A Microfluidics Approach for the Functional Investigation of Signaling Oscillations Governing Somitogenesis

The research presented here has been approved by the EMBL ethics committee¹⁰ and the Dutch committee for animal research (dierenexperimentencommissie, DEC), and follows the Hubrecht guidelines for animal care.

Wear gloves while working with liquid PDMS. Cover surfaces and equipment. Remove any spills immediately, as cleaning becomes difficult once it's hardened.

1. Generation of the chip

NOTE: Microfluidic chips are generated by casting PDMS (Polydimethylsiloxane) in a mold. Molds can be designed using CAD software (e.g., uFlow or 3DµF³⁰). A repository of free designs is provided by the MIT (Metafluidics.org). Molds are either printed with a 3D printer or can be generated by soft lithography using a photomask that contains the desired design (for further information see, e.g., Qin et al., 2010³¹). Here, a design for a mold is provided that can be printed with a 3D printer for the on-chip culture of mouse embryo tissue (Figure 1B,C, Supplementary Files 1 and 2). To allow printing with lower resolution 3D printers, the design has been modified compared to the previously published one¹⁰. A mold without air bubble traps and one with air bubble traps is provided (Supplementary Files 1 and 2, respectively). As the procedure for printing is dependent on the printer available, the details for printing will not be described. However, one must make sure that all unpolymerized resin is entirely removed. It is often required to perform extra washing with solvents to remove unpolymerized resin from the small <200 µm holes within the microfluidic chamber. These holes will form PDMS pillars to trap the embryonic tissue within the chip during the experiment. PDMS is generally used for microfluidics, as it is cheap, biocompatible, and transparent, and has low autofluorescence. After curing, the PDMS chip is cut out of the mold and bonded onto a glass slide. Plasma treatment of both the glass and the PDMS chip activates the surfaces and allows the formation of covalent bonds, when brought into contact.

1. Prepare the required amount of PDMS by mixing the monomer with the catalyst in a 9:1 ratio (w:w) to induce polymerization. Use disposable tools to mix, such as plastic cups and forks. Ensure that the mixing is properly achieved.
2. Place the PDMS mixture in a desiccator and apply vacuum for approximately 30 min to remove air from the PDMS mixture. Due to the vacuum within the desiccator, air will be removed from the PDMS mixture.
   NOTE: Cover the inside of the desiccator with tissues in case PDMS flows over.
3. Pour the PDMS mixture into the chip mold (a PDMS layer of approximately 3-5 mm is enough).
4. Place the mold filled with PDMS back into the desiccator and apply vacuum to remove any remaining bubbles. Make sure that air is removed from all smaller structures of the mold.
5. Cure the mold by placing it in a 65 °C (or lower depending on the mold material) oven overnight. Cut the chip out of the mold using a scalpel. Cut the hardened PDMS out of the mold with sufficient space (1-2 cm) around the design to allow for later bonding. Make sure to cut the PDMS completely to prevent breaking of the chip when taking it out. Lift the PDMS chip off carefully, first with the blunt end of the scalpel, and when possible, with hands.
6. Punch inlet and outlet holes, starting from the inside of the microfluidic chamber using a 1 mm biopsy punch.
7. Clean the PDMS chip briefly with compressed air and stick adhesive tape on both sides to remove any stuck bits of PDMS and dust and keep it clean until further use.
   NOTE: The chip can be kept like this until further use. 
8. Clean the glass slide with compressed air. Be careful that it does not break. Remove adhesive tape from the PDMS chip.
9. Bond the chip to the glass slide using a plasma oven. The following instructions are optimized for air plasma.
   NOTE: Consult the manual of the lab's plasma oven and optimize the procedure for the specific experiment.
   1. Place the chip and glass slide into the plasma oven with the sides to be bonded facing up.
   2. Place the lid onto the plasma oven and hold it in place while applying vacuum and wait until vacuum has established.
   3. Turn the gas on by pressing the Gas button and wait for approximately 1 min (pressure should be at around 0.37 mbar).
   4. Generate plasma by pressing Generator (Power: 9.0, Time: 1 min).
   5. When finished, turn the pump and gas off; ventilate and open the door.
   6. Bond the chip to the glass by placing the activated surfaces onto each other and applying pressure evenly. Be careful that the glass does not break.
      NOTE: A water test can be performed to confirm that plasma treatment has worked. Place a drop of water on the glass surface. If plasma treatment has worked, the water should spread out immediately instead of forming a droplet.
10. Place the bonded chip in an oven at 65 °C for 5 min.
11. Seal the exposed side of the chip with adhesive tape to prevent dust from entering the inlets.
    ​NOTE: The chip can be kept until use. Openings should be closed with tape until then.

