The authors acknowledge technical or editorial assistance from Taylor Wilt (Xona), Smita Paranjape (UNC-Chapel Hill), Joyce Ciechanowski (Xona) and Brad Taylor (Xona). The authors acknowledge support from Xona Microfluidics, LLC and the National Institute of Mental Health (R42 MH097377). Imaging was partially supported by the Confocal and Multiphoton Imaging Core Facility of NINDS Center Grant P30 NS045892 and NICHD Center Grant (U54 HD079124). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

NOTE: A schematic of the plastic multicompartment chip is shown in Figure 1A, B. The chip is the size of a standard microscope slide (75 mm &#215; 25 mm). The features of the chip, including main channels or compartments, wells, and microgrooves are labeled and are provided for future reference. Figure 1C is a photograph of the chip demonstrating the fluidic isolation of the compartments.
1. Preparation and Coating of the Multicompartment Chips
<ol>
	<li>In a bio-safety cabinet, place the chip into a Petri dish or other suitable sterile container.</li>
	<li>Add 100 &#181;L of pre-coating solution to the upper left well of the chip and allow it to flow through the main channel into the adjoining well. 
	NOTE: The pre-coating solution is used to pre-coat the microfluidic channels to eliminate the potential for trapping air bubbles within the chip.</li>
	<li>Fill the lower left well with 100 &#181;L of pre-coating solution. Wait 5 min to allow the solution to flow through the microgrooves.</li>
	<li>Add 100 &#181;L of pre-coating solution to the upper right well and allow it to flow through the main channel into the adjoining well. Fill the lower right well with 100 &#181;L of pre-coating solution.</li>
	<li>Aspirate the solution from each well. Aspirate away from the main channels to avoid removing liquid from the main channels (Figure 2A). Immediately add 150 &#181;L of phosphate-buffered saline (PBS) to the upper left well. Wait 1.5 min. 
	CAUTION: Do not aspirate all liquid from the enclosed main channels.</li>
	<li>Add 150 &#181;L PBS to the lower left well. Wait 5 min to allow liquids to flow through the microgrooves. Add 150 &#181;L PBS to the upper right well. Add 150 &#181;L PBS to lower right well. Wait 10 min.</li>
	<li>Repeat steps 1.5-1.6 for a second PBS wash.</li>
	<li>Check the chip under a tissue culture microscope for bubbles in the main channels. If bubbles are present, perform the procedures below. If no bubbles are present, skip to step 1.9.
	<ol>
		<li>Aspirate PBS from the wells angling the pipet tip away from the channel opening (Figure 2A).</li>
		<li>Dispense 100 &#181;L of pre-coating solution into the upper well, angling the tip of the pipet near the channel opening (Figure 2B). The bubbles should move through the channel into the lower well. Wait 1.5 min.</li>
		<li>Repeat steps 1.3-1.8.</li>
	</ol>
	</li>
	<li>Aspirate PBS from the wells angling the pipet tip away from the channel opening (Figure 2A).</li>
	<li>Add 100 &#181;L of 0.5 mg/mL poly d-lysine (PDL) to the upper left well of the chip. Wait 1.5 min. Fill the lower left well with 100 &#181;L of PDL.</li>
	<li>Add 100 &#181;L of PDL to the upper right well of the chip. Wait 1.5 min. Add 100 &#181;L to the lower right well.</li>
	<li>Close the petri dish&#160;and place the chip in an incubator at 37 &#176;C for 1 h.</li>
	<li>Repeat PBS wash steps 1.5-1.6 twice to remove excess PDL.</li>
	<li>Aspirate the PBS from the device.</li>
	<li>Immediately add 100 &#181;L of cell culture media to the upper left well of the chip. Wait 1.5 min. Add media to the lower left well. Add media to the upper right well. Wait 1.5 min. Add 100 &#181;L medium to the lower right well of the chip.</li>
	<li>Place the chip in the 37&#176;C incubator until ready to plate cells.</li>
</ol>
2. Seeding Neurons into the Multicompartment Chips
<ol>
	<li>Prepare cell suspension of dissociated rat hippocampal neurons according to established protocols21,22 to yield a density of ~12 &#215; 106 cells/mL. 
	NOTE: Use of cell suspension densities between 3 and 12 &#215; 106 cells/mL is possible. If a lower density is used, the volume of cell suspension to be added to the chip may be increased (see below). The procedure described below is applicable for murine dissociated cortical or hippocampal neurons. Optimal cell densities for other neuron types may vary.</li>
	<li>Remove the majority of media in each well of the chip, leaving approximately 5&#160;&#181;L in each well. Aspirate away from the main channels to avoid removing liquid from the main channels (Figure 2A). 
	CAUTION: Do not aspirate liquid from the enclosed main channels. Air bubbles may become trapped in the chip if fluid is aspirated from the main channels.</li>
	<li>Load 5 &#181;L of cell suspension in the upper right well and another 5 &#181;L of cell suspension in the lower right well (~120,000 cells total). Load the cells by dispensing close to the main channel (Figure 2B). Check under a microscope to ensure the neurons are in the main channel. Wait for 5 min to allow the cells to attach. 
	NOTE: Neurons can be loaded into either compartment. For explanation purposes, the somatic compartment is the main channel on the right side, but either compartment can be used as the somatic compartment. Use of lower cell densities down to 60,000 cells per chip is possible. Up to 10 &#181;L of cell suspension may be added to each well of the somatic compartment in combination with a cell suspension with fewer cells than described above.</li>
	<li>Add approximately 150 &#181;L of neuronal culture media to each of the upper and lower right wells, and then add 150 &#181;L of media to each of the upper and lower left wells. Place the chip into humidified tray in a 5% CO2 37 &#176;C incubator.</li>
	<li>After 24 h, perform a media change by removing media from the wells. Make sure the main channel remains filled. Add 150 &#181;L of media to each top well, and then fill the bottom wells.</li>
	<li>Place the chip back in the incubator for the desired number of days. 
	NOTE: Monitor the media every couple of days to make sure it remains light pink. If the media is yellowish, replace 50% of it with fresh media. If the fluid level is low, make sure there is adequate humidity and appropriate secondary containment of the chips to prevent evaporation. Minimizing, or even eliminating, media changes is possible using secondary containment and/or covering the dish containing the chip with polytetrafluoroethylene (PTFE)-FEP film.</li>
</ol>
3. Retrograde Labeling of Neurons within the Chip
NOTE: Retrograde labeling can be performed using multiple techniques, including using modified cholera toxin and rabies virus. Below are instructions for labeling neurons using G-deleted Rabies-mCherry or -eGFP virus. Handle potentially infectious materials according to the local organization&#39;s guidelines. Additional training may be required.
