This work was supported by MEXT KAKENHI on Innovative Areas Mesoscopic Neurocircuitry 23115102 and Dynamic Regulation of Brain Function by Scrap and Build System 16H06460 to N.Y.&#160;We also thank Gabriel Hand for critical reading.

All experiments were performed according to the guidelines established by the animal welfare committees of Osaka University and the Japan Neuroscience Society.
1. Organotypic cocultures of the thalamus and cerebral cortex
Note: For the detailed procedure, refer to the original publications7,8,13. All procedures should be performed under sterile conditions. Sprague-Dawley (SD) rats are used for neuronal cultures.
<ol>
	<li>Preparations of reagents 
	Note: Volumes of the following reagents are for one rat brain.
	<ol>
		<li>Prepare 10x modified N2 Supplement13: Insulin (50 &#956;g/mL), progesterone (200 nM), hydrocortisone (200 nM), sodium selenite (300 nM), transferrin (1 mg/mL), putrescine (1 mM), glucose (60 mg/mL)</li>
		<li>Prepare serum-containing culture medium (10 mL): 85% DMEM/F12, 10% 10x modified N2 supplement, 5% fetal bovine serum (FBS)</li>
		<li>Prepare serum-free culture medium (20 mL): 90% DMEM/F12, 10% 10x modified N2 supplement</li>
		<li>Prepare rat tail collagen solution (0.5 mL). In brief, take tender fibers from a rat tail under sterile conditions and incubate in acetic acid solution overnight. After centrifugation, collect the supernatant (a few mg/mL) and store in a refrigerator13.</li>
	</ol>
	</li>
	<li>Preparation of culture dishes
	<ol>
		<li>Place a membrane insert into a 35 mm Petri dish.</li>
		<li>Add 40 - 50 &#956;L of rat tail collagen solution to the membrane, and spread the collagen solution around the center.</li>
		<li>Air dry the membrane completely in a clean bench.</li>
		<li>Put 1.5 - 2 mL of the serum-containing culture medium into the dish, so that the membrane is soaked with the medium.</li>
		<li>Place the culture dish in an incubator (37 &#176;C, humidified 95% air, and 5% CO2) before plating.</li>
	</ol>
	</li>
	<li>Preparation of cortical slices
	<ol>
		<li>Sterilize all surgical instruments with 70% ethanol for 10 min or longer.</li>
		<li>Dissect the whole brain quickly from a postnatal day (P) 2 rat under ice-cold anesthesia (immersed in ice-chips for a few minutes).</li>
		<li>Place the brain into a 100 mm Petri dish containing 100 mL of cold Hanks&#8217; balanced salt solution.</li>
		<li>Remove the pia matter from the brain with fine forceps, and cut cortical slices (300 - 400 &#956;m thickness) from the visual and somatosensory cortex with microsurgical scissors (for detail, see Figure 1). 
		Note: The thickness of cortical slices is roughly estimated by the scale of grids under a binocular microscope. 
		Note: Alternatively, cortical slices can be made with a vibratome under sterile conditions.</li>
		<li>Transfer a few cortical slices onto the membrane insert with a disposable plastic pipette.</li>
		<li>Adjust the positions of the slices near the center of the culture insert with a fire-polished Pasteur pipette (for details, see8,13) and remove excess medium from the insert. 
		Note: Adjust the level of the medium slightly below the surface of the slices so that the slices can receive a sufficient supply of both the gas and medium. This is crucial for cell viability.</li>
		<li>Maintain the slices in the serum-containing culture medium at 37 &#176;C in an environment of humidified 95% air and 5% CO2, and culture alone for one day before a thalamic block is taken and placed next to it.</li>
	</ol>
	</li>
	<li>Preparation of thalamic blocks
	<ol>
		<li>Sterilize all surgical instruments with 70% ethanol for 10 min or longer.</li>
		<li>Anesthetize a pregnant rat (embryonic day 15) by intraperitoneal injection of pentobarbital (50 mg/kg) containing anesthetic.</li>
		<li>Dissect each embryo from the uterine horns, and place the embryos into a 100 mm Petri dish containing 100 mL of cold Hanks&#8217; balanced salt solution.</li>
		<li>Under a binocular microscope, remove the brain from each embryo and cut thalamic blocks which mainly contain the ventrobasal complex and lateral geniculate nucleus (approximately 0.5 mm x 3)7 using microsurgical scissors (see Figure 1).</li>
	</ol>
	</li>
	<li>Culture and maintain organotypic cocultures
	<ol>
		<li>Using a Pasteur pipette, place the thalamic block next to the ventricular surface of the cortical slice which has been placed on the membrane filter.</li>
		<li>Maintain the cocultures in the serum-containing culture medium at 37 &#176;C in an environment of humidified 95% air and 5% CO2.</li>
		<li>Exchange half of the medium with the serum-free culture medium after two days in culture. Thereafter, exchange the medium every 2 - 3 days. 
		Note: The level of the medium should be checked after the medium exchange, as it is important for cell survival.</li>
	</ol>
	</li>
</ol>
2. Electroporation
All procedures should be performed under sterile conditions.
Note: Perform electroporation one day after the preparation of the thalamic blocks to prevent detachment of the blocks.
