Revealing Electromechanical Control of Tissue Homeostasis Using a Two-Layer Microfluidic Device

Summary

Here, we present a protocol to fabricate a unique two-layer microfluidic device to study the electromechanical regulation of epithelial tissue homeostasis. The device applies static physiological electric currents perpendicular to the tissue plane, impacting cell-cell adhesion, proliferation, and extrusion. Live-cell imaging and mechanical stress measurements reveal mechanisms of these processes.

Abstract

Cell behavior and cell fate are impacted by electric currents or fields that are endogenous or externally applied. Static electric signals can be applied in customized microfluidic devices to mimic the electric environment in slow physiological processes such as development, wound healing, and homeostasis. An important class of cellular electric studies is the control of cell migration by static in-plane galvanic currents in simple microfluidic channels mimicking wound currents, with current densities of ~0.1-1000 A/m². However, due to incompatible geometry, these devices are not appropriate to study electric effects in tissue homeostasis, where cells adopt apico-basal polarity and a transepithelial potential difference (TEPD). Here, we detail a unique microfluidic-based device that applies physiological ion currents perpendicular to the plane of confluent epithelial cell layers to perturb the TEPD and investigate electrical regulation of tissue steady states. The setup is made from a two-layer UV-curable polymer embedded with soft, polyacrylamide gel substrate coated with extracellular-matrix protein of choice. This microfluidic device provides the correct geometry and permeable substrate to induce a relatively uniform ion current across the cell layers of centimetric-scale. The setup is compatible with confocal live-cell imaging and Traction Force Microscopy to infer mechanical stresses induced by the transepithelial currents. Strikingly, the proliferation, extrusion and migration of cells are collectively influenced within the confluent epithelium depending on the direction of ion current, inducing a new tissue state characterized by different cell-cell interaction strengths, cell events (death and proliferation), and tissue structures. The electrically controlled cell behaviors can be understood as an electrically induced mechanical stress and cell response. This novel microfluidic device and protocol provide the tool and documentation required for the mechanobiology and bioengineering communities to study electric effects in tissue homeostasis and develop novel tissue engineering applications.

Introduction

Bioelectric signals are important biophysical factors contributing to governing cell behavior. The most famous example is the action potential in neurons and heart cells, which transmit electrical signals over long distances (up to a meter) and at high frequencies (up to hundreds of Hertz)¹. Electrical pulses applied with electrodes can be used to stimulate action potential firing and is useful in research and medical applications². Endogenous bioelectric signals in slow physiological processes such as development, wound healing, and homeostasis^3,4,5,6,7,8 have also been measured^9,10 over periods of hours to days. An important example is confluent epithelial layers which actively maintain a transepithelial potential difference (TEPD) ranging from a few to tens of millivolts^4,11. A cut or wound in these confluent layers generates a short-circuit current, driven by the electrical activity of the tissue^12,13. Wound currents up to tens of µA/cm² and electric fields up to hundreds of mV/mm have been reported with different techniques. Studies suggest that these electric signals produce a directional cue for cells to migrate toward wound or infection sites that is important for the healing process^3,6,14. In general, both static (or slow-varying) and fast-varying electric signals are important and trigger different cell or tissue responses.

Within the realm of slow-varying signals, one important phenomenon is the electrically-driven migration process termed "galvanotaxis". Controlled in-plane galvanic currents can be applied in microfluidic devices with simple channels to mimic the wound currents while cell migration is investigated as a function of current density^15,16. Electric fields of 0.1-1000 V/m (corresponding current density of 0.1-1000 A/m²) are found to drive cell migration in many cell types in this in vitro setting¹⁷, while signaling pathways canonical to chemotaxis (cell migration guided by chemical gradients), such as PI3K, have been implicated in galvanotaxis^7,18. "Bare-bones" microfluidic devices for galvanotaxis study can be fabricated with standard soft lithography and microfabrication techniques¹⁹. These devices typically comprise a single-channel polydimethyl-siloxane (PDMS) substrate bonded to a glass slide (channel cross-sectional size: hundreds of micrometers, length: centimeters). Electrodes used to supply the static current are installed far away from the cell region, and connected with or without salt bridges. The salt bridge connection may be made by punched holes through the PDMS substrate, microfluidic connectors, and tubings filled with agar. The simple glass slide (typically 170 µm thick) beneath the channel also supports high-resolution live-cell imaging during the galvanotaxis experiments, which is a crucial feature for studying the process.

The role of endogenous electric signals in the maintenance of tissue homeostasis is much less understood than that in wound healing. One important reason is the lack of appropriate tools to study these effects in intact tissues. The microfluidic device for galvanotaxis is not suitable for this purpose due to its geometry, which only supports in-plane electric currents. In the context of epithelial homeostasis, a new device is needed to control and perturb the TEPD or out-of-plane currents across a confluent epithelial layer. To achieve the aim of applying a relatively uniform ion current density across a monolayer, we have designed and fabricated a microfluidic device with innovations in device geometry, material usage, and in situ biomechanical measurement capabilities⁴. The latter capability is important as electric effects may be affecting cell behavior through coupling with biomechanics and mechanobiology pathways, which can be studied with this feature.

