Chips and Tips

Eriko Kamata, Momoko Maeda, Kanako Yanagisawa, Kae Sato*

Department of Chemical and Biological Sciences, Faculty of Science, Japan Women’s University, Bunkyo, Tokyo 112-8681, Japan

*E-mail: satouk@fc.jwu.ac.jp

Why is this useful?

There have been many reports on microfluidic devices for cell culture having upper and lower microchannels separated by a thin PDMS membrane. In these devices, the lower channel often interferes with the microscopic observation of cells cultured in the upper channel. To avoid interference, a microdevice with a detachable lower channel was developed.

What do I need?

Materials:

PDMS (SILPOT 184w/c, Dow Corning Toray)

Hexane

PMMA sheet (56 × 76 × 2 mm)

Glass slides (52 × 76 mm and 26 × 76 mm)

Cover slip (24 × 60 mm)

1 kg weight

PTFE tubing (1 × 2 mm and 0.46 × 0.92 mm)

Tygon tubing (1.59 × 3.18 mm)

Biopsy Punches (1 mm and 2 mm, Kai corporation)

Equipment:

Vacuum desiccator

Oven (65 ˚C and 100˚C)

Spin coater

Plasma generator

Vacuum pump

What do I do?

1. PDMS molding

Mix the elastomer and curing agent at a 10:1 mass ratio. De-gas the mixture under vacuum until no bubbles remain (20 min). Pour the degassed PDMS mixture onto a master, which has an upper channel (1 × 1 × 10 mm) or a lower channel (0.5 × 2 × 15 mm) structure, and then place it in an oven at 65˚C for 1 h. Peel off the PDMS replica from the master and adhere it to a glass slide (26 × 76 mm). Place it in an oven at 100˚C for 1 h (Fig.1).

2. Preparation of a thin PDMS membrane

Spin coat 600 µL of the PDMS prepolymer (at a 10:1 mass ratio) on a PMMA sheet at 500 rpm for 20 s followed by 2,400 rpm for 600 s. Bake it at 65˚C for 1.5 h.

Fig.1 PDMS sheet with the upper channel, that with the lower channel, PDMS membrane, and tubing.

3. Permanent bonding of the PDMS membrane and the sheet with the upper channel

Punch the inlet and outlet holes at both the ends of the upper channel with a 2-mm biopsy punch. Expose both the bonding surfaces of the PDMS membrane and the PDMS sheet with the upper channel (upper sheet) to plasma at 100 W, 35 s (Fig.2a and 2b). Laminate and bake them at 65˚C for 1 h (Fig.2c). Remove the PMMA sheet, and punch a hole to connect the sheet with the lower channel by using a 1-mm biopsy punch from the membrane side.

Fig.2 (a) Schematic diagram of bonding the PDMS membrane and the upper sheet. (b) Plasma treatment. (c) Bonded PDMS membrane and sheet.

4. Detachable bonding of the PDMS membrane and the PDMS sheet with the lower channel

The PDMS sheet with the lower channel (lower sheet) is bonded to the PDMS membrane by using a PDMS prepolymer diluted with hexane (a dilution ratio of 1:3) as a glue¹ (Fig.3a). Spin coat the diluted PDMS prepolymer (600 µL) at 2,000 rpm for 30 s on the surface of a glass slide (52 × 76 mm) to cover the slide with a thin layer of the glue and incubate it for 10 min to dry the solvent (Fig.3b). Place the lower sheet on the coated glass slide (Fig.3c). Apply glue at the four corners of the PDMS membrane bonded with the upper sheet (Fig.3d). Peel off the lower sheet from the glass slide and place the glue-coated surface of the sheet on the PDMS membrane (Fig.3e). After 30 min of incubation, bond the lower sheet to a cover slip by plasma bonding. Place a 1-kg weight and a glass slide on the device and bake it at 100˚C for 1 h (Fig.3f).

Fig.3 (a) Schematic diagram of bonding of the PDMS membrane and lower sheet. (b) Spin coat the diluted PDMS prepolymer (glue). (c) Lower sheet is placed on the thin film of the diluted PDMS prepolymer. (d) The diluted PDMS prepolymer is applied on the four corners of the PDMS membrane. (e)The glue-coated surface of the lower sheet is placed on the PDMS membrane. (f) A weight and a glass slide are placed on the device to bake it at 100˚C.