2. Preparing microfluidics experiment

NOTE: The following preparatory steps are necessary to perform the microfluidics experiment: First, when performing the microfluidics experiment, a fluid flow is created from the inlets to the outlets. Any holes in the chip will function as outlets due to the pressure build-up from the inlets. Therefore, all holes that are not used must be closed; for instance, holes that are used to load tissue onto the chip. If these are not closed, the tissue might flow out with the fluid. To close these holes, PDMS-filled tubing is used. The PDMS-filled tubing can be kept indefinitely and, therefore, will not need to be prepared for each microfluidics experiment separately but can be made in larger batches. Second, before the start of the microfluidic experiment, tubing, PDMS-filled tubing, and the chip, required for the experiment, are prepared and UV-sterilized. Finally, the chip has to be coated with fibronectin, so that the embryonic tissue can attach to the glass²⁵.

1. Making PDMS-filled tubing.
   1. Take ±1 m of tubing. More tubing can be filled with PDMS by taking multiple pieces of tubing.
   2. Prepare the required amount of PDMS by mixing the monomer with the catalyst in a 9:1 ratio (w:w) to induce polymerization.
      NOTE: Use disposable tools to mix, such as plastic cups and forks. Mix properly.
   3. Place the PDMS mixture in a desiccator and apply vacuum for approximately 30 min to remove air from the PDMS mixture.
      NOTE: Cover the inside of the desiccator with tissues in case PDMS flows over.
   4. Fill a 3 mL syringe with PDMS. Fill carefully to prevent the formation of air bubbles in the mixture.
   5. Use forceps to insert the tip of a 22 G needle into the tubing.
      NOTE: Be careful not to puncture the tubing or your fingers with the needle.
   6. Attach the tubing to the PDMS-filled syringe via the needle. Use the syringe pump to fill the tubing, with a flow rate of approximately 500 µL/h. Be careful not to apply too high pressure to prevent breaking of the pump.
   7. Once the tubing is entirely filled with PDMS, place it in a 15 cm dish and cure at room temperature (at least overnight).
2. Prepare the tubing and chip for the experiment.
   NOTE: The following tubing is required for the experiment: first, approximately 50 cm of tubing per outlet and second, approximately 2-3 m per inlet (the exact size depends on the spatial organization of the pumps, incubator or microscope used later on). The inlet tubing needs to be long to allow the culture medium to equilibrate with CO₂ for embryo culture during the experiment.
   1. Cut the tubing at a 45° angle so that pointed ends are created; this will make it easier to insert the tubing into the holes of the chip. Attach one needle to each of the inlet tubing. See step 2.1.5.
   2. Cut PDMS-filled tubing into ±1 cm plugs. Cut at a 45° angle so that pointed ends are created; this will make it easier to insert the tubing into the holes of the chip. Enough plugs are needed to fill each hole that is not attached to any tubing.
   3. Remove the adhesive tape from the chip.
   4. Place all the tubing with needles attached, the chip, and the plugs in a dish and sterilize by exposing to UV light for ±15 min. Any cell culture hood with the possibility for UV sterilization can be used.
3. Coating of chip with fibronectin.
   NOTE: For culture of 2D ex vivo cultures of posterior PSM, coat the chip with fibronectin.
   1. Place the chip in a beaker containing PBS + 1% Penicillin/Streptomycin at room temperature. Make sure the chip is completely covered with PBS. Flush the chip with PBS to remove air with a P200 pipette.
      NOTE: All further handling of the chip will be done in this beaker until the start of the experiment to prevent entrance of air into the chip. Be careful not to break the glass slide of the chip.
   2. Prepare 2 mL of 1:20 Fibronectin in PBS and fill into a beaker.
   3. Load a 3 mL syringe for each chamber of the chip.With forceps, insert a 22 G needle into the outlet tubing. Attach the needle to the syringe containing fibronectin.
   4. Connect the syringe to the syringe pump. Flush the tubing, until there is no air left in the tubing.
   5. Attach the outlet tubing to the outlet of the chip. Make sure to push the tubing all the way to the bottom. Set a low flow rate to coat the chip. All other openings can remain open. Let the syringe pump run for at least 2 h, can also run overnight.
   6. Stop the pump. Cut off the tubing right after the needle. The remaining tubing will serve as the outlet during the experiment later.

