Microfluidic Chip for Axonal Injury Models Construction and Enabling Multi-Omics Analysis

We have developed large-scale microfluidic chips to enable multiomics analysis, with the aim of for illustrating the metabolic mechanism of neurons following axonal injury. We mainly utilize microfluidic technology, metabolic flux analysis, and the transcriptomics to advance the research on the metabolic mechanisms of neurons during axonal injury and regeneration. We found that there are metabolic differences among cortical neurons at different developmental disease, and that young neurons undergo metabolic remodeling after external injury.

The core advantage of our protocol lies in the larger-scale microfluidic platform design and operational standards, which enable accurate multiomics and size of millions of cells. To begin, transfer the PDMS base and curing agent into a centrifuge tube. Place the centrifuge tube containing the mixture into a centrifugal stirrer mixer.

Centrifuge at 2, 000 G for four minutes twice to mix and to degas. Pour 13 to 15 grams of the degassed PDMS into the microfluidic mold, ensuring the bottom of the mold is completely covered. Now, place the mold in a vacuum desiccator, then use a vacuum pump to evacuate air for five to 10 minutes to thoroughly remove bubbles from the PDMS.

Using a rubber bulb, gently tap the surface of the PDMS to break any remaining surface bubbles. Transfer the mold to a convection oven and bake at 80 degrees Celsius for 100 minutes to cure the polydimethylsiloxane. Once the PDMS is fully cured, gently slide the tip of a scalpel under the edge of the PDMS to carefully lift and separate it from the mold.

Using a biopsy punch with a diameter of 2.0 to 2.5 millimeters, create holes in the PDMS for culture medium infusion and cell loading. Remove any surface impurities from the PDMS using adhesive tape, then place it in a clean glass dish and wrap it with aluminum foil for storage. Prior to the experiment, autoclave the microfluidic device at 121 degrees Celsius and 101 kilopascals for five minutes to ensure sterility.

Place a cover slip intended for conventional microfluidic devices in a 35 millimeter culture dish. Add two milliliters of 0.1 milligram per milliliter poly-D-lysine solution to the dish. Incubate the dish at 37 degrees Celsius for six hours or overnight.

After incubation, carefully retrieve the poly-D-lysine solution for reuse or proper disposal. Now, add two milliliters of sterile ultrapure water to the dish. Rotate it 50 times clockwise, followed by 50 times counterclockwise.

Aspirate the water using a vacuum pump connected to a sterile pipette tip, collecting waste in a liquid waste container. After all washes are complete, allow the cover slips to air dry naturally in a biosafety cabinet before use. Using sterile fine tip forceps, place the autoclaved microfluidic chip at the center of the glass surface with microchannels facing downward.

Gently press the chip onto the glass surface using a 200 microliter pipette tip to ensure complete adhesion. Next, mark the left chamber with a dot using a permanent marker to designate the soma compartment. Aspirate 10 microliters of neuronal suspension with a 10 microliter pipette, then slowly dispense the suspension into the loading port above the marked soma compartment.

Confirm proper fluid flow into the lower reservoir of the left chamber. Transfer the culture dish to a humidified incubator set at 37 degrees Celsius and 5%carbon dioxide for 20 minutes. After incubation, place the dish under a 20X microscope to confirm media profusion from the soma compartment to the axonal compartment through the microchannels.

Now, pipette 150 microliters of neuronal basal medium to the upper port of the axonal compartment. Similarly, add 150 microliters of complete neuronal medium to the upper port of the soma compartment. Next, pour 20 milliliters of ultrapure water into a 150 millimeter culture dish to create a humidity chamber.

Place the 35 millimeter culture dish inside the large dish, then transfer the entire culture system to the incubator and maintain for the required duration. Use a 10 centimeter wide Petri dish for placement of the large-scale microfluidic device. Choose either full channel or interval channel seeding based on experimental needs, as each method requires different axonal injury protocols.

After seeding is completed, add five milliliters of complete neuron culture medium to the Petri dish to maintain neuron growth. Transfer an appropriate amount of neuronal basal medium into a new 15 milliliter centrifuge tube for later use. Place a microfluidic device containing cortical neurons on a clean bench.

Using lint-free paper, dry the bottom of the 35 millimeter culture dish, then use a 200 microliter pipette to aspirate the medium from the two right side holes of the microfluidic device. Next, connect a vacuum pump tubing to a sterile, filter-free 200 microliter pipette tip. Turn on the vacuum pump at a suction rate of 60 liters per minute and aim the tip at the connection point between the lower hole of the axon terminal chamber and the chamber to aspirate the old medium, causing axon breakage due to negative pressure.

Replace the pipette tip and draw 150 microliters of neuronal basal medium, slowly adding it through the lower hole of the right chamber. After the performing addition and aspiration four times, replace with fresh complete neuronal medium, then aspirate the old medium from the cell body side hole and add fresh complete neuronal medium containing drugs if needed. Place the microfluidic device along with the culture dish in a 150 millimeter culture dish and continue culturing for the required time, such as 24 hours.

For manual axonal injury, seed neurons into all chambers of the large-scale microfluidic device. Incubate the device at 37 degrees Celsius in a humidified incubator with 5%carbon dioxide for three days. On day seven in vitro, remove the culture dish containing the microfluidic device from the incubator and place it on a sterile stage.

Prepare a microscope with 20X magnification and sterilize the work area using 75%ethanol. Hold a 10 microliter sterile filtered pipette tip and identify axon bundles in the original microchannels under microscope guidance. Perform longitudinal scratches along each axon bundle using the pipette tip.

Observe the axon breakage under the microscope in real time. The microchannels of a standard microfluidic device permit the growth of axons, but not soma, as confirmed by beta three tubulin staining at seven days in vitro. Neuronal density showed no significant difference following axonal injury, indicating no observable neuronal death.

A large-scale microfluidic device with a three-dimensional expandable design was developed for multiomics analyses. Both unpunched and perforated PDMS replicas adhered securely to 10 centimeter dishes or PC boards. Axonal damage induced by pipette tip scratching led to clearly severed axons with disrupted morphology, in contrast to intact axons prior to injury.

Axonal regeneration was observed at both six hours and 24 hours following injury, while uninjured axons remained stable. Neuronal protein concentration rose significantly from 1420.4 micrograms per milliliter at baseline to 1748.9 per milliliter at six hours, and 1823.7 micrograms per milliliter at 24 hours post-injury. RNA yields from control and axon injured groups were nearly identical.

RNA sequencing revealed 595 injury-specific genes and 609 control-specific genes, with 17, 471 genes shared between groups. KEGG pathway analysis identified significant upregulation of oxidative phosphorylation, reactive oxygen species response, autophagy, and TCA cycle pathways post-injury.