The authors are grateful to Ulrich Marggraf (ISAS) for SU-8 fabrication and Maria Becker (ISAS) for SEM imaging. The research was financially supported by the Deutsche Forschungsgemeinschaft (DFG WE3737/3-1), a Bundesministerium f&#252;r Bildung und Forschung grant (BMBF 0101-31P6541) and by the Ministerium f&#252;r Innovation, Wissenschaft und Forschung des Landes Nordrhein-Westfalen. Heike Hardelauf thanks the International Leibniz Graduate School "Systems Biology Lab-on-a-Chip" for financial support.

The protocols are intended for neuroscientists, and are summarized in Figure 2. As such it is recommend that microfabrication of the SU-8 masters used for PDMS replication is outsourced to commercial or institutional facilities. The two layer mask designs are freely available as supplementary information (ZIP) on the Royal Society of Chemistry website12. Importantly, the first SU-8 layer should be fabricated to a depth of 2.5-3.0 &#956;m, and the second to a depth of 25-30 &#956;m. These are key dimensions necessary for effective neuron arraying. The 3 micron height limit is also necessary to prevent neurons being transported or migrating between the two compartments12.
1. Microfluidic Device Replication in PDMS
<ol>
	<li>Mix the PDMS prepolymer thoroughly with the curing agent (184) at a ratio of 10:1 and degas the mixture in a vacuum desiccator for typically 20 min. Alternatively, put the mixture in a 50 ml Falcon tube and use low g centrifugation for degassing.</li>
	<li>Place the 2-layer microstructured SU-8 wafer on an 80 &#176;C hotplate and align polymer frames to each device on the wafer. Pour PDMS into the frames to a depth of &#8805;5 mm and allow &#62;1 hr to ensure complete thermal curing.</li>
	<li>Once cured, remove the wafer from the hotplate and allow to cool on a flat surface.</li>
	<li>Wear protective eyewear and work from one corner (away from the SU-8 microstructures) with a rigid scalpel to gently prize the frame and PDMS from the wafer.</li>
	<li>Remove the PDMS mold from the frame and use a pair of scissors to trim the device of excess PDMS. This so-called flashing is often produced when a thin film of PDMS spreads between the polymer frame and the wafer surface.</li>
	<li>Remaining PDMS can be applied to the entire wafer and cured to protect the wafer during storage. Remove this prior to subsequent device molding, and in doing so remove any particulate contaminants from the surface.</li>
</ol>
2. Device Assembly and Interconnection
<ol>
	<li>Prepare the 6 interface ports using a 3 mm diameter biopsy punch. Work on a dark surface, in good light conditions and with the microstructured features facing upwards to aid alignment of the ports with the microfluidic channels.</li>
	<li>Inspect the device for particulates. Remove these using reversible adhesive tape.</li>
	<li>The microfluidic circuit is encapsulated using a thin PDMS layer mounted on a glass coverslip: These are prepared by pouring a small volume (~0.5 ml) of the PDMS-curing agent mixture (10:1, 601) onto the base of a flexible, bacteriological grade Petri dish. Gently press a coverslip onto the PDMS layer, place on the hotplate and thermally cure for a few minutes. Quickly after cooling, use a scalpel to cut out and prize the coverslip-PDMS bilayer from the Petri dish. Trim with scissors if necessary.</li>
	<li>Plasma bond the PDMS device and support layer to produce a good seal: Optimum conditions are instrument specific. For example, a plasma oven operating at 70 W and 40 kHz in a 0.2 mbar oxygen atmosphere for 40 sec produces an excellent PDMS&#8211;PDMS bond. Once plasma treated, press both parts together and leave for a minute to fully bond.</li>
	<li>Use silicone tubing (1.6 mm ID; 3.2 mm OD) to interface with pumps and syringes.</li>
</ol>
3. Biomaterial Patterning and Microfluidic Operations
The protocols for in situ biomaterial patterning, neuron culture and fluidic treatments are illustrated in Figure 2.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51389/51389fig2highres.jpg" width="500" /> 