<ol>
	<li>Warm fresh neuronal culture media to 37 &#176;C. Estimate ~400 &#181;L of media per chip.</li>
	<li>Dilute 100,000 viral units of modified rabies virus in a total of 50 &#181;L using media taken from either well of the axonal compartment. 
	NOTE: Dispose of tips and tubes in contact with the virus according to the organization-approved protocol.</li>
	<li>Gently pipet the remaining media from the wells of the axonal compartment and store in a centrifuge tube at 37 &#176;C.</li>
	<li>Add 150 &#181;L of fresh warm media and the 50 &#181;L of diluted virus to the axonal compartment. Incubate for 2 h at 37 &#176;C incubator.</li>
	<li>Remove media containing virus and dispose of it properly. 
	NOTE: Air bubbles may become trapped in the chip if fluid is aspirated from the main channels.</li>
	<li>Gently add 75 &#181;L of fresh media to one axon well and allow it to flow to the other axon well.</li>
	<li>Remove flow-through from the second axon well and dispose properly.</li>
	<li>Repeat steps 3.6 and 3.7 once.</li>
	<li>Add back stored media to the axonal compartment. Add approximately 50 &#181;L fresh media, if necessary, to maintain adequate volume and return the cells to the incubator. 
	NOTE: Fluorescent protein expression is visible by 48 h and persists for up to 8 days. Neurons can be imaged for up to 30 min at room temperature in neuronal culture media. Culture media can also be replaced with warmed CO2-independent hibernate E with B27 and imaged for longer. Neurons can also be imaged within a well-humidified environmental chamber at 37 &#176;C and 5% CO2. In this case, humidification is critical for minimizing evaporative losses within the chips, which is exacerbated by heating and can compromise neuron health.</li>
</ol>
4. Fluidic Isolation of the Axonal Compartment within the Chip
<ol>
	<li>Remove 20 &#181;L from the lower left well of the axonal compartment and place into the upper right well of the somatic compartment. Wait 2 min for flow within each channel to equilibrate.</li>
	<li>Remove 50 &#181;L of media from the axonal compartment. Add 0.3 &#181;L of 1 mM Alexa Fluor 488 hydrazide to this media, mix via pipet and return back to the axonal compartment. The chip is ready for imaging. 
	NOTE: Other compounds of interest can be added. Adding a fluorescent dye with a similar molecular weight as the compound of interest is recommended in order to monitor fluidic isolation over time.</li>
</ol>
5. Performing Axotomy Within the Chip
<ol>
	<li>Remove media from the axonal compartment keeping the pipet tip away from the entrance of the main channel (Figure 2A) and store it in a centrifuge tube.</li>
	<li>Aspirate the axonal compartment completely, placing the aspiration pipet near either entrance of the main channel of the axonal compartment (Figure 2B). Continue aspiration for 1-2 min. Make sure that solution is completely removed from the compartment. 
	NOTE: The vacuum pressure for aspiration must be at least 18 inch-Hg for the axotomy procedure to work properly.</li>
	<li>Replace the axonal compartment with the stored media and confirm that the axons are&#160; severed by looking at the chip under a microscope. 
	NOTE: If bubbles form in the axonal compartment when replacing the media, repeat steps 5.1-5.2.</li>
	<li>Return the chip to the incubator.</li>
</ol>
6. Fluorescence Immunostaining within the Chip
<ol>
	<li>Prepare 4% formaldehyde fixation solution in PBS (4% formaldehyde, 1 &#181;M MgCl2, 0.1 &#181;m CaCl2, 120 mM sucrose)</li>
	<li>Remove most of the media in the wells of the chip (do not dry interior compartments).</li>
	<li>Immediately add 100 &#181;L of fixation solution to the top wells of the axonal and somatic compartments.</li>
	<li>After 1 min, add 100 &#181;L of fixation solution to the bottom wells. Fix for 30 min at room temperature.</li>
	<li>Remove most of the solution from the wells of the chip (do not dry interior compartments). Immediately add 150 &#181;L of PBS to each of the top wells of the axonal and somatic compartments. Wait 2 min for the PBS to flow into the bottom wells.</li>
	<li>Repeat step 6.5 twice.</li>
	<li>Remove most of the PBS from the wells of the chip. Immediately add 150 &#181;L of PBS with 0.25% TritonX-100 to each of the top wells of the axonal and somatic compartments. Wait for 15 min.</li>
	<li>Remove most of the liquid from the wells of the chip and immediately add 150 &#181;L of blocking solution (10% normal goat serum in PBS) to each of the top wells of the axonal and somatic compartments. Wait for 15 min. 
	NOTE: Effective blocking solutions should be specific to the secondary antibody, e.g., for a donkey anti-sheep secondary antibody, use donkey serum in the blocking solution.</li>
	<li>Remove most of the liquid from the wells of the chip and immediately add 100 &#181;L of primary antibody (or antibodies) in 1% normal goat serum in PBS to each of the top wells of the axonal and somatic compartments. Cover to minimize evaporation and wait for 1 h at room temperature or 4 &#176;C overnight.</li>
	<li>Remove most of the solution from the wells of the chip (do not dry interior compartments). Immediately add 150 &#181;L of PBS to each of the top wells of the axonal and somatic compartments. Wait 5 min for the PBS to flow into the bottom wells.</li>
	<li>Repeat step 6.10 twice.</li>
	<li>Remove most of the liquid from the wells of the chip and immediately add 100 &#181;L of secondary antibody (or antibodies) in PBS to each of the top wells of the axonal and somatic compartments. Cover to minimize evaporation and wait for 1 h at room temperature. 
	NOTE: Refer to the manufacturer&#39;s instructions for the recommended dilution of secondary antibodies.</li>
	<li>Repeat steps 6.10-6.11.</li>
	<li>If imaging within 1 day of immunostaining, keep the chip filled with PBS. If chip will be stored longer than 1 day before imaging, wrap the dish containing the chip in paraffin film to prevent evaporation and store at 4 &#176;C until ready to image.</li>
	<li>For longer term storage of samples, mounting media (e.g., Fluoromount-G) can be used.
	<ol>
		<li>Remove most of the liquid from the wells of the chip. Use a 1 mL disposable plastic pipet to add 2 drops of mounting media to each of the top wells of the axonal and somatic compartments.</li>
		<li>Tilt the chip to encourage the flow of the mounting media through the channels. After 5 min add 2 drops to the bottom wells. Wait for 1 h before imaging. 
		NOTE: After using mounting media it will not be possible to re-probe for other targets.</li>
	</ol>
	</li>
</ol>

