The overall goal of this procedure is to analyze single bacteria and evolving isogenic microcolonies with full spatial and temporal resolution under constant environmental conditions using a microfluidic cultivation device. This is accomplished by first designing a microfluidic cultivation system using CAD software. Next, disposable microfluidic cultivation devices are fabricated out of polydimethylsiloxane which contain picoliter bioreactors, one micron and height to restrict bacteria to monolayer growth.

Microbial culture is then infused into the microfluidic device to inoculate single mother cells within the bioreactors. These cells are expanded by continuous infusion of growth medium through the system. Results are obtained from automated time-lapse microscopy showing the cultivation of an industrially relevant strain of Corynebacterium glutamicum, growth data, and cell-cell heterogeneity.

In conventional biotechnology, most analytics are based on average data. Our microfluidics device facilitate single cell analysis of individual bacteria with spatial and temporal resolution. It is a useful tool to look close at cell to cell heterogeneity of isogenic microcolonies.

We apply common SU-8 photolithography and polydimethylsiloxane chip molding to manufacture single use microfluidic devices with sub micro media resolution. We have successfully demonstrated our technology with several biotechnical microorganisms such as Escherichia coli and Corynebacterium glutamicum. To begin, design the microfluidic device using CAD software.

The design presented in this protocol consists of two seeding inlets, a gradient generator for mixing two different substrates, one outlet, and six arrays consisting of five bio-reactors each. In a clean room, prepare a four-inch silicon wafer and spin code one micron of SU-8 2000.5 photoresist onto the wafer as described in the accompanying text protocol. Place the coated wafer on a hot plate at 95 degrees Celsius to drive off excess solvent, and then insert the first layer photo mask consisting of the trapping regions of the picoliter reactors and the coated wafer inside the mask aligner.

Then, expose the wafer in vacuum contact mode to 350 to 400 nanometer light for three seconds at an intensity of seven milliwatts per square centimeter. Perform a post-exposure bake on a level hot plate at 95 degrees Celsius to initiate the polymerization of SU-8. Next, place the wafer in an SU-8 developer bath for one minute, and then transfer the wafer into a second container with fresh SU-8 developer for a few seconds.

Rinse the wafer in isopropanol to remove the SU-8 developer and to dry the wafer using nitrogen flow or a wafer spinner. Then, hard bake the wafer for 10 minutes at 150 degrees Celsius. Next, fabricate the second layer as described in the accompanying text protocol in order to complete the master mold.

Begin PDMS chip fabrication by mixing the base and carrying agent in a 10 to one ratio. Degas the PDMS mixture for approximately 30 minutes inside a vacuum desiccator until all bubbles have disappeared. Next, place the SU-8 wafer into a molding device and pour a three millimeter layer of PDMS mixture onto the wafer.

Then, bake the PDMS for three hours at 80 degrees Celsius in an oven. Once it is cooled, carefully peel the PDMS slab from the wafer and cut the slab into single chips using a clean and sharp scalpel. Wash the chips in an n-pentane bath for 90 minutes, followed by two acetone washing baths for 90 minutes each.

Dry the chips overnight to remove any solvent residue. Just before the experiment, punch the inlet and outlet holes into the PDMS chip using a needle or hole puncher with a slightly smaller diameter than the connectors that are used to connect tubing with PDMS chip. Next, clean the microfluidic PDMS chip carefully with isopropanol and use scotch tape to remove any sticking dust particles.

Also, clean a 170 micron thin glass slide first with acetone, then isopropanol, followed by deionized water, and dry the slide with a stream of nitrogen. Once clean and dry, plasma oxidize the glass slide and PDMS chip for 25 seconds using a power of 50 Watts and an oxygen flow rate of 20 sccm. Then, carefully align the PDMS and glass chip before placing the PDMS onto the glass, which will cause it to bond in seconds.

Secure the bond by baking the setup for 10 seconds at 80 degrees Celsius. To prepare the bacteria for a chip experiment, inoculate a single colony of a desired bacterial strain, such as C.glutamicum into 20 milliliters of fresh BHI medium, and incubate the culture overnight at 30 degrees Celsius on a rotary shaker at 120 RPMs. At the next morning, transfer 10 microliters of the starter culture into a second culture containing 20 milliliters of the desired medium and let the cells grow overnight under the same conditions.

The next day, prewarm the incubator of an inverted microscope to 30 degrees Celsius. This may take up to two hours depending on the individual setup. While the setup is warming, open the incubator and select the desired objective and prepare it with any required mounting medium.

Then, mount the chip inside the chip holder and secure it in place. Center the sample on the microscope and focus into the picoliter bioreactor arrays. With the chip in focus, connect the inlets and outlets with appropriate tubing and then place the outlet tubing into a waste reservoir.

Next, fill a syringe with fresh media, place it into a syringe pump, and rinse the microfluidic channels for one hour at a flow rate of 200 nanoliters per minute to prime the chip. While the cells are still in the early exponential growth phase, transfer one milliliter of the bacterial culture into a sterile syringe and replace the syringe and tubing containing media with the cell suspension and fresh tubing. Infuse the cell suspension into the channels at a volumetric flow rate of 200 nanoliters per minute until most of the picoliter bioreactors are filled.

If only a small number of bioreactors are filled, increase the flow rate to 800 to 1200 nanoliters per minute. When the desired amount of cells have been seeded, disconnect the cell suspension, reconnect the growth medium to the chip, and perfuse with fresh media at 100 nanoliters per minute. Next, scan the chip and select specific bioreactors for time-lapse imaging.

Then, configure a time-lapse microscopy sequence in order to image the bioreactors from start to finish and to begin the experiment. Once all of the picoliter bioreactors are overgrown, stop the experiment and discard the chips appropriately. To begin analysis, first, determine which bioreactors fulfill all desired criteria for the experiment such as number and position of the mother cells.

Using an analysis program, such as image J, determine the growth rate of each microcolony by counting the number of cells at each time point. Calculate the maximum growth rate by plotting time versus the log of the cell number. The system presented here can be applied to study various bacterial species with respect to different biological parameters such as growth, morphology, or a fluorescent signal.

In this example, C.glutamicum was cultured under standard cultivation conditions and the resulting growth curves derived from three isogenic microcolonies are shown here. The picoliter bioreactors have also proven useful in accessing protein production through the use of time-lapse fluorescence microscopy as shown here. By exposing the cells to a low concentration of the protein inducer IPTG, the cell to cell variations of the protein can be studied at the single cell level within colonies starting from a single mother cell.

The demonstrated technique can be applied to monitor and analyze single bacterial cells with perfect environmental control. This is hardly possible using conventional bench scale cultivation systems. After watching this video, you should have a good understanding of how to prepare microfluidic cultivation chips to perform comparable single cell analysis.