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>

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	<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. 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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 st...

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 border="0" cellpadding="0" cellspacing="0" width="1038">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">Name of Material/ Equipment</td>
			<td style="width:255px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:339px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">Flourescent Microscope</td>
			<td style="width:255px;">Nikon</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">LB agar</td>
			<td style="width:255px;">Fisher</td>
			<td style="width:119px;">BP1425-500</td>
			<td>suspend 40 g in 1 L of purified water</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">LB broth</td>
			<td style="width:255px;">Fisher</td>
			<td style="width:119px;">BP1427-500</td>
			<td>suspend 20 g in 1 L of purified water</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">Biosafety hood</td>
			<td style="width:255px;">Microzone corporation</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">Petri-dish</td>
			<td style="width:255px;">Fisher</td>
			<td style="width:119px;">875712</td>
			<td>sterile 100mmx15mm polystyrene petri dish</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:325px;">Incubator shaker</td>
			<td>New Brunswick Scientific</td>
			<td style="width:119px;"></td>
			<td style="width:339px;">Excella E24incubator shaker series</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">50 mL sterilized centrifuge tube</td>
			<td style="width:255px;">Corning</td>
			<td style="width:119px;">430828</td>
			<td>Polypropylene Rnase-/Dnase-free</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">Tetracycline free base</td>
			<td style="width:255px;">MP Biomedicals</td>
			<td style="width:119px;">103012</td>
			<td>50 ug/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">SYLGARD 184 silicone</td>
			<td style="width:255px;">Dow Corning Corporation</td>
			<td style="width:119px;">68037-59-2</td>
			<td>Elastomer Base and curing agent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">Positive photoresist (AZ4620)</td>
			<td style="width:255px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">Plastic tube</td>
			<td style="width:255px;">Cole- Parmer</td>
			<td style="width:119px;"></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 d...

Watch this Scientific Journal Video about Protocol for Biofilm Streamer Formation in a Microfluidic Device with Micro-pillars at JoVE.com

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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Bioengineering

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

Microfluidic Device for Recreating a Tumor Microenvironment in Vitro

We have developed a microfluidic device that mimics the delivery and systemic clearance of drugs to heterogeneous three-dimensional tumor tissues in vitro. Nutrients delivered by vasculature fail to reach all parts of tumors, giving rise to heterogeneous microenvironments consisting of viable, quiescent and necrotic cell types. Many cancer drugs fail to effectively penetrate and treat all types of cells because of this heterogeneity. Monolayers of cancer cells do not mimic this heterogeneity, making it difficult to test cancer drugs with a suitable in vitro model. Our microfluidic devices were fabricated out of PDMS using soft lithography. Multicellular tumor spheroids, formed by the hanging drop method, were inserted and constrained into rectangular chambers on the device and maintained with continuous medium perfusion on one side. The rectangular shape of chambers on the device created linear gradients within tissue. Fluorescent stains were used to quantify the variability in apoptosis within tissue. Tumors on the device were treated with the fluorescent chemotherapeutic drug doxorubicin, time-lapse microscopy was used to monitor its diffusion into tissue, and the effective diffusion coefficient was estimated. The hanging drop method allowed quick formation of uniform spheroids from several cancer cell lines. The device enabled growth of spheroids for up to 3 days. Cells in proximity of flowing medium were minimally apoptotic and those far from the channel were more apoptotic, thereby accurately mimicking regions in tumors adjacent to blood vessels. The estimated value of the doxorubicin diffusion coefficient agreed with a previously reported value in human breast cancer. Because the penetration and retention of drugs in solid tumors affects their efficacy, we believe that this device is an important tool in understanding the behavior of drugs, and developing new cancer therapeutics.

Microfluidic Device for Recreating a Tumor Microenvironment in Vitro

We present the procedure for fabrication and operation of a microfluidic device that recreates heterogeneous tumor microenvironments in vitro. The variability in apoptosis within tumor tissue was quantified using fluorescent stains and the effective diffusion coefficient of the chemotherapeutic drug doxorubicin into tumor tissue was evaluated.

We have developed a microfluidic device that mimics the delivery and systemic clearance of drugs to heterogeneous three-dimensional tumor tissues in ...