Figure 2. Illustrated microfluidic protocols. A) PL and PLL-g-PEG patterning; addition of the cell adhesion layer (e.g., PL, green), with evaporation producing the aligned water mask. B) Atmospheric air plasma treatment of the exposed PL; two antenna pins are used, with the plasma introduction antenna illustrated with a pink star. C) PBS (blue) washing by aspiration is used to avoid contaminating the plasma-treated region with PL. D) Delivery of PLL-g-PEG (red) by aspiration to coat the plasma-treated regions. E) Cell loading; simultaneous microfluidic arraying of neurons to both flanking compartments by aspiration, F) media (pink) perfusion using a hydrostatic-driven flow. G) Isolated fluidic treatments; peripheral treatment of a test agent (black) delivered from the bottom flanking inlet by aspiration and H) central treatment delivered from the bottom central inlet by aspiration. Figure and legend reproduced with permission of the Royal Society of Chemistry (RSC)12. <a href="https://www.jove.com/files/ftp_upload/51389/51389fig2highres.jpg" target="_blank">Click here to view larger image.</a>
Immediately following plasma bonding the PDMS microfluidic channels are highly hydrophilic (contact angle &#8804;5&#176;). Such surfaces are suitable for the adhesion and culture of cell lines such as differentiated human SH-SY5Y neuron-like cells. For the ex vivo culture of primary neurons and the culture of neuronal precursor cell lines such as the Lund human mesencephalic cell line (LUHMES) a polyamine coating, either polylysine (PL) or polyornithine (PO), is required as an electrostatic anchor. In addition adhesion proteins such as laminin or fibronectin are often required as the integrin-interfacing coating. However, the PL and PO coatings readily bind neurons, hindering the microfluidic delivery to the array sites and also imposing unwanted shear stresses. To solve this problem in-chip biomaterial patterning is required to localize adhesion materials at the trap sites and the outgrowth microchannels. This also promotes the development of inter-compartment connections and reduces local entanglement. The protocol also involves the addition of PEG materials to neighboring areas to ensure proteins and neurons are restricted to the desired locations during lengthy culture. The following protocol is documented in Figure 3:
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51389/51389fig3highres.jpg" width="500" /> 
Figure 3. Water masking for biomaterial patterning by plasma stenciling. A) Illustration of the water masking process with the PDMS micropillars pinning the water meniscus in place for plasma stenciling. Curvature of the meniscus in the vertical plane has been neglected. B) Water mask visualized with a red dye and positioned by meniscus pinning. C) The patterned PL-FITC coating imaged after plasma stenciling. D) Addition of protein rejecting and cell repellent PLL-g-PEG&#8211;TRITC to a plasma-patterned PL-FITC coating. This was used for the registration of&#160;SH-SY5Y cells and also for patterning an additional fibronectin layer for LUHMES cells. Figure and legend reproduced with permission of the Royal Society of Chemistry (RSC)12. <a href="https://www.jove.com/files/ftp_upload/51389/51389fig3highres.jpg" target="_blank">Click here to view larger image.</a>
<ol>
	<li>Water Masking for In situ Biomaterial Patterning by Plasma Stenciling
	<ol>
		<li>Shortly after plasma bonding pipette a 0.5 &#956;l&#160;volume of PL or PO (100&#160;&#956;g/ml in 1x PBS) into the bottom center port. Within a matter of seconds the entire microfluidic circuit is filled by capillary action. Leave for a few minutes to fully coat the hydrophilic PDMS surfaces with the polyamines.</li>
		<li>Remove unbound polyamine molecules by aspiration-driven washing with 1x PBS (see Figure 2C) for 1 min.</li>
		<li>Remove all PBS from the ports and heat the device by illumination with a microscope. This increases evaporation from the ports for rapid water mask formation. The water mask (illustrated in Figure 3A and documented in Figure 3B) is established within ~5 min.</li>
		<li>Inspect the device by microscopy to ensure water masking is complete. Please note reservoir structures upstream of the array site are included in the mask design to prolong the stability of the water mask (~15 min).</li>
		<li>Insert stainless steel pins into the ports (Figure 2B). These are used to couple the plasma generated by an atmospheric pressure hand-held corona discharge or Tesla generator (e.g.,&#160;operating at 2 MHz, 30 kV signal)19,20 into the vacant microfluidic channels. Point the tip of the plasma source close to the tip of one pin and plasma treat the microchannel for &#8804;1 sec. Use low light conditions to observe plasma patterning.</li>
