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

We demonstrate that through use of eco chip devices, physical structures and pressure heads can be held constant or varied. While the influence of surface chemistry fluid properties or the characteristics of the particles or microbes are investigated systematically through transport experiments using a non-pathogenic green fluorescent protein expressing VIO bacterial strain, we illustrate the importance of habitat structure, flow conditions, and inoculum size on fundamental transport phenomena. Microfluidic software compelling view of a hidden world.

Hi, our Dimitri Marker from Vanderbilt Institute for Integrative Biosystem Restoration Education at Vanderbilt University. And I’m David Schafer, a research and development engineer here at the Viro Laboratories. Hi, I am Leslie Sho from the Department of Chemical Materials and Biomolecular Engineering in the Center for Environmental Science and Engineering at the University of Connecticut, And I’m added Dicot also from Zebra.

Today we’re going to show you a procedure for creating modular microfluidic habitat arrays for basic environmental science and engineering research. We use this procedure in our laboratory to investigate the effects of micrometer scale structures on microbial transporting interactions in forest media. So let’s get started.

The first step in creating a microfluidic device is to draw a two dimensional layout of the device in a computer assisted drawing or CAD program. We’ve used AutoCAD, but other drawing programs are also available, such as clue in or coral draw. The software enables the designer to faithfully replicate physical structures throughout a macroscopic field, while individual features are defined at resolution of less than five microns.

From the CAD drawing, a photographic mask is created. We use a chromium mask manufactured by Advanced Reproductions Corporation, north end of Massachusetts. The next several steps of microfluidic device production take place in a clean room to create a master.

A clean silicon wafer is coated with photoresist in a spin coder. First, the wafer is attached to the spin coder platform and a few grams of SU 8 20 25 photoresist are poured onto the center of the wafer. Then the wafer is spun for an initial 15 second spread cycle at 500 RPM, followed by a 4, 000 RPM thin coating cycle for 35 seconds.

After spinning, the wafer is placed on a hot plate for 10 minutes at 95 degrees Celsius to volatilize excess solvent. The last step in preparing the wafer is to remove the edge bead. This is done by returning the wafer to the spin coder and applying a steady stream of SUA edge speed remover to the outside edge of the coated wafer.

The next step is to pattern the photoresist by selective exposure to UV light. First, the coated wafer is placed on a flat surface and the chrome mask is placed directly on top. Rubylith red masking film blocks the transfer of patterns outside the desired area.

Next, a 300 milli JUUL per centimeter square dose of UV light is delivered to the coated wafer through the mask using a spot carrying system. Finally, the wafer is baked for 15 minutes at 95 degrees Celsius to cross-link the light exposed regions of the photoresist rendering them insoluble. The last step in creating the master is to remove the non cross-linked regions of the photoresist leaving behind raised relief structures.

First, the wafer is returned once more to the spin coder and is covered with the developer solution. The developer is left on the coated wafer approximately five minutes and is then spun off, resulting in a clearly evident pattern on the wafer. Any remaining residue is washed off by squirting isopropyl alcohol onto the spinning wafer.

We use a spin speed of 2200 RPM for this step for about five to 10 seconds. Finally, the finished master is dried using a nitrogen gas stream and stored in a dish until it is needed for casting a device. We fabricate microfluidic devices using the silicone elastomer polymethyl alane.

First, the base material and curing agent are delivered in a 10 to one weight ratio into a plastic cup. Next, the mixture is mixed. Andy bubbled using a thinky AR 100 conditioning mixer.

Once the cycle is complete, liquid PDMS is poured evenly over the master. The dish containing the master and the uncured PDMS is then placed into a vacuum chamber to degas. The polymer bubbles appear almost immediately.

Degassing continues until all bubbles disappear. The dish is placed into a level oven at 65 degrees Celsius for at least four hours to fully allow polymer cross-linking. Then the cured PDMS is carefully removed from the reusable master.

First four edges are cut well outside the patterned region using a scalpel, the device is peeled away from the patterned wafer using tweezers. The edges of the device may be trimmed using shears if desired. A subsection of the device may be separated from the rest of the molded PDMS device.

Next, access holes are punched through the PDMS. We use biopsy punches that are commercially available in several different sizes mounted to a small arbor. Press PDMS cores are removed using forceps.

Finally, the trimmed and punched device is cleaned using isopropyl alcohol and a foam tipped polypropylene swab. The device is dried using nitrogen then placed in the oven to remove all traces of solvent. The last step in making a microfluidic device is bonding the PDMS to a glass slide substrate, then filling with fluid first.

The PDMS device and a glass slide are placed with the bonding surface spacing up into a plasma cleaner. In this experiment, we use a PDC 32 G herrick plasma cleaner. Once the vacuum is established and the blue violet plasma is visible, a timer is set for 30 seconds.

The plasma generator is shut off. The chamber is re pressurized, and the device and slider taken from the chamber and the bonding surfaces are placed into contact. In removing the activated components from the chamber, it’s important to move quickly so that surfaces remain activated.

Gentle pressure helps to ensure a good bond. The quality of the bond is checked by pulling on a corner with forceps. The hydrophilic interior can then be easily filled with an aqueous solution such as buffer media or deionized.