A Microfluidic Device for Studying Multiple Distinct Strains

The study of cell responses to environmental changes poses many experimental challenges: cells need to be imaged under changing conditions, often in a comparative manner. Multiwell plates are routinely used to compare many different strains or cell lines, but allow limited control over the environment dynamics. Microfluidic devices, on the other hand, allow exquisite dynamic control over the surrounding conditions, but it is challenging to image and distinguish more than a few strains in them. Here we describe a method to easily and rapidly manufacture a microfluidic device capable of applying dynamically changing conditions to multiple distinct yeast strains in one channel. The device is designed and manufactured by simple means without the need for soft lithography. It is composed of a Y-shaped flow channel attached to a second layer harboring microwells. The strains are placed in separate microwells, and imaged under the exact same dynamic conditions. We demonstrate the use of the device for measuring protein localization responses to pulses of nutrient changes in different yeast strains.

We present a simple method to produce microfluidic devices capable of applying similar dynamic conditions to multiple distinct strains, without the need for a clean room or soft lithography.

The study of cell responses to environmental changes poses many experimental challenges: cells need to be imaged under changing conditions, often in a ...

Microfluidic Fabrication of Polymeric and Biohybrid Fibers with Predesigned Size and Shape

A &#8220;sheath&#8221; fluid passing through a microfluidic channel at low Reynolds number can be directed around another &#8220;core&#8221; stream and used to dictate the shape as well as the diameter of a core stream.&#160;Grooves in the top and bottom of a microfluidic channel were designed to direct the sheath fluid and shape the core fluid.&#160;By matching the viscosity and hydrophilicity of the sheath and core fluids, the interfacial effects are minimized and complex fluid shapes can be formed.&#160;Controlling the relative flow rates of the sheath and core fluids determines the cross-sectional area of the core fluid.&#160;Fibers have been produced with sizes ranging from 300 nm to ~1 mm, and fiber cross-sections can be round, flat, square, or complex as in the case with double anchor fibers. Polymerization of the core fluid downstream from the shaping region solidifies the fibers.&#160;Photoinitiated click chemistries are well suited for rapid polymerization of the core fluid by irradiation with ultraviolet light.&#160;Fibers with a wide variety of shapes have been produced from a list of polymers including liquid crystals, poly(methylmethacrylate), thiol-ene and thiol-yne resins, polyethylene glycol, and hydrogel derivatives. Minimal shear during the shaping process and mild polymerization conditions also makes the fabrication process well suited for encapsulation of cells and other biological components.

Two adjacent fluids passing through a grooved microfluidic channel can be directed to form a sheath around a prepolymer core; thereby&#160;determining both shape and cross-section. Photoinitiated polymerization, such as thiol click chemistry, is well suited for rapidly solidifying the core fluid into a microfiber with predetermined size and shape.

A &#8220;sheath&#8221; fluid passing through a microfluidic channel at low Reynolds number can be directed around another &#8220;core&#8221; stream and ...

Analyzing Mixing Inhomogeneity in a Microfluidic Device by Microscale Schlieren Technique

In this paper, we introduce the use of microscale schlieren technique to measure mixing inhomogeneity in a microfluidic device. The microscale schlieren system is constructed from a Hoffman modulation contrast microscope, which provides easy access to the rear focal plane of the objective lens, by removing the slit plate and replacing the modulator with a knife-edge. The working principle of microscale schlieren technique relies on detecting light deflection caused by variation of refractive index1-3. The deflected light either escapes or is obstructed by the knife-edge to produce a bright or a dark band, respectively. If the refractive index of the mixture varies linearly with its composition, the local change in light intensity in the image plane is proportional to the concentration gradient normal to the optical axis. The micro-schlieren image gives a two-dimensional projection of the disturbed light produced by three-dimensional inhomogeneity.
To accomplish quantitative analysis, we describe a calibration procedure that mixes two fluids in a T-microchannel. We carry out a numerical simulation to obtain the concentration gradient in the T-microchannel that correlates closely with the corresponding micro-schlieren image. By comparison, a relationship between the grayscale readouts of the micro-schlieren image and the concentration gradients presented in a microfluidic device is established. Using this relationship, we are able to analyze the mixing inhomogeneity from associate micro-schlieren image and demonstrate the capability of microscale schlieren technique with measurements in a microfluidic oscillator4. For optically transparent fluids, microscale schlieren technique is an attractive diagnostic tool to provide instantaneous full-field information that retains the three-dimensional features of the mixing process.

Herein, we describe a procedure that employs microscale schlieren technique to measure mixing inhomogeneity in a microfluidic device. Through calibration, distribution of concentration gradient can be derived from the micro-schlieren image.

In this paper, we introduce the use of microscale schlieren technique to measure mixing inhomogeneity in a microfluidic device. The microscale schlieren ...

