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>

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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>(3-Aminopropyl) triethoxysilane (APTES)</td>
			<td>Sigma-Aldrich</td>
			<td>A3648</td>
			<td>&#62;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>&#62;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><a href="https://www.3dprintingstudios.com/" target="_blank">https://www.3dprintingstudios.com/</a></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 &#181;m pore size, 19 % open area, 24 &#181;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&#34; x 0.043&#34; / 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. 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.

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

Micropatterned Surfaces to Study Hyaluronic Acid Interactions with Cancer Cells

Cancer invasion and progression involves a motile cell phenotype, which is under complex regulation by growth factors/cytokines and extracellular matrix (ECM) components within the tumor microenvironment. Hyaluronic acid (HA) is one stromal ECM component that is known to facilitate tumor progression by enhancing invasion, growth, and angiogenesis1. Interaction of HA with its cell surface receptor CD44 induces signaling events that promote tumor cell growth, survival, and migration, thereby increasing metastatic spread2-3. HA is an anionic, nonsulfated glycosaminoglycan composed of repeating units of D-glucuronic acid and D-N-acetylglucosamine. Due to the presence of carboxyl and hydroxyl groups on repeating disaccharide units, native HA is largely hydrophilic and amenable to chemical modifications that introduce sulfate groups for photoreative immobilization 4-5. Previous studies involving the immobilizations of HA onto surfaces utilize the bioresistant behavior of HA and its sulfated derivative to control cell adhesion onto surfaces6-7. In these studies cell adhesion preferentially occurs on non-HA patterned regions.
To analyze cellular interactions with exogenous HA, we have developed patterned functionalized surfaces that enable a controllable study and high-resolution visualization of cancer cell interactions with HA. We utilized microcontact printing (uCP) to define discrete patterned regions of HA on glass surfaces. A "tethering" approach that applies carbodiimide linking chemistry to immobilize HA was used 8. Glass surfaces were microcontact printed with an aminosilane and reacted with a HA solution of optimized ratios of EDC and NHS to enable HA immobilization in patterned arrays. Incorporating carbodiimide chemistry with mCP enabled the immobilization of HA to defined regions, creating surfaces suitable for in vitro applications. Both colon cancer cells and breast cancer cells implicitly interacted with the HA micropatterned surfaces. Cancer cell adhesion occurred within 24 hours with proliferation by 48 hours. Using HA micropatterned surfaces, we demonstrated that cancer cell adhesion occurs through the HA receptor CD44. Furthermore, HA patterned surfaces were compatible with scanning electron microscopy (SEM) and allowed high resolution imaging of cancer cell adhesive protrusions and spreading on HA patterns to analyze cancer cell motility on exogenous HA.

A novel approach that allows the high-resolution analysis of cancer cell interactions with exogenous hyaluronic acid (HA) is described. Patterned surfaces are fabricated by combining carbodiimide chemistry and microcontact printing.

Cancer invasion and progression involves a motile cell phenotype, which is under complex regulation by growth factors/cytokines and extracellular matrix ...

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Synthesis of an Intein-mediated Artificial Protein Hydrogel

We present the synthesis of a highly stable protein hydrogel mediated by a split-intein-catalyzed protein trans-splicing reaction. The building blocks of this hydrogel are two protein block-copolymers each containing a subunit of a trimeric protein that serves as a crosslinker and one half of a split intein. A highly hydrophilic random coil is inserted into one of the block-copolymers for water retention. Mixing of the two protein block copolymers triggers an intein trans-splicing reaction, yielding a polypeptide unit with crosslinkers at either end that rapidly self-assembles into a hydrogel. This hydrogel is very stable under both acidic and basic conditions, at temperatures up to 50 &#176;C, and in organic solvents. The hydrogel rapidly reforms after shear-induced rupture. Incorporation of a &#34;docking station peptide&#34; into the hydrogel building block enables convenient incorporation of &#34;docking protein&#34;-tagged target proteins. The hydrogel is compatible with tissue culture growth media, supports the diffusion of 20 kDa molecules, and enables the immobilization of bioactive globular proteins. The application of the intein-mediated protein hydrogel as an organic-solvent-compatible biocatalyst was demonstrated by encapsulating the horseradish peroxidase enzyme and corroborating its activity.

We present the synthesis of a split-intein-mediated protein hydrogel. The building blocks of this hydrogel are two protein copolymers each containing a subunit of a trimeric protein that serves as a crosslinker and one half of a split intein. Mixing of the two protein copolymers triggers an intein trans-splicing reaction, yielding a polypeptide unit that self-assembles into a hydrogel. This hydrogel is highly pH- and temperature-stable, compatible with organic solvents, and easily incorporates functional globular proteins.

We present the synthesis of a highly stable protein hydrogel mediated by a split-intein-catalyzed protein trans-splicing reaction. The building blocks of ...

