Microfluidic devices used to compartmentalize neurons have become a standard tool in neuroscience, enabling researchers to physically and chemically manipulate subcellular regions of neurons including somata, axons, dendrites and synapses. These compartmentalized culture devices can help answer numerous questions in the neuroscience field, such as, how axons extend during development to form synapses, how synapses remodel and reform following axon damage and how pathological conditions can trans-synaptically propagate in diseases such as Alzheimer's. This protocol describes the use of preassembled plastic multicompartment chip for compartmentalizing culture primary rat neurons.
The main advantage of using these chips is that they provide an easy-to-use and highly reproducible method to compartmentalize neurons. Another advantage is that these chips yield healthier, long-term growth of neurons and isolated axons than previous silicon-based compartmentalized devices and are equally compatible with high resolution microscopy. Other model systems including human stem cells are also able to be cultured within the preassembled compartmentalize chip.
To begin, place a sterile, multicompartment chip into a Petri dish. Add 100 microliters of a precoding solution to the upper-left well of the chip and allow it to flow through the main channel into the adjoining well. Next, fill the lower-left well of the chip with 100 microliters of the precoding solution.
This time, wait five minutes to allow the solution to flow through the micro-grooves. Now, add 100 microliters of the precoding solution to the upper-right well and allow it to flow through the main channel into the adjoining well. After 90 seconds, fill the lower-right well with 100 microliters of the precoding solution and incubate the chip for 10 minutes.
Carefully angle the pipette tip, away from the main channel and aspirate the solution from each well. Then, immediately add 150 microliters of PBS to the upper-left well and wait one and a half minutes. Next, add 150 microliters of PBS to the lower-left well and wait five minutes to allow the liquid to flow through the micro-grooves.
Then, add 150 microliters of PBS to the upper-right and lower-right wells, in that order. After 10 minutes, again aspirate the PBS and repeat the wash steps a second time. Following the second wash, aspirate the PBS from the wells as before by angling the pipette tip, away from the channel opening.
Then, add 100 microliters of 0.5 mix per mil Poly-D-Lysine solution, to the upper-left well of the chip. Wait one and a half minutes and then fill the lower-left well with 100 microliters of the Poly-D-Lysine solution. Repeat this process on the right half of the chip.
Close the Petri dish. Then, place the chip in an incubator at 37 degrees Celsius for one hour. After incubation, rinse the chip twice with PBS as previously described.
Next, aspirate the PBS from the device. Immediately, add 100 microliters of cell culture media to the upper-left well of the chip. After one and a half minutes, add media to the lower-left well.
Wait five minutes to allow media to flow through the micro-grooves and then repeat the filling process on the right side of the chip. Prepare a cell suspension of dissociated rat hippocampal neurons according to established protocols. Remove the majority of media in each well of the chip but leave the channels filled.
Next, immediately load five microliters of the cell suspension into the upper-right well and another five microliters of cell suspension in the lower-right well. When loading the cells, point the tip of the pipette towards the main channel. After loading the cells, transfer the chip to a microscope stage and to make sure that the neurons are in the main channel.
After waiting five minutes for the cells to attach, add approximately, 150 microliters of neuronal culture media to each of the upper and lower-right wells. Then, add 150 microliters of media to each of the upper and lower-left wells. Cover the Petri dish and place the chip in a humidified tray in a 5%carbon dioxide, 37 degrees Celsius incubator.
First, remove 20 microliters from the lower-left well of the axonal compartment and place it into the upper-right well of the somatic compartment. Wait two minutes for flow within each channel to equilibrate. Next, remove 50 microliters of media from the axonal compartment, and add 0.3 microliters of one millimolar Alexa Fluor 488 hydrazide to this media.
Pipette up and down to mix the solution and then return the media containing the fluorescent dye back into the axonal compartment. At this point, place the chip onto the microscope and image as desired. To perform axotomy within the chip, first, remove the media from the axonal compartment, keeping the pipette tip away from the entrance of the main channel.
Store the removed media in a centrifuge tube for now. Next, place the aspiration pipette near, either entrance of the main channel of the axonal compartment and aspirate the axonal compartment completely. Continue to aspirate the area for one to two minutes.
Replace the axonal compartment with the stored media. Make sure that solution is completely removed from the compartment and the axons are severed by looking at the sample under a microscope. Then, cover the chip and return it to the incubator.
To begin fluorescence immunostaining, remove most of the media from the chip, keeping the interior compartments hydrated. Immediately, add 100 microliters of fixation solution to the top wells of the axonal and somatic compartments. After one minute, add 100 microliters of fixation solution to the bottom wells and fix the cells for 30 minutes at room temperature.
Remove most of the solution from the wells and immediately, add 150 microliters of PBS to each of the top wells of the axonal and somatic compartments. Wait two minutes for the PBS to flow into the bottom wells, then, remove the PBS and rinse the wells twice more using this same process. Following the last rinse, remove most of the PBS from the wells of the chip and immediately, add 150 microliters of PBS with 0.25%triton x-100 to each of the top wells of the axonal and somatic compartments.
Wait for 15 minutes and then, remove most of the liquid from the wells. Immediately, add 150 microliters of blocking solution to each of the top wells of the axonal and somatic compartments. After 15 minutes, remove most of the liquid from the wells and immediately, add 100 microliters of the primary antibody solution, to each of the top wells of the axonal and somatic compartments.
Cover the chip to minimize evaporation and incubate the cells in the primary antibody for one hour at room temperature. Next, rinse the chip by removing most of the solution and immediately adding 150 microliters of PBS to each of the top wells of the axonal and sematic compartments. After five minutes, remove the PBS from both wells and repeat the rinse two more times.
Now, remove most of the liquid from the wells and immediately, add 100 microliters of secondary antibody in PBS, to each of the top wells of the axonal and somatic compartments. Cover the chip to minimize evaporation and incubate the chip for one hour at room temperature. As previously shown, rinse the chip three times with PBS then, mount and image the chips as described in the accompanying text protocol.
Shown here, is a side-by-side comparison of neurons grown on plastic chips and PDMS devices. Neuronal growth is comparable within the two platforms up to 15 days in culture, but at longer culture ages, isolated axons within the plastic chip, appear healthier with less beating. To further visualize axons within the axonal compartments, these samples were also immunostained for beta tubulin III, which shows healthy axonal growth within the plastic chips at 22 days in culture.
Further staining with 24-day samples shows neurons grown in the plastic chips to express both the excitatory and inhibitory synaptic markers, vGlut1 and vGat. Here are the two synaptic markers combined with retrograde labeled mCherry neurons. To demonstrate the suitability of this chip for axon injury and regeneration studies, retrograde labeled neurons were imaged before and 24 hours after axotomy.
A retraction bulb and regenerating axon are both evident following axotomy. These results are equivalent to published data using PDMS based devices. Classic multicompartment chips provide an easy-to-use option for compartmentalizing neurons providing long-term neuron cultures, which remain viable for longer than three weeks.
While attempting this procedure, it is important to remember to seed appropriate number of neurons and make sure your incubator is well humidified during culture.