5. Tubing

Connect polytetrafluoroethylene (PTFE) tubes (1 × 2 × 100 mm) with Tygon tubes (1.59 × 3.18 × 10 mm) to the holes present at both ends of the upper microchannel. Connect a PTFE tube (46 × 0.92 × 150 mm) to the hole of the lower channel. Apply PDMS prepolymer at the root of the tubes, and then bake it at 100˚C for 1 h for firm connection (Fig.4a and b).

6. Cell culture

Introduce a cell suspension into the upper microchannel, which is manually precoated with 0.1 mg/mL of fibronectin. Incubate the device at 37˚C with 5% CO₂ for 16 h to allow cells to adhere to the bottom of the upper channel (surface of the PDMS membrane).

7. Detachment of the lower sheet for cell observation

Remove the lower sheet from the device carefully (Fig.4c and d). Place the rest of the device on a cover slip for observation with an inverted microscope (Fig.4 f and h).

Fig.4 (a) The complete microdevice. (b) Side view of the microdevice. The cell culture channel (upper) is filled with water containing a red food color, while the lower channel is filled with water containing a blue food color. (c) and (d) The lower sheet is peeled off from the microdevice carefully. Phase contrast images of cells (e) before and (f) after detachment of the lower sheet. Fluorescent images of cells stained with CellTracker Red CMTPX (g) before and (h) after detachment.

Conclusion

We developed a microfluidic device with a detachable lower microchannel. It is important that different bonding techniques be used for each side of the PDMS membrane. If the lower channel is filled with air and the device is incubated in a CO₂ incubator, dew condensation is often observed in the lower channel when the device is taken out from the incubator. The condensation in the lower channel makes observation difficult (Fig.4e and g). This problem was solved with the detachable device.

Acknowledgment

This work was supported in part by the Japan Society for the Promotion of Science (JSPS) KAKENHI Grant Number JP16H04170.

Reference

1. Chueh, B.H., Huh, D., Kyrtsos, C.R., Houssin, T., Futai, N., Takayama, S. (2007). Leakage-free bonding of porous membranes into layered microfluidic array systems. Analytical Chemistry 79 (9), 3504–3508.

Greiner, A.^1,2, Tekwa, E.W.^1,3, Gonzalez, A.¹, Nguyen, D.⁴

¹Department of Biology, McGill University, 1205 Dr. Penfield, Montreal, QC, H3A 1B1, Canada.
²Department of Ecology and Evolution, University of Toronto, 25 Willcocks Street, Toronto, ON, M5S 3B2, Canada
³Department of Ecology, Evolution, and Natural Resources, Rutgers University, 14 College Farm Road, New Brunswick, NJ, 08901, USA.
⁴Meakins Christie Laboratories, Research Institute of the McGill University Health Centre, and Department of Medicine, McGill University, 1001 Decarie Blvd, Montreal, QC, H4A 3J1, Canada.

Why is this useful?

Microfluidic devices are used for many different types of experiments across the medical, ecological and evolutionary disciplines (Park et al., 2003; Keymer et al., 2008; Connell et al., 2013; Hol & Dekker, 2014). For example, microfluidic devices for microbial experiments require inoculation into smaller chambers that simulate natural microbial environments such as porous soils (Or et al., 2007) and biological hosts (Folkesson et al., 2012). These devices often involve complicated pump setups and irreversible seals. We developed a technique that requires only common lab equipment and makes the device reusable while also allowing the microbes to grow undisturbed (based on Tekwa et al., 2015; Tekwa et al., in review). Here, we provide a detailed guide for the assembly and the previously undocumented non-destructive disassembly of polydimethylsiloxane (PDMS) experimental devices to recover microbes in situ, which can then be plated for relative counts and further molecular analyses of population changes. This is complemented by videos for each step.

Figure 1: Microfluidic device containing 14 habitats on an elastomer (PDMS) layer pressed onto a 60mm x 24mm glass cover slip. The habitats are 10 or 20µm in depth, range from 1400 µm to 2670 µm in diameter and take the shape of a ring or network of patches. This device is used to test the effects of habitat patchiness on microbe dynamics. Habitats were dyed blue for visualization. For more information see Tekwa et al. (2015).

What do I need?

