The authors would like to thank Professor Howard Ceri from the Biological Sciences Department of the University of Calgary for providing bacterial strains. A. Kumar acknowledges support from NSERC. T. Thundat acknowledges financial support from the Canada Excellence Research Chair (CERC) program. The authors would also like to acknowledge help from Ms. Zahra Nikakhtari for help with videography.

Perform the experimental protocols here in the order described below. Microfabrication protocols for creating the microfluidic platform are discussed in Step 1. Step 2 describes the bacterial culture protocol (Figure 2), and Step 3 pertains to assembly of the experimental setup (Figure 3). Finally, the actual experimental step is described in Step 4.
1. Chip Fabrication Procedure
NOTE: Proper safety procedures must be followed for the processes described below. Consult the institutional safety officer for details.
<ol>
	<li>Design the mask with an appropriate software (e.g., L-Edit). The channel design consists of a main micro-channel of width 625 &#956;m. The central region of the channel contains an array of micro-posts 50 &#956;m in diameter, spaced 25 &#956;m apart (See Supplemental Files).</li>
	<li>Print this design on glass (5&#34; x 5&#34; soda lime glass), which has a thickness of 0.09&#34; and is coated by approximately 70 nm thick layer of Chromium (Cr), using masking in order to prepare a photo mask. Use AZ400K developer for 1 min. Then, etch the Cr layer using Cr etchant for about 1 min. Use acetone to strip the resist and clean it with cold piranha solution (H2SO4 and H2O2 in a ratio of 3:1).</li>
	<li>Photolithography
	<ol>
		<li>Clean a standard 4&#34; silicon wafer chemically with piranha solution for 20 min.</li>
		<li>Rinse the wafer with DI water and dry it.</li>
		<li>Heat the wafer on a hot plate (200 &#176;C for 15 min).</li>
		<li>Coat the silicon wafer with photoresist. Here, the positive photoresist AZ4620 was spin-coated on a silicon wafer at 2,000 rpm for 25 sec to obtain a 12.5 &#956;m thick layer.</li>
		<li>Remove all the solvent by soft baking of the wafer on a hot plate by floating the wafer for 90 sec&#160;on nitrogen flow at 100 &#176;C. Then, keep it in vacuum at the same temperature for 60 sec.</li>
		<li>Place the wafer in a dark box for 24 hr for dehydration.</li>
		<li>Expose the wafer to UV light in order to transfer the designed pattern to the photoresist.</li>
		<li>Immerse the wafer in photoresist developer solution (AZ400K) for 240 sec. Then, rinse the wafer with isopropyl alcohol and dry it by placing in a stream of nitrogen gas.</li>
	</ol>
	</li>
	<li>ICP-DRIE (Inductively Coupled Plasma - Deep Reactive Ion Etching) Process
	<ol>
		<li>Apply DRIE etching. Choose the appropriate etch depth according to final depth required for device (50 &#956;m in this investigation). Photoresist acts as a masking layer during this process.</li>
		<li>Remove the remaining photoresist with acetone and clean the wafer.</li>
	</ol>
	</li>
	<li>PDMS (polydimethylsiloxane) Casting
	<ol>
		<li>Use trichloromethylsilane (TCMS) for silanizing the silicon master mold. Pour 2 or 3 drops of trichloromethylsilane in a vial and place it in a desiccator beside the silicon master mold. Allow 2-3 hr for the silanizing process to complete.</li>
		<li>In a separate container, mix the Sylgard 184 silicone base with curing agent by weight ratio of 10:1 to prepare PDMS. Degas the PDMS by subjecting it to vacuum conditions (about 2 hr).</li>
		<li>Put the silicon master mold in a holder. Then, pour the PDMS on the silicon master mold to form the PDMS stamp. Ensure that bubbles do not form in the PDMS during this process.</li>
		<li>Cure the PDMS for 2 hr at 80 &#176;C.</li>
		<li>Peel off the PDMS stamp from the master mold. Then, cut the PDMS stamp into separate microchips. Finally, use a cutting core to drill holes for the inlet(s) and outlet(s).</li>
	</ol>
	</li>
	<li>Bonding of PDMS to Glass
	<ol>
		<li>Expose the cover slip and PDMS stamp to oxygen plasma for 30 sec. Bond PDMS stamp to the cover slip.</li>
		<li>To achieve proper sealing between the PDMS stamp and the cover slip, anneal the device by putting it in oven at 70 &#176;C for 10 min.</li>
	</ol>
	</li>
</ol>
2. Bacterial Culture