This new device exhibits multiple innovative features. It is designed with two layers instead of the single channel found in the galvanotaxis devices (innovation in geometry), while keeping the connection to the electrode chambers similar to that in the latter. The first microfluidic layer has channels to direct ion currents to (or away from) the tissue region, while the second layer has an opening that will direct this current perpendicularly across the epithelium sitting on top of this second microfluidic layer (Figure 1A,B). Microfabrication techniques are used for producing sufficiently thin layers such that high-resolution confocal imaging is possible despite the stacking of microfluidic layers that increases sample thickness. For the other important innovations, a polyacrylamide (PA) synthetic hydrogel is incorporated into the middle of the device, which plays a crucial role (Figure 1A,B). The physico-chemical properties of this gel, such as available chemistry to coat extracellular matrix proteins, permeability to ions, as well as known elasticity, are respectively essential for cell attachment and growth, application of out-of-plane currents, and measurement of mechanical stress²⁰ during electric stimulation. Barrier structures are incorporated in Layer 1 of the tissue region for confinement of the PA gel precursor solution, as done in previous microfluidic devices²¹ (Figure 1A,B). Last but not least, since PA gel polymerization is oxygen-inhibited at PDMS surfaces permeable to oxygen gas, another material²² is used for device fabrication, which can allow polymerization and tight bonding of PA gel. All the innovative features are crucial for the new device to work.

Overall, the in-depth study of static or slowly varying bioelectric signals in the homeostasis of intact tissues is made possible with this new device, the protocol for which will be detailed below. The protocol and results referred to here are focused on the Madin Darby Canine Kidney II (MDCK II) epithelial cell line, including the cell events (proliferation, death, and extrusion, and migration) and tissue structural changes that are induced by the electric current. These studies are also instructive for researchers who are interested in applying this technique to study the electrical regulation of epithelial transport and induction of cell events and cell fate changes in cell layers and more complex systems, such as primary cells and 2D organoids.

Protocol

NOTE: This protocol provides steps to construct a setup that applies static electric currents and perturbs the TEPD of confluent epithelia and to study its effects through live-cell imaging and mechanobiology techniques (Figure 1). The setup consists of devices made of microfluidic chips bonded to cartridges, inserts, and holders to house multiple devices for multiplexed experiments on a microscope, a pair of electrode chambers connected to a source meter and devices, and a live-cell chamber lid for the maintenance of proper live-cell conditions (Figure 1C). The design principles and CAD file drawings of the cartridge and other important components of the setup can be found in Supplementary File 1. The steps include: 1) PDMS soft lithography; 2) Chip fabrication and device assembly; 3) Setup completion; and 4) Monolayer growth and live-cell experiments. Detailed information regarding the reagents and materials utilized in this protocol is given in the Table of Materials.

1. PDMS soft lithography

NOTE: This section details producing PDMS elastomers from SU-8 wafers (Figure 2A, B) to mold UV-curable adhesive-based layers of the microfluidic chip.

1. PDMS soft lithography
   1. Fabricate a separate SU-8-based wafer for each microfluidic layer, Layer 1 and 2, using photolithography techniques (Supplementary File 2 and Supplementary File 3).
      NOTE: Wafer fabrication may take a month if a request is sent to a microfabrication/clean room facility. Silicon etching technique for wafer fabrication may also be used.
   2. Silanize the silicon wafer to allow PDMS unmolding¹⁹.
      1. Clean the wafer with a pressurized nitrogen air blow gun. Activate the surface of the wafer by applying a short oxygen plasma process (30 s, 30 W in 20 sccm O₂ gas flow at a reactor pressure of 1-3 mbar)²³.
      2. Pipette 200 µL of Tridecafluoro-1,1,2,2-tetrahydrooctyl-1-trichlorosilane in a clean plastic cap (diameter 60 mm, height 15 mm). Put the cap next to the wafer in the vacuum desiccator (only for silanization). Turn on the vacuum pump and wait for 1-2 h, while the vaporized silane is deposited as a monolayer on the silicon wafer.
         CAUTION: Silane is corrosive and toxic. Wear gloves and safety goggles, and operate in a fume hood to ensure safe operation.
         NOTE: A newly silanized wafer can be used to fabricate 10-20 pieces of PDMS molds. Vacuum time depends on the condition of the vacuum pump. Excessive silanization causes uneven thickness and results in a cloudy appearance, which is to be avoided. Store wafer in humidity-controlled chambers when not in use. The PDMS structures made from new wafers can be checked using scanning electron microscopy (SEM).
   3. Mix sufficient base PDMS resin and crosslinking agent thoroughly (w:w = 10:1, and 70-80 g) for a few minutes in a Petri dish (diameter 150 mm, height 25 mm). Ensure small bubbles form uniformly in the mixture, which confirms a good mixture. Transfer the mixture into 50 mL centrifuge tubes and centrifuge at 550 x g for 3 min to remove air bubbles thoroughly.
      NOTE: PDMS liquid is sticky and difficult to clean; be careful when pouring. Clean with isopropanol if bench surfaces are contaminated by spilled PDMS mixture.
   4. Gently pour the PDMS mixture on the wafer in a plastic or glass dish (diameter 150 mm, height 25 mm) with a flat bottom, then push down the wafer to purge PDMS from the bottom of the wafer. Degas mixture with a vacuum pump.
      NOTE: Purging of liquid PDMS from the bottom of the wafer is important; a thick PDMS layer at the bottom makes it difficult to remove the wafer from the plastic dish, increasing the probability for the wafer to shatter. Alternatively, one can cut the PDMS along the edge of the wafer and keep the wafer in the petri dish. Ensure PDMS thickness of roughly 1.5 mm for Layer 1 to ensure flexibility of the mold in later fabrication steps. Ensure PDMS thickness of roughly 2.5 mm for Layer 2 to reduce the probability of sagging in later fabrication steps. The PDMS thickness is measured by a ruler after the PDMS molds are cut into the shapes needed.
   5. Cure the PDMS in an oven at 80 °C for 2 h, and ensure the dish is placed as horizontally as possible during the process. Allow the PDMS mold and wafer to cool down to room temperature. Carefully remove the PDMS mold in its entirety from the wafer using a sharp surgical blade and padded tweezers.
      NOTE: Be careful not to scratch the surface of the wafer. Check if PDMS remnants are formed on the wafer surface when PDMS is peeled off. This signals that the wafer surface should be re-silanized.
   6. Cut the PDMS mold into blocks that contain the features of interest and store these blocks in clean Petri dishes (diameter 150 mm, height 25 mm) or a vacuum chamber when not in use. Manipulate the PDMS mold without touching the essential features.