• Single-layer PDMS devices with habitats on one side
• Pipette + sterile pipette tips
• Sterile petri dishes (1/device)
• Sterile tweezers
• Inoculum
• Filtered water
• Kimwipes
• Autoclavable plastic container
• Ethanol
• Tinfoil
• Sterile 1μL inoculating loop (1/habitat)
• Sterile eppendorfs
• Phosphate-buffered saline (PBS)
• Biological safety cabinet (BSC)

How do I do it?

1. Clean the PDMS devices: The PDMS device should be pre-treated once with 0.01N HCl for one hour and plasma-treated to keep it hydrophilic and amenable to bonding to glass or plastic substrates (Cho et al., 2007; Tekwa et al., 2015). Fill autoclavable plastic container 1/3rd full of 70% ethanol and PDMS devices and then cover with tinfoil, let them sit in this for >30 minutes before carefully disposing of the ethanol down the sink. Fill and empty the container with water 10 times in order to rinse the devices. Lastly, fill the container with filtered water, seal with tin foil and autoclave in order to sterilize the devices.
2. Inoculating the devices (perform in a Biological Safety Cabinet, BSC): Set the devices to dry in the BSC for 30 minutes. Place the devices with features facing up on the lid of a petri dish and place a small amount (i.e. 0.7 µL) of inoculum onto each (of 14 in sample device Fig. 1) habitat. The amount of liquid must be sufficient to fill the habitats, but not too much as to prevent bonding between PDMS and the cover glass/petri dish (Fig. 2). Using sterile tweezers, pick up the device and place face down into the centre of a petri dish or cover glass, sealing it to the surface by pushing on the back with gloved fingers repeatedly, using a kimwipe to wick excess liquid away from the side. Then surround, but not touch, the device with kimwipes soaked in filtered water (Fig. 3) to ensure that the device does not dry out in the incubator, before closing the petri dish. Place upright in incubator for the desired amount of time. The experiment can now proceed untouched for up to 24 hours (see Supplementary video).

Figure 2. Device with bacteria droplets, 1 droplet per habitat.

Figure 3. An ‘upright’ petri dish + kimwipes + device + coverslip ready to be incubated.

3. Recovering from the devices (perform in a BSC): Open petri dish, carefully remove and discard kimwipes and then use sterile tweezers to gently unseal the device and place face up in the lid of the petri dish. By around 12 hours, the spaces between the habitats will be void of liquid from PDMS absorption, preventing microbes from being mixed across chambers during disassembly. Habitats that have dried out will appear white (Fig. 4) and cannot be used.  Dip sterile inoculating loop into an eppendorf with PBS, then use that loop to scrape one of the habitats (Fig. 5).  Dip that inoculating loop back into the eppendorf media again, which can then be grown overnight for further analyses such as plating for relative cell count (if there are different strains) and other molecular analyses. Repeat for the rest of the habitats that you are interested in, using new inoculating loops. The PDMS device can now be cleaned as in Step 1 and reused again.

Figure 4. View of an inoculated and incubated device, looking through the bottom of a petri dish.

Figure 5. Recovering bacteria from a habitat in the disassembled PDMS device.

What else should I know?

The recovery technique can be used to estimate relative proportions of different types of microbes (e.g. morph frequencies) which is useful when performing competition assays and evolutionary experiments. Unlike in Tekwa et al. (2015), this technique forgoes the use of a confocal microscope; assessment of the contents of the device is instead performed through direct microbe recovery and standard plating procedures.

Links to Videos

These videos go through the specific procedure that we used to perform experiments on competition and cooperation in Pseudomonas aeruginosa and may be useful in determining specific amounts of media, growth times, etc. that may be utilized for experiments with similar PDMS microfluidic devices.

Part 1 – Intro + Washing the Devices https://youtu.be/Bne3ZN3wU4Q

Part 2 – Inoculating the Devices https://youtu.be/p-UYIRreYyM

Part 3 – Recovering from the Devices https://youtu.be/YLnmGxqMYgE

Supplementary – Fluorescent Bacteria Experiment https://youtu.be/lgMtryS62pA

Acknowledgements

AGr was supported by an NSERC Undergraduate Student Research Award and by an NSERC Discovery Grant. EWT was supported by the Fonds Québécois de la Recherche sur la Nature et les Technologies and the Québec Centre for Biodiversity Science. AGo was supported by the Canada Research Chair program and NSERC Discovery grants. DN was supported by a CFI Leaders Opportunity Fund (25636), a Burroughs Wellcome Fund CAMS award (1006827.01) and a CIHR salary award.