NOTE: Proper biosafety protocols must be followed for Steps 2-4. Consult the institutional safety officer for details.
<ol>
	<li>Prepare LB Agar Plates
	<ol>
		<li>Add 20 g Luria-Bertani (LB) agar (Miller) powders and 500 ml&#160;of ultrapure water to a 1 L flask. Stir to dissolve the powder.</li>
		<li>Sterilize by autoclaving at 15 psi, 121 &#176;C for 15 min.</li>
		<li>Allow the flask to cool down to 50-55 &#176;C on a bench or in a water bath in the biosafety hood.</li>
		<li>Add the antibiotic Tetracycline to achieve a final concentration of 50 &#956;g/ml. Mix well by swirling.</li>
		<li>Pour the mixture into plates. Fill each plate till 1/2 - 2/3 full.</li>
		<li>Flame air bubbles briefly to pop them if they form. Solidified air bubbles are difficult to spread bacterial culture over.</li>
		<li>Allow the plates to cool at room temperature overnight.</li>
		<li>When they are cool, put plates back into their sleeve, seal the bag, label (antibiotic and date), and store at 4 &#176;C. 
		NOTE: Cover the stock of plates with tin foil, as light deactivates many antibiotics.</li>
	</ol>
	</li>
	<li>LB Broth Preparation
	<ol>
		<li>Add 20 g Luria-Bertani (LB) broth (Miller) powders and 1 L&#160;of ultrapure water to a&#160;flask. Stir to dissolve the powder.</li>
		<li>Sterilize by autoclaving at 15 psi, 121 &#176;C for 15 min.</li>
		<li>Allow flask to cool down to 50-55 &#176;C on a bench or in a water bath in the biosafety hood.</li>
		<li>Add the antibiotic tetracycline to achieve a final concentration of 50 &#956;g/ml. Mix well by swirling.</li>
		<li>When cool, place the labeled bottle at 4 &#176;C. 
		NOTE: Cover the bottle with tin foil, as light deactivates many antibiotics.</li>
	</ol>
	</li>
	<li>Culture Bacteria on an LB Agar Plate (This protocol uses Pseudomonas fluorescens)
	<ol>
		<li>Take the bacterial stock from the freezer (-80 &#176;C) and place it on ice.</li>
		<li>Place the -80 &#176;C bacterial stock and an LB agar plate inside a biosafety hood.</li>
		<li>Streak the bacterial strain onto an LB agar plate in a zigzag pattern. Cover the agar plate and incubate it at 30 &#186;C overnight. Finally, store the plate in the refrigerator at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare the Bacterial Solution (S1)
	<ol>
		<li>Pour 50 ml LB broth media to an autoclaved flask. Perform this operation inside a biosafety hood.</li>
		<li>Transfer a single bacterial colony from the LB agar plate to the flask. This operation should also be performed inside a biosafety hood.</li>
		<li>Put the flask in a shaker incubator at 30 &#176;C and 150 rpm for adequate time (4 hr).</li>
	</ol>
	</li>
	<li>Prepare the Dilute Bacterial Solution (S2)
	<ol>
		<li>Pour 5 ml LB broth media into a sterilized plastic tube.</li>
		<li>Dilute S1 by mixing with LB broth media. Then, vortex the solution. Dilute the solution achieve the desired optical density (OD measured at 600 nm = 0.1). 
		NOTE: Biofilm experiments typically employ OD values in this vicinity.</li>
	</ol>
	</li>
</ol>
3. Prepare the Experimental Setup
<ol>
	<li>Using tweezers connect flexible plastic tubes (0.20&#34; ID) into the inlet(s) and outlet(s) of the microchip. The inlets and outlets were previously drilled into the PDMS portion of the microchip (step 1.5.5). In this investigation, the microchip consists of two inlets and one outlet.</li>
	<li>Fill syringe(s) with bacterial solution (S2 solution) and remove all the bubbles in the syringe(s).</li>
	<li>Connect syringe tip(s) (30 G 0.5&#34; blunt needle) into inlet tube(s). Then, connect outlet tube(s) to waste container.</li>
</ol>
4. Run the Experiment
<ol>
	<li>Connect the syringe(s) to the syringe tip(s).</li>
	<li>Place and fix syringe(s) onto the syringe pump. Then place the microchip under an optical microscope with objective lenses of desired magnification (e.g., 40X). Cover the microchip with a live cell chamber device to maintain a constant temperature environment for bacterial growth (30 &#176;C for P. fluorescens).</li>
	<li>Set the pump to the desired flow rate level (say 10 &#956;l/hr) and initiate fluid pumping.</li>
	<li>Once bacteria are introduced into the chamber, biofilm formation is also initiated. Biofilm formation and maturation typically occur over a period of several hours or even days. Observe and take images of biofilm growth through the microscope.</li>
</ol>