		<li>Remove the water mask with a wash step. Use parallel aspiration from the upper 3 ports. Use 4 equal lengths of silicone tubing connected by a 4-way adaptor to an aspiration pump to achieve parallel aspiration. Start aspiration and dispense PBS into the bottom 3 ports to draw PBS through the microfluidic circuit. In this manner a PDMS-polyamine material contrast is produced (see Figure 3C). Evacuate PBS from the microfluidic circuit and leave for several hours to restore the native, hydrophobic state. This renders the PDMS fully non-permissive to cell adhesion.</li>
		<li>Should additional adhesion materials, such as laminin or fibronectin, be needed the following steps are required and should be done immediately after removing the water mask: Pipette a 10&#160;&#956;l volume of the graft co-polymer poly-L-lysine&#8211;poly(ethylene glycol) (PLL-g-PEG, 100&#160;&#956;g/ml, in PBS) into both bottom flanking channels, followed by aspiration from the upper flanking channels for 2 min.</li>
		<li>Without interrupting the flow, remove excess PLL-g-PEG by exchange with a PBS buffer. This selectively coats the non-polyamine-coated regions with the PEG moiety acting to prevent protein adsorption and cell adhesion17,21.</li>
		<li>Adhesion proteins such as laminin or fibronectin can then be co-localized with the poly-amine coating: Pipette 10&#160;&#956;l volumes of these ECM proteins (typically 10&#160;&#956;g/ml) into all three bottom ports. Aspirate from the top three ports until the device is filled with the protein solution and incubate for 1 hr to produce a good quality coating.</li>
		<li>Remove excess proteins by aspirating PBS from the bottom 3 ports.</li>
	</ol>
	</li>
	<li>Microfluidic Single Neuron Arraying
	<ol>
		<li>Prime the devices with appropriate media. Add 20&#160;&#956;l volumes to the bottom 3 ports and by parallel aspiration from the upper three ports rapidly fill the microfluidic circuit with media. Use a 4-way tubing connector to couple the 3 upper ports via equal-length silicone tubing to an aspiration pump.</li>
		<li>Add 20&#160;&#956;l of a disaggregated cell suspension (1 x 106 cells/ml) to each of the flanking inlet ports (0, in Figure 1).</li>
		<li>Equip an aspiration pump with a flow regulator or use a syringe for gentle aspiration from the upper 3 ports. Complete neuron arraying typically requires 1 min.</li>
		<li>Harvest excess neurons remaining in the 4 flanking ports with a pipette for arraying in subsequent microfluidic devices.</li>
		<li>Remove the tubing. Fill all ports with media and place the device in the incubator. Within a few hours the cells become fully adherent on the biomaterial pattern.</li>
	</ol>
	</li>
	<li>Neuron Culture, Selective Fluidic Treatment, and Immunostaining
	<ol>
		<li>Exchange the media by periodic perfusion using hydrostatic feed (a 2 mm column height difference is appropriately slow).</li>
		<li>Alternatively, submerge the microfluidic device in media, exchanging the media periodically (2&#8211;3 days) as with standard cell cultures.</li>
		<li>Selective fluidic treatments to either neuron culture compartment or the central neurite outgrowth compartment are achieved as follows: Dispense 20&#160;&#956;l of the test substance into the bottom port of the channel of interest, and aspirate from the port directly above. For lengthy treatments reduce the flow rate using a flow regulator and replenish the test substance as required.</li>
		<li>Immunostaining protocols vary for each molecular target and associated reagents. In order to counter the potential risk of shear-induced damage to the neurons a slow flow rate was used. This results in increased application times to fully perfuse the system, extending normal protocol times. Hydrostatic perfusion using ports with differing column heights (e.g. ,by 2 mm) was used to deliver reagents. More rapid reagent addition and removal using a pump can be used to reduce experimental timescales. Connect an aspiration pump to the upper 3 ports to draw reagents from the bottom 3 ports.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