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A.M.T. is an inventor of the microfluidic chamber/chip (US 7419822 B2) and is a member of Xona Microfluidics, LLC. V.P. is an employee of Xona Microfluidics, LLC. J.H. is a member of Xona Microfluidics, LLC. T.N. declares no competing financial interests.&#160;Open access publication of this article is sponsored by Xona Microfluidics.

Plastic multicompartment chips provide an easy-to-use option for compartmentalizing neurons, providing long-term neuronal cultures (&gt;3 weeks). This protocol details how to culture cortical and hippocampal murine neurons within these chips. The creation of soluble microenvironments and how to retrograde label neurons, perform axotomy, and perform immunocytochemistry were also discussed. Importantly, these chips are compatible with high-resolution, fluorescence, and live imaging. 
Plastic multicompartment chips provide many of the same functions as the PDMS-based compartmentalized devices, but have advantages, disadvantages, and some distinguishing features. Table 1 provides a feature comparison of both plastic chip and PDMS-based devices. First and foremost, the chips are pre-assembled and made permanently hydrophilic, which facilitates wetting, making them easier to use. The plastic is not gas permeable, unlike PDMS, so if bubbles unexpectedly form within the channels, they do not readily escape and must be removed. A pre-coating solution containing mainly ethanol and some other proprietary agents eliminates bubble formation. 
Live imaging of projections following transduction of fluorescent proteins was performed within the chips (Figure 4) and there was no detectable autofluorescence of the plastic. A caveat is that immersion oils used with the chip for high numerical aperture objectives must be silicone oil based, and not mineral oil based. Mineral oil can cause an adverse reaction with the cyclic olefin copolymer. For brightfield imaging, it is important to note that the microgrooves in the chip are rounded at the ends and there is a gradual tapering of the z-direction of the main channels towards the microgroove barrier causing some light refraction at either end of the microgrooves during brightfield imaging (Figure 3). Because the chip is pre-assembled, antibody penetration into the micron-sized microgrooves may be uneven (as with permanently bonded PDMS-based devices); thus, quantitative analysis following immunostaining should be performed in the channels/compartments. Immunostaining of neuronal projections within the microgrooves can be improved by creating a volume difference between the compartments to aid the flow of antibodies and fluorophores into the microgrooves.

Traditional neuron culture approaches result in random outgrowth of axons and dendrites, which prevent the study of neurons in their unique polarized morphology. Microfabricated multicompartment devices have become well-established and well-used research tools for neuroscientists in the last 10-15 years (selected high-profile publications are referenced1,2,3,4,5,6,7,8,9,10,11,12,13,14,15,16,17). These devices compartmentalize neurons and provide a method to physically and chemically manipulate subcellular regions of neurons, including somata, dendrites, axons, and synapses18,19. They also provide multiple experimental paradigms that are not possible using random cultures, including studies of axonal transport, axonal protein synthesis, axon injury/regeneration, and axon-to-soma signaling. The basic 2-compartment configuration consists of two parallel microfluidic channels separated by a series of smaller perpendicular microgrooves. Primary or stem cell-derived neurons are plated into one of the microfluidic channels, settle and attach to the bottom surface of the device, and extend neurites over the course of days. Many growth cones find their way into the microgrooves, which are small enough that they prevent cell bodies from entering. Because growth cones are physically restricted and unable to turn around within the microgrooves, they grow straight into the adjacent compartment (axonal compartment) where they are isolated.
Historically, these devices have been molded using poly(dimethylsiloxane) (PDMS) from a photolithographically patterned master mold and are either made in-house in investigators&#39; laboratories or purchased commercially. One of the main drawbacks of using PDMS is its hydrophobicity20. PDMS can be made hydrophilic temporarily, but then quickly becomes hydrophobic within hours in a non-aqueous environment20. Because of this, the devices must be attached to a glass coverslip or other suitable substrate at the time of use. Pre-assembled plastic multicompartment chips are now commercially available (e.g., XonaChips) in injection molded plastic. These chips are made permanently hydrophilic, simplifying device wetting and allowing the pre-assembly of the chip with a thin film of cyclic olefin copolymer (COC) enclosing the microfluidic channels on the bottom. These chips are fabricated in an optically transparent plastic suitable for high-resolution fluorescence imaging.
The purpose of this protocol is to demonstrate the use of the pre-assembled plastic microfluidic chips for multiple experimental paradigms performed using murine hippocampal or cortical neurons. This protocol describes how to retrograde label neurons using a modified rabies virus within the chip. Axotomy for studies of axon injury and regeneration are also described. Lastly, this protocol shows how to perform fluorescence immunostaining with the device.