2. Chip fabrication and device assembly

NOTE: Use a low-viscosity, UV-curable polymer (maximum UV absorption range 350-380 nm) to fabricate the microfluidic chip through PDMS molding. UV-curable adhesive allows integration of polyacrylamide (PA) hydrogel as a transparent, ion-permeable, and elastic cell substrate. The properties of PA gel support it as a tissue mechanical stress sensor that allows live-cell imaging and static current stimulation. The chip will finally be attached to a cartridge that houses culture medium and connects to electrode chambers.

1. Glass coverslip base preparation
   1. Score the required amount of rectangular glass coverslips (170 µm) with a diamond cutter to a size of 37 × 24 mm², to fit cartridge dimensions (Supplementary File 4).
      NOTE: The coverslip has 170 µm thickness compatible with high-resolution imaging, but is fragile and requires careful handling. Wear gloves and eye protection. Dispose of the rectangular glass coverslips immediately if any visible cracks appear, as chips from these rectangular glass coverslips may pose a risk of leaking. Alternatively, order coverslips of the right size from coverslip manufacturers. Avoid touching the surfaces of rectangular glass coverslips with bare hands and leave stains that can deteriorate imaging quality.
   2. Put the rectangular coverslips in a clean Petri dish without overlapping, and transfer them to a plasma chamber. Induce a vacuum condition, but let in a bit of air at the end. Activate the surfaces of the rectangular glass coverslips using air plasma (30 s to 5 min, 29.6 W, depending on the condition of the equipment)
      NOTE: Only the sides fully exposed to plasma (i.e., upper surfaces) should be used in the subsequent step (step 2.1.3). Complete step 2.1.3 within 30 min after plasma exposure. The chamber should appear as Pink-Purple at this stage if air plasma has been generated. If the latter bonding steps are successful, reduce the plasma duration for convenience.
   3. Prepare 10 mL of surface-treatment solution made of 0.3% acetic acid and 0.5% 3-(trimethoxysilyl)propyl methacrylate dissolved in 100% ethanol. Treat rectangular glass coverslips with this solution in a Petri dish (diameter 150 mm, height 25 mm) for 5 min on a shaker, to allow good bonding with UV-curable adhesive-based microfluidic layers. Ensure the rectangular glass coverslips are fully immersed in the solution, and both sides are treated.
      CAUTION: The surface-treatment solution is corrosive. Handle it in a fume hood.
   4. Rinse rectangular glass coverslips with 100% ethanol three times, without letting them dry during the process. Dry rectangular glass coverslips one by one with nitrogen gas and a blow gun. Ensure a sufficient blow rate that cleanly removes liquid droplets from the coverslip base surfaces, preventing liquid stains. Store rectangular glass coverslips in a clean Petri dish (diameter 150 mm, height 25 mm).
      NOTE: Use a blow gun to direct air flow parallel to coverslip bases and avoid using excessive blow rate to prevent glasses from shattering.
2. Layer 1 fabrication (Figure 2C)
   1. Place and flatten a clean Layer 1 PDMS mold, face-up, on a clean plastic dish cap (diameter 60 mm, height 15 mm). Pipette 500 µL liquid UV-curable adhesive onto the mold slowly, carefully wetting all the protruding structures with the aid of a spatula. Remove air bubbles due to the wetting process.
      NOTE: Clean PDMS molds can be prepared by using tape to remove dust or UV-curable adhesive remnants, or by cleaning with ethanol. For additional precaution, keep PDMS molds in vacuum chambers when not in use.
   2. Gently press down the coverslip onto the PDMS mold. Prevent excess UV-curable adhesive from wetting the top side of the coverslip, as UV-curable adhesive remnants can prevent a clean and flat coverslip. Remove excess UV-curable adhesive with laboratory wipers.
      NOTE: It is critical to ensure that the features of the PDMS mold transferred to the UV-curable adhesive layer will align with the cartridge to which it will be bonded. To do so, mark the coverslip with a soluble marker pen where the centers of the cartridge wells (three of them) will be situated. Use these marks to align the coverslip with the PDMS mold features when pressing the rectangular glass coverslip down.
   3. Partially cure the structure under a uniformly illuminated UV light (363-370 nm, 224 mW/cm²) for 10 s. Wipe off the marks with 75% ethanol later. Hold the structure firmly on a flat surface, and peel Layer 1 PDMS mold slowly from it.
      NOTE: Partial curing enables the formation of a sufficiently solid UV-curable adhesive layer, allowing for the peeling of the PDMS mold at this stage. This also facilitates bonding with a partially cured Layer 2 in subsequent steps for the fabrication of a two-layer UV-curable adhesive structure. The Layer 1 PDMS mold, which was prepared with a thickness of 1.5 mm (step 1.2.4), ensures sufficient PDMS flexibility that reduces the probability of the thin coverslip from shattering during the peeling process. PDMS mold can be reused for up to ten to twenty times, after which there may be a whitish layer of UV-curable adhesive remnants on the PDMS surface, reducing the structure thickness. Use new PDMS molds in this case.
3. Layer 2 fabrication (Figure 2D)
   1. Place a clean Layer 2 PDMS mold, face-down, on a flat PDMS slab (slab thickness of > 2.5 mm to ensure high rigidity). Gently tap the features from above using a tweezer. Observe the interface for darkening or optical contrast beneath the features - this indicates proper contact.
      NOTE: The support pillars surrounding the main features (two round openings and a slit) are crucial to prevent the PDMS mold surface from sagging.