<ol>
	<li>Plasmids
	<ol>
		<li>Make a mixture of plasmids as follows: pCAGGS-DsRed 2 &#956;g/&#956;L; pCAG-SYP-EGFP 3.5 &#956;g/&#956;L. 
		Note: Purify the both plasmids using an endotoxin-free Maxiprep kit and dissolve in Hanks&#8217; balanced salt solution.</li>
	</ol>
	</li>
	<li>Electroporation equipment setup 
	The electrical instruments such as a stimulator and an isolator should be connected as shown in Figure 2.
	<ol>
		<li>Connect the stimulator to the biphasic isolator.</li>
		<li>Connect an electrode (a silver wire 0.2 mm diameter) to the negative terminal of the isolator.</li>
		<li>Connect an electrode (a silver wire 1 mm diameter) in series to a 100 &#937; resistor and the positive terminal of the isolator.</li>
		<li>Connect the amplifier to the oscilloscope to monitor whether electrical currents are passed across the resistor by measuring the voltage with the amplifier.</li>
	</ol>
	</li>
	<li>Preparation of glass pipettes 
	Note: Two types of glass pipettes are used for ejection of the plasmid solution and application of electrical pulses.
	<ol>
		<li>Make the pipettes from glass capillaries with an electrode puller.</li>
		<li>Break the tip for the ejection pipette (the tip diameter should be 20 - 50 &#956;m).</li>
		<li>Grind and polish the tip of the electrode pipette with sandpaper followed by fire-polishing (the inner diameter should be 50 - 200 &#956;m). 
		Note: The electrode tip should be flat and smooth, as it directly touches the tissue.</li>
	</ol>
	</li>
	<li>Procedure of electroporation
	<ol>
		<li>Connect the ejection and electrode pipettes to the manipulators.</li>
		<li>Insert a silver wire (0.2 mm diameter) into the electrode pipette.</li>
		<li>Attach the ejection pipette to a 1 mL syringe with a plastic tube.</li>
		<li>Take up &#62;5 &#956;L of plasmid solution into the ejection pipette by syringe suction.</li>
		<li>Put the coculture on the electroporation stage, and insert the 1 mm diameter electrode into the culture medium.</li>
		<li>Place the ejection pipette on the thalamic explant.</li>
		<li>Push the syringe manually after the tip touches the surface of the thalamic block. 
		Note: About 0.5 &#956;L of the plasmid solution is usually used per one thalamic block.</li>
		<li>Place the electrode pipette on the thalamic block immediately after retraction of the ejection pipette.</li>
		<li>Deliver electrical pulses (50 - 400 &#956;A, 3 to 5 trains of 200 square pulses of 1 ms duration at 200 Hz). Monitor the amplitude and the number of trains on the oscilloscope. 
		Note: The amplitude of electrical currents depends on the inner diameter of an electrode pipette13.</li>
		<li>Retract the electrode pipette and return the dish to the incubator.</li>
	</ol>
	</li>
	<li>Microscopic observation
	<ol>
		<li>Put the culture dish on a microscopic stage of an upright confocal microscope equipped with two filter sets for EGFP (excitation, 488 nm; emission, 515 nm) and DsRed (excitation, 514 nm; emission, 565 nm).</li>
		<li>Take images of 10 - 25 optical sections at 1 - 7 &#181;m-step sizes with a 20X&#160;long-working distance objective lens. Select individually distinguishable labeled axons which express both DsRed and SYP-EGFP. 
		Note: The observed area is approximately 0.7 x 0.7 mm (corresponding to 1024 x 1024 pixels, that is, 0.68 &#956;m/pixel).</li>
		<li>Capture images every 24 h for daily imaging. Complete within 10 min at room temperature, and return the cultures to the incubator. For short-interval time lapse imaging, keep the culture in a culture chamber, which maintains the environment of humidified 95% air and 5% CO2 at 37 &#176;C.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The current protocol is also a powerful tool to study developmental aspects of growing axons other than of the TC projection11. For instance, a combination of cortical slice culture and the electroporation technique allows visualizing individual axonal morphology of cortical neurons and long term observation9,18.
By using the current protocol, the roles of interesting genes in axon branching and synapse formation can also be analyzed by co-expression of fluorescent proteins and the interest genes. Typically, over 90% of fluorescent protein expressing cells are co-transfected with a second plasmid when the molar ratio of the plasmid solutions is adjusted to 1:218,19. However, the optimal ratio may be different as co-transfection efficacy and expression level are varied among vectors.
Although the current protocol is efficient for time-lapse experiments, there are some technical problems. For daily imaging, a coculture was placed on a microscopic stage every day. Although axonal development did not seem to be affected seriously by daily imaging10, each observation should be completed within 10 min to minimize evaporation of culture solution and changes in pH of the culture medium. Alternatively, it would be better to maintain the temperature, humidity and pH of the cultures on the microscopic stage12.