<ol>
	<li>Valiei, A., Kumar, A., Mukherjee, P. P., Liu, Y., Thundat, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+web+of+streamers:+biofilm+formation+in+a+porous+microfluidic+device.">A web of streamers: biofilm formation in a porous microfluidic device.</a> Lab Chip. 12, 5133-5137 (2012).</li><li>Costerton, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bacterial+Biofilms:+A+Common+Cause+of+Persistent+Infections.">Bacterial Biofilms: A Common Cause of Persistent Infections.</a> Science. 284, 1318-1322 (1999).</li><li>Flemming, H. C., Wingender, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+biofilm+matrix.">The biofilm matrix.</a> Nat Rev Microbiol. 8, 623-633 (2010).</li><li>Wong, G. C. L., O&#8217;Toole, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=All+together+now:+Integrating+biofilm+research+across+disciplines.">All together now: Integrating biofilm research across disciplines.</a> MRS Bulletin. 36, 339-342 (2011).</li><li>Yazdi, S., Ardekani, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bacterial+aggregation+and+biofilm+formation+in+a+vortical+flow.">Bacterial aggregation and biofilm formation in a vortical flow.</a> Biomicrofluidics. 6, 044114 (2012).</li><li>Rusconi, R., Lecuyer, S., Guglielmini, L., Stone, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laminar+flow+around+corners+triggers+the+formation+of+biofilm+streamers.">Laminar flow around corners triggers the formation of biofilm streamers.</a> J R Soc Interface. 7, 1293-1299 (2010).</li><li>Drescher, K., Shen, Y., Bassler, B. L., Stone, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biofilm+streamers+cause+catastrophic+disruption+of+flow+with+consequences+for+environmental+and+medical+systems.">Biofilm streamers cause catastrophic disruption of flow with consequences for environmental and medical systems.</a> P Natl Acad Sci USA. 110, 4345-4350 (2013).</li><li>Marty, A., Roques, C., Causserand, C., Bacchin, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Formation+of+bacterial+streamers+during+filtration+in+microfluidic+systems.">Formation of bacterial streamers during filtration in microfluidic systems.</a> Biofouling. 28, 551-562 (2012).</li><li>Taherzadeh, D., 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=Computational+Study+of+the+Drag+and+Oscillatory+Movement+of+Biofilm+Streamers+in+Fast+Flows.">Computational Study of the Drag and Oscillatory Movement of Biofilm Streamers in Fast Flows.</a> Biotechnol Bioeng. 105, 600-610 (2010).</li><li>Vrouwenvelder, J. S., 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=Impact+of+flow+regime+on+pressure+drop+increase+and+biomass+accumulation+and+morphology+in+membrane+systems.">Impact of flow regime on pressure drop increase and biomass accumulation and morphology in membrane systems.</a> Water Res. 44, 689-702 (2010).</li><li>Mitchell, A. C., 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=Biofilm+enhanced+geologic+sequestration+of+supercritical+CO2.">Biofilm enhanced geologic sequestration of supercritical CO2.</a> International Journal of Greenhouse Gas Control. 3, 90-99 (2009).</li><li>Soleimani, S., Van Geel, P. J., Isgor, O. B., Mostafa, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+of+biological+clogging+in+unsaturated+porous+media.">Modeling of biological clogging in unsaturated porous media.</a> J Contam Hydrol. 106, 39-50 (2009).</li><li>Kumar, A., 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=Microscale+confinement+features+can+affect+biofilm+formation.">Microscale confinement features can affect biofilm formation.</a> Microfluid Nanofluid. 14, 895-902 (2013).</li><li>Neethirajan, S., et al., Bhushan, B., 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=.">.</a> Encylopedia of Nanotechnology. , (2012).</li><li>Barathi, S., Vasudevan, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Utilization+of+petroleum+hydrocarbons+by+Pseudomonas+fluorescens+isolated+from+a+petroleum-contaminated+soil.">Utilization of petroleum hydrocarbons by Pseudomonas fluorescens isolated from a petroleum-contaminated soil.</a> Environ Int. 26, 413-416 (2001).</li><li>Das, S., Kumar, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Formation+and+post-formation+dynamics+of+bacterial+biofilm+streamers+as+highly+viscous+liquid+jets.">Formation and post-formation dynamics of bacterial biofilm streamers as highly viscous liquid jets.</a> arXiv preprint arXiv:1312.6056. , (2013).</li><li>Shaw, T., Winston, M., Rupp, C. J., Klapper, I., Stoodley, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Commonality+of+elastic+relaxation+times+in+biofilms.">Commonality of elastic relaxation times in biofilms.</a> Phys Rev Lett. 93, (2004).</li><li>Berejnov, V., Djilali, N., Sinton, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lab-on-chip+methodologies+for+the+study+of+transport+in+porous+media:+energy+applications.">Lab-on-chip methodologies for the study of transport in porous media: energy applications.</a> Lab Chip. 8, 689-693 (2008).</li></ol>

The authors have nothing to disclose.