The microfluidic arraying technique is the first of its kind that enables precision single neuron handling for establishing minimalistic cultures. Coupled with the in situ biomaterial patterning method, a powerful approach to align cell patterns with microfluidic structures, these minimalistic cultures have high intercompartment connectivity levels with reduced local entanglement. These features can be used for the efficient study of inter-compartment transmission, and with simple design modifications have the p...

Neuronal tissue is highly complex; a heterogeneous cellular mixture that is spatially ordered within defined layers and compartments and with plastic connectivity via cell contacts and especially via axon and dendrite outgrowths. New techniques are required to confer greater experimental freedom to gain deeper insights and unravel mechanisms central to disease, development, and healthy function. The Campenot chamber1,2 and more recently microfabricated embodiments3,4 can be used for the ex vivo preparation of networked neuronal co-cultures with the ability to selectively perturb the different somatic populations and also their neurite outgrowths. These microfluidic devices have for example been used to study axon degeneration and regeneration following chemical5,6 or laser axotomy6-8, tauopathy9, viral dissemination10,11, and mRNA localization in axons4.

To extend the reach of the neurobiologists, technological developments are required to prepare minimalistic neuronal co-cultures. This enables disentanglement of the neuronal network for the investigation of the system with single cell and sub-cellular precision. The requirement for minimal cell numbers opens the possibility to analyze rare cell types, including dopaminergic substantia nigra cells relevant to Parkinson&#39;s disease, spiral ganglia from the ear, peripheral neurons, and stem cells. Beyond this, cellular economy is relevant to the 3Rs initiative. Using these microfluidic platforms, large-scale toxicity screens or other high throughput, data rich experimental series requiring animal neurons can now be considered.

In this paper protocols for the fabrication and use of a microfluidic device are described. Microfluidic arraying in combination with an in situ biomaterial patterning method can be used for the registration of highly interconnected neuronal co-cultures using minimal cell numbers. Microfluidic arraying is based on a differential flow approach12&#8211;15, whereby microstructured traps are positioned within a fluidic circuit (illustrated along with SEM images in Figure 1). The path 0 &#8594; 1 has the lower fluidic resistance (R2 &#62; R1) for transporting neurons to a linear array of microstructured apertures &#8211; the inlets to the neurite outgrowth channels. Occupancy of the trap by a single cell locally impedes the flow to divert the streamlines for trapping subsequent cells in neighboring traps. Complete occupancy of the traps in the array switches the fluidic ratio (R1 &#62; R2) to divert the streamlines into the serpentine path (0 &#8594; 2) to produce a bypass mode of operation for the removal of excess neurons.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51389/51389fig1highres.jpg" width="500" /> 
Figure 1. Microfluidic Circuit. A) The differential resistance fluidic circuit for single neuron arraying, with flanking culture chambers interconnected by neurite outgrowth channels. B,C) SEM images of the bilayer compartmentalized neuron co-culture array with meniscus pinning micropillars. With this design, trident-shaped neuron trapping structures were used to promote fasciculation of the neurite outgrowths. Figure and legend reproduced with permission of the Royal Society of Chemistry (RSC)12. <a href="https://www.jove.com/files/ftp_upload/51389/51389fig1highres.jpg" target="_blank">Click here to view larger image.</a>
The preparation of micropatterned neuronal networks on planar substrates can readily be achieved (for examples from our group, see Frimat et al.16 and Heike et al.17). However, encapsulating bioactive material patterns within PDMS devices and with the requirement of micrometer-scale alignment of these to the microfluidic channels poses a major technical challenge. In section 3.1 a protocol for the in-chip, or in situ, preparation of biomaterial patterns is presented. These patterns enable neuron registration during lengthy culture timescales and promote outgrowths between compartments. Meniscus-pinning microstructures are used to align a so-called water mask with the neuron arraying sites and neurite outgrowth channels. The water mask protects adhesion molecule coatings during plasma treatment, whereas exposed surfaces are disintegrated to define the biomaterial pattern. In addition, protocols are provided for cell culture and for fluidic isolation necessary for the selective treatment of the different co-culture compartments.
The protocols are designed to leverage the user-friendly principles of soft lithography for the replication of poly(dimethylsiloxane) (PDMS) microfluidic devices18. Similarly, in situ biomaterial patterning is straightforward, exploiting evaporation and surface tension phenomena, and only requiring an inexpensive hand-held plasma source. The microfluidic circuit effectively programs cell loading and compartment specific treatments making these operations simply a matter of dispensing materials into the correct bottom port and aspirating from above. In this manner, it is intended to give the neurobiologists the freedom to prepare and use microfluidic devices in their own laboratories.