   2. Fill the space in between the PDMS mold and slab with liquid UV-curable adhesive via capillary effect. Partially cure the structure under UV light (363-370 nm, 224 mW/cm²) for 10 s.
   3. Gently peel the composite UV-curable adhesive layer and PDMS mold from the PDMS slab with a soft-padded tweezer. Trim excess edges of the UV-curable adhesive layer with a sharp pair of scissors if needed.
4. Layer 1 and 2 bonding (Figure 2E)
   1. Place Layer 1 attached to a rectangular glass coverslip on a flat surface. Align Layer 2 (still attached to its PDMS mold) with Layer 1 and press both layers firmly, ensuring "shadows" show up under the features.
   2. Cure the structure under UV light (363-370 nm, 224 mW/cm²) for 10 s to bond Layers 1 and 2, which form the two-layer structure of the microfluidic chip. Gently peel the PDMS mold from the chip.
      NOTE: To ensure the successful peeling of PDMS mold from the chip, use tweezers to separate each PDMS support pillar from the UV-curable adhesive Layer 2 in step 2.4.1. This action helps to reduce the overall adhesion between the PDMS mold and Layer 2, which is important for peeling.
5. Polyacrylamide gel integration (Figure 2F)
   NOTE: Before this section, prepare as many chips as required for a multiplexed experiment.
   1. Prepare PA gel precursor solution by gently mixing the ingredients in the table below, corresponding to the PA gel stiffness of choice²⁴ (check Table 1). Be careful not to introduce air bubbles during pipetting as oxygen inhibits gelation.
      CAUTION: TEMED and APS are toxic. Work in a fume hood and wear gloves.
      NOTE: Careful pipetting and handling are required as small volumes of liquids are involved (1 µL TEMED for 1 mL of PA gel precursor solution). Gel stiffness can influence cell behavior as shown in many mechanobiology studies. Choose the stiffness most appropriate for your experiments. Fluorescent beads serve as fiducial markers for gel deformation and mechanical stress inference. Vortex bead solution before pipetting the solution into the PA gel precursor solution. It is recommended not to mix the small volume TEMED last since PA gel precursor solutions starts to gel when both initiators are added and small volumes are difficult to mix thoroughly during this short time.
   2. Immediately pipette 10 µL of the PA gel precursor solution in the middle of the tissue region and on top of the slit of Layer 2 (Fig. 1B, 2F). Gently press down with a round glass coverslip (diameter 10 mm) to form a flat gel. Observe the PA solution flow into the Layer 1 through the slit and stop at the row of barrier pillars.
      NOTE: The 10 µL volume of PA gel precursor solution produces a 100 µm-thick gel above Layer 2. Change this volume depending on the gel thickness required. Pipette the solution on the chip as soon as possible, as the PA solution starts to polymerize once initiators are added. Press the solution with the right amount of force to prevent the solution from overflowing across the barrier pillars and blocking the channel. Discard the chip if the channels are blocked. For soft gels with 5 kPa stiffness or below, use glass water repellent to treat the round glass coverslips to render the coverslips hydrophobic. In particular, immerse the coverslips in glass water repellent for 5 min, rinse with 100% ethanol 3 times, and dry with nitrogen gas. This facilitates peeling the coverslip from the polymerized gel later without destroying the gel.
   3. Immediately cure the microfluidic chip under UV light (363-370 nm, 224 mW/cm²) for 5 min. Wait for at least ~1 h for the PA gel to fully solidify.
      NOTE: This will fully cure the chip while allowing the polymerized PA gel to bond to the structure. Steps 2.5.1, 2.5.2, and 2.5.3 should be completed in quick succession.
   4. Incubate the chip thoroughly in HEPES (0.1 M, pH 7.4) for at least 1 h or more to wet the gel and loosen the adhesion between the coverslip and the gel. Gently remove the cover glass with a sharp pair of tweezers.
      NOTE: For softer gels with 5 kPa stiffness or lower, increase incubation period up to 10 h on a shaker to ensure clean removal of cover glass from the gel without disrupting gel structure. Sometimes, structural patterns of several tens and hundreds of micrometers may form between the rectangular coverslip base and the UV-curable adhesive-based Layer 1. These patterns can interfere with bright-field imaging but have minimal effects on fluorescence imaging. This can be improved by plasma activation of rectangular coverslips as in step 2.1.2.
6. Chip and cartridge bonding (Figure 2F)
   1. Place a cleaned, medical-grade polycarbonate cartridge (Supplementary File 4) on a flat surface, with the bottom face up. Dry the gel on the chip with laboratory wipes from the side. Align the chip, face down, with the cartridge.
      NOTE: When testing new cartridge designs, one can use machining or 3D printing methods. When the design is set, one can use injection molding to produce large quantities of the cartridge.
   2. Pipette UV-curable adhesive into the crevices between the chip and cartridge, which will be filled via capillary effect. Cure with UV light (363-370 nm, 224 mW/cm²) for 5 min.
      NOTE: Careful to avoid UV-curable adhesive overflowing into the channels, or to the other side of the coverslip base.
7. Removal of unpolymerized PA solution
   1. Use a PDMS stopper and a syringe to draw approximately 5 mL of HEPES (0.1 M, pH 7.4) through the waste removal channels to flush out the unpolymerized PA gel precursor solution that is toxic to cells.
   2. Incubate the gel in the device with 2 mL of HEPES (0.1 M, pH 7.4) per device, and put it on a shaker for 1-2 days, to ensure complete removal of unpolymerized PA gel precursor solution.