We demonstrated a simple microfluidic device that mimics porous media for studying biofilm development in complex habitats. There are several critical steps that dictate the outcome of the experiments. They include device geometry. While the post geometry can vary, adequate pore-space for streamers to form is necessary. Moreover, Valiei et al.1 have demonstrated that streamer formation occurs only in a certain flow rate range. At flow rates lower than a threshold value, deformation of biofilms into streamers may not be observed. Yet above a certain another threshold flow rate value, biofilm fracture can dominate and not allow formation of streamers. Another issue that can plague these experiments is gas bubbles that can become trapped in the micro-pillar array. Usually these bubbles have to be removed by increasing the flow rate initially and then gradually decreasing it to the desired value.
Microfluidic platforms such as these offer several advantages and few limitations. The platform enables us to work with small culture volumes, and has the flexibility of incorporating user-defined features. For example, different porous structures can be simulated by altering the geometrical parameters of the micro-pillar array. Even structures that mimic the random structure of real porous media can be fabricated on microfluidic platforms18. Moreover, several such channels can be implemented on a single device allowing for collection of significant pertinent data for accurate statistical analysis. However, microfluidic systems typically mimic two-dimensional structure of porous media. Devices that can mimic the three-dimensional nature of porous media are usually quite challenging to fabricate.
Formation and evolution of streamers are not well understood yet, and further research is required in this direction. Understanding of how streamers form and lead to the formation of mature biofilm structures will be relevant to a wide variety of scenarios including clogging of biomedical devices such as heart stents, biofilms in soil, and filtration systems. Our microfluidic platform is a step in that direction.

Recently, we demonstrated bacterial biofilm formation dynamics in a microfluidic device that mimics porous media1. Bacterial biofilms are essentially colonies of surface aggregated bacteria that are encased by extracellular polymeric substances (EPS)2-4. These thin films of bacteria can form in almost every conceivable niche ranging from smooth surfaces to the much more complex habitat of porous media. Valiei et al.1 used a microfluidic device with an array of micro-pillars to simulate a porous media structure and studied biofilm formation in this device as a function of fluid flow rate. They found that in a certain flow regime, filamentous biofilms known as streamers began to emerge between different pillars. Streamers can be tethered at one or both ends to solid surfaces, but the rest of the structure is suspended in liquid. Streamer formation typically starts after an initial layer of biofilm has formed and its formation can dictate the long-term evolution of biofilm in such complex habitats. Recently, several researchers have investigated the dynamics of streamer formation. Yazdi et al.5 showed that the streamers can form in vortical flows originating from an oscillating bubble. In another experiment, Rusconi et al.6 investigated the effect of channel curvature and channel geometry on the formation of streamers. They found that the streamers can form in curved sections of microchannels, and streamer morphology is related to motility. Recent research has demonstrated that streamers can have wide repercussions in various natural and artificial scenarios as they can act as precursors to the formation of mature structures in porous interfaces, lead to rapid and catastrophic biofilm proliferation in a biomedical systems, and also cause substantial flow-structure interactions, etc1,7-9.
Biofilm streamers often form in complex habitats such as porous media. Understanding biofilm growth in porous media environment is relevant to several environmental and industrial processes such as biological wastewater treatment10, maintaining well-bore integrity in situations such as CO2 capture11 and plugging of pores in soil12. Observing biofilm formation in such complex habitats can often be challenging due to the opacity of porous media. In such situations, microfluidics based porous media platforms can prove extremely advantageous as they allow real-time and in&#160;situ monitoring. Another advantage of microfluidics is the ability to build multiple bioreactors on a single bio-microfluidic platform and simultaneously allow for online monitoring and/or incorporation of sensors. The flexibility to implement multiple laboratory experiments in one device and the ability to collect significant pertinent data for accurate statistical analysis is an important advantage of microfluidic systems13,14.
In the context of the above discussion, understanding streamer formation dynamics in a porous media environment would be beneficial to several applications. In this study, we develop the protocol for investigating streamer formation in a device that mimics porous media. Fabrication of the microfluidic platform, necessary steps for cell culture and experimentation are described. In our experiments, the wild type bacterial strain of Pseudomonas fluorescens was employed. P. fluorescens, found naturally in soil, plays a key role in maintaining soil ecology15. The bacterial strain employed had been genetically engineered to express green fluorescent protein (GFP) constitutively.