Fully Automated Centrifugal Microfluidic Device for Ultrasensitive Protein Detection from Whole Blood

Enzyme-linked immunosorbent assay (ELISA) is a promising method to detect small amount of proteins in biological samples. The devices providing a platform for reduced sample volume and assay time as well as full automation are required for potential use in point-of-care-diagnostics. Recently, we have demonstrated ultrasensitive detection of serum proteins, C-reactive protein (CRP) and cardiac troponin I (cTnI), utilizing a lab-on-a-disc composed of TiO2 nanofibrous (NF) mats. It showed a large dynamic range with femto molar (fM) detection sensitivity, from a small volume of whole blood in 30 min. The device consists of several components for blood separation, metering, mixing, and washing that are automated for improved sensitivity from low sample volumes. Here, in the video demonstration, we show the experimental protocols and know-how for the fabrication of NFs as well as the disc, their integration and the operation in the following order: processes for preparing TiO2 NF mat; transfer-printing of TiO2 NF mat onto the disc; surface modification for immune-reactions, disc assembly and operation; on-disc detection and representative results for immunoassay. Use of this device enables multiplexed analysis with minimal consumption of samples and reagents. Given the advantages, the device should find use in a wide variety of applications, and prove beneficial in facilitating the analysis of low abundant proteins.

This protocol demonstrates how to achieve femto molar detection sensitivity of proteins in 10 &#181;L of whole blood within 30 min. This can be achieved by using electrospun nanofibrous mats integrated in a lab-on-a-disc, which offers high surface area as well as effective mixing and washing for enhanced signal-to-noise ratio.

Enzyme-linked immunosorbent assay (ELISA) is a promising method to detect small amount of proteins in biological samples. The devices providing a platform ...

Co-culture of Living Microbiome with Microengineered Human Intestinal Villi in a Gut-on-a-Chip Microfluidic Device

Here, we describe a protocol to perform long-term co-culture of multi-species human gut microbiome with microengineered intestinal villi in a human gut-on-a-chip microphysiological device. We recapitulate the intestinal lumen-capillary tissue interface in a microfluidic device, where physiological mechanical deformations and fluid shear flow are constantly applied to mimic peristalsis. In the lumen microchannel, human intestinal epithelial Caco-2 cells are cultured to form a &#39;germ-free&#39; villus epithelium and regenerate small intestinal villi. Pre-cultured microbial cells are inoculated into the lumen side to establish a host-microbe ecosystem. After microbial cells adhere to the apical surface of the villi, fluid flow and mechanical deformations are resumed to produce a steady-state microenvironment in which fresh culture medium is constantly supplied and unbound bacteria (as well as bacterial wastes) are continuously removed. After extended co-culture from days to weeks, multiple microcolonies are found to be randomly located between the villi, and both microbial and epithelial cells remain viable and functional for at least one week in culture. Our co-culture protocol can be adapted to provide a versatile platform for other host-microbiome ecosystems that can be found in various human organs, which may facilitate in vitro study of the role of human microbiome in orchestrating health and disease.

We describe an in vitro protocol to co-culture gut microbiome and intestinal villi for an extended period using a human gut-on-a-chip microphysiological system.

Here, we describe a protocol to perform long-term co-culture of multi-species human gut microbiome with microengineered intestinal villi in a human ...

Microfluidic Platform with Multiplexed Electronic Detection for Spatial Tracking of Particles

Microfluidic processing of biological samples typically involves differential manipulations of suspended particles under various force fields in order to spatially fractionate the sample based on a biological property of interest. For the resultant spatial distribution to be used as the assay readout, microfluidic devices are often subjected to microscopic analysis requiring complex instrumentation with higher cost and reduced portability. To address this limitation, we have developed an integrated electronic sensing technology for multiplexed detection of particles at different locations on a microfluidic chip. Our technology, called Microfluidic CODES, combines Resistive Pulse Sensing with Code Division Multiple Access to compress 2D spatial information into a 1D electrical signal. In this paper, we present a practical demonstration of the Microfluidic CODES technology to detect and size cultured cancer cells distributed over multiple microfluidic channels. As validated by the high-speed microscopy, our technology can accurately analyze dense cell populations all electronically without the need for an external instrument. As such, the Microfluidic CODES can potentially enable low-cost integrated lab-on-a-chip devices that are well suited for the point-of-care testing of biological samples.

We demonstrate a microfluidic platform with an integrated surface electrode network that combines resistive pulse sensing (RPS) with code division multiple access (CDMA), to multiplex detection and sizing of particles in multiple microfluidic channels.

Microfluidic processing of biological samples typically involves differential manipulations of suspended particles under various force fields in order to ...