The thalamocortical (TC) projection in the mammalian brain is a suitable system to investigate axon guidance and targeting mechanisms. During development, sensory TC axons grow in the cortical plate, and form branches and synapses preferentially in layer IV of the primary sensory areas in the cerebral cortex1,2. Even after establishment of fundamental connections, axonal arbors and synaptic terminals are remodeled depending on environmental changes3,4. However, how TC axon morphology is dynamically altered is poorly understood. One of the main reasons is the lack of an adequate technique to observe structural changes at a single cell level. Although recent developments in microscopy, such as two-photon microscopy, have allowed direct observation of living cortical neurons in vivo, there are still technical limitations for capturing the overall TC trajectories5,6. Therefore, in vitro methods for live imaging of TC axons would provide powerful tools for structural analyses of axon branching and synapse formation.
Our group for the first time established a static slice culture method with permeable membrane7. Using this method, a rat cortical slice was cocultured with a sensory thalamic block, and lamina-specific TC connections were recapitulated in this organotypic cocultures7,8. Sparse labeling with a fluorescent protein further allowed us to observe TC axon growth and branch formation9,10,11. Recently, we have developed a novel method for simultaneous imaging of branching and synapse formation of individual TC axons in the organotypic cocultures12. To visualize TC axons and presynaptic sites simultaneously, DsRed and EGFP-tagged synaptophysin (SYP-EGFP) were co-transfected into a small number of thalamic neurons by electroporation of the organotypic coculture. The current method facilitates morphological analysis of TC axons and allows for long-term observation, which can be used to show the causal relationship between axon branching and synapse formation.

<table><tbody><tr><td>DMEM/F12</td><td>GIBCO</td><td>11320-033</td><td></td></tr><tr><td>Hanks&amp;rsquo; balanced salt solution (HBSS)</td><td>Nissui</td><td>5905</td><td></td></tr><tr><td>Fetal bovine serum (FBS)</td><td>Thermo Scientific</td><td>SH30396-03</td><td>Hyclone</td></tr><tr><td>Insulin</td><td>Sigma</td><td>I6634</td><td></td></tr><tr><td>Progesterone</td><td>Sigma</td><td>P8783</td><td></td></tr><tr><td>Hydrocortisone</td><td>Sigma&amp;nbsp;</td><td>H0888</td><td></td></tr><tr><td>Sodium selenite</td><td>Wako Pure&lt;br/&gt; Chemical Industries</td><td>192-10843</td><td></td></tr><tr><td>Transferrin&amp;nbsp;</td><td>Sigma</td><td>T1147</td><td></td></tr><tr><td>Putrescine&amp;nbsp;</td><td>Sigma</td><td>P5780</td><td></td></tr><tr><td>Glucose</td><td>Wako&lt;br/&gt; Pure Chemical Industries</td><td>16806-25</td><td></td></tr><tr><td>35 mm petri dishes</td><td>Falcon</td><td>351008</td><td></td></tr><tr><td>Millicell-CM insert</td><td>Millipore</td><td>PICMORG50</td><td></td></tr><tr><td>100 mm petri dishes</td><td>BIO-BIK</td><td>I-90-20</td><td>petri dish sterrile</td></tr><tr><td>HiPure Plasmid Maxiprep Kit</td><td>Invitrogen</td><td>K210006</td><td></td></tr><tr><td>Disposable sterile plastic pipettes</td><td></td><td>202-IS</td><td>transfer pipets sterile</td></tr><tr><td>Glass capillary: OD 1.2 mm</td><td>Narishige</td><td>&amp;nbsp;G-1.2</td><td>inner diameter, 1.2 mm</td></tr><tr><td>Silver wire: 0.2 and 1 mm&amp;nbsp;</td><td>Nilaco</td><td>AG-401265 (diameter, 0.2 mm), AG-401485 (diameter, 1.0 mm)</td><td></td></tr><tr><td>1 mL syringe</td><td>Terumo</td><td>SS-01T</td><td></td></tr><tr><td>Stimulator&amp;nbsp;</td><td>A.M.P.I</td><td>Master 8</td><td></td></tr><tr><td>Biphasic isolator&amp;nbsp;</td><td>BAK ELECTRONICS</td><td>BSI-2</td><td></td></tr><tr><td>Amplifier&amp;nbsp;</td><td>A-M Systems</td><td>Model 1800</td><td></td></tr><tr><td>Oscilloscope</td><td>Hitachi</td><td>VC-6723</td><td></td></tr><tr><td>Manipulator</td><td>Narishige</td><td>SM-15</td><td></td></tr><tr><td>Micromanipulator</td><td>Narishige</td><td>MO-10</td><td></td></tr><tr><td>Stereomicroscope&amp;nbsp;</td><td>Olympus</td><td>SZ40</td><td></td></tr><tr><td>Universal stand&amp;nbsp;</td><td>Olympus</td><td>SZ-STU2</td><td></td></tr><tr><td>Light illumination system&amp;nbsp;</td><td>Olympus</td><td>LG-PS2, LG-DI, HLL301</td><td></td></tr><tr><td>Electrode puller&amp;nbsp;</td><td>Narishige</td><td>PC-10</td><td></td></tr><tr><td>Confocal microscope</td><td>Nikon</td><td>Digital eclipse C1 laser</td><td></td></tr><tr><td>x20 objective</td><td>Nikon</td><td>ELWD 20x/0.45</td><td></td></tr><tr><td>Culture chamber</td><td>Tokai Hit</td><td>UK A16-U</td><td></td></tr><tr><td>Sprague-Dawley (SD) rat</td><td>Japan SLC and Nihon-Dobutsu</td><td></td><td></td></tr><tr><td>Microsurgery scissors</td><td>Natsume&amp;nbsp;</td><td>MB-54-1</td><td></td></tr></tbody></table>