<table><tbody><tr><td>XonaChip</td><td>Xona Microfluidics, LLC</td><td>XC150</td><td>150 &amp;micro;m length microgroove barrier</td></tr><tr><td></td><td>Xona Microfluidics, LLC</td><td>XC450</td><td>450 &amp;micro;m length microgroove barrier</td></tr><tr><td></td><td>Xona Microfluidics, LLC</td><td>XC900</td><td>900 &amp;micro;m length microgroove barrier</td></tr><tr><td>XC pre-coat&amp;nbsp;</td><td>Xona Microfluidics, LLC</td><td>XC Pre-Coat</td><td>included with XonaChips</td></tr><tr><td>XonaPDL&amp;nbsp;</td><td>Xona Microfluidics, LLC</td><td>XonaPDL</td><td></td></tr><tr><td>E17/E18 timed pregnant Sprague Dawley rats</td><td>Charles River</td><td>24100564</td><td></td></tr><tr><td>&lt;em&gt;neuronal culture media:&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>- Neurobasal medium</td><td>ThermoFisher Scientific</td><td>21103049</td><td></td></tr><tr><td>-B-27 Plus Supplement (50x)</td><td>ThermoFisher Scientific</td><td>A3582801</td><td></td></tr><tr><td>-GlutaMAX Supplement</td><td>ThermoFisher Scientific</td><td>35050061</td><td></td></tr><tr><td>-Antibiotic-Antimycotic (100x)</td><td>ThermoFisher Scientific</td><td>15240112</td><td></td></tr><tr><td>fluorinated ethylene propylene film</td><td>American Durafilm</td><td>50A</td><td>0.5 mil thickness</td></tr><tr><td>modified rabies virus</td><td>Salk Institute for Biological Studies</td><td>G-deleted Rabies-eGFP</td><td>Material Transfer Agreement required</td></tr><tr><td></td><td>Salk Institute for Biological Studies</td><td>G-deleted Rabies-mCherry</td><td>Material Transfer Agreement required</td></tr><tr><td>Alexa Fluor hydrazide 488</td><td>ThermoFisher Scientific</td><td>A10436</td><td></td></tr><tr><td>Fluoromount G</td><td>ThermoFisher Scientific</td><td>00-4958-02</td><td></td></tr><tr><td>Glass Pasteur pipettes</td><td>Sigma-Aldrich</td><td>CLS7095D5X SIGMA</td><td>5.75 in length</td></tr><tr><td>hibernate-E Medium</td><td>ThermoFisher Scientific</td><td>A1247601</td><td></td></tr><tr><td>Pierce 16% formaldehyde</td><td>ThermoFisher Scientific</td><td>28906</td><td></td></tr><tr><td>PBS (10x)</td><td>ThermoFisher Scientific</td><td>QVC0508</td><td></td></tr><tr><td>normal goat serum</td><td>ThermoFisher Scientific</td><td>16210064</td><td></td></tr><tr><td>triton X-100</td><td>ThermoFisher Scientific</td><td>28314</td><td></td></tr><tr><td>anti-vGlut1 antibody</td><td>NeuroMab</td><td>75-066</td><td>clone N28/9, 1:100</td></tr><tr><td>anti-vGAT antibody</td><td>Synaptic Systems</td><td>131 003</td><td>1:1000</td></tr><tr><td>anti_beta-tubulin III</td><td>Aves</td><td>TUJ</td><td>1:1000</td></tr><tr><td>Alexa Fluor secondary antibodies</td><td>ThermoFisher Scientific</td><td></td><td>1:1000</td></tr><tr><td>Incubator, 5% CO&lt;sub&gt;2&lt;/sub&gt; 37 &amp;deg;C</td><td></td><td></td><td></td></tr><tr><td>Epifluorescence imaging system</td><td>EVOS Fluorescence imaging system</td><td>AMF4300</td><td>10x objective</td></tr><tr><td>Spinning disk confocal imaging system</td><td>Andor Technology</td><td>CSU-X1, iXon X3 EMCCD</td><td>60x silicone oil &amp;amp; 20x objectives</td></tr><tr><td>Laser scanning confocal imaging system</td><td>Olympus</td><td>FV3000RS</td><td>30x silicone oil objective</td></tr></tbody></table>

use of pre-assembled plastic microfluidic chips for compartmentalizing primary murine neurons

Microfabricated methods to compartmentalize neurons have become essential tools for many neuroscientists. This protocol describes the use of a commercially available pre-assembled plastic chip for compartmentalizing cultured primary rat hippocampal neurons. These plastic chips, contained within the footprint of a standard microscope slide, are compatible with high-resolution, live, and fluorescence imaging. This protocol demonstrates how to retrograde label neurons via isolated axons using a modified rabies virus encoding a fluorescent protein, create isolated microenvironments within one compartment, and perform axotomy and immunocytochemistry on-chip. Neurons are cultured for &gt;3 weeks within the plastic chips, illustrating the compatibility of these chips for long-term neuronal cultures.