References

Cho, H., Jönsson, H., Campbell, K., Melke, P., Williams, J. W., Jedynak, B., … & Levchenko, A. (2007). Self-organization in high-density bacterial colonies: efficient crowd control. PLoS biology, 5(11), e302.

Connell, J. L., Ritschdorff, E. T., Whiteley, M., & Shear, J. B. (2013). 3D printing of microscopic bacterial communities. Proceedings of the National Academy of Sciences, 110(46), 18380-18385.

Folkesson, A., Jelsbak, L., Yang, L., Johansen, H. K., Ciofu, O., Høiby, N., & Molin, S. (2012). Adaptation of Pseudomonas aeruginosa to the cystic fibrosis airway: an evolutionary perspective. Nature reviews. Microbiology, 10(12), 841.

Hol, F. J., & Dekker, C. (2014). Zooming in to see the bigger picture: Microfluidic and nanofabrication tools to study bacteria. Science, 346(6208), 1251821.

Keymer, J. E., Galajda, P., Lambert, G., Liao, D., & Austin, R. H. (2008). Computation of mutual fitness by competing bacteria. Proceedings of the National Academy of Sciences, 105(51), 20269-20273.

Or, D., Smets, B. F., Wraith, J. M., Dechesne, A., Friedman, S. P. (2007). Physical constraints affecting bacterial habitats and activity in unsaturated porous media – a review. Advances in Water Resources, 30(6), 1505-1527.

Park, S., Wolanin, P. M., Yuzbashyan, E. A., Silberzan, P., Stock, J. B., & Austin, R. H. (2003). Motion to form a quorum. Science, 301(5630), 188-188.

Tekwa, E. W., Nguyen, D., Juncker, D., Loreau, M., & Gonzalez, A. (2015). Patchiness in a microhabitat chip affects evolutionary dynamics of bacterial cooperation. Lab on a Chip, 15(18), 3723-3729.

Tekwa, E.W., Nguyen, D., Loreau, M., Gonzalez, A. Defector clustering is linked to cooperation in a pathogenic bacterium. In review.

Dmitry A. Markov,^1,2 Elizabeth M. Lillie,^1,2 Philip C. Samson,^2,3 John P. Wikswo,^1,2,3,4 Lisa J. McCawley^2,5
1 Department of Biomedical Engineering, Vanderbilt University, Nashville, TN
² Vanderbilt Institute for Integrative Biosystems Research and Education (VIIBRE), Vanderbilt University, Nashville, TN
³ Department of Physics and Astronomy, Vanderbilt University, Nashville, TN
⁴ Department of Molecular Physiology and Biophysics, Vanderbilt University Medical Center, Nashville, TN
⁵ Department of Cancer Biology, Vanderbilt University Medical Center, Nashville, TN

Why is this useful?

Allows for sterile sample collection during long-term slow perfusion experiments. Current developments in microfluidic devices and their applications in biology during long-term cell and tissue culture have introduced additional challenges associated with periodic sterile collections of effluent for storage and further analysis. Pumps (either mechanical or electroosmotic) and gravity-fed systems work well for delivering media at very slow flow rates (< 1 µL/min). However, sample collection at these rates becomes a non-trivial task, especially when one-, two-, or three-week-long experiments are required. For example, it would take 24 hours to collect 1 mL of effluent at a flow of 0.7µL/min. Two important parameters to consider are sample evaporation during the collection step and sterility maintenance during multiple periodic collections. These are especially critical for the gravity-fed systems where the output port of the cell culture device has to be opened to the atmospheric pressure. To overcome these challenges, we have tested use of Breathe-Easy gas permeable adhesive membranes from Electron Microscopy Sciences (cat. # 70536-10) as aseptic filters on top of sample collection vials. These membranes are inexpensive, easy to use, and gas permeable, allowing for unimpeded fluid flow into the sample collection reservoir, while maintaining sterility of the whole system during sample collection. During our three-week-long cell culture experiments, we were able to perform multiple 24-hour effluent collections from our bioreactor with a 5-day period while maintaining sterility of the whole system. Upchurch valves were used to switch the flow between the waste reservoir (4 days) and the sample collection vials (1 day). The evaporation was reduced by keeping the complete system at 100% humidity within a CO2 cell culture incubator.

