Microfluidics-Assisted Selective Depolarization of Axonal Mitochondria

All animal experiments were performed following the relevant guidelines and regulations of the Government of Upper Bavaria. The primary neurons were prepared from E16.5 C57BL/6 wild-type mouse embryos of both sexes following standard methods as previously described⁶.

1. Assembly of the microfluidic device

1. Coat one six-well glass-bottom tissue culture plate with a final concentration of 20 µg/mL of Poly-D-Lysine and 3.4 µg/mL of Laminin in PBS (phosphate-buffered saline) (see Table of Materials).
2. Incubate the coated plate overnight in the dark at room temperature.
   NOTE: Coating can also be performed at 37 °C for 1-2 h. The choice of the coating depends on the cell type used.
3. After coating, bring the six-well plates to the sterile hood and wash them twice with sterile ddH₂O.
   NOTE: Do not wash with salt-containing buffers such as PBS, as salt crystals will interfere with seal formation.
4. Allow the plate to dry in a tilted position for 3-5 min. Remove excess water by vacuum suction or pipetting.
5. Soak the microfluidic chamber in 80% ethanol.
   NOTE: The microfluidic chamber was obtained from commercial sources (see Table of Materials).
6. Allow the microfluidic chamber to dry for 3-5 min in a tilted position. Remove excess ethanol by vacuum, suction, or pipetting.
7. When completely dry, place the microfluidic chamber in the center of the well.
   NOTE: The formation of the seal can be observed as a change in reflective properties upon exclusion of air from the interface between the plate and the silicone device. Both the plate and microfluidic chamber must be completely dry before assembly to create a good seal. However, drying time needs to be minimized as much as possible to ensure proper adherence to the cells. Therefore, the wells and chambers must be visually inspected for remaining liquid droplets. If no droplets are visible, immediately assemble the chambers.
8. Gently tap the microfluidic chamber at its borders and microgroove section in the middle for proper attachment to the glass plate.

2. Seeding and maintaining of neurons

1. Collect the desired number of dissociated hippocampal neurons per chamber in a 1.5 mL reaction tube.
   NOTE: For the present study, 1.5 x 10⁵ neurons were seeded per device for imaging-based applications.
2. Centrifuge neurons at 1000 x g for 4 min at room temperature.
3. Discard the supernatant with a pipette and resuspend the pellet in 8 µL of B27-Neurobasal media (see Table of Materials).
4. Pipette the cell solution into the channel entrance of the microfluidic chamber (b in Figure 1A).
5. Tap on the back of the plate to assist flow through the channel.
   NOTE: Tapping can be done quite forcefully without worry that the seal will break.
6. Aspirate with a pipette any remaining cell suspension at the exit of the channel (g in Figure 1A) to decrease the number of cells outside the channel.
7. Incubate the microfluidic chamber with cells for 15-20 min at 37 °C and 5% CO_2.
8. Fill the top axonal well with 50 µL of B27-Neurobasal media in (c in Figure 1A).
9. Tap on the back of the plate to assist flow through the channel.
10. Fill wells on the soma side (a and h in Figure 1A) with 150 µL of B27-Neurobasal media each, creating a tension bubble on top. The volume of the bubble is about 20 µL.
    NOTE: Creating a tension bubble is not necessary to achieve fluidic isolation, but it provides a clear visual of the higher volume on the soma side.
11. Fill both wells of the axonal side (c and i in Figure 1A) with 100 µL of B27-Neurobasal media each.
    NOTE: To promote axonal growth through the microgrooves, it is recommended that the wells on the soma side contain more media than the wells on the axon side. However, axons will by chance grow through the microgrooves even without volume differences.
12. Incubate the microfluidic chamber at 37 °C and 5% CO₂ for 7-8 days.
    NOTE: Growth of axons into the axonal chamber can usually be observed starting at day in vitro (DIV) 5. Longer culturing times are possible if more mature cultures are desired.
13. Feed neurons every 2-3 days by removing the medium from the two upper wells (a and c) and replacing it with fresh B27-Neurobasal medium until a tension bubble is formed.