After approximately 5-7 days of neuron growth within the chip, axonal growth is evident. The chips are compatible with phase-contrast imaging as demonstrated in Figure 3, which shows neuronal growth at 24 days. The chips are also compatible with fluorescence imaging (Figure 4, Figure 5, Figure 6, and Figure&#160;7). Three days after rabies virus infection via the axonal compartment, mCherry-positive neurons with axons extending into the axonal compartment were imaged in the chip (Figure 4). To demonstrate the ability to fluidically isolate the compartments, a low molecular weight fluorescent dye (Alexa Fluor 488 hydrazide) was added to the axonal compartment. These results are comparable to PDMS-based chamber17 and demonstrate the suitability of the plastic chip for phase-contrast and fluorescence imaging.
To illustrate neuronal growth with the plastic chips and PDMS devices, we cultured neurons in both platforms and monitored neuronal growth over time. Figure 5 shows neuronal growth from 3 to 22 days in culture; these results are representative of 3 independent experiments. Neuronal growth is comparable within the two platforms up to 15 days in culture, but at longer culture ages (&#62;21 days) isolated axons within the plastic chip appear healthier with less beading (Figure 5A). To further visualize axons within the axonal compartments, we immunostained for &#946;-tubulin III which shows healthy axonal growth within the plastic chips at 22 days in culture (Figure 5B).
Axon injury and regeneration studies are common using microfluidic compartmentalized devices. To demonstrate the suitability of these studies using the chips, retrograde labeled neurons were imaged before and 24 h after axotomy (Figure 6). A retraction bulb and regenerating axon are both evident following axotomy. These results are equivalent to published data using PDMS-based devices14,17.
Immunocytochemistry is a common technique performed within multi-compartment devices to visualize protein localization. After 24 days in culture, neurons within the chips were fixed and stained for both excitatory and inhibitory synaptic markers, vGlut1 and vGat, respectively (Figure 7). Retrograde labeled mCherry neurons were also imaged (Figure 7C). Imaging was performed using spinning disk confocal with a 60&#215; silicone oil immersion objective, demonstrating the ability to perform high-resolution imaging. Importantly, dendritic spines were evident within a magnified region, demonstrating that the neurons cultured within the chips were forming mature synapses.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58421/58421fig1.jpg" /> 
Figure 1: A pre-assembled, plastic two-compartment microfluidic chip for compartmentalizing neurons. (A) Schematic representation of the multicompartment chip showing the locations of the upper and lower wells. (B) An enlarged schematic of the chip showing the main channels (or compartments) and microgrooves which connect the compartments. The main channels are approximately 1.5 mm &#215; 7 &#215; 0.060 mm (W &#215; L &#215; H). The width and height of the microgrooves are approximately 0.01 mm &#215; 0.005 mm, respectively. The length of the microgrooves varies depending on the configuration, 0.15 mm to 0.9 mm. (C) A photograph of a representative multicompartment chip containing food coloring dye in each main channel or compartment demonstrating the ability to fluidically isolate each channel. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/58421/58421fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58421/58421fig2.jpg" /> 
Figure 2: Pipetting techniques needed when using plastic multicompartment chips. (A) When adding and aspirating media for washes, the pipet tip should be angled away from the entrance of the main channel as shown. (B) When loading neurons or performing axotomy, the pipet tip should be angled towards the entrance of the main channel. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/58421/58421fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58421/58421fig3.jpg" /> 
Figure 3: A phase contrast micrograph showing typical neuronal growth within the chip at 24 days in culture. Embryonic hippocampal neurons were seeded into the right somatic compartment. Axon growth is visible in the axonal compartment beginning at 5-7 days. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/58421/58421fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58421/58421fig4.jpg" /> 
Figure 4: Retrograde labeled neurons express mCherry fluorescent protein and extend axons into a fluidically isolated axonal compartment. (A) A merged fluorescence micrograph showing live retrograde labeled neurons infected via a modified mCherry rabies virus briefly applied to the axonal compartment. Neurons were imaged 3 days post-infection at 21 days in culture. Creating an isolated microenvironment within the axonal compartment is demonstrated by application of a low molecular weight dye, Alexa Fluor 488 hydrazide. (B) A merged image of (A) including a differential interference contrast (DIC) image to visualize the microgrooves region of the chip. Images were acquired with laser scanning confocal microscope using a 30&#215;/1.05 N.A. silicone oil (ne = 1.406) objective lens. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/58421/58421fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58421/58421fig5.jpg" /> 
Figure 5: Side-by-side comparison of neuronal growth within multicompartment PDMS devices and plastic chips. (A) Phase contrast micrographs of both platforms taken at 3, 7, 15, and 22 days in culture. At the bottom, a higher magnification region taken from the images at 22 days is included to illustrate axonal growth at this age within both platforms. Axons within the chip are more continuous and appear healthier than in the PDMS device at this age. (B) An invert immunofluorescence micrograph of &#946;-tubulin III stained axons within the axonal compartment of both the PDMS device and plastic chip at 22 days in culture. Images were acquired with a spinning disk confocal imaging system using a 20x/0.45 N.A. objective lens. All scale bars are 100 &#181;m. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/58421/58421fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/58421/58421fig6.jpg" /> 
Figure 6: Axotomy and regeneration of hippocampal neurons within the plastic multicompartment chip. (Top) Neurons were retrograde labeled using a modified mCherry rabies virus and then imaged before axotomy at 24 days in culture. Images were pseudocolored using the &#39;Fire&#39; color look-up table. (Bottom) The same neuron imaged in the top panel was imaged 24 h post-axotomy. White dashed lines show the edges of the microgroove barrier. Axotomy occurred at the location of the left dashed line. The white arrowhead shows a retraction bulb. The white arrow indicates a regenerating axon. Images were acquired with a spinning disk confocal imaging system using a 20&#215;/0.45 N.A. objective lens. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/58421/58421fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/58421/58421fig7.jpg" /> 
Figure 7: Synapses form between hippocampal neurons cultured within plastic multicompartment chips. Immunostaining was performed at 24 days in culture and imaged within the somatic compartment using a 60&#215; silicone oil immersion lens. Neurons express (A) the excitatory synapse marker, vGlut1 (green) and (B) the inhibitory synapse marker, vGAT (blue). (C) Retrograde labeled mCherry neurons (red) were infected via modified rabies virus applied to the axonal compartment. (D) A merged fluorescence micrograph of vGlut1, vGat, and mCherry. (E) The magnified region in (D) indicated with a white box shows dendritic spines, the sites that receive synaptic input from other neurons. Images were acquired with a spinning disk confocal imaging system using a 60&#215;/1.3 N.A. silicone oil (ne = 1.406) objective lens. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/58421/58421fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Plastic multicompartment chips</td>
			<td>PDMS multicompartment devices</td>
		</tr>
		<tr>
			<td>isolate axons</td>
			<td>isolate axons</td>
		</tr>
		<tr>
			<td>establish microenvironments</td>
			<td>establish microenvironments</td>
		</tr>
		<tr>
			<td>axotomize neurons</td>
			<td>axotomize neurons</td>
		</tr>
		<tr>
			<td>optically transparent</td>
			<td>optically transparent</td>
		</tr>
		<tr>
			<td>compatible with high resolution imaging</td>
			<td>compatible with high resolution imaging</td>
		</tr>
		<tr>
			<td>compatible with fluorescence microscopy</td>
			<td>compatible with fluorescence microscopy</td>
		</tr>
		<tr>
			<td>fully assembled</td>
			<td>assembly to substrate required</td>
		</tr>
		<tr>
			<td>healthy axons &#62;21 days</td>
			<td>healthy axons &#62;14 days</td>
		</tr>
		<tr>
			<td>hydrophilic culturing surface</td>
			<td>hydrophobic</td>
		</tr>
		<tr>
			<td>gas impermeable</td>
			<td>gas permeable</td>
		</tr>
		<tr>
			<td>rounded microgrooves and channels</td>
			<td>straight microgrooves</td>
		</tr>
		<tr>
			<td>fewer preparation steps</td>
			<td>top is removable for staining within microgrooves</td>
		</tr>
		<tr>
			<td>not compatible with laser ablation</td>
			<td>absorption of small molecules &#38; organic solvents</td>
		</tr>
		<tr>
			<td>not compatible with mineral oil-based immersion oils (silicone-based oils are fine)</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 1:&#160;Comparison of plastic and PDMS multicompartment platforms for culturing neurons.