<table><tbody><tr><td>Flourescent Microscope</td><td>Nikon</td><td></td><td></td></tr><tr><td>LB agar</td><td>Fisher</td><td>BP1425-500</td><td>suspend 40 g in 1 L of purified water</td></tr><tr><td>LB broth</td><td>Fisher</td><td>BP1427-500</td><td>suspend 20 g in 1 L of purified water</td></tr><tr><td>Biosafety hood</td><td>Microzone corporation</td><td></td><td></td></tr><tr><td>Petri dish</td><td>Fisher</td><td>875712</td><td>sterile 100 mm x 15 mm polystyrene Petri dish</td></tr><tr><td>Incubator shaker</td><td>New Brunswick Scientific</td><td></td><td>Excella E24 incubator shaker series</td></tr><tr><td>50 ml sterilized centrifuge tube</td><td>Corning</td><td>430828</td><td>Polypropylene RNase-/DNase-free</td></tr><tr><td>Tetracycline free base</td><td>MP Biomedicals</td><td>103012</td><td>50 &amp;#956;g/ml</td></tr><tr><td>SYLGARD 184 silicone</td><td>Dow Corning Corporation</td><td>68037-59-2</td><td>Elastomer Base and curing agent</td></tr><tr><td>Positive photoresist (AZ4620)</td><td></td><td></td><td></td></tr><tr><td>Plastic tube</td><td>Cole-Parmer</td><td></td><td></td></tr></tbody></table>

protocol for biofilm streamer formation in a microfluidic device with micro-pillars

Several bacterial species possess the ability to attach to surfaces and colonize them in the form of thin films called biofilms. Biofilms that grow in porous media are relevant to several industrial and environmental processes such as wastewater treatment and CO2 sequestration. We used Pseudomonas fluorescens, a Gram-negative aerobic bacterium, to investigate biofilm formation in a microfluidic device that mimics porous media. The microfluidic device consists of an array of micro-posts, which were fabricated using soft-lithography. Subsequently, biofilm formation in these devices with flow was investigated and we demonstrate the formation of filamentous biofilms known as streamers in our device. The detailed protocols for fabrication and assembly of microfluidic device are provided here along with the bacterial culture protocols. Detailed procedures for experimentation with the microfluidic device are also presented along with representative results.

Using the above mentioned microfabrication protocol, a PDMS based microfluidic device was constructed. Figure 1 shows the scanning electron microscope (SEM) images of the PDMS device. Figure 1a shows the entrance section of the device. A fork-like entrance is created to equalize pressure head across the device. Further SEM imaging also showed that the pillar walls are almost vertical (Figure&#160;1b). The cultured bacterial solution (Figure&#160;2) was diluted and its optical density was adjusted to a value of 0.1. We examined biofilm formation in the microfluidic device as a function of input flow rate. When P. fluorescens was injected into the device at a low flow-rate of 0.8 &#956;l/hr, bacterial attachment and biofilm formation occurred at the walls of the device. Even after a prolonged period of time (&#62;20 hr), no other bacterial structures other than surface-hugging biofilms were observed. Next, the same experiment was repeated at a flow rate of 8 &#956;l/hr. In this case, biofilm formation again started after a few minutes of infusion of the diluted bacterial culture. However, after a few hours, appearance of filamentous structures extending between micro-pillars was observed near the mid-section of the device (Figure&#160;4). These filamentous structures could be visualized through the presence of immobile bacteria. These structures are known as streamers and they are filamentous biofilms that are only tethered at one or both ends to surfaces. The rest of the structure is often suspended in the liquid medium (as in this case). Figure 4 shows the time-evolution of biofilm streamer structure. Streamers usually form due to the effect of fluid shear on the visco-elastic biofilm. Figure 5 shows the streamlines and velocity contours for flow past a series of pillars. The simulation shows that the streamers that form in our microfluidic system are essentially aligned along the fluid flow streamlines. The correlation between the flow structures and formation of biofilm streamers is not yet well understood. However, Das and Kumar16 have recently proposed that these streamers form as highly viscous liquid state of the intrinsically viscoelastic biofilms. They based their conjecture on the observation that the time-scale of biofilm streamer formation typically far exceeds the viscoelastic relaxation time scales of biofilms. Biofilms are known to behave as viscoelastic liquids and hence at time-scales much larger than the viscoelastic relaxation time scale, they essentially behave as highly viscous liquids17. According to this formulation, streamers can be expected to originate at locations of high shear stresses. Figure 5 shows the locations of high velocity in the channel, and these locations coincide with locations of high shear stresses. In the initial phase of growth, streamers are observed to originate near these locations (Figure&#160;4).
<img alt="Figure 1" src="/files/ftp_upload/51732/51732fig1highres.jpg" /> 
Figure 1.&#160;Scanning electron microscope (SEM) images of the microfluidic channel (top-view). a)&#160;Inlet section, b)&#160;Region containing micro-pillars. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/51732/51732fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/51732/51732fig2highres.jpg" /> 
Figure 2.&#160;Sequential steps involved in bacterial culture. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/51732/51732fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/51732/51732fig3highres.jpg" /> 
Figure 3.&#160;Set up for microfluidic experiments. 1&#8212;Optical microscope (inverted), 2&#8212;Syringe pump, 3&#8212;Image and data acquisition, 4&#8212;Syringe containing dye (optional), 5&#8212;Syringe containing bacteria, 6&#8212;Waste reservoir. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/51732/51732fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/51732/51732fig4highres.jpg" /> 
Figure 4.&#160;Time-lapse confocal imaging of evolution of streamers. Image plane corresponds to z = 25 &#956;m i.e. middle of device. Dashed ellipses demonstrate biofilm streamers. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/51732/51732fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/51732/51732fig5highres.jpg" /> 
Figure 5.&#160;Computational fluid mechanical simulations showing streamlines and velocity contours of flow past micro-pillars. Fluid flow is from top to bottom and velocity scale is in m/sec. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/51732/51732fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Protocol for Biofilm Streamer Formation in a Microfluidic Device with Micro-pillars