3. Staining with mitochondrial membrane potential sensitive dye

1. At DIV 7-8, assess that the axons have grown through the microgrooves and extend into the axonal compartment underneath a light microscope.
   NOTE: Different DIV stages are also possible depending on the desired age.
2. Perform two washes with pre-warmed imaging medium (37 °C) by removing the B27-Neurobasal medium from all wells and adding about 100 µL of the imaging medium to the top wells of both the axonal and soma channels (a and c). Let the medium flow through to the lower wells (h and i).
   1. Remove the medium from the lower wells and any leftover medium in the top wells, and repeat with fresh medium, leaving the wells empty at the end of the wash.
      NOTE: Due to the high capillary force in the channels (d and f), there will be a remaining wash medium in the channels that should not be removed to prevent the neurons from drying. Any phenol-red-free imaging medium can be used here, for instance, Hibernate E (see Table of Materials). If the microscope is not set up with a CO₂ supply, ensure that the imaging medium uses an alternative buffering system to carbonate/CO₂ (e.g., HEPES)⁸.
3. Dilute 1 mM tetramethylrhodamine ethyl ester (TMRE, red, see Table of Materials) stock to 5 nM in the imaging medium.
   NOTE: Do not exceed 1% DMSO in the final dilution to avoid toxicity.
4. Add 100 µL of the 5 nM TMRE dilution to both the somatic and axonal top wells (a and c), and let the medium flow through both the somatic and axonal compartments until there is an equal volume in all wells. Fill up with the TMRE-containing medium.
5. Place the neurons back into a controlled environment chamber (37 °C and 5% CO₂) for 25 min.
6. Perform two washes with the pre-warmed imaging medium (37 °C) as described before (step 3.2). On the last wash, fill up with 100 µL in each well and create a tension bubble of the medium on the somatic wells (a and c).

4. Live-cell imaging

NOTE: Still images shown were acquired on a spinning disk confocal microscope, using a 40x NA 1.25 immersion objective (see Table of Materials). 200 ms exposure time and 10% laser power for the red channel and 500 ms exposure time for brightfield were chosen. However, regular confocal or widefield inverted microscopes can also be used to study TMRE intensity.

1. Image the cells with an inverted microscope.
2. Ensure that the microscope is equipped with a stage incubator to maintain the neuronal culture at 37 °C throughout the experiment.
3. Select a region of interest in the somatic compartment, and follow it through the microgrooves into the axonal compartment. Ensure that the microgrooves imaged match those in the somatic and the axonal compartments (for easier baseline and post-treatment comparison).
4. Capture fluorescence images using excitation at 561 nm and emission at 625 nm for red fluorescence signal.
5. Acquire the images.
6. To induce mitochondrial depolarization, add 20 µM Antimycin A (see Table of Materials) in the imaging medium to the axonal compartment. To ensure a volume difference between the soma and axonal chambers, check the presence of a tension bubble on the somatic side (equals volume >110 µL).
   1. Remove all imaging medium from the lower well of the axonal side (i), except for the medium inside the channel (f). Add 160 µL of 20 µM Antimycin A to the top axonal well (c) and let it flow through the channel until there is an equal volume in both axonal wells. Volumes in axonal wells are roughly 80 µL per well.
      NOTE: Ensure that the volume in the axonal wells is smaller than in the somatic wells to ensure proper fluid pressure. A difference of at least 10 µL (10% of the well volume) is recommended, but also higher differences are possible.
7. Incubate neurons in the controlled environment chamber (37 °C) for 20-30 min.
8. Repeat imaging as described above (steps 4.3-4.5). Ensure that the same positions are imaged (easily identified by the microgrooves) as during the baseline acquisition.