<table border="0" cellpadding="0" cellspacing="0" width="857">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">PDMS Sylgard 184</td>
			<td style="width:163px;">Dow Corning</td>
			<td style="width:143px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PDMS Elastosil RT 601</td>
			<td style="width:163px;">Wacker</td>
			<td style="width:143px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">Coverslips</td>
			<td style="width:163px;">VWR</td>
			<td>630-1590</td>
			<td>130-160 mm thick</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">3 mm Biopsy Punches</td>
			<td style="width:163px;">Kai Medical</td>
			<td style="width:143px;"></td>
			<td>Handle with care - extremely sharp</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tygon Tubing</td>
			<td style="width:163px;">Fisher Scientific</td>
			<td style="width:143px;">S-50-HL</td>
			<td>1.65 mm ID; 3.35 mm OD</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">1 mL Syringe (Inkjekt and Omnifix)</td>
			<td style="width:163px;">Braun</td>
			<td style="width:143px;">6064204</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:235px;">4-Way Tubing Connector</td>
			<td style="width:163px;">VWR or Fisher Scientific</td>
			<td style="width:143px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">Flow Regulator</td>
			<td style="width:163px;">Harvard Apparatus</td>
			<td style="width:143px;">722645</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:235px;">0.5 mm Pins</td>
			<td style="width:163px;">Dressmaking Departments</td>
			<td style="width:143px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:235px;">PLL-g-PEG</td>
			<td style="width:163px;">SuSoS</td>
			<td style="width:143px;">PLL(20)-g[3.5]-PEG(5)</td>
			<td>Stability in storage can be an issue</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">poly-D-lysine</td>
			<td style="width:163px;">Sigma-Aldrich</td>
			<td style="width:143px;">P6407</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">poly-lysine-FITC</td>
			<td style="width:163px;">Sigma-Aldrich</td>
			<td style="width:143px;">P3069</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">poly-ornithine</td>
			<td style="width:163px;">Sigma-Aldrich</td>
			<td style="width:143px;">P4957</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">fibronectin</td>
			<td style="width:163px;">Sigma-Aldrich</td>
			<td style="width:143px;">F2006</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">laminin</td>
			<td style="width:163px;">Sigma-Aldrich</td>
			<td style="width:143px;">L2020</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">PBS</td>
			<td style="width:163px;">Sigma-Aldrich</td>
			<td style="width:143px;">P4417</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">Inverted Fluorescent Microscope</td>
			<td style="width:163px;"></td>
			<td style="width:143px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:235px;">Example Aspiration Pump</td>
			<td style="width:163px;">KNF Neuberger, Laboport</td>
			<td style="width:143px;">N811KVP</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:235px;">Hand Held Corona Discharge Device</td>
			<td style="width:163px;">Leybold-Heraeus, USA</td>
			<td style="width:143px;">VP23</td>
			<td>May not comply with your country&#39;s safety standards</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">Femto Plasma Oven</td>
			<td style="width:163px;">Diener Electronic</td>
			<td style="width:143px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:235px;">Vacuum Dessicator or Centrifuge</td>
			<td style="width:163px;"></td>
			<td style="width:143px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Water masking exploits surface tension. The pressure tolerance at the air-liquid interface scales inversely with the radius of curvature [<img fo:content-width="1in" src="/files/ftp_upload/51389/51389eq1.jpg" width="113" />, producing highly stable interfaces at microfluidic dimensions. The micropillars effectively anchor the water mask in place. Water mask formation is driven by evaporation from the microfluidic ports, with the rate increased by heating. Using the heat from low magnification (4X - 10X) microscopy illumi...

Watch this Scientific Journal Video about Preparation of Neuronal Co-cultures with Single Cell Precision at JoVE.com

Preparation of Neuronal Co-cultures 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 ...

preparation-of-neuronal-co-cultures-with-single-cell-precision

Research

JoVE Journal

Neuroscience

13.6K Views.  ISAS.  Sono descritti protocolli per singolo neurone microfluidica arraying e mascheramento acqua per il patterning plasma in-chip di rivestimenti biomateriale. Co-colture altamente interconnessi possono essere preparati utilizzando gli ingressi di cellule minimali. 

Video: Preparation of Neuronal Co-cultures with Single Cell Precision

Derivation of Enriched Oligodendrocyte Cultures and Oligodendrocyte/Neuron Myelinating Co-cultures from Post-natal Murine Tissues