Controlled Microfluidic Environment for Dynamic Investigation of Red Blood Cell Aggregation

Blood, as a non-Newtonian biofluid, represents the focus of numerous studies in the hemorheology field. Blood constituents include red blood cells, white blood cells and platelets that are suspended in blood plasma. Due to the abundance of the RBCs (40% to 45% of the blood volume), their behavior dictates the rheological behavior of blood especially in the microcirculation. At very low shear rates, RBCs are seen to assemble and form entities called aggregates, which causes the non-Newtonian behavior of blood. It is important to understand the conditions of the aggregates formation to comprehend the blood rheology in microcirculation. The protocol described here details the experimental procedure to determine quantitatively the RBC aggregates in microcirculation under constant shear rate, based on image processing. For this purpose, RBC-suspensions are tested and analyzed in 120 x 60 &#181;m poly-dimethyl-siloxane (PDMS) microchannels. The RBC-suspensions are entrained using a second fluid in order to obtain a linear velocity profile within the blood layer and thus achieve a wide range of constant shear rates. The shear rate is determined using a micro Particle Image Velocimetry (&#181;PIV) system, while RBC aggregates are visualized using a high speed camera. The videos captured of the RBC aggregates are analyzed using image processing techniques in order to determine the aggregate sizes based on the images intensities.

The protocol described details an experimental procedure to quantify Red Blood Cell (RBC) aggregates under a controlled and constant shear rate, based on image processing techniques. The goal of this protocol is to relate RBC aggregate sizes to the corresponding shear rate in a controlled microfluidic environment.

Blood, as a non-Newtonian biofluid, represents the focus of numerous studies in the hemorheology field. Blood constituents include red blood cells, white ...

A Novel Bioreactor for High Density Cultivation of Diverse Microbial Communities

A novel reactor design, coined a high density bioreactor (HDBR), is presented for the cultivation and study of high density microbial communities. Past studies have evaluated the performance of the reactor for the removal of COD1 and nitrogen species2-4 by heterotrophic and chemoautotrophic bacteria, respectively. The HDBR design eliminates the requirement for external flocculation/sedimentation processes while still yielding effluent containing low suspended solids. In this study, the HDBR is applied as a photobioreactor (PBR) in order to characterize the nitrogen removal characteristics of an algae-based photosynthetic microbial community. As previously reported for this HDBR design, a stable biomass zone was established with a clear delineation between the biologically active portion of the reactor and the recycling reactor fluid, which resulted in a low suspended solid effluent. The algal community in the HDBR was observed to remove 18.4% of total nitrogen species in the influent. Varying NH4+ and NO3- concentrations in the feed did not have an effect on NH4+ removal (n=44, p=0.993 and n=44, p=0.610 respectively) while NH4+ feed concentration was found to be negatively related with NO3- removal (n=44, p=0.000) and NO3- feed concentration was found to be positively correlated with NO3- removal (n=44, p=0.000). Consistent removal of NH4+, combined with the accumulation of oxidized nitrogen species at high NH4+ fluxes indicates the presence of ammonia- and nitrite-oxidizing bacteria within the microbial community.

A novel reactor design, coined a high density bioreactor (HDBR), is presented for the cultivation and study of high density microbial communities. Here, the HDBR is successfully applied in a photobioreactor (PBR) configuration for the study of nitrogen metabolism by a mixed high density algal community.

A novel reactor design, coined a high density bioreactor (HDBR), is presented for the cultivation and study of high density microbial communities. Past ...

Alternating Magnetic Field-Responsive Hybrid Gelatin Microgels for Controlled Drug Release

Magnetically-responsive nano/micro-engineered biomaterials that enable a tightly controlled, on-demand drug delivery have been developed as new types of smart soft devices for biomedical applications. Although a number of magnetically-responsive drug delivery systems have demonstrated efficacies through either in vitro proof of concept studies or in vivo preclinical applications, their use in clinical settings is still limited by their insufficient biocompatibility or biodegradability. Additionally, many of the existing platforms rely on sophisticated techniques for their fabrications. We recently demonstrated the fabrication of biodegradable, gelatin-based thermo-responsive microgel by physically entrapping poly(N-isopropylacrylamide-co-acrylamide) chains as a minor component within a three-dimensional gelatin network. In this study, we present a facile method to fabricate a biodegradable drug release platform that enables a magneto-thermally triggered drug release. This was achieved by incorporating superparamagnetic iron oxide nanoparticles and thermo-responsive polymers within gelatin-based colloidal microgels, in conjunction with an alternating magnetic field application system.

We present a facile method to fabricate a biodegradable gelatin-based drug release platform that is magneto-thermally responsive. This was achieved by incorporating superparamagnetic iron oxide nanoparticles and poly(N-isopropylacrylamide-co-acrylamide) within a spherical gelatin micro-network crosslinked by genipin, in conjunction with an alternating magnetic field application system.

Magnetically-responsive nano/micro-engineered biomaterials that enable a tightly controlled, on-demand drug delivery have been developed as new types of ...