Protocols for the study of biofilm formation in a microfluidic device that mimics porous media are discussed. The microfluidic device consists of an array of micro-pillars and biofilm formation by Pseudomonas fluorescens in this device is investigated.

Several bacterial species possess the ability to attach to surfaces and colonize them in the form of thin films called biofilms. Biofilms that grow in ...

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12.1K Views.  University of Alberta. Protocols for the study of biofilm formation in a microfluidic device that mimics porous media are discussed. The microfluidic device consists of an array of micro-pillars and biofilm formation by Pseudomonas fluorescens in this device is investigated. 

Video: Protocol for Biofilm Streamer Formation in a Microfluidic Device with Micro-pillars

Fabrication and Visualization of Capillary Bridges in Slit Pore Geometry

A procedure for creating and imaging capillary bridges in slit-pore geometry is presented. High aspect ratio hydrophobic pillars are fabricated and functionalized to render their top surfaces hydrophilic. The combination of a physical feature (the pillar) with a chemical boundary (the hydrophilic film on the top of the pillar) provides both a physical and chemical heterogeneity that pins the triple contact line, a necessary feature to create stable long but narrow capillary bridges. The substrates with the pillars are attached to glass slides and secured into custom holders. The holders are then mounted onto four axis microstages and positioned such that the pillars are parallel and facing each other. The capillary bridges are formed by introducing a fluid in the gap between the two substrates once the separation between the facing pillars has been reduced to a few hundred micrometers. The custom microstage is then employed to vary the height of the capillary bridge. A CCD camera is positioned to image either the length or the width of the capillary bridge to characterize the morphology of the fluid interface. Pillars with widths down to 250 &#181;m and lengths up to 70 mm were fabricated with this method, leading to capillary bridges with aspect ratios (length/width) of over 1001.

A procedure for creating and imaging capillary bridges in slit-pore geometry is presented. The creation of capillary bridges relies on the formation of pillars to provide a directional physical and chemical heterogeneity to pin the fluid. Capillary bridges are formed and manipulated using microstages and visualized using a CCD camera.

A procedure for creating and imaging capillary bridges in slit-pore geometry is presented. High aspect ratio hydrophobic pillars are fabricated and ...

Engineering

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 ...

A Microfluidic Technique to Probe Cell Deformability

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an efficient manner. Typically, data for ~102 cells can be acquired within a 1 hr experiment. An automated image analysis program enables efficient post-experiment analysis of image data, enabling processing to be complete within a few hours. Our device geometry is unique in that cells must deform through a series of micron-scale constrictions, thereby enabling the initial deformation and time-dependent relaxation of individual cells to be assayed. The applicability of this method to human promyelocytic leukemia (HL-60) cells is demonstrated. Driving cells to deform through micron-scale constrictions using pressure-driven flow, we observe that human promyelocytic (HL-60) cells momentarily occlude the first constriction for a median time of 9.3 msec before passaging more quickly through the subsequent constrictions with a median transit time of 4.0 msec per constriction. By contrast, all-trans retinoic acid-treated (neutrophil-type) HL-60 cells occlude the first constriction for only 4.3 msec before passaging through the subsequent constrictions with a median transit time of 3.3 msec. This method can provide insight into the viscoelastic nature of cells, and ultimately reveal the molecular origins of this behavior.

We demonstrate a microfluidics-based assay to measure the timescale for cells to transit through a sequence of micron-scale constrictions.

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an ...

Combining Fluidic Devices with Microscopy and Flow Cytometry to Study Microbial Transport in Porous Media Across Spatial Scales

Understanding the transport, dispersion and deposition of microorganisms in porous media is a complex scientific task comprising topics as diverse as hydrodynamics, ecology and environmental engineering. Modeling bacterial transport in porous environments at different spatial scales is critical to better predict the consequences of bacterial transport, yet current models often fail to up-scale from laboratory to field conditions. Here, we introduce experimental tools to study bacterial transport in porous media at two spatial scales. The aim of these tools is to obtain macroscopic observables (such as breakthrough curves or deposition profiles) of bacteria injected into transparent porous matrices. At the small scale (10-1000 &#181;m), microfluidic devices are combined with optical video-microscopy and image processing to obtain breakthrough curves and, at the same time, to track individual bacterial cells at the pore scale. At larger scale, flow cytometry is combined with a self-made robotic dispenser to obtain breakthrough curves. We illustrate the utility of these tools to better understand how bacteria are transported in complex porous media such as the hyporheic zone of streams. As these tools provide simultaneous measurements across scales, they pave the way for mechanism-based models, critically important for upscaling. Application of these tools may not only contribute to the development of novel bioremediation applications but also shed new light on the ecological strategies of microorganisms colonizing porous substrates.