visualization of thalamocortical axon branching and synapse formation in organotypic cocultures

Axon branching and synapse formation are crucial processes for establishing precise neuronal circuits. During development, sensory thalamocortical (TC) axons form branches and synapses in specific layers of the cerebral cortex. Despite the obvious spatial correlation between axon branching and synapse formation, the causal relationship between them is poorly understood. To address this issue, we recently developed a method for simultaneous imaging of branching and synapse formation of individual TC axons in organotypic cocultures.
This protocol describes a method which consists of a combination of an organotypic coculture and electroporation. Organotypic cocultures of the thalamus and cerebral cortex facilitate gene manipulation and observation of axonal processes, preserving characteristic structures such as laminar configuration. Two distinct plasmids encoding DsRed and EGFP-tagged synaptophysin (SYP-EGFP) were co-transfected into a small number of thalamic neurons by an electroporation technique. This method allowed us to visualize individual axonal morphologies of TC neurons and their presynaptic sites simultaneously. The method also enabled long-term observation which revealed the causal relationship between axon branching and synapse formation.

The experiment described here aims to reveal the relationship between TC axon branching and synapse formation. To simultaneously visualize axonal trajectories and locations of presynaptic sites, single or a few thalamic cells in organotypic cocultures were transfected with two plasmids encoding SYP-EGFP and DsRed using electroporation. During the second week in culture, individually distinguishable TC axons were clearly labeled by DsRed (Figure 3). Only the axons that exhibited DsRed and SYP-EGFP were selected for observation. Labeled TC axons invaded the cortical slice and formed branches primarily in the upper layers, indicating that thalamic axons in the organotypic cocultures form branches with laminar specificity resembling that found in vivo7,8,10,14,15,16,17. At the same time, punctate aggregations of SYP-EGFP (SYP-EGFP puncta) could be observed along DsRed-labeled TC axons (Figure 3D-3F). Since these SYP-EGFP puncta were mostly colocalized with immunopositive signals for VGLUT2, which is a presynaptic marker of thalamic cells, and for PSD95, which is a general postsynaptic marker, it is likely that SYP-EGFP puncta mostly represent presynaptic sites on TC axons12.
Time-lapse imaging of TC axons further showed that axon branching and synapse formation of thalamic neurons were continuous and dynamic with addition and elimination (Figure 4)12. Moreover, most branches were found to emerge from SYP-EGFP puncta, indicating that presynaptic sites trigger branch formation (for details, see12).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56553/56553fig1.jpg" /> 
Figure 1. The procedure for preparation of an organotypic cocultures of the thalamus and cerebral cortex.&#160;(A) The top view of a neonatal rat brain. The first cut is made parallel to the midline (1). Then, 300 - 500-&#956;m-thick coronal sections are cut from the primary visual and somatosensory cortex (2). Finally, make a parasagittal cut to obtain the cortical slices (3). (B) A schematic representation of an organotypic coculture of the thalamus and cerebral cortex. Cortical slices from postnatal day 2 rat and thalamic blocks from embryonic day 15 are cocultured on the membrane filter. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56553/56553fig1large.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/56553/56553fig2.jpg" /> 
Figure 2. A schematic diagram of electrical instruments for electroporation. The stimulator, the isolator, and the electrode pipette are connected in series. To deliver negatively charged DNA, the electrode pipette is connected to the negative terminal of the isolator. The electrical currents for electroporation can be monitored with the oscilloscope. TH: thalamus. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56553/56553fig2large.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/56553/56553fig3.jpg" /> 
Figure 3. Thalamocortical axons in a coculture after 2 weeks in culture. (A-F) Coexpression of SYP-EGFP with DsRed in a thalamocortical (TC) axon. DsRed plus SYP-EGFP plasmids were introduced into cultured thalamic cells at 1 day in vitro (DIV) using electroporation. (A) An organotypic coculture of thalamic and cortical slices after 12 DIV. (B) A DsRed-labeled thalamic cell extends an axon into the cortical slice and forms branches in upper layers. (C) A magnified image of the boxed region in (A). (D-F) show the DsRed-labeled TC axon (D) and SYP-EGFP puncta (E) in the boxed region of (C). Discrete accumulation of SYP-EGFP was observed along a single TC axon. Th: thalamus. Scale bars: (B) 1 mm, (C) 100 &#181;m, and (F) 20 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56553/56553fig3large.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/56553/56553fig4v2.jpg" /> 
Figure 4. Time-lapse imaging of a thalamocortical axon expressing SYP-EGFP and DsRed in an organotypic coculture. DsRed plus SYP-EGFP plasmids were introduced into cultured thalamic cells at 1 DIV using electroporation. This axon was imaged daily over 4 days (11 DIV to 14 DIV). Scale bar: 50 &#181;m.&#160;<a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56553/56553fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Visualization of Thalamocortical Axon Branching and Synapse Formation in Organotypic Cocultures at JoVE.com