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications for understanding the pathogenesis of demyelinating diseases such as Multiple Sclerosis (MS). Cellular development is commonly studied with primary cell culture models. Primary cell culture facilitates the evaluation of a given cell type by providing a controlled environment, free of the extraneous variables that are present in vivo. While OL cultures derived from rats have provided a vast amount of insight into OL biology, similar efforts at establishing OL cultures from mice has been met with major obstacles. Developing methods to culture murine primary OLs is imperative in order to take advantage of the available transgenic mouse lines. 
Multiple methods for extraction of OPCs from rodent tissue have been described, ranging from neurosphere derivation, differential adhesion purification and immunopurification 1-3. While many methods offer success, most require extensive culture times and/or costly equipment/reagents. To circumvent this, purifying OPCs from murine tissue with an adaptation of the method originally described by McCarthy &amp; 
de Vellis 2 is preferred. This method involves physically separating OPCs from a mixed glial culture derived from neonatal rodent cortices. The result is a purified OPC population that can be differentiated into an OL-enriched culture. This approach is appealing due to its relatively short culture time and the unnecessary requirement for growth factors or immunopanning antibodies. 
While exploring the mechanisms of OL development in a purified culture is informative, it does not provide the most physiologically relevant environment for assessing myelin sheath formation. Co-culturing OLs with neurons would lend insight into the molecular underpinnings regulating OL-mediated myelination of axons. For many OL/neuron co-culture studies, dorsal root ganglion neurons (DRGNs) have proven to be the neuron type of choice. They are ideal for co-culture with OLs due to their ease of extraction, minimal amount of contaminating cells, and formation of dense neurite beds. While studies using rat/mouse myelinating xenocultures have been published 4-6, a method for the derivation of such OL/DRGN myelinating co-cultures from post-natal murine tissue has not been described. Here we present detailed methods on how to effectively produce such cultures, along with examples of expected results. These methods are useful for addressing questions relevant to OL development/myelinating function, and are useful tools in the field of neuroscience.

This article describes methods to derive enriched populations of murine oligodendrocyte precursor cells (OPCs) in primary culture, which differentiate to produce mature oligodendrocytes (OLs). In addition, this report describes techniques to produce murine myelinating co-cultures by seeding mouse OPCs onto a neurite bed of mouse dorsal root ganglion neurons (DRGNs).

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications ...

Single Cell Measurement of Dopamine Release with Simultaneous Voltage-clamp and Amperometry

After its release into the synaptic cleft, dopamine exerts its biological properties via its pre- and post-synaptic targets1. The dopamine signal is terminated by diffusion2-3, extracellular enzymes4, and membrane transporters5. The dopamine transporter, located in the peri-synaptic cleft of dopamine neurons clears the released amines through an inward dopamine flux (uptake). The dopamine transporter can also work in reverse direction to release amines from inside to outside in a process called outward transport or efflux of dopamine5. More than 20 years ago Sulzer et al. reported the dopamine transporter can operate in two modes of activity: forward (uptake) and reverse (efflux)5. The neurotransmitter released via efflux through the transporter can move a large amount of dopamine to the extracellular space, and has been shown to play a major regulatory role in extracellular dopamine homeostasis6. Here we describe how simultaneous patch clamp and amperometry recording can be used to measure released dopamine via the efflux mechanism with millisecond time resolution when the membrane potential is controlled. For this, whole-cell current and oxidative (amperometric) signals are measured simultaneously using an Axopatch 200B amplifier (Molecular Devices, with a low-pass Bessel filter set at 1,000 Hz for whole-cell current recording). For amperometry recording a carbon fiber electrode is connected to a second amplifier (Axopatch 200B) and is placed adjacent to the plasma membrane and held at +700 mV. The whole-cell and oxidative (amperometric) currents can be recorded and the current-voltage relationship can be generated using a voltage step protocol. Unlike the usual amperometric calibration, which requires conversion to concentration, the current is reported directly without considering the effective volume7. Thus, the resulting data represent a lower limit to dopamine efflux because some transmitter is lost to the bulk solution.

The amperometric technique measures dopamine release from a single cell by detecting the oxidative current produced by spontaneous dopamine oxidization. Simultaneous voltage clamp and amperometry methodology reveal the mechanistic relationship between the overall "activity" of dopamine transporter and the regulatory role of this activity on the reverse transport of dopamine.

After its release into the synaptic cleft, dopamine exerts its biological properties via its pre- and post-synaptic targets1. The dopamine signal is ...

Optical Recording of Suprathreshold Neural Activity with Single-cell and Single-spike Resolution

Signaling of information in the vertebrate central nervous system is often carried by populations of neurons rather than individual neurons. Also propagation of suprathreshold spiking activity involves populations of neurons. Empirical studies addressing cortical function directly thus require recordings from populations of neurons with high resolution. Here we describe an optical method and a deconvolution algorithm to record neural activity from up to 100 neurons with single-cell and single-spike resolution. This method relies on detection of the transient increases in intracellular somatic calcium concentration associated with suprathreshold electrical spikes (action potentials) in cortical neurons. High temporal resolution of the optical recordings is achieved by a fast random-access scanning technique using acousto-optical deflectors (AODs)1. Two-photon excitation of the calcium-sensitive dye results in high spatial resolution in opaque brain tissue2. Reconstruction of spikes from the fluorescence calcium recordings is achieved by a maximum-likelihood method. Simultaneous electrophysiological and optical recordings indicate that our method reliably detects spikes (&gt;97% spike detection efficiency), has a low rate of false positive spike detection (&lt; 0.003 spikes/sec), and a high temporal precision (about 3 msec) 3. This optical method of spike detection can be used to record neural activity in vitro and in anesthetized animals in vivo3,4.