Expanding Nanopatterned Substrates Using Stitch Technique for Nanotopographical Modulation of Cell Behavior

Substrate nanotopography has been shown to be a potent modulator of cell phenotype and function. To dissect nanotopography modulation of cell behavior, a large area of nanopatterned substrate is desirable so that enough cells can be cultured on the nanotopography for subsequent biochemical and molecular biology analyses. However, current nanofabrication techniques have limitations to generate highly defined nanopatterns over a large area. Herein, we present a method to expand nanopatterned substrates from a small, highly defined nanopattern to a large area using stitch technique. The method combines multiple techniques, involving soft lithography to replicate poly(dimethylsiloxane) (PDMS) molds from a well-defined mold, stitch technique to assemble multiple PDMS molds to a single large mold, and nanoimprinting to generate a master mold on polystyrene (PS) substrates. With the PS master mold, we produce PDMS working substrates and demonstrate nanotopographical modulation of cell spreading. This method provides a simple, affordable yet versatile avenue to generate well-defined nanopatterns over large areas, and is potentially extended to create micro-/nanoscale devices with hybrid components.

A protocol for producing a large area of nanopatterned substrate from small nanopatterned molds for study of nanotopographical modulation of cell behavior is presented.

Substrate nanotopography has been shown to be a potent modulator of cell phenotype and function. To dissect nanotopography modulation of cell behavior, a ...

Modeling Ovarian Cancer Multicellular Spheroid Behavior in a Dynamic 3D Peritoneal Microdevice

Ovarian cancer is characterized by extensive peritoneal metastasis, with tumor spheres commonly found in the malignant ascites. This is associated with poor clinical outcomes and currently lacks effective treatment. Both the three-dimensional (3D) environment and the dynamic mechanical forces are very important factors in this metastatic cascade. However, traditional cell cultures fail to recapitulate this natural tumor microenvironment. Thus, in vivo-like models that can emulate the intraperitoneal environment are of obvious importance. In this study, a new microfluidic platform of the peritoneum was set up to mimic the situation of ovarian cancer spheroids in the peritoneal cavity during metastasis. Ovarian cancer spheroids generated under a non-adherent condition were cultured in microfluidic channels coated with peritoneal mesothelial cells subjected to physiologically relevant shear stress. In summary, this dynamic 3D ovarian cancer-mesothelium microfluidic platform can provide new knowledge on basic cancer biology and serve as a platform for potential drug screening and development.

To study ovarian tumor progression in a physiologically relevant model, multicellular spheroids were cultured in a microdevice under simulated fluid flow. This dynamic 3D model emulates the intraperitoneal environment with the cellular and mechanical components where ovarian cancer metastasis occurs.

Ovarian cancer is characterized by extensive peritoneal metastasis, with tumor spheres commonly found in the malignant ascites. This is associated with ...

Environmental Dynamic Mechanical Analysis to Predict the Softening Behavior of Neural Implants

When using dynamically softening substrates for neural implants, it is important to have a reliable in vitro method to characterize the softening behavior of these materials. In the past, it has not been possible to satisfactorily measure the softening of thin films under conditions mimicking body environment without substantial effort. This publication presents a new and simple method that allows dynamic mechanical analysis (DMA) of polymers in solutions, such as phosphate buffered saline (PBS), at relevant temperatures. The use of environmental DMA allows measurement of the softening effects of polymers due to plasticization in various media and temperatures, which therefore allows a prediction of the materials behavior under in vivo conditions.

To allow reliable predictions of the softening of polymeric substrates for neural implants in an in vivo environment, it is important to have a reliable in vitro method. Here, the use of dynamic mechanical analysis in phosphate buffered saline at body temperature is presented.

When using dynamically softening substrates for neural implants, it is important to have a reliable in vitro method to characterize the softening behavior ...

Stepwise Cell Seeding on Tessellated Scaffolds to Study Sprouting Blood Vessels

The cardiovascular system is a key player in human physiology, providing nourishment to most tissues in the body; vessels are present in different sizes, structures, phenotypes, and performance depending on each specific perfused tissue. The field of tissue engineering, which aims to repair or replace damaged or missing body tissues, relies on controlled angiogenesis to create a proper vascularization within the engineered tissues. Without a vascular system, thick engineered constructs cannot be sufficiently nourished, which may result in cell death, poor engraftment, and ultimately failure. Thus, understanding and controlling the behavior of engineered blood vessels is an outstanding challenge in the field. This work presents a high-throughput system that allows for the creation of organized and repeatable vessel networks for studying vessel behavior in a 3D scaffold environment. This two-step seeding protocol shows that vessels within the system react to the scaffold topography, presenting distinctive sprouting behaviors depending on the compartment geometry in which the vessels reside. The obtained results and understanding from this high throughput system can be applied in order to inform better 3D bioprinted scaffold construct designs, wherein fabrication of various 3D geometries cannot be rapidly assessed when using 3D printing as the basis for cellularized biological environments. Furthermore, the understanding from this high throughput system may be utilized for the improvement of rapid drug screening, the rapid development of co-cultures models, and the investigation of mechanical stimuli on blood vessel formation to deepen the knowledge of the vascular system.

Engineered tissues heavily rely on proper vascular networks to provide vital nutrients and gases and remove metabolic waste. In this work, a stepwise seeding protocol of endothelial cells and support cells creates highly organized vascular networks in a high-throughput platform for studying developing vessel behavior in a controlled 3D environment.

The cardiovascular system is a key player in human physiology, providing nourishment to most tissues in the body; vessels are present in different sizes, ...