What do I need?

• 1.5 mL microcentrifuge tube
• Butane torch or Bunsen burner
• Small 25 gauge heated needle for puncturing
• Short piece of small 23 gauge stainless steel tubing to be inserted into the microcentrifuge tube
• 90 second epoxy (Araldite 2043 or similar)
• Small bore Tygon tubing (OD = 1/16th “, ID=0.020”)
• Drill bit #53
• Breathe-Easy membrane (Electron Microscopy Sciences, cat. # 70536-10)

How do I do it?

Tube insertion using stainless steel interface connector:

• Heat the puncturing needle with a cigarette lighter, Bunsen burner, or a small butane torch and carefully make a hole at the bottom of the microcentrifuge tube.
• Use a 4-6 mm long piece of small gauge stainless steel interface tubing, and insert it into the punctured hole.  Outer diameter of this tubing should be larger by 1 or 2 gauges than the puncturing needle (23 ga. tubing has larger diameter than 25 ga. tubing). The fit should be snug.
• Apply a small bead of 90 second curing epoxy to affix the interface tube to the microcentrifuge tube.
• After the epoxy dries, feed tygon tubing onto the stainless steel interface tube.
• In our case, we used a 25 ga needle to create a hole and a 23 ga stainless steel tubing (OD=0.022″) as an interface between a 1.5 ml microcentrifuge tube and flexible Tygon tubing (OD=1/16th, ID=0.020″).

Alternative procedure for direct flexible tube insertion:

• Drill a small hole in the bottom of the microcentrifuge tube
• Insert the Tygon tube and apply 90 second epoxy to glue the two parts together.
• We used a #53 drill bit (Ø = 0.0595″) for 1/16th OD tygon tubing.

To apply the filter:
The following procedure should be performed wearing gloves and under aseptic conditions.  Biosafety cabinets or laminar flow hoods with proper filtration are adequate.  The Breathe-Easy membranes typically come sterile from the supplier and should be opened inside the biosafety hood and handled with sterile tweezers.  The modified microcentrifuge tube with attached Tygon tubing can be sterilized using common procedures involving autoclave, alcohol, or gamma radiation sterilization (gas sterilization was not tested). All mentioned approaches performed reasonably well and were quite adequate; however, we have found that multiple autoclave sterilization cycles often softened the epoxied connections and alcohol sterilization required excessive handling of the components.  In our opinion, gamma ray sterilization was found to be the easiest, simplest, and the most convenient approach.

• Start with sterile membrane and microcentrifuge tube with proper tubing attached.
• Cut off the snap lid of the microcentrifuge tube.
• Peel one half of the protective film from one side of Breathe-Easy filter and attach exposed adhesive side of filter to the top rim of the tube.
• Then, remove the second half of the protective film and fully attach filter (make certain the seal is good around the perimeter with firm pressure applied with flat edge of forceps).
• When ready to use the assembly, carefully remove the top protective layer.

We used this method to periodically sample effluent from our bioreactor cartridges during long term cell culture.  Each cartridge contains 4 reactors that are connected to a single manifold shown in Figure 4D.

Figure 1. Tools and supplies needed

Figure 2. Preparing the microcentrifuge tube – Method 1. A) Hole punched with a hot needle. B) A piece of stainless steel needle inserted into the punctured hole. C) Inserted tube fixed in place with a 90 second epoxy. D) Flexible Tygon tubing attached to the interface tube.

Figure 3. Preparing the microcentrifuge tube – Method 2. A) Hole being drilled at the bottom. B) View of the drilled hole. C) Insertion of the small diameter Tygon tubing. D) Application of the 90 second epoxy to fix the inserted tube in place.

Figure 4. Preparation of the sterile cover. A) Protective plastic is removed from the adhesive side of the filter. B) Filter membrane is attached to the tube rim. C) Use caution to remove top protective plastic from the membrane attached to the tube. D) View of the tube with the filter mounted in the sample holder and attached to the collection network.

Acknowledgements

We would like to acknowledge support provided by the following grants: NIH/NCI R21 CA126728-01A1 and DOD/BCRP W81XWH-09-1-0444.