A Microfluidic Flow Chamber Model for Platelet Transfusion and Hemostasis Measures Platelet Deposition and Fibrin Formation in Real-time

Microfluidic models of hemostasis assess platelet function under conditions of hydrodynamic shear, but in the presence of anticoagulants, this analysis is restricted to platelet deposition only. The intricate relationship between Ca2+-dependent coagulation and platelet function requires careful and controlled recalcification of blood prior to analysis. Our setup uses a Y-shaped mixing channel, which supplies concentrated Ca2+/Mg2+ buffer to flowing blood just prior to perfusion, enabling rapid recalcification without sample stasis. A ten-fold difference in flow velocity between both reservoirs minimizes dilution. The recalcified blood is then perfused in a collagen-coated analysis chamber, and differential labeling permits real-time imaging of both platelet and fibrin deposition using fluorescence video microscopy. The system uses only commercially available tools, increasing the chances of standardization. Reconstitution of thrombocytopenic blood with platelets from banked concentrates furthermore models platelet transfusion, proving its use in this research domain. Exemplary data demonstrated that coagulation onset and fibrin deposition were linearly dependent on the platelet concentration, confirming the relationship between primary and secondary hemostasis in our model. In a timeframe of 16 perfusion min, contact activation did not take place, despite recalcification to normal Ca2+ and Mg2+ levels. When coagulation factor XIIa was inhibited by corn trypsin inhibitor, this time frame was even longer, indicating a considerable dynamic range in which the changes in the procoagulant nature of the platelets can be assessed. Co-immobilization of tissue factor with collagen significantly reduced the time to onset of coagulation, but not its rate. The option to study the tissue factor and/or the contact pathway increases the versatility and utility of the assay.

This paper describes an experimental model of hemostasis that simultaneously measures platelet function and coagulation. Platelet and fibrin fluorescence is measured in real-time, and platelet adhesion rate, coagulation rate, and onset of coagulation are determined. The model is used to determine platelet procoagulant properties under flow in concentrates for transfusion.

Microfluidic models of hemostasis assess platelet function under conditions of hydrodynamic shear, but in the presence of anticoagulants, this analysis is ...

Perfusable Vascular Network with a Tissue Model in a Microfluidic Device

A spheroid (a multicellular aggregate) is regarded as a good model of living tissues in the human body. Despite the significant advancement in the spheroid cultures, a perfusable vascular network in the spheroids remains a critical challenge for long-term culture required to maintain and develop their functions, such as protein expressions and morphogenesis. The protocol presents a novel method to integrate a perfusable vascular network within the spheroid in a microfluidic device. To induce a perfusable vascular network in the spheroid, angiogenic sprouts connected to microchannels were guided to the spheroid by utilizing angiogenic factors from human lung fibroblasts cultured in the spheroid. The angiogenic sprouts reached the spheroid, merged with the endothelial cells co-cultured in the spheroid, and formed a continuous vascular network. The vascular network could perfuse the interior of the spheroid without any leakage. The constructed vascular network may be further used as a route for supply of nutrients and removal of waste products, mimicking blood circulation in vivo. The method provides a new platform in spheroid culture toward better recapitulation of living tissues.

The protocol describes how to engineer a perfusable vascular network in a spheroid. The spheroid's surrounding microenvironment is devised to induce angiogenesis and connect the spheroid to the microchannels in a microfluidic device. The method allows the perfusion of the spheroid, which is a long-awaited technique in three-dimensional cultures.

A spheroid (a multicellular aggregate) is regarded as a good model of living tissues in the human body. Despite the significant advancement in the ...

A Microfluidic Platform for Stimulating Chondrocytes with Dynamic Compression

Mechanical stimuli are known to modulate biological functions of cells and tissues. Recent studies have suggested that compressive stress alters growth plate cartilage architecture and results in growth modulation of long bones of children. To determine the role of compressive stress in bone growth, we created a microfluidic device actuated by pneumatic pressure, to dynamically (or statically) compress growth plate chondrocytes embedded in alginate hydrogel cylinders. In this article, we describe detailed methods for fabricating and characterizing this device. The advantages of our protocol are: 1) Five different magnitudes of compressive stress can be generated on five technical replicates in a single platform, 2) It is easy to visualize cell morphology via a conventional light microscope, 3) Cells can be rapidly isolated from the device after compression to facilitate downstream assays, and 4) The platform can be applied to study mechanobiology of any cell type that can grow in hydrogels.

This article provides detailed methods for fabricating and characterizing a pneumatically actuating microfluidic device for chondrocyte compression.

Mechanical stimuli are known to modulate biological functions of cells and tissues. Recent studies have suggested that compressive stress alters growth ...