Visualization of Thalamocortical Axon Branching and Synapse Formation in Organotypic Cocultures

This protocol describes a method for simultaneous imaging of thalamocortical axon branching and synapse formation in organotypic cocultures of the thalamus and cerebral cortex. Individual thalamocortical axons and their presynaptic terminals are visualized by a single cell electroporation technique with DsRed and GFP-tagged synaptophysin.

Axon branching and synapse formation are crucial processes for establishing precise neuronal circuits. During development, sensory thalamocortical (TC) ...

visualization-thalamocortical-axon-branching-synapse-formation

Nanyang Technological University

Research

JoVE Journal

Neuroscience

6.4K Views.  Kanazawa University. Dit protocol beschrijft een methode voor gelijktijdige beeldvorming van thalamocortical axon vertakking en de synapse formatie in organotypic cocultures van de thalamus en cortex cerebri. Individuele thalamocortical axonen en hun presynaptische terminals worden gevisualiseerd door een eencellige electroporation techniek met DsRed en synaptophysin van GFP-gelabeld.

Video: Visualization of Thalamocortical Axon Branching and Synapse Formation in Organotypic Cocultures

Coculture System with an Organotypic Brain Slice and 3D Spheroid of Carcinoma Cells

Patients with cerebral metastasis of carcinomas have a poor prognosis. However, the process at the metastatic site has barely been investigated, in particular the role of the resident (stromal) cells. Studies in primary carcinomas demonstrate the influence of the microenvironment on metastasis, even on prognosis1,2. Especially the tumor associated macrophages (TAM) support migration, invasion and proliferation3. Interestingly, the major target sites of metastasis possess tissue-specific macrophages, such as Kupffer cells in the liver or microglia in the CNS. Moreover, the metastatic sites also possess other tissue-specific cells, like astrocytes. Recently, astrocytes were demonstrated to foster proliferation and persistence of cancer cells4,5. Therefore, functions of these tissue-specific cell types seem to be very important in the process of brain metastasis6,7.
Despite these observations, however, up to now there is no suitable in vivo/in vitro model available to directly visualize glial reactions during cerebral metastasis formation, in particular by bright field microscopy. Recent in vivo live imaging of carcinoma cells demonstrated their cerebral colonization behavior8. However, this method is very laborious, costly and technically complex. In addition, these kinds of animal experiments are restricted to small series and come with a substantial stress for the animals (by implantation of the glass plate, injection of tumor cells, repetitive anaesthesia and long-term fixation). Furthermore, in vivo imaging is thus far limited to the visualization of the carcinoma cells, whereas interactions with resident cells have not yet been illustrated. Finally, investigations of human carcinoma cells within immunocompetent animals are impossible8.
For these reasons, we established a coculture system consisting of an organotypic mouse brain slice and epithelial cells embedded in matrigel (3D cell sphere). The 3D carcinoma cell spheres were placed directly next to the brain slice edge in order to investigate the invasion of the neighboring brain tissue. This enables us to visualize morphological changes and interactions between the glial cells and carcinoma cells by fluorescence and even by bright field microscopy. After the coculture experiment, the brain tissue or the 3D cell spheroids can be collected and used for further molecular analyses (e.g. qRT-PCR, IHC, or immunoblot) as well as for investigations by confocal microscopy. This method can be applied to monitor the events within a living brain tissue for days without deleterious effects to the brain slices. The model also allows selective suppression and replacement of resident cells by cells from a donor tissue to determine the distinct impact of a given genotype. Finally, the coculture model is a practicable alternative to in vivo approaches when testing targeted pharmacological manipulations.

The organotypic brain slice coculture with carcinoma cells enables visualizing morphological changes by fluorescence as well as bright field (video) microscopy during the process of carcinoma cell invasion of brain tissue. This model system also allows for cell exchange and replenishment approaches and offers a wide variety of manipulations and analyses.