Watch this Scientific Journal Video about Use of Pre-Assembled Plastic Microfluidic Chips for Compartmentalizing Primary Murine Neurons at JoVE.com

Use of Pre-Assembled Plastic Microfluidic Chips for Compartmentalizing Primary Murine Neurons

This protocol describes the use of plastic chips to culture and compartmentalize primary murine neurons. These chips are preassembled, user-friendly, and compatible with high-resolution, live, and fluorescence imaging. This protocol describes how to plate rat hippocampal neurons within these chips and perform fluidic isolation, axotomy and immunostaining.

Microfabricated methods to compartmentalize neurons have become essential tools for many neuroscientists. This protocol describes the use of a ...

use-pre-assembled-plastic-microfluidic-chips-for-compartmentalizing

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

50.8K Views.  UNC Neuroscience Center. This protocol describes the use of plastic chips to culture and compartmentalize primary murine neurons. These chips are preassembled, user-friendly, and compatible with high-resolution, live, and fluorescence imaging. This protocol describes how to plate rat hippocampal neurons within these chips and perform fluidic isolation, axotomy and immunostaining. 

Video: Use of Pre-Assembled Plastic Microfluidic Chips for Compartmentalizing Primary Murine Neurons

Fabrication of a Microfluidic Device for the Compartmentalization of Neuron Soma and Axons

In this video, we demonstrate the technique of soft lithography with polydimethyl siloxane (PDMS) which we use to fabricate a microfluidic device for culturing neurons. Previously, a silicon wafer was patterned with the design for the neuron microfluidic device using SU-8 and photolithography to create a master mold, or what we simply refer to as a "master". Next, we pour the silicon polymer PDMS on top of the master which is then cured by heating the PDMS to 80&deg;C for 1 hour. The PDMS forms a negative mold of the device. The PDMS is then carefully cut and lifted away from the master. Holes are punched where the reservoirs will be and the excess PDMS trimmed away from the device. Nitrogen is used to blow away any excess debris from the device. At this point the devices are now ready for use and can either bonded to corning No. 1 cover glass with a plasma sterilizer/cleaner or can be reversibly bound to the cover glass by simply placing the device on top of the cover glass. The reversible bonding of the device to glass is covered in a separate video and requires first that the device be sterilized either with 70% ethanol or by autoclaving. Plasma treating sterilizes the devices so no further treatment is necessary. It is, however, important, when plasma-treating the devices, to add liquid to the devices within 10 minutes of the plasma treatment while the surfaces are still hydrophilic. Waiting longer than 10 minutes to add liquid to the device makes it difficult for the liquid to enter the device. The neuron devices are typically plasma-bound to cover glass and 0.5 mg/ml poly-L-lysine (PLL) in pH 8.5 borate buffer is immediately added to the device. After a minimum of 3 hours incubating with PLL, the devices are washed with dH2O water a minimum of 3 times with at least 15 minutes between each wash. Next, the water is removed and fresh media is added to the device. At this point the device is ready for use. It is important to remember at this point to never remove all the media from the device. Always leave media in the main channel.

In this video we demonstrate the technique of soft lithography with polydimethyl siloxane (PDMS) which we use to farbricate a microfluidic device for culturing neurons.

In this video, we demonstrate the technique of soft lithography with polydimethyl siloxane (PDMS) which we use to fabricate a microfluidic device for ...

Biology

Preparing E18 Cortical Rat Neurons for Compartmentalization in a Microfluidic Device

In this video, we demonstrate the preparation of E18 cortical rat neurons. E18 cortical rat neurons are obtained from E18 fetal rat cortex previously dissected and prepared. The E18 cortex is, upon dissection, immediately dissociated into individual neurons. It is possible to store E18 cortex in Hibernate E buffer containing B27 at 4&deg;C for up to a week before the dissociation is performed. However, there will be a drop in cell viability. Typically we obtain our E18 Cortex fresh. It is transported to the lab in ice cold Calcium free Magnesium free dissection buffer (CMFM). Upon arrival, trypsin is added to the cortex to a final concentration of 0.125%. The cortex is then incubated at 37&deg;C for 8 minutes. DMEM containing 10% FBS is added to the cortex to stop the reaction. The cortex is then centrifuged at 2500 rpm for 2 minutes. The supernatant is removed and 2 ml of Neural Basal Media (NBM) containing 2% B27 (vol/vol) and 0.25% Glutamax (vol/vol) is added to the cortex which is then re-suspended by pipetting up and down. Next, the cortex is triturated with previously fire polished glass pipettes, each with a successive smaller opening. After triturating, the cortex is once again centrifuged at 2500 rpm for 2 minutes. The supernatant is then removed and the cortex pellet re-suspended with 2 ml of NBM containing B27 and Glutamax. The cell suspension is then passed through a 40 um nylon cell strainer. Next the cells are counted. The neurons are now ready for loading into the neuron microfluidic device.

