeting live cells and ...

The Use of Chemostats in Microbial Systems Biology

Cells regulate their rate of growth in response to signals from the external world. As the cell grows, diverse cellular processes must be coordinated including macromolecular synthesis, metabolism and ultimately, commitment to the cell division cycle. The chemostat, a method of experimentally controlling cell growth rate, provides a powerful means of systematically studying how growth rate impacts cellular processes - including gene expression and metabolism - and the regulatory networks that control the rate of cell growth. When maintained for hundreds of generations chemostats can be used to study adaptive evolution of microbes in environmental conditions that limit cell growth. We describe the principle of chemostat cultures, demonstrate their operation and provide examples of their various applications. Following a period of disuse after their introduction in the middle of the twentieth century, the convergence of genome-scale methodologies with a renewed interest in the regulation of cell growth and the molecular basis of adaptive evolution is stimulating a renaissance in the use of chemostats in biological research.

Cell growth rate is a regulated process and a primary determinant of cell physiology. Continuous culturing using chemostats enables extrinsic control of cell growth rate by nutrient limitation facilitating the study of molecular networks that control cell growth and how those networks evolve to optimize cell growth.

Cells regulate their rate of growth in response to signals from the external world. As the cell grows, diverse cellular processes must be coordinated ...

Design and Use of Multiplexed Chemostat Arrays

Chemostats are continuous culture systems in which cells are grown in a tightly controlled, chemically constant environment where culture density is constrained by limiting specific nutrients.1,2 Data from chemostats are highly reproducible for the measurement of quantitative phenotypes as they provide a constant growth rate and environment at steady state. For these reasons, chemostats have become useful tools for fine-scale characterization of physiology through analysis of gene expression3-6 and other characteristics of cultures at steady-state equilibrium.7 Long-term experiments in chemostats can highlight specific trajectories that microbial populations adopt during adaptive evolution in a controlled environment. In fact, chemostats have been used for experimental evolution since their invention.8 A common result in evolution experiments is for each biological replicate to acquire a unique repertoire of mutations.9-13 This diversity suggests that there is much left to be discovered by performing evolution experiments with far greater throughput. 
We present here the design and operation of a relatively simple, low cost array of miniature chemostats&mdash;or ministats&mdash;and validate their use in determination of physiology and in evolution experiments with yeast. This approach entails growth of tens of chemostats run off a single multiplexed peristaltic pump. The cultures are maintained at a 20 ml working volume, which is practical for a variety of applications. It is our hope that increasing throughput, decreasing expense, and providing detailed building and operation instructions may also motivate research and industrial application of this design as a general platform for functionally characterizing large numbers of strains, species, and growth parameters, as well as genetic or drug libraries.

We developed and validated a small-footprint array of miniature chemostats built from readily available parts for low cost. Physiological and experimental evolution results were similar to larger volume chemostats. The ministat array provides a compact, inexpensive, and accessible platform for traditional chemostat experiments, functional genomics, and chemical screening applications.

Chemostats are continuous culture systems in which cells are grown in a tightly controlled, chemically constant environment where culture density is ...

Generating Controlled, Dynamic Chemical Landscapes to Study Microbial Behavior

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