3. Microfluidics experiment

NOTE: Here, a protocol for the entrainment of the mouse segmentation clock in 2D ex vivo cultures by applying pulses of the Notch signaling inhibitor DAPT is presented. Applying pulses with a period of 130 min, close to the natural period of the mouse segmentation clock (137 min²⁵), allows efficient entrainment. Pulses of 100 min of medium and 30 min of 2 µM DAPT are applied (Figure 2A). The presence of drug within the chip is monitored using a fluorescent dye. The excitation and emission spectra of this dye and the fluorescent signaling reporter must be different enough to prevent bleed-through during fluorescence real-time imaging. When yellow fluorescent proteins are used as signaling reporter, such as the Notch signaling reporter LuVeLu⁵ (Lunatic fringe-Venus-Lunatic fringe), the dye, Cascade Blue, can be applied. To identify other possible combinations of fluorescent dye and reporter, freely available spectra viewer can be used. The effect of entrainment can be detected by real-time imaging of dynamic signaling reporters^5,10,32. Each chip has two incubation chambers. This allows the direct comparison of drug pulses (DAPT) to control pulses (DMSO).

1. Make medium and degas.
   1. On the day of the experiment, prepare 50 mL of culture medium (DMEM-F12 supplemented with 0.5 mM glucose, 2 mM glutamine, and 1% BSA, for further information of mouse tailbud culture see²⁵).
   2. Prepare syringes filled with the medium for the experiment. Prepare culture medium in beakers: To entrain Notch signaling oscillations, prepare 6 mL of medium with 1.2 µL of DMSO + 10 µM Cascade Blue; 6 mL of culture medium with 2 µM DAPT + 10 µM Cascade Blue; two tubes with 6 mL of medium each. Use at least 6 mL of medium per syringe. Load syringes with the medium. Make sure to label the syringes containing drug and DMSO.
   3. Degas the chip within PBS and the syringes containing medium in a desiccator. Place the syringes into a beaker with the tip facing up, so that the air can escape at the top. It might occur that the chip floats and is still filled with bubbles. These will be removed subsequently.
   4. Try to flush out/suck up most of the air bubbles from the chip using a P200 pipette. If some air remains, this will be removed during the following experiment.
   5. Install syringes in the pumps and attach inlet tubing to the syringes. To entrain Notch signaling, use two syringe pumps, one carrying the medium syringes, one carrying the drug/DMSO syringes. Let this run at a higher flow rate (0.5 mL/h) until the start of the experiment to remove air bubbles from the tubing. Be careful to set the syringe details (diameter) properly in the pump.
2. Loading of tissue onto the chip.
   1. Dissect mouse embryo tissue. Dissect the most posterior tip of the tail (tailbud). Place the dissected tissue into culture medium + 25 µM HEPES.
   2. Flush the chip with culture medium + 25 µM HEPES using a P200 to remove PBS.
   3. Load the tissue into the chip using a P200 pipette (Figure 1C). Make sure not to introduce air bubbles into the chip.
      NOTE: Efficient loading requires some practice. If the tissue is not oriented properly, it might be possible to turn it by carefully sucking out and flushing in again. However, this often results in quick cell death.
   4. After each tissue-loading step, close the corresponding tissue-loading inlet using a piece of PDMS-filled tubing. Make sure the plugs are sufficiently pushed down to the bottom of the chip using blunt tweezers.
   5. After all samples have been loaded, close any unused inlets/outlets with pieces of PDMS-filled tubing using blunt forceps.
3. Assembly of microfluidic setup.