Understanding the function of the vertebrate central nervous system requires recordings from many neurons because cortical function arises on the level of populations of neurons. Here we describe an optical method to record suprathreshold neural activity with single-cell and single-spike resolution, dithered random-access scanning. This method records somatic fluorescence calcium signals from up to 100 neurons with high temporal resolution. A maximum-likelihood algorithm deconvolves the underlying suprathreshold neural activity from the somatic fluorescence calcium signals. This method reliably detects spikes with high detection efficiency and a low rate of false positives and can be used to study neural populations in vitro and in vivo.

Signaling of information in the vertebrate central nervous system is often carried by populations of neurons rather than individual neurons. Also ...

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

Much information about the coupling of presynaptic ionic currents with the release of neurotransmitter has been obtained from invertebrate preparations, most notably the squid giant synapse1. However, except for the preparation described here, few vertebrate preparations exist in which it is possible to make simultaneous measurements of neurotransmitter release and presynaptic ionic currents. Embryonic Xenopus motoneurons and muscle cells can be grown together in simple culture medium at room temperature; they will form functional synapses within twelve to twenty-four hours, and can be used to study nerve and muscle cell development and synaptic interactions for several days (until overgrowth occurs). Some advantages of these co-cultures over other vertebrate preparations include the simplicity of preparation, the ability to maintain the cultures and work at room temperature, and the ready accessibility of the synapses formed2-4. The preparation has been used widely to study the biophysical properties of presynaptic ion channels and the regulation of transmitter release5-8. In addition, the preparation has lent itself to other uses including the study of neurite outgrowth and synaptogenesis9-12, molecular mechanisms of neurotransmitter release13-15, the role of diffusible messengers in neuromodulation16,17, and in vitro synaptic plasticity18-19.

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

This video demonstrates the procedures used to grow primary cultures of embryonic Xenopus nerve and muscle cells and the usefulness of this preparation for making simultaneous pre- and post-synaptic patch clamp recordings.

Much information about the coupling of  presynaptic ionic currents with the release of neurotransmitter has been obtained  from invertebrate preparations, ...

A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges.&#160;Methods for performing this technique and the equipment necessary are described.&#160;Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

Investigating Functional Regeneration in Organotypic Spinal Cord Co-cultures Grown on Multi-electrode Arrays

Adult higher vertebrates have a limited potential to recover from spinal cord injury. Recently, evidence emerged that propriospinal connections are a promising target for intervention to improve functional regeneration. So far, no in vitro model exists that grants the possibility to examine functional recovery of propriospinal fibers. Therefore, a representative model that is based on two organotypic spinal cord sections of embryonic rat, cultured next to each other on multi-electrode arrays (MEAs) was developed. These slices grow and, within a few days in vitro, fuse along the sides facing each other. The design of the used MEAs permits the performance of lesions with a scalpel blade through this fusion site without inflicting damage on the MEAs. The slices show spontaneous activity, usually organized in network activity bursts, and spatial and temporal activity parameters such as the location of burst origins, speed and direction of their propagation and latencies between bursts can be characterized. Using these features, it is also possible to assess functional connection of the slices by calculating the amount of synchronized bursts between the two sides. Furthermore, the slices can be morphologically analyzed by performing immunohistochemical stainings after the recordings. 
Several advantages of the used techniques are combined in this model: the slices largely preserve the original tissue architecture with intact local synaptic circuitry, the tissue is easily and repeatedly accessible and neuronal activity can be detected simultaneously and non-invasively in a large number of spots at high temporal resolution. These features allow the investigation of functional regeneration of intraspinal connections in isolation in vitro in a sophisticated and efficient way.

Here we present a protocol that is based on spinal cord slices cultured on multi-electrode arrays to study functional regeneration of propriospinal connections in vitro.

Adult higher vertebrates have a limited potential to recover from spinal cord injury. Recently, evidence emerged that propriospinal connections are a ...

Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and awake animals. Therefore, methods allowing the manipulation of synaptic systems in awake animals are required in order to determine the contribution of synaptic inputs to neuronal processing unaffected by anesthetics. Here, we present methodology for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. By combining these procedures, we performed microiontophoretic injections of gabazine to selectively block GABAA receptors in neurons of the inferior colliculus of head-restrained mice. Gabazine successfully modified neural response properties such as the frequency response area and stimulus-specific adaptation. Thus, we demonstrate that our methods are suitable for recording single-unit activity and for dissecting the role of specific neurotransmitter receptors in auditory processing.
The main limitation of the described procedure is the relatively short recording time (~3 hr), which is determined by the level of habituation of the animal to the recording sessions. On the other hand, multiple recording sessions can be performed in the same animal. The advantage of this technique over other experimental procedures used to manipulate the level of neurotransmission or neuromodulation (such as systemic injections or the use of optogenetic models), is that the drug effect is confined to the local synaptic inputs to the target neuron. In addition, the custom-manufacture of electrodes allows adjustment of specific parameters according to the neural structure and type of neuron of interest (such as the tip resistance for improving the signal-to-noise ratio of the recordings).

We present methods for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. This technique allows the detailed analysis of putative local synaptic inputs to the neuron of interest.

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and ...

In Utero Electroporation Approaches to Study the Excitability of Neuronal Subpopulations and Single-cell Connectivity

The nervous system is composed of an enormous range of distinct neuronal types. These neuronal subpopulations are characterized by, among other features, their distinct dendritic morphologies, their specific patterns of axonal connectivity, and their selective firing responses. The molecular and cellular mechanisms responsible for these aspects of differentiation during development are still poorly understood.
Here, we describe combined protocols for labeling and characterizing the structural connectivity and excitability of cortical neurons. Modification of the in utero electroporation (IUE) protocol allows the labeling of a sparse population of neurons. This, in turn, enables the identification and tracking of the dendrites and axons of individual neurons, the precise characterization of the laminar location of axonal projections, and morphometric analysis. IUE can also be used to investigate changes in the excitability of wild-type (WT) or genetically modified neurons by combining it with whole-cell recording from acute slices of electroporated brains. These two techniques contribute to a better understanding of the coupling of structural and functional connectivity and of the molecular mechanisms controlling neuronal diversity during development. These developmental processes have important implications on axonal wiring, the functional diversity of neurons, and the biology of cognitive disorders.

In Utero Electroporation Approaches to Study the Excitability of Neuronal Subpopulations and Single-cell Connectivity

This manuscript provides protocols that use in utero electroporation (IUE) to describe the structural connectivity of neurons at the single-cell level and the excitability of fluorescently labeled neurons. Histology is used to characterize dendritic and axonal projections. Whole-cell recording in acute slices is used to investigate excitability.

The nervous system is composed of an enormous range of distinct neuronal types. These neuronal subpopulations are characterized by, among other features, ...

Neuron-Macrophage Co-cultures to Activate Macrophages Secreting Molecular Factors with Neurite Outgrowth Activity

There is strong evidence that macrophages can participate in the regeneration or repair of injured nervous system. Here, we describe a protocol in which macrophages are induced to produce conditioned medium (CM) that promotes neurite outgrowth. Adult dorsal root ganglion (DRG) neurons are acutely dissociated and plated. After the neurons are stably attached, peritoneal macrophages are co-cultured on a cell culture insert overlaid on the same well. Dibutyryl cyclic AMP (db-cAMP) is applied to the co-cultures for 24 h, after which the cell culture insert containing the macrophages is moved to another well to collect CM for 72 h. The CM from the co-cultures treated with db-cAMP, when applied to a separate adult DRG neuron culture, exhibits robust neurite outgrowth activity. The CM obtained from the db-cAMP-treated cultures consisting of single cell type alone, either DRG neuron or peritoneal macrophage, did not exhibit neurite outgrowth activity. This indicates that the interaction between neurons and macrophages is indispensable for the activation of macrophages secreting molecular factors with neurite outgrowth activity into CM. Thus, our co-culture paradigm will also be useful to study intercellular signaling in the neuron-macrophage interaction to stimulate the macrophages to be endowed with a pro-regenerative phenotype.

The current protocol presents experimental procedures to stimulate cultured macrophages to be endowed with capacity to release molecular factors that promote neurite outgrowth. Treatment of cAMP to the neuron-macrophage co-cultures induces the macrophages to produce conditioned medium that possesses strong neurite outgrowth activity.

There is strong evidence that macrophages can participate in the regeneration or repair of injured nervous system. Here, we describe a protocol in which ...