Axonal Transport of Organelles in Motor Neuron Cultures using Microfluidic Chambers System

Axonal transport is vital for neuronal health and viability. Our protocol describes an elegant method for monitoring and analyzing axonal transport. And can be adapted for various neuronal disease models.

The use of microfluidic chambers enable precise spatial temporal control, which is crucial for studying axonal biology. Axonal transport abnormalities are implicated with several neurodegenerative disease such as ALS. The microfluidic platform enables study of basic mechanisms as well as testing therapies to repair axonal transport defects.

The microfluidic chamber platform can be easily adapted to suit different aspects of axonal research, including axonal growth and degeneration of various neural subtypes. This method is complex and usually taught hands-on. The visual demonstration will increase the accessibility of this procedure to any scientist interested in studying axonal transport.

Begin by casting PDMS in primary molds. Use pressurized air to remove any dirt from the wafer platform, making sure that the wafers look smooth and clear before proceeding. Next, fill a container with liquid nitrogen and prepare a 10 milliliter syringe with a 23 gauge needle.

In a chemical fume hood, place the wafer containing plate in a sealable container, and fill the syringe with two milliliters of liquid nitrogen per each wafer containing plate. Open a chlorotrimethylsilane bottle. Pierce the rubber cap with the syringe and inject the nitrogen into the bottle.

Without pulling out the needle, Turn the bottle upside down and drawback two milliliters of chlorotrimethylsilane for each wafer containing plate. Spread the chlorotrimethylsilane in the container, but not directly on the wafer containing plate. Close the container.

And incubate it for five minutes per wafer. Next weigh 47.05 grams of PDMS base in a 50 milliliter tube, and add PDMS curing agent at a ratio of 16 to 1 base to curing agent for a total of 50 grams. Mix the PDMS for 10 minutes on a low speed rotator, then pour it into each wafer containing plate to the desired height.

Place the plates inside of a vacuum desiccator for two hours. Which will remove air trapped within the PDMS, and form a clear uniform mold. Afterwards incubate the plates in an oven for three hours or overnight at 70 Celsius.

To create the microfluidic chamber or MFC. Cut and remove the PDMS molds from the plate with a scalpel, making sure not to force the molds because they're fragile. Follow the instructions in the text manuscript to punch and cut the chambers, depending on the experimental setup.

For spinal cord explant culture, punch two seven millimeter wells in the distal side of a large MFC, making sure that they will overlap with the channel edges. On the proximal side, punch one seven millimeter well in the middle of the channel with minimal overlap, so that sufficient space is left for the explants. Punch two additional one millimeter holes in the two edges of the proximal channel.

And cut the PDMS into single chambers. Then turn the MFC facing upwards, and use a 20 gauge needle to carve at three small explant caves on the punched seven millimeter well. For dissociated motor neuron culture, punch four six millimeter wells, in the edges of the two channels of a small MFC and cut the PDMS into single chambers.

To sterilize the MFC, spread 50 centimeter long sticky tape bands on the bench. Then press both faces of the chamber on the tape and pull it back. Place the clean chambers in a new plate and incubate them in analytical grade, of 70%ethanol for 10 minutes on an orbital shaker.

Dispose of the ethanol and dry the chambers in a tissue culture hood or in an oven at 70 degrees Celsius. Then place the chamber in the center of a tissue culture grade glass bottom ditch, and apply minor force on the edges to bind the PDMS to the dish bottom. Incubate the dish for 10 minutes at 70 degrees Celsius.

Then press on the chambers to strengthen the adherence. Incubate the dish under UV for 10 minutes, then proceed to coding the MFC. Add 1.5 nanograms per milliliter PLO to both compartments.

Making sure that the PLO is running through the channels. By pipetting the coating media directly in the channel entrance. Examine the MFC under a light microscope, to check for bubbles.

If air bubbles are blocking the micro grooves, place the MFC in a vacuum desiccators for two minutes. Remove excess air from the MFC channels. Then incubate it overnight with PLO.

Replace the PLO with laminin and incubate the MFC for an additional night. Prior to plating, wash the laminin with culture medium. Dissect an ICR E12.5 pregnant mouse, and separate the embryos from the amniotic sac.

Put the embryo in HBSS Silicon dish. Remove the head and tail and flip it onto his belly. Gently peel off superficial skin from the embryos back, exposing the spinal cord.

Separate the spinal cord from the body from head to tail, using gentle tweezers. Remove the meninges, then remove the dorsal horns and cut the spinal cord into one millimeter thick transverse sections. Dispose of all medium from the proximal compartment of the MFC.

Pick up a single spinal cord explant with a pipette, in a total volume of four microliters. And inject it as close as possible to the cave. Draw out any excess liquid from the proximal well via the lateral outlets.

Repeat the previous step two more times and make sure that the explants are embedded in the proximal channel. Slowly add 150 microliters of SCEX medium, to the proximal well. One day after plating.

Replaced the SCEX medium in the proximal compartment and add rich SCEX medium to the distal compartment. Maintaining a volume gradient of at least 15 microliters per well between the distal and proximal wells. After four to six days, the axons should cross to the distal compartment and be ready for axonal transport imaging.

To label the mitochondria and acidic compartments, prepare fresh SCEX medium with 100 nanomolar MitoTracker Deep Red FM, and 100 nanomolar LysoTracker Red. And add it to the microfluidic chambers. Incubate them for 30 to 60 minutes at 37 degrees Celsius.

Then wash three times with warm SCEX medium. Proceed with live imaging of axonal transport. Acquiring a 100 time lapse image series at three second intervals.

With a total of five minutes per movie. After seven days in microfluidic chambers. Mouse embryonic HB9 GFP spinal cord explants.

We're stained with MitoTracker Deep Red and LysoTracker Red dyes. To label the mitochondria and acidic compartments. Axons in the distal grooves were imaged and Kymograph analysis was used to determine the general movement distribution.

This revealed a bias in the retrograde direction only in acidic city compartments, but not in mitochondrial transport. The particle density was also quantified, revealing a higher number of mitochondrial particles compared to acidic compartments in the HB9 GFP spinal cord explant axons. Next, single particle transport analysis was conducted using semi-automated software followed by in-house code.

This analysis revealed that acidic components and mitochondria have similar particle velocities. But only acidic compartments display a bias towards retrograde movement. Isolating the embryonic spinal cord can be difficult at first.

Practice this procedure several times, making sure to use the correct embryonic age and the appropriate dissection tools. The use of microfluidic chambers changes the way we study axonal biology. It allows us to visualize localized axonal processes, which we're once hypothesized to occur only in the cell body.