3. Setup completion

NOTE: With the devices made, complete the setup (Figure 1C) by coating PA gel with ECM protein for cell seeding, connecting the devices to the electrode chamber for electric stimulation, and placing the devices in inserts, holders, and a live-cell chamber lid for multiplexed, live-cell imaging experiments.

1. Preparation of sterilized components
   1. All components of the setup other than the device (Table of Materials) should be sonicated in the following order: first in soapy water for 30 min, then in MQ water for 30 min, and finally in 75% ethanol for 30 min. After that, dry them in an oven at 60 °C and treat them with UV light at 200-280 nm in a biosafety cabinet for 30 min to achieve sterilization.
2. PA gel functionalization with ECM protein
   1. Sterilize the devices with UV light (200-280 nm) in a biosafety cabinet.
   2. Prepare the required amount of 50 µg/mL Collagen I working solution in 1x DPBS on ice.
      NOTE: Maintain Collagen I stock on ice to prevent gelation. 100-150 µL of Collagen I working solution is required for each device to fully immerse the gel. Gently pipette Collagen I solutions to avoid fiber formation.
   3. Prepare the required amount of Sulfo-SANPAH (SS) working solution on ice. Each gel treatment requires approximately 80 µL of SS working solution, where 2 µL SS stock solution (dissolved in anhydrous DMSO) is diluted in 80 µL cold HEPES (from 4 °C fridge, 0.1 M, pH 7.4).
      NOTE: SS is moisture-sensitive; avoid moisture condensation onto the product by equilibrating stock vials to room temperature before opening. Use No-weigh SS for best performance, and prepare stocks in anhydrous DMSO. Use it as soon as possible. A cold HEPES buffer is used to reduce the hydrolysis rate of SS in preparation for the SS working solution. Each gel requires two treatments with the SS working solution to ensure the PA gel surface will be functionalized well.
   4. Dry the gel in the device by dabbing with laboratory wipes on the edges of the gel. Pipette 80 µL of SS working solution on the gel, ensuring the gel surface is completely immersed with SS solution. Treat the gel with UV (363-370 nm, 24.5 mW/cm²) for 5 min to activate SS. Rinse 3 times with cold HEPES (0.1 M, pH 7.4).
      NOTE: Carefully pipette SS in the middle of the cartridge well to prevent SS from being attracted to the wall.
   5. Repeat step 3.2.4, but rinse three times with cold 1x DPBS at the end.
      NOTE: Steps 3.2.3 to 3.2.5 should be done in quick succession to prevent SS hydrolysis. The rinsed gel should appear as a darker shade of red at this stage if surface activation by SS is successful.
   6. Pipette 100 µL Collagen I solution onto PA gel, making sure to fully immerse the gel. Incubate at room temperature for 1 h, then rinse off unattached collagen with 1x DPBS three times. Immerse the gel in 1x DPBS before use.
      NOTE: Collagen I-coated sample can be stored in 4 °C fridge for some days.
3. Connection of devices to electrode chambers
   1. Attach devices to the inserts using screws (Figure 1C, screw type: M3 X 10 mm). Then, secure up to four device-insert assemblies to the holder designed for mounting on a microscope stage (screw type: M3 X 5 mm).
      NOTE: If the maximum number of device-insert assemblies is not used, it is recommended to fill all the other holder slots with empty devices to seal the space so that good live-cell conditions can be preserved for the imaging experiments later. Empty devices can simply be made by bonding cartridges to bare rectangular glass coverslips.
   2. Fill devices with culture medium (DMEM++, 2 mL in each of the main wells). Further pull 2 mL of medium through the waste removal channels to flush out and replace the 1x DPBS.
   3. Prepare two 50 mL tubes with caps that have through holes to function as electrode chambers. Create holes of appropriate size and quantity to securely fit the platinum electrodes and Tygon tubings, connecting the electrode chambers to the devices. Fill each electrode chamber with 45 mL of culture medium.
      NOTE: Holes can be easily created on the caps using sharp surgical blades.
   4. Prepare the connection between electrode chambers and devices: Attach two tubing connectors to each device. Slot a tubing clip onto each Tygon tubing (Figure 1C). Pull culture medium through the tubings with a syringe and clip them tightly. Connect one end of the medium-filled tubing to the electrode chamber and the other to the device.
      NOTE: Reduce the probability of emergence of bubbles in tubing by using prewarmed medium and/or degasing the medium in a vacuum chamber. Prevent air bubbles from forming at the interface between Tygon tubing and the tubing connector, pinch on the tubing ends to bulge the medium surface out before immersing into the medium in the tube, or make a physical attachment to the tubing connectors.
   5. Adjust the relative heights between the culture medium surfaces in the devices and electrode chambers using a lab jack. Release the tubing clips and let all medium surfaces equilibrate to the same height.
      NOTE: Remember to fasten the tubing clips whenever the whole setup is being moved to avoid overflowing of fluids.
   6. Perform quality control by measuring the resistance of each device separately using an electric source meter.
      NOTE: Make sure to release only the tubing clip of the device for which the resistance is being measured, while keeping the clips of the other devices connected in parallel securely fastened.