In this video we demonstrate the preparation of E18 Cortical Rat Neurons.

In this video, we demonstrate the preparation of E18 cortical rat neurons. E18 cortical rat neurons are obtained from E18 fetal rat cortex previously ...

Culturing and Electrophysiology of Cells on NRCC Patch-clamp Chips

Due to its exquisite sensitivity and the ability to monitor and control individual cells at the level of ion channels, patch-clamping is the gold standard of electrophysiology applied to disease models and pharmaceutical screens alike 1. The method traditionally involves gently contacting a cell with a glass pipette filled by a physiological solution in order to isolate a patch of the membrane under its apex 2. An electrode inserted in the pipette captures ion-channel activity within the membrane patch or, when ruptured, for the whole cell. In the last decade, patch-clamp chips have been proposed as an alternative 3, 4: a suspended film separates the physiological medium from the culture medium, and an aperture microfabricated in the film replaces the apex of the pipette. Patch-clamp chips have been integrated in automated systems and commercialized for high-throughput screening 5. To increase throughput, they include the fluidic delivery of cells from suspension, their positioning on the aperture by suction, and automated routines to detect cell-to-probe seals and enter into whole cell mode. We have reported on the fabrication of a silicon patch-clamp chip with optimized impedance and orifice shape that permits the high-quality recording of action potentials in cultured snail neurons 6; recently, we have also reported progress towards interrogating mammalian neurons 7. Our patch-clamp chips are fabricated at the Canadian Photonics Fabrication Centre 8, a commercial foundry, and are available in large series. We are eager to engage in collaborations with electrophysiologists to validate the use of the NRCC technology in different models. The chips are used according to the general scheme represented in Figure 1: the silicon chip is at the bottom of a Plexiglas culture vial and the back of the aperture is connected to a subterranean channel fitted with tubes at either end of the package. Cells are cultured in the vial and the cell on top of the probe is monitored by a measuring electrode inserted in the channel.The two outside fluidic ports facilitate solution exchange with minimal disturbance to the cell; this is an advantage compared to glass pipettes for intracellular perfusion.
<img alt="Figure 1" src="/files/ftp_upload/3288/3288fig1.jpg" />
Figure 1. Principle of measurement using the NRCC patch-clamp chip
We detail here the protocols to sterilize and prime the chips, load them with medium, plate them with cells, and finally use them for electrophysiological recordings.

We show how planar patch-clamp chips fabricated at the National Research Council of Canada are sterilized, primed, loaded with medium, plated with cells, and used for electrophysiological recordings.

Due to its exquisite sensitivity and the ability to monitor and control individual cells at the level of ion channels, patch-clamping is the gold standard ...

A Multi-compartment CNS Neuron-glia Co-culture Microfluidic Platform

We present a novel multi-compartment neuron co-culture microsystem platform for in vitro CNS axon-glia interaction research, capable of conducting up to six independent experiments in parallel for higher-throughput. We developed a new fabrication method to create microfluidic devices having both micro and macro scale structures within the same device through a single soft-lithography process, enabling mass fabrication with good repeatability.
The multi-compartment microfluidic co-culture platform is composed of one soma compartment for neurons and six axon/glia compartments for oligodendrocytes (OLs). The soma compartment and axon/glia compartments are connected by arrays of axon-guiding microchannels that function as physical barriers to confine neuronal soma in the soma compartment, while allowing axons to grow into axon/glia compartments. OLs loaded into axon/glia compartments can interact only with axons but not with neuronal soma or dendrites, enabling localized axon-glia interaction studies. The microchannels also enabled fluidic isolation between compartments, allowing six independent experiments to be conducted on a single device for higher throughput.
Soft-lithography using poly(dimethylsiloxane) (PDMS) is a commonly used technique in biomedical microdevices. Reservoirs on these devices are commonly defined by manual punching. Although simple, poor alignment and time consuming nature of the process makes this process not suitable when large numbers of reservoirs have to be repeatedly created. The newly developed method did not require manual punching of reservoirs, overcoming such limitations. First, seven reservoirs (depth: 3.5 mm) were made on a poly(methyl methacrylate) (PMMA) block using a micro-milling machine. Then, arrays of ridge microstructures, fabricated on a glass substrate, were hot-embossed against the PMMA block to define microchannels that connect the soma and axon/glia compartments. This process resulted in macro-scale reservoirs (3.5 mm) and micro-scale channels (2.5 &mu;m) to coincide within a single PMMA master. A PDMS replica that served as a mold master was obtained using soft-lithography and the final PDMS device was replicated from this master.
Primary neurons from E16-18 rats were loaded to the soma compartment and cultured for two weeks. After one week of cell culture, axons crossed microchannels and formed axonal only network layer inside axon/glia compartments. Axons grew uniformly throughout six axon/glia compartments and OLs from P1-2 rats were added to axon/glia compartments at 14 days in vitro for co-culture.

We developed a novel multi-compartment neuron co-culture microsystem platform for in vitro CNS axon-glia interaction research. The platform is capable of conducting up to six independent experiments in parallel and was fabricated using a newly developed macro/micro hybrid fabrication method.

We present a novel multi-compartment neuron co-culture microsystem platform for in vitro CNS axon-glia interaction research, capable of conducting up to ...