Breakthrough curves (BTCs) are efficient tools to study the transport of bacteria in porous media. Here we introduce tools based on fluidic devices in combination with microscopy and flow cytometric counting to obtain BTCs.

Understanding the transport, dispersion and deposition of microorganisms in porous media is a complex scientific task comprising topics as diverse as ...

Environment

Use of a High-throughput In Vitro Microfluidic System to Develop Oral Multi-species Biofilms

There are few high-throughput in vitro systems which facilitate the development of multi-species biofilms that contain numerous species commonly detected within in vivo oral biofilms. Furthermore, a system that uses natural human saliva as the nutrient source, instead of artificial media, is particularly desirable in order to support the expression of cellular and biofilm-specific properties that mimic the in vivo communities. We describe a method for the development of multi-species oral biofilms that are comparable, with respect to species composition, to supragingival dental plaque, under conditions similar to the human oral cavity. Specifically, this methods article will describe how a commercially available microfluidic system can be adapted to facilitate the development of multi-species oral biofilms derived from and grown within pooled saliva. Furthermore, a description of how the system can be used in conjunction with a confocal laser scanning microscope to generate 3-D biofilm reconstructions for architectural and viability analyses will be presented. Given the broad diversity of microorganisms that grow within biofilms in the microfluidic system (including Streptococcus, Neisseria, Veillonella, Gemella, and Porphyromonas), a protocol will also be presented describing how to harvest the biofilm cells for further subculture&#160;or DNA extraction and analysis. The limits of both the microfluidic biofilm system and the current state-of-the-art data analyses will be addressed. Ultimately, it is envisioned that this article will provide a baseline technique that will improve the study of oral biofilms and aid in the development of additional technologies that can be integrated with the microfluidic platform.

Use of a High-throughput In Vitro Microfluidic System to Develop Oral Multi-species Biofilms

The goal of this methods paper is to describe the use of a microfluidic system for the development of multi-species biofilms that contain species typically identified in human supragingival dental plaque.&#160;Methods to describe biofilm architecture, biofilm viability, and an approach to harvest biofilm for culture-dependent or culture-independent analyses are highlighted.

There are few high-throughput in vitro systems which facilitate the development of multi-species biofilms that contain numerous species commonly detected ...

Methods for Characterizing the Co-development of Biofilm and Habitat Heterogeneity

Biofilms are surface-attached microbial communities that have complex structures and produce significant spatial heterogeneities. Biofilm development is strongly regulated by the surrounding flow and nutritional environment. Biofilm growth also increases the heterogeneity of the local microenvironment by generating complex flow fields and solute transport patterns. To investigate the development of heterogeneity in biofilms and interactions between biofilms and their local micro-habitat, we grew mono-species biofilms of Pseudomonas aeruginosa and dual-species biofilms of P. aeruginosa and Escherichia coli under nutritional gradients in a microfluidic flow cell. We provide detailed protocols for creating nutrient gradients within the flow cell and for growing and visualizing biofilm development under these conditions. We also present protocols for a series of optical methods to quantify spatial patterns in biofilm structure, flow distributions over biofilms, and mass transport around and within biofilm colonies. These methods support comprehensive investigations of the co-development of biofilm and habitat heterogeneity.

Biofilms have complex interactions with their surrounding environment. To comprehensively investigate biofilm-environment interactions, we present here a series of methods to create heterogeneous chemical environment for biofilm development, to quantify local flow velocity, and to analyze mass transport in and around biofilm colonies.

Biofilms are surface-attached microbial communities that have complex structures and produce significant spatial heterogeneities. Biofilm development is ...

A Microfluidic Platform to Study Bioclogging in Porous Media

Bacterial biofilms are found in several environmental and industrial porous media, including soils and filtration membranes. Biofilms grow under certain flow conditions and can clog pores, thereby redirecting the local fluid flow. The ability of biofilms to clog pores, the so-called bioclogging, can have a tremendous effect on the local permeability of the porous medium, creating a pressure buildup in the system, and impacting the mass flow through it. To understand the interplay between biofilm growth and fluid flow under different physical conditions (e.g., at different flow velocities and pore sizes), in the present study, a microfluidic platform is developed to visualize biofilm development using a microscope under externally-imposed, controlled physical conditions. The biofilm-induced pressure buildup in the porous medium can be measured simultaneously using pressure sensors and, later, correlated with the surface coverage of the biofilm. The presented platform provides a baseline for a systematic approach to investigate bioclogging caused by biofilms in porous media under flow conditions and can be adapted to studying environmental isolates or multispecies biofilms.