4. Monolayer growth and live-cell experiments

NOTE: To form a proper confluent monolayer and to achieve a uniformly distributed electric current density that depends on the confluency and electrical resistance of the monolayer. Then, ensure proper live-cell conditions on a microscope for long-term imaging and electric stimulation experiments.

1. Monolayer growth
   1. Detach MDCK cells using Trypsin from T25 flasks. Centrifuge down and count the medium volume needed to obtain 0.1 million cells. Suspend this number of cells and mix them well in 2 mL medium in centrifuge tubes.
   2. Pipette all the cell-laden medium into the well onto the tissue region. Let the cells sediment for 15 min before slowly transferring the whole setup into a tissue culture incubator.
      NOTE: For each device, a seeding density of 0.1 million cells corresponds to 0.06 million cells per square centimeter (6 × 10⁴ cells/cm²). This seeding density is sparse enough, and the cells grow for 2 days before reaching a confluent state. This process ensures a properly polarized and homogenous monolayer, without needing to rinse off excessive unattached cells that risk perturbing the tissue. The sedimentation step is important to be left undisturbed so that there is even seeding of the cells.
   3. Allow cells to grow in the incubator (37 °C, 5% CO₂, 90% humidity) for 72 h to reach a cell density of >~50 cells/100 μm². Check daily on a bench-top microscope, with phase contrast, to monitor the growth process.
      NOTE: With this number of cells seeded, MDCK cells will have reached 70%-80% confluency after 24 h, full confluency after 48 h, and further cell density increase and tissue maturation till 72 h. After 24 h, the average cell length is about 10-20 µm. At 48 h, cell-cell junctions and the cell body have a clear contrast under phase contrast imaging. This contrast drastically reduces after 72 h.
2. Live-cell imaging
   1. Choose an epifluorescence or a confocal microscope depending on experimental needs. When cells express or are stained with fluorescent markers, choose confocal for three-dimensional observation of cell shape and gel substrate deformations. For two-dimensional observation, choose phase contrast and epifluorescence, which produce less phototoxicity. Turn on the live-cell system 1 h prior to bringing the setup to the microscope, allowing the microscope stage-top incubator to stabilize to 37 °C, 5% CO₂, and 90% humidity.
      NOTE: The live-cell system controller shows the conditions in the stage-top incubator and will sound a warning when the conditions are sub-optimal. Since the stage-top incubator lid is different from the commercial one due to the need to accommodate device tubings that extend outward, the system needs recalibration to ensure good live-cell conditions for long-time experiments (a few days up to a week). Calibration can be done based on the user's manual, and the positions of the sensors need to be maintained the same in experiments as in the calibration conditions. Check the evaporation rate of medium in the stage-top incubator, and pH conditions with phenol red-infused medium.
   2. Mount the setup onto the microscope, with the electrode chambers on the sides. Ensure that the microscope electronic stage has sufficient space to move and not bump into the electrode chambers. Ensure the medium surface heights are equilibrated and at the same height between those in the devices and the electrode chambers.
      NOTE: The medium surface in the device wells, where the tissue region is in (Figure 1C), has a significant impact on bright-field and phase contrast imaging. Meniscus will cause unwanted light focusing. Ensure the medium surface is flat at the brim of the well, but note that medium evaporation in the live-cell chamber will lead to the distortion of this flat surface after some time. To overcome this problem, one solution is to place a flat glass coverslip in contact with the medium surface to flatten this surface. This coverslip should have a smaller area than the well so it does not obstruct gas exchange, which is important for cell growth. Another option is to layer cell-culture grade mineral oil on the medium.
   3. Set up software.
      1. Choose 10x or 20x objective lenses for whole-tissue imaging with an epifluorescence microscope, and 30x objective lenses (WD 0.8 mm, NA 1.05) for higher resolution confocal imaging.
      2. For whole-tissue imaging, choose multiple positions to cover the rectangular area in the tissue region (Figure 1A, e.g., 5 rows by 5 columns, the middle row covering the slit, while the others outside of the slit). Choose the fluorescent and bright-field channels according to the cell lines used in the result part (channel 488 for green fluorescence and channel 561 for red fluorescence). Choose 1 h time interval to reduce phototoxicity in this long-term imaging experiment for up to a few days.
         NOTE: Other objective lenses, such as 4x, can be chosen if a larger view of the whole tissue is needed.
3. Electric stimulation
   1. Check the number of devices, relative device resistance, and target electric current of each device. Decide the total electric current to be sourced for a few hours up to a week in the overall setup.
   2. Source an electric signal under the current-clamp mode. For 3 devices with the same channel resistance, with one control group (no electric current, tubing clip fastened), one apical-to-basal (AtB) current condition, and one basal-to-apical (BtA) current condition (Figure 1D), supply a total current of 20 µA with a source-meter. This leads to 10 µA supplied for each device with an estimated current density ~10 µA/cm² that is physiologically relevant (more details in discussion).
      NOTE: The total voltage is around tens of millivolts, depending on the length of the Tygon tubing. Such an electric current can be supplied continuously for a few days up to a week. The phenol red indicator will change color in the electrode chambers (more acidic in the chamber with the positive electrode), but not in the stage-top incubator, which is well calibrated. The cells will remain healthy as long as the live-cell conditions are well maintained. Researchers can change the medium in the stage-top incubator once every other day, and the medium in the electrode chambers once every few days. Fasten tubing clips when switching the electrode chambers to avoid overflow of medium. As mentioned in step 3.1.1, remember to have an empty device to cover the holder space. This ensures the live-cell chamber is in the proper condition.