Preparation of Neuronal Co-cultures with Single Cell Precision

Microfluidic embodiments of the Campenot chamber have attracted great interest from the neuroscience community. These interconnected co-culture platforms can be used to investigate a variety of questions, spanning developmental and functional neurobiology to infection and disease propagation. However, conventional systems require significant cellular inputs (many thousands per compartment), inadequate for studying low abundance cells, such as primary dopaminergic substantia nigra, spiral ganglia, and Drosophilia melanogaster neurons, and impractical for high throughput experimentation. The dense cultures are also highly locally entangled, with few outgrowths (&#60;10%) interconnecting the two cultures. In this paper straightforward microfluidic and patterning protocols are described which address these challenges: (i) a microfluidic single neuron arraying method, and (ii) a water masking method for plasma patterning biomaterial coatings to register neurons and promote outgrowth between compartments. Minimalistic neuronal co-cultures were prepared with high-level (&#62;85%) intercompartment connectivity and can be used for high throughput neurobiology experiments with single cell precision.

Protocols for single neuron microfluidic arraying and water masking for the in-chip plasma patterning of biomaterial coatings are described. Highly interconnected co-cultures can be prepared using minimal cell inputs.

Microfluidic embodiments of the Campenot chamber have attracted great interest from the neuroscience community. These interconnected co-culture platforms ...

Compartmentalization of Human Stem Cell-Derived Neurons within Pre-Assembled Plastic Microfluidic Chips

Use of microfluidic devices to compartmentalize cultured neurons has become a standard method in neuroscience. This protocol shows how to use a pre-assembled multi-compartment chip made in a cyclic olefin copolymer (COC) to compartmentalize neurons differentiated from human stem cells. The footprint of these COC chips are the same as a standard microscope slide and are equally compatible with high resolution microscopy. Neurons are differentiated from human neural stem cells (NSCs) into glutamatergic neurons within the chip and maintained for 5 weeks, allowing sufficient time for these neurons to develop synapses and dendritic spines. Further, we demonstrate multiple common experimental procedures using these multi-compartment chips, including viral labeling, establishing microenvironments, axotomy, and immunocytochemistry.

This protocol demonstrates the use of compartmentalized microfluidic chips, injection molded in a cyclic olefin copolymer to cultured neurons differentiated from human stem cells. These chips are preassembled and easier-to-use than traditional compartmentalized poly(dimethylsiloxane) devices. Multiple common experimental paradigms are described here, including viral labeling, fluidic isolation, axotomy, and immunostaining.

Use of microfluidic devices to compartmentalize cultured neurons has become a standard method in neuroscience. This protocol shows how to use a ...

Microfluidics-Assisted Selective Depolarization of Axonal Mitochondria

Mitochondria are the primary suppliers of ATP (adenosine triphosphate) in neurons. Mitochondrial dysfunction is a common phenotype in many neurodegenerative diseases. Given some axons' elaborate architecture and extreme length, it is not surprising that mitochondria in axons can experience different environments compared to their cell body counterparts. Interestingly, dysfunction of axonal mitochondria often precedes effects on the cell body. To model axonal mitochondrial dysfunction in vitro, microfluidic devices allow treatment of axonal mitochondria without affecting the somal mitochondria. The fluidic pressure gradient in these chambers prevents diffusion of molecules against the gradient, thus allowing for analysis of mitochondrial properties in response to local pharmacological challenges within axons. The current protocol describes the seeding of dissociated hippocampal neurons in microfluidic devices, staining with a membrane-potential sensitive dye, treatment with a mitochondrial toxin, and the subsequent microscopic analysis. This versatile method to study axonal biology can be applied to many pharmacological perturbations and imaging readouts, and is suitable for several neuronal subtypes.

The present protocol describes the seeding and staining of neuronal mitochondria in microfluidic chambers. The fluidic pressure gradient in these chambers allows for the selective treatment of mitochondria in axons to analyze their properties in response to pharmacological challenges without affecting the cell body compartment.

Mitochondria are the primary suppliers of ATP (adenosine triphosphate) in neurons. Mitochondrial dysfunction is a common phenotype in many ...

Fluorescence Immunostaining of Rat Hippocampal Neurons within a Microfluidic Chip

Source: Nagendran, T., et al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/58421">Use of Pre-Assembled Plastic Microfluidic Chips for Compartmentalizing Primary Murine Neurons.</a> J. Vis. Exp. (2018)
This video demonstrates immunofluorescence staining of rat hippocampal neurons cultured within a microfluidic chip to analyze neuronal maturation and connectivity.

1. Fluorescence Immunostaining within the Chip

	Prepare 4% formaldehyde fixation solution in PBS (4% formaldehyde, 1 &#181;M MgCl2, 0.1 &#181;m CaCl2, ...

JoVE Encyclopedia of Experiments

Neuropathology

Trauma

Compartmentalized Primary Murine Neuron Culture Using Plastic Microfluidic Chips

Source: Nagendran, T., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/58421">Use of Pre-Assembled Plastic Microfluidic Chips for Compartmentalizing Primary Murine Neurons.</a> J. Vis. Exp. (2018).
This video demonstrates the utilization of a pre-assembled plastic microfluidic chip for the compartmentalized culture of primary rat hippocampal neurons. The neurons are plated into one of the microfluidic compartments, where they attach and grow, extending their axons through the chip&#39;s microgrooves into the adjacent compartment. The narrow microgrooves prevent neuronal cell bodies from entering, thereby enabling physical separation between the somatic and axonal compartments.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Neuronal Culture Techniques

Primary Neuronal Culture

Differentiation of Neural Stem Cells to Neurons Using a Compartmentalized Microfluidic Device

Source: Paranjape, S. R., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/59250">Compartmentalization of Human Stem Cell-Derived Neurons within Pre-Assembled Plastic Microfluidic Chips.</a> J. Vis. Exp. (2019)
The video demonstrates the differentiation of neural stem cells (NSCs) using a microfluidic device comprising a somatic and an axonal compartment separated by a microgroove barrier. NSCs are introduced into the somatic compartment and are allowed to adhere. The NSCs differentiate into neurons inside the somatic compartment and extend axons through the microgroove barrier to the axonal compartment.

NOTE:&#160;Figure 1A, B&#160;shows a schematic of the pre-assembled cyclic olefin copolymer (COC) chip, including the locations of the main channels or ...