   1. Lower the flow rate of the microfluidic pumps. 60-100 µL/h works well for mouse embryo tailbuds. At higher flow rates, cell death was observed – presumably due to too high cell shearing. The cumulative flow rate of multiple pumps should not exceed these 60-100 µL/h per microfluidic chamber at a given time.
   2. Attach tubing to the microfluidic chip. Do this without getting air bubbles inside the chip (this can be achieved by ensuring that there is a drop of medium present at the end of the tubing). Outlets are still present from the fibronectin coating (see 2.3).
   3. Once all tubing is attached to the chip, take it out of the beaker, dry it properly, place it in a dish, and put this together with approximately 1.5 m of inlet tubing in an incubator (37 °C, 20% O₂, 5% CO₂) for overnight culture. The tubing is gas permeable; this will allow equilibration of O₂ and CO₂ in the medium. Alternatively, for simultaneous fluorescence real-time imaging, the chip and approximately 1.5 m inlet tubing are placed into the microscope directly. A holder for the chip, which fits into standard 96-well-plate holders, is provided in Supplementary File 3.
      ​NOTE: Make sure the humidity is high enough during incubation. If necessary, add a moist tissue to the imaging chamber or incubator to prevent evaporation in the microfluidics set-up. If humidity in the microscope incubator is too low, this results in air bubble formation in the tubing and the microfluidic chip. A closed imaging box, in which chip and tubing are placed, can then be used. The simplest way of making such an imaging box is by placing a big lid over the chip and adding a wet tissue, it does not have to be closed off entirely. Make sure that the height difference between pump and chip is not too big and, if necessary, change heights slowly to prevent the formation of air bubbles due to gravity.
   4. Let all pumps flow at a flow rate of 20 µL/h for 20 min, after the setup has been placed to its final location. Because the chamber and pump have most likely moved in height, this is necessary to re-establish the correct pressure in the tubing.
   5. Turn the drug pump off, let only the medium pump run at 60 µL/h until the start of experiment/real-time imaging.
4. Start of the experiment.
   1. Start the planned pumping program for the experiment. To entrain Notch signaling, use a pumping program of 100 min medium and 30 min drug pulses, which is repeated until the end of the experiment, typically for 24 h.
      NOTE: Any programmable microfluidic pump can be used. In the simplest format, the pumps can be programmed and started manually. Alternatively, the pumps can be controlled by a computer software.
   2. In case real-time imaging is performed, start imaging after a period of at least 30 min. This will allow the temperature of the chip to adjust to the temperature within the incubator and prevent a too strong drift during imaging. Autofocus is still beneficial.
   3. Perform standard confocal imaging via the glass slide of the chip using an inverted microscope. Use an imaging interval of 10 min to sufficiently detect signaling oscillations with a period of 130 min.
   4. For shorter periods, shorten the interval for sufficient sampling. Include a low-resolution imaging track for detection of Cascade Blue to visualize the presence of drug on the chip.
   5. For excitation of a Venus reporter, use a 515 nm laser or a 2-photon laser at a wavelength of 960 nm.
   6. Acquire a z stack of 6-8 planes with a distance of 8 µm through a 20x plan objective (resolution 512 x 512 pixels, 1.38 µm/pixel). Use a motorized stage to image multiple samples within one experiment.
   7. Excite Cascade Blue with a 405 nm laser and acquire a single z plane every 10 min (resolution 32 x 32 pixel, 22.14 µm/pixel).
      NOTE: Further information on imaging and image analysis are provided in the Representative Results and can be found in reference¹⁰.