The present protocol describes a microfluidic platform to study biofilm development in quasi-2D porous media by combining high-resolution microscopy imaging with simultaneous pressure difference measurements. The platform quantifies the influence of pore size and fluid flow rates in porous media on bioclogging.

Bacterial biofilms are found in several environmental and industrial porous media, including soils and filtration membranes. Biofilms grow under certain ...

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 ...

Biology

Microfluidic Co-culture of Epithelial Cells and Bacteria for Investigating Soluble Signal-mediated Interactions

The human gastrointestinal (GI) tract is a unique environment in which intestinal epithelial cells and non-pathogenic (commensal) bacteria coexist. It has been proposed that the microenvironment that the pathogen encounters in the commensal layer is important in determining the extent of colonization. Current culture methods for investigating pathogen colonization are not well suited for investigating this hypothesis as they do not enable co-culture of bacteria and epithelial cells in a manner that mimics the GI tract microenvironment. Here we describe a microfluidic co-culture model that enables independent culture of eukaryotic cells and bacteria, and testing the effect of the commensal microenvironment on pathogen colonization. The co-culture model is demonstrated by developing a commensal Escherichia coli biofilm among HeLa cells, followed by introduction of enterohemorrhagic E. coli (EHEC) into the commensal island, in a sequence that mimics the sequence of events in GI tract infection.

This protocol describes a microfluidic co-culture model for simultaneous and localized culture of epithelial cells and bacteria. This model can be used for investigating the role of different soluble molecular signals on pathogenesis as well as screen the effectiveness of putative probiotic bacterial strains.

The human gastrointestinal (GI) tract is a unique environment in which intestinal epithelial cells and non-pathogenic (commensal) bacteria coexist. It has ...

Patterning of Microorganisms and Microparticles through Sequential Capillarity-assisted Assembly

Controlled patterning of microorganisms into defined spatial arrangements offers unique possibilities for a broad range of biological applications, including studies of microbial physiology and interactions. At the simplest level, accurate spatial patterning of microorganisms would enable reliable, long-term imaging of large numbers of individual cells and transform the ability to quantitatively study distance-dependent microbe-microbe interactions. More uniquely, coupling accurate spatial patterning and full control over environmental conditions, as offered by microfluidic technology, would provide a powerful and versatile platform for single-cell studies in microbial ecology. 
This paper presents a microfluidic platform to produce versatile and user-defined patterns of microorganisms within a microfluidic channel, allowing complete optical access for long-term, high-throughput monitoring. This new microfluidic technology is based on capillarity-assisted particle assembly and exploits the capillary forces arising from the controlled motion of an evaporating suspension inside a microfluidic channel to deposit individual microsized objects in an array of traps microfabricated onto a polydimethylsiloxane (PDMS) substrate. Sequential depositions generate the desired spatial layout of single or multiple types of micro-sized objects, dictated solely by the geometry of the traps and the filling sequence. 
The platform has been calibrated using colloidal particles of different dimensions and materials: it has proven to be a powerful tool to generate diverse colloidal patterns and perform surface functionalization of trapped particles. Furthermore, the platform was tested on microbial cells, using Escherichia coli cells as a model bacterium. Thousands of individual cells were patterned on the surface, and their growth was monitored over time. In this platform, the coupling of single-cell deposition and microfluidic technology allows both geometric patterning of microorganisms and precise control of environmental conditions. It thus opens a window into the physiology of single microbes and the ecology of microbe-microbe interactions, as shown by preliminary experiments.

We present a technology that uses capillarity-assisted assembly in a microfluidic platform to pattern micro-sized objects suspended in a liquid, such as bacteria and colloids, into prescribed arrays on a polydimethylsiloxane substrate.

Controlled patterning of microorganisms into defined spatial arrangements offers unique possibilities for a broad range of biological applications, ...

Protocol for Biofilm Streamer Formation in a Microfluidic Device with Micro-pillars

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Fabrication and Visualization of Capillary Bridges in Slit Pore Geometry

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

A Microfluidic Technique to Probe Cell Deformability

Combining Fluidic Devices with Microscopy and Flow Cytometry to Study Microbial Transport in Porous Media Across Spatial Scales

Use of a High-throughput In Vitro Microfluidic System to Develop Oral Multi-species Biofilms

Methods for Characterizing the Co-development of Biofilm and Habitat Heterogeneity

A Microfluidic Platform to Study Bioclogging in Porous Media

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

Microfluidic Co-culture of Epithelial Cells and Bacteria for Investigating Soluble Signal-mediated Interactions

Patterning of Microorganisms and Microparticles through Sequential Capillarity-assisted Assembly