Patients with cerebral metastasis of carcinomas have a poor prognosis. However, the process at the metastatic site has barely been investigated, in ...

Medicine

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 ...

Biology

Synaptic Microcircuit Modeling with 3D Cocultures of Astrocytes and Neurons from Human Pluripotent Stem Cells

A barrier to our understanding of how various cell types and signals contribute to synaptic circuit function is the lack of relevant models for studying the human brain. One emerging technology to address this issue is the use of three dimensional (3D) neural cell cultures, termed &#39;organoids&#39; or &#39;spheroids&#39;, for long term preservation of intercellular interactions including extracellular adhesion molecules. However, these culture systems are time consuming and not systematically generated. Here, we detail a method to rapidly and consistently produce 3D cocultures of neurons and astrocytes from human pluripotent stem cells. First, pre-differentiated astrocytes and neuronal progenitors are dissociated and counted. Next, cells are combined in sphere-forming dishes with a Rho-Kinase inhibitor and at specific ratios to produce spheres of reproducible size. After several weeks of culture as floating spheres, cocultures (&#39;asteroids&#39;) are finally sectioned for immunostaining or plated upon multielectrode arrays to measure synaptic density and strength. In general, it is expected that this protocol will yield 3D neural spheres that display mature cell-type restricted markers, form functional synapses, and exhibit spontaneous synaptic network burst activity. Together, this system permits drug screening and investigations into mechanisms of disease in a more suitable model compared to monolayer cultures.

In this protocol, we aim to describe a reproducible method for combining dissociated human pluripotent stem cell derived neurons and astrocytes together into 3D sphere cocultures, maintaining these spheres in free floating conditions, and subsequently measuring synaptic circuit activity of the spheres with immunoanalysis and multielectrode array recordings.

A barrier to our understanding of how various cell types and signals contribute to synaptic circuit function is the lack of relevant models for studying ...

Developmental Biology

Three-dimensional Tissue Engineered Aligned Astrocyte Networks to Recapitulate Developmental Mechanisms and Facilitate Nervous System Regeneration

Neurotrauma and neurodegenerative disease often result in lasting neurological deficits due to the limited capacity of the central nervous system (CNS) to replace lost neurons and regenerate axonal pathways. However, during nervous system development, neuronal migration and axonal extension often occur along pathways formed by other cells, referred to as &#34;living scaffolds&#34;. Seeking to emulate these mechanisms and to design a strategy that circumvents the inhibitory environment of the CNS, this manuscript presents a protocol to fabricate tissue engineered astrocyte-based &#34;living scaffolds&#34;. To create these constructs, we employed a novel biomaterial encasement scheme to induce astrocytes to self-assemble into dense three-dimensional bundles of bipolar longitudinally-aligned somata and processes. First, hollow hydrogel micro-columns were assembled, and the inner lumen was coated with collagen extracellular-matrix. Dissociated cerebral cortical astrocytes were then delivered into the lumen of the cylindrical micro-column and, at a critical inner diameter of &#60;350 &#181;m, spontaneously self-aligned and contracted to produce long fiber-like cables consisting of dense bundles of astrocyte processes and collagen fibrils measuring &#60;150 &#181;m in diameter yet extending several cm in length. These engineered living scaffolds exhibited &#62;97% cell viability and were virtually exclusively comprised of astrocytes expressing a combination of the intermediate filament proteins glial-fibrillary acidic protein (GFAP), vimentin, and nestin. These aligned astrocyte networks were found to provide a permissive substrate for neuronal attachment and aligned neurite extension. Moreover, these constructs maintain integrity and alignment when extracted from the hydrogel encasement, making them suitable for CNS implantation. These preformed constructs structurally emulate key cytoarchitectural elements of naturally occurring glial-based &#34;living scaffolds&#34; in vivo. As such, these engineered living scaffolds may serve as test-beds to study neurodevelopmental mechanisms in vitro or facilitate neuroregeneration by directing neuronal migration and/or axonal pathfinding following CNS degeneration in vivo.

We showcase the development of self-assembled, three-dimensional bundles of longitudinally aligned astrocytic somata and processes within a novel biomaterial encasement. These engineered &#34;living scaffolds&#34;, exhibiting micron-scale diameter yet extending centimeters in length, may serve as test-beds to study neurodevelopmental mechanisms or facilitate neuroregeneration by directing neuronal migration and/or axonal pathfinding.

Neurotrauma and neurodegenerative disease often result in lasting neurological deficits due to the limited capacity of the central nervous system (CNS) to ...