Results

All results presented here are adapted from the previous publication⁴; for more details, refer to this publication.

This protocol allows the application of an external ion current or electric field perpendicular to the plane of cells or tissues. This electric stimulation perturbs the endogenous transepithelial electric potential difference (TEPD) and allows one to study the role of TEPD in governing tissue behavior. Following the protocol, wild-type and genetically edit...

Discussion

This protocol provides detailed steps and design logic to fabricate a novel microfluidic device to perform transepithelial electric potential perturbation experiments and to study bioelectric effects through live-cell imaging and mechanobiology methods. The significance of this method is obvious when compared to other related methods. For example, one related method uses a simple microfluidic chip with a single channel design that applies a stationary in-plane current to study galvanotaxis (cell migration as a function o...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1x DPBS	Gibco	14190144
3-(trimethoxysilyl)propyl methacrylate, TPM	Sigma	2530850	Store under nitrogen after opening.
50 mL Falcon tube	Corning	430829	Punch 4 holes in the 50 mL falcon tube cap, to enable electrode and tubing pass through the cap and immersed in the medium. For a set of microfluidic device, at least 2 caps is needed.
Acetic acid	Sinopharm	10000208
Acrylamide solution, 40%	Sigma	79061	Toxic
Air pump	OKO lab	OKO-AP	Component of microscope stage-top incubator
Ammonium Persulfate, APS	Aladdin	A112447
Cartridge	-	-	Cartridge is custom-made, and the design refer to supplementary material. Raw material is PC Makrolon 2858 Medical Grade,and here you can find more details: https://solutions.covestro.com/zh/products/makrolon/2858_000000000000508397?SelectedCountry=US
Clips	-	-	Robert Clip, for tubing  of outer 4.5 mm diameter. https://item.taobao.com/item.htm?id=18717491819&pisk=gAsj0ailDjcjqv-tCE2rVYReT5-1h8rFWA9OKOnqBnKv6A1RBjpxDchRepvBWczD0Ft1wQS2gt-v6c1NQi5NBGJOfnjPucR4ih6OThNUTkrFntYBXWPETzI4tHpikchwH3nJbds8Mz2dntxMXYlT8aBmCOO1qfp968tJCdc9XKnte89wBdK9MFh-ypdJXhdvkYpJBpn9HCnYF3pDKKhv6qK-edJXXCC9X86JZdt96bH_3dwJgtw2FbzzkyxCHQitXiEMlB_i7ci1TKLfXtd5zaSWhEOpWac1uiCPBiSDrzgJjT7C1NCLaqRARp1W8ZNIc61DB6tV4jeJXFQP8E_7McOP0dTpXUMtXTtPPFsWvkhWEaBVWifjCcpc0M86tUwtjF-RYeQdGA2henpOsejatmAAB9jh8HZjtK_Rd3syxDRBL50sFem6FBy7FV0GhSGPYc782SYvEK__F8GxLEpkFBy7FV0MkLvf18wSMv5..&spm=a21xtw.29978518.0.0&skuId=3344063585144
Collagen I	Corning	354236
DI Milli-Q water	Milli-Q	-
Dulbecco's Modified Eagle's Medium culture medium,DMEM (high glucose)	Gibco	11965092	DMEM++ is DMEM with additional 10%  FBS (Sigma-Aldrich), and 100 units/mL Penicillin-Streptomycin (Gibco).
Enclosure black cage	OKO lab	H201	Component of microscope stage-top incubator
Epi-Fluoresence Inverted Microscope	Olympus	IX83
Ethanol absolute	Sinopharm	10009218
Fetal Bovine Serum	Sigma	F8318
Fluorescent beads, 0.1 μL, red (580/605)	Thermo	F8801
Gas chamber	OKO lab	H201-PRIOR-SP200/400/600	Component of microscope stage-top incubator
Gas Manual Mixer	OKO lab	2GF-MIXER	Component of microscope stage-top incubator
Geneticin	Gibco	10131027	GFP–actin and H1–GFP MDCK were maintained using media supplemented with 0.5 mg/mL geneticin (Gibco) to sustain their gene expression
HEPES	Sigma	736459
Isopropanol	Sinopharm	80109218
Kimwipes	Kimberly-Clark Professional	LH-70155	Laboratory wiper