Water fluid is administered into each source well and through the corresponding microporous flow cell. Once all chambers are full, a thin layer of cured PDMS is placed over the entire device to help retard evaporation and maintain sterility. To calibrate pressure driven flow in the device, preload the micro structured flow cells with deionized water, place a flow module, which acts as a fluid delivery interface and a sterile seal on top of the device.

This flow module can be used on its own or connected to an external fluid containing reservoir, such as a plastic syringe. Connect a liquid containing reservoir such as a plastic syringe or the flow module to the upstream inlet. Well maintain the height of fluid in the reservoir above the height of the downstream.

Well, the difference in height between the upstream reservoir and each downstream outlet well defines the driving force for pressure driven flow to quantify flow rates through the devices. A blunt end 25 gauge needle is used to collect samples from the downstream wells at regular intervals. For example, we collect samples every 15 to 20 minutes carefully noting the actual elapsed time between samples.

The weight of the syringe is recorded before and after sampling on an analytical balance. Aggregate volume versus aggregate time is charted with the slope equal to the average mass flow rate. The large cross section of the inlet reservoir versus the outlet wells helps to prevent flow rates from changing significantly as fluid moves through the devices plot the sampled volume on the Y axis versus sample time on the x axis.

In a simple graphing program such as Excel to obtain the slope of the line, the slope is used for the average volumetric flow rate. A dilute solution of three micromolar fluorescent latex beads with carboxy, YG, and plain YG surfaces is used to visualize the flow and study surface interactions. First, a solution of beads is prepared in deionized water at a concentration of 0.01%solids.

The solution is added to the upstream wells and the flow module is reattached to zu. Flow bead flow is visualized over the course of one hour and recorded approximately every 10 minutes on a Zeiss AIOt 25 fluorescent microscope, equipped with a MITx BCEB 0 1 3 U monochrome camera. Individual frames are captured in rapid succession and assembled into movies using a software program such as Image J with fluorescence illumination and transmitted light illumination.

The beads and structures are both visible. When transmitted light is blocked, only the bright fluorescent beads are visible. Stationary beads deposited along the bottom surface or attached to the collectors are evident throughout the device.

For experiments using live bacteria, a culture such as Vibrio species, a non-pathogenic green fluorescent protein expressing bacterium is prepared. First, a stock culture is removed from deep free storage. A flask with sterile LB growth media is inoculated with the stock culture.

Using a disposable inoculating loop, the flask is incubated overnight with shaking to result in a stationary phase culture with a concentration of approximately 10 to the ninth cells per milliliter. Depending on the application, the initial concentration of bacteria may be quantified more precisely. A microfluidic device filled with sterile buffer is prepared.

Then the sterile liquid in the upstream wells is emptied and replaced with 20 microliters of the bacteria culture. Using a pipette, the bacteria are seeded into the device using gentle suction with a syringe on the output wells After incubation, the quantity of bacteria in the wells can be determined by quantifying the fluorescence intensity of each well following colonization. The upstream inlet well is emptied as before and replaced with buffer flushed with 25 parts per thousand salts.

Artificial seawater gentle pressure is applied to the upstream wells with a syringe flushing individual flow cells with 20 to 40 microliters of the buffer solution. This is approximately 100 to 200 times the poor volume of the microfluidic soil portion of the flow cell after the initial flush. Long-term perfusion of bacteria through the porous media is achieved by use of the flow module exactly as shown before.

Fluorescent and white light images of the bacterial growth and transport can be measured over time. In this experiment, images are taken every 15 to 20 hours for a period of seven days with a color five cooled CCD camera from Q imaging ink. Our devices can also be used to investigate effects of physical habitat structure on providing bacteria refuge from predation by protozoa.Seen.

Here are bacteria in a microporous flow cell with the much larger ciliated protozoa eulo visible in the device grazing on bacteria. In analysis of fluid flow in the microfluidic device, we find the average flow rate to be 0.71 microliters per minute for a 40 millimeter pressure head. By increasing the pressure head to 60 millimeters, the flow rate increased proportionally to 1.04 microliters per minute.

When using beads with various surface chemistries in the device, we observe a greater accumulation of un carboxylated beads at the device surfaces. While beads of six to 10 millimeters in diameter were much more likely to become entrained in the smaller pore openings. Faster flow rates caused by greater hydrostatic pressure resulted in decreased bead retention and entrainment, as would be expected to analyze bacterial growth in the eco chipp microfluidic device.

We note the qualitative differences in image intensity in different sectors of the device. Fluorescence intensity can be quantified as a proxy for microbial biomass in the channel. In ongoing experiments, we observe that the continuous shearing forces of low flow conditions seems to cause bacterial aggregation and flock formation as is evident in these images, We’ve just showed you how to create and use simple microfluidic devices to measure particle scale, collide transport through porous media, or to investigate microbial interactions in micro structured habitats.

When performing studies using flow cells, it’s important to consider the buoyancy of the beads and the surface chemistry of the colloids and collectors. When scaling down to the microfluidic regime, surface effects can dominate observed phenomena. Practically any physical structure can be incorporated into microfluidic flow cell design.

So that’s it. Thanks for watching and good luck with your experiments.