Bioengineering

2D and 3D Human Induced Pluripotent Stem Cell-Based Models to Dissect Primary Cilium Involvement during Neocortical Development

Primary cilia (PC) are non-motile dynamic microtubule-based organelles that protrude from the surface of most mammalian cells. They emerge from the older centriole during the G1/G0 phase of the cell cycle, while they disassemble as the cells re-enter the cell cycle at the G2/M phase boundary. They function as signal hubs, by detecting and transducing extracellular signals crucial for many cell processes. Similar to most cell types, all neocortical neural stem and progenitor cells (NSPCs) have been shown harboring a PC allowing them to sense and transduce specific signals required for the normal cerebral cortical development. Here, we provide detailed protocols to generate and characterize two-dimensional (2D) and three-dimensional (3D) cell-based models from human induced pluripotent stem cells (hIPSCs) to further dissect the involvement of PC during neocortical development. In particular, we present protocols to study the PC biogenesis and function in 2D neural rosette-derived NSPCs including the transduction of the Sonic Hedgehog (SHH) pathway. To take advantage of the three-dimensional (3D) organization of cerebral organoids, we describe a simple method for 3D imaging of in toto immunostained cerebral organoids. After optical clearing, rapid acquisition of entire organoids allows detection of both centrosomes and PC on neocortical progenitors and neurons of the whole organoid. Finally, we detail the procedure for immunostaining and clearing of thick free-floating organoid sections preserving a significant degree of 3D spatial information and allowing for the high-resolution acquisition required for the detailed qualitative and quantitative analysis of PC biogenesis and function.

We present detailed protocols for the generation and characterization of 2D and 3D human induced pluripotent stem cell (hIPSC)-based models of neocortical development as well as complementary methodologies enabling qualitative and quantitative analysis of primary cilium (PC) biogenesis and function.

Primary cilia (PC) are non-motile dynamic microtubule-based organelles that protrude from the surface of most mammalian cells. They emerge from the older ...

Modification of a Colliculo-thalamocortical Mouse Brain Slice, Incorporating 3-D printing of Chamber Components and Multi-scale Optical Imaging

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study these two systems and how they interact has been with the use of in vivo preparations. Major advances have been made with acute brain slices containing the thalamocortical and cortico-thalamic pathways in the somatosensory, visual, and auditory systems. With key refinements to our recent modification of the auditory thalamocortical slice1, we are able to more reliably capture the projections between most of the major auditory midbrain and forebrain structures: the inferior colliculus (IC), medial geniculate body (MGB), thalamic reticular nucleus (TRN), and the auditory cortex (AC). With portions of all these connections retained, we are able to answer detailed questions that complement the questions that can be answered with in vivo preparations. The use of flavoprotein autofluorescence imaging enables us to rapidly assess connectivity in any given slice and guide the ensuing experiment. Using this slice in conjunction with recording and imaging techniques, we are now better equipped to understand how information processing occurs at each point in the auditory forebrain as information ascends to the cortex, and the impact of descending cortical modulation. 3-D printing to build slice chamber components permits double-sided perfusion and broad access to networks within the slice and maintains the widespread connections key to fully utilizing this preparation.

Using multiple angles to cut the mouse pup brain, we improve upon a previously-described acute brain slice which captures the connections between most of the major auditory midbrain and forebrain structures.

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study ...

Visualization of Cortical Modules in Flattened Mammalian Cortices

The cortex of mammalian brains is parcellated into distinct substructures or modules. Cortical modules typically lie parallel to the cortical sheet, and can be delineated by certain histochemical and immunohistochemical methods. In this study, we highlight a method to isolate the cortex from mammalian brains and flatten them to obtain sections parallel to the cortical sheet. We further highlight selected histochemical and immunohistochemical methods to process these flattened tangential sections to visualize cortical modules. In the somatosensory cortex of various mammals, we perform cytochrome oxidase histochemistry to reveal body maps or cortical modules representing different parts of the body of the animal. In the medial entorhinal cortex, an area where grid cells are generated, we utilize immunohistochemical methods to highlight modules of genetically determined neurons which are arranged in a grid-pattern in the cortical sheet across several species. Overall, we provide a framework to isolate and prepare layer-wise flattened cortical sections, and visualize cortical modules using histochemical and immunohistochemical methods in a wide variety of mammalian brains.

This article describes a detailed methodology to obtain flattened tangential sections from mammalian cortices and visualize cortical modules using histochemical and immunohistochemical methods.

The cortex of mammalian brains is parcellated into distinct substructures or modules. Cortical modules typically lie parallel to the cortical sheet, and ...