Longer screw	-	-	Nickel-plated round head cross precision machine screw, M3 x 10 mm. https://item.taobao.com/item.htm?id=604899872494&pisk=gbX7s64OI40STNJKAQqqhx58y8d9NoyapDtdjMHrvLpJOvIP5HkeT7xIdEsTqUP3Zw1fJZAyyeRedH_wogkUqgrQdMI9yp5yzHQVRZcPz98FuJsG5QkPv9oliNStUTPkLv9kKpUa7RyZqgvHpfPM7c0okhx_LYdKpLvx7Uiz6RyNqiiy2lSYQ9Sp5stS9vQpwIpvqEhKe3QpMKKBkYKKeH3YcEYv24HKwxnvvh0K9wKdDnKkV03KeBKYHhYvJppdJiEXYELK1OzXm7TRVu-iUlqUdgsJlvHdeJAv5LhELvJw2Q6d2EsV0ttWNFRjAbyCeZSdUTA09X9h01_C9NUqaUC6fZONNPM55axdAHQ7IcTAFM6XZsoZBisfv1Rf3leMFLLRTLjEfzSfFaSlhMoKML9PGC6WBR0J8sS1dH63--_1vGfpG9g54UMw5pb-OMiidnTacoGntEWrjhPgtYw2wnxVQoZjPXAJmn1UcoGntQKDm2rbc4Gh.&spm=tbpc.boughtlist.suborder_itemtitle.1.494a2e8d0lg31n
Microfluidic connector	-	-	Polypropylene Straight Union Fitting, 1/16"-10-32UNF. https://item.taobao.com/item.htm?abbucket=16&detail_redpacket_pop=true&id=555839592609<k2=1752310108435i7ldxj7ekdgu9qhnphlmj&ns=1&priceTId=2150470617523101021033708e1a4b&query=10-32unf%E8%9E%BA%E7%BA%B9%E7%9B%B4%E9%80%9A%E6%8E%A5%E5%A4%B4%20%E5%A1%91%E6%96%99%E5%A4%96%E8%9E%BA%E7%BA%B9%E6%8E%A5%E5%A4%B4&skuId=3600497427939&spm=a21n57.1.hoverItem.1&utparam=%7B%22aplus_abtest%22%3A%2245243a364caf5527e8392ded0656451a%22%7D&xxc=taobaoSearch
Multiwell plates holder	OKO lab	H201-MW-HOLDER-NZ500	Component of microscope stage-top incubator
N',N'-Methylenebisacrylamide, 2%	Sigma	110269	Toxic
NOA 73	Norland Products	17-345	UV-curable adhesive
Penicillin-Streptomycin, PS (10,000 U/mL)	Gibco	15140122
Petri dish (150 mm ´ 25 mm)	Corning	430599	Resist to at least 80 °C for 2 h
Pico Plasma tool	Diener Electronic GmbH + Co. KG	Pico Plasma	For O2 plasma treatment
Plasma cleaner	Harrick Plasma	PDC-002	For air plasma treatment
Plastic dish (diameter 60 mm, height 15 mm)	Nest	705001
Platinum electrodes	Tianjin Aida Hengsheng	Pt005	Diameter 0.5 mm, length 37 mm
Rain-X Original Glass Water Repellent	ITW Chemical Products	800002242
Rectangle coverslip	Marienfeld	0107222	1.5H, 24 ´ 50 mm2, 170 μm thickness
Round glass coverslip	Marienfeld	0111500	1.5H, round 10 mm diameter
Shorter screw	-	-	Nickel-plated round head cross precision machine screw, M3 x 4 mm. https://item.taobao.com/item.htm?id=604899872494&pisk=gbX7s64OI40STNJKAQqqhx58y8d9NoyapDtdjMHrvLpJOvIP5HkeT7xIdEsTqUP3Zw1fJZAyyeRedH_wogkUqgrQdMI9yp5yzHQVRZcPz98FuJsG5QkPv9oliNStUTPkLv9kKpUa7RyZqgvHpfPM7c0okhx_LYdKpLvx7Uiz6RyNqiiy2lSYQ9Sp5stS9vQpwIpvqEhKe3QpMKKBkYKKeH3YcEYv24HKwxnvvh0K9wKdDnKkV03KeBKYHhYvJppdJiEXYELK1OzXm7TRVu-iUlqUdgsJlvHdeJAv5LhELvJw2Q6d2EsV0ttWNFRjAbyCeZSdUTA09X9h01_C9NUqaUC6fZONNPM55axdAHQ7IcTAFM6XZsoZBisfv1Rf3leMFLLRTLjEfzSfFaSlhMoKML9PGC6WBR0J8sS1dH63--_1vGfpG9g54UMw5pb-OMiidnTacoGntEWrjhPgtYw2wnxVQoZjPXAJmn1UcoGntQKDm2rbc4Gh.&spm=tbpc.boughtlist.suborder_itemtitle.1.494a2e8d0lg31n
Sulfo-SANPAH, no weigh	Thermo	A35395	SS stock solution: 20 mg/mL in anhydrous DMSO, 2 μL for each; SS working solution: 0.5 mg/mL SS in HEPES (0.1 M, pH 7.4).
SYLGARD 184 Silicone Elastomer Kit	Dow Corning	H052KCM147	The final product is polydimethylsiloxane, PDMS. This Kit includes base PDMS resin and crosslinking agent.
Temperature Unit	OKO lab	H201-T-UNIT-BL	Component of microscope stage-top incubator
Tetramethylethylenediamine, TEMED	Sigma	110189	Toxic
Touch Screen Display	OKO lab	OKO-TOUCH	Component of microscope stage-top incubator
Tridecafluoro-1,1,2,2-tetrahydrooctyl-1-trichlorosilane	Sigma	78560459
Trypsin-EDTA (0.25%)	gibco	25200072
Tygon tubings	Tygon	AJK00002	Outer diameter (OD) = 3.2mm, Inner diameter (ID) = 1.6 mm
Ultrasonic cleaner	Supmile	KD-300DE
UV LED curing machine	GHUV	GHS-MFLG	UV-LED light source, Emision range: 363–370 nm, maxsium power density: 230 mW/cm2
Vibration Free Humidifying Module	OKO lab	HM-VF	Component of microscope stage-top incubator
Visiview	Visitron Systems GmbH	-	Imaging softer ware
Wafer-layer1	National University of Singapore, Mechanobiology institute microfabrication facility	-	The wafer is custom-made, and the design refer to supplementary material.
Wafer-layer2	National University of Singapore, Mechanobiology institute microfabrication facility	-	The wafer is custom-made, and the design refer to supplementary material.