A Model of Epileptogenesis in Rhinal Cortex-Hippocampus Organotypic Slice Cultures

Organotypic slice cultures have been widely used to model brain disorders and are considered excellent platforms for evaluating a drug&#8217;s neuroprotective and therapeutic potential. Organotypic slices are prepared from explanted tissue and represent a complex multicellular ex vivo environment. They preserve the three-dimensional cytoarchitecture and local environment of brain cells, maintain the neuronal connectivity and the neuron-glia reciprocal interaction. Hippocampal organotypic slices are considered suitable to explore the basic mechanisms of epileptogenesis, but clinical research and animal models of epilepsy have suggested that the rhinal cortex, composed of perirhinal and entorhinal cortices, play a relevant role in seizure generation.
Here, we describe the preparation of rhinal cortex-hippocampus organotypic slices. Recordings of spontaneous activity from the CA3 area under perfusion with complete growth medium, at physiological temperature and in the absence of pharmacological manipulations, showed that these slices depict evolving epileptic-like events throughout time in culture. Increased cell death, through propidium iodide uptake assay, and gliosis, assessed with fluorescence-coupled immunohistochemistry, was also observed. The experimental approach presented highlights the value of rhinal cortex-hippocampus organotypic slice cultures as a platform to study the dynamics and progression of epileptogenesis and to screen potential therapeutic targets for this brain pathology.

Here, we describe the preparation of rhinal cortex-hippocampus organotypic slices. Under a gradual and controlled deprivation of serum, these slices depict evolving epileptic-like events and can be considered an ex vivo model of epileptogenesis. This system represents an excellent tool for monitoring the dynamics of spontaneous activity, as well as for assessing the progression of neuroinflammatory features throughout the course of epileptogenesis.

Organotypic slice cultures have been widely used to model brain disorders and are considered excellent platforms for evaluating a drug&#8217;s ...

In Situ Visualization of Axon Growth and Growth Cone Dynamics in Acute Ex Vivo Embryonic Brain Slice Cultures

During neuronal development, axons navigate the cortical environment to reach their final destinations and establish synaptic connections. Growth cones -the sensory structures located at the distal tips of developing axons- execute this process. Studying the structure and dynamics of the growth cone is crucial to understanding axonal development and the interactions with the surrounding central nervous system (CNS) that enable it to form neural circuits. This is essential when devising methods to reintegrate axons into neural circuits following injury in fundamental research and pre-clinical contexts. Thus far, the general understanding of growth cone dynamics is primarily founded on studies of neurons cultured in two dimensions (2D). Although undoubtedly fundamental to the current knowledge of growth cone structural dynamics and response to stimuli, 2D studies misrepresent the physiological three-dimensional (3D) environment encountered by neuronal growth cones in intact CNS tissue. More recently, collagen gels were employed to overcome some of these limitations, enabling the investigation of neuronal development in 3D. However, both synthetic 2D and 3D environments lack signaling cues within CNS tissue, which direct the extension and pathfinding of developing axons. This protocol provides a method for studying axons and growth cones using organotypic brain slices, where developing axons encounter physiologically relevant physical and chemical cues. By combining fine-tuned in utero and ex utero electroporation to sparsely deliver fluorescent reporters along with super-resolution microscopy, this protocol presents a methodological pipeline for the visualization of axon and growth cone dynamics in situ. Furthermore, a detailed toolkit description of the analysis of long-term and live-cell imaging data is included.

In Situ Visualization of Axon Growth and Growth Cone Dynamics in Acute Ex Vivo Embryonic Brain Slice Cultures

This protocol demonstrates a straightforward and robust method to study in situ axon growth and growth cone dynamics. It describes how to prepare ex vivo physiologically relevant acute brain slices and provides a user-friendly analysis pipeline.

During neuronal development, axons navigate the cortical environment to reach their final destinations and establish synaptic connections. Growth cones ...

Neuronal Activity Mapping in Developing Mouse Sensory Cortex

In Vivo Visualization of Spontaneous Activity in Neonatal Mouse Sensory Cortex at a Single-Neuron Resolution

The mammalian brain undergoes dynamic developmental changes at both the cellular and circuit levels throughout prenatal and postnatal periods. Following the discovery of numerous genes contributing to these developmental changes, it is now known that neuronal activity also substantially modulates these processes. In the developing cerebral cortex, neurons exhibit synchronized activity patterns that are specialized to each primary sensory area. These patterns markedly differ from those observed in the mature cortex, emphasizing their role in regulating area-specific developmental processes. Deficiencies in neuronal activity during development can lead to various brain diseases. These findings highlight the need to examine the regulatory mechanisms underlying activity patterns in neuronal development. This paper summarizes a series of protocols to visualize primary sensory areas and neuronal activity in neonatal mice, to image the activity of individual neurons within the cortical subfields using two-photon microscopy in vivo, and to analyze subfield-related activity correlations. We show representative results of patchwork-like synchronous activity within individual barrels in the somatosensory cortex. We also discuss various potential applications and some limitations of this protocol.

Author Spotlight: Deciphering Neural Circuit Formation from Two-Photon Microscopy and Single Neuron Imaging

Primary sensory areas in the neocortex exhibit unique spontaneous activities during development. This article describes how to visualize individual neuron activities and primary sensory areas to analyze area-specific synchronous activities in neonatal mice in vivo.

The mammalian brain undergoes dynamic developmental changes at both the cellular and circuit levels throughout prenatal and postnatal periods. Following ...

Visualization of Thalamocortical Axon Branching and Synapse Formation in Organotypic Cocultures

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