Supported by MIUR National Technology Clustersproject IRMI (CTN01_00177_888744), and Regione Emilia-Romagna, Mat2Rep, POR-FESR 2014-2020.
Special thanks to IRET Foundation for hosting the experimental work.

All animal protocols described herein were carried out according to European Community Council Directives (86/609/EEC) and comply with the guidelines published in the NIH Guide for the Care and Use of Laboratory Animals.
1. Solutions and reagents
<ol>
	<li>Prepare standard medium: DMEM/F12 GlutaMAX 1x; 8 mmol/L HEPES; 100 U/100 &#956;g Penicillin/Streptomycin (1% P/S); 1x B27; 1x N-2.</li>
	<li>Prepare neurosphere medium: add 10 ng/mL bFGF; 10 ng/mL EGF to standard medium.</li>
	<li>Prepare oligosphere/OPC medium: add 10 ng/mL bFGF; 10 ng/mL PDGF-AA to standard medium.</li>
	<li>Prepare oligodendroctye differentiation medium: add 50 nM T3; 10 ng/mL CNTF; 1x N-acetyl-L-cysteine (NAC) to standard medium.</li>
	<li>Prepare non-enzymatic dissociation buffer: add 1% P/S to non-enzymatic dissociation buffer and keep ice cold.</li>
	<li>Prepare sucrose solution: HBSS, 0.3 g/mL sucrose.</li>
	<li>Prepare BSA washing solution: EBSS, 40 mg/mL BSA, 0.02 mL/l HEPES.</li>
	<li>Prepare enzymatic dissociation buffer: HBSS, 5.4 mg/mL D-glucose, 15 mmol/L HEPES, 1.33 mg/mL Trypsin, 0.7 mg/mL Hyaluronidase, 80 U/mL DNase.</li>
	<li>Prepare cytokine mix: TGF-&#946;1, TNF-&#945;, IL-1&#946;, IL-6, IL-17, and IFN-&#947; (20 ng/mL each).</li>
	<li>Prepare cytokine mix vehicle: 0.04% of the stock (10% glycerol/100 nM glycine/25 nM Tris, pH 7.3).</li>
	<li>Prepare oxygen-glucose deprivation medium: standard medium using DMEM w/o glucose.&#160;Depending on the stringency of the desired glucose deprivation condition, it is possible to remove also B27 and/or N2 from the medium to avoid glucose-related compounds (e.g., D-Galactose in B27).</li>
</ol>
2. Dissection and NSC isolation
NOTE: Fetal and adult NSCs were isolated from E13.5 fetal forebrain or 2.5-month-old adult sub-ventricular zone (SVZ), following the Ahlenius and Kokaia protocol9 with modifications.
<ol>
	<li>Fetal NSC cultures 
	NOTE: Before starting the dissections, prepare 1.5 mL tubes containing 150 &#181;L of non-enzymatic dissociation buffer each; clean Petri dishes and add ice cold HBSS.
	<ol>
		<li>Collect the embryos at E13.5 - 14.5 from timed pregnant mice and place in a Petri dish containing cold HBSS.</li>
		<li>Decapitate the embryos using forceps.</li>
		<li>Place the heads of the embryos in a clean Petri dish containing ice cold PBS and remove the skin from the skull with forceps, using magnifying glasses or a stereoscope.</li>
		<li>Once the brain is visible and cleared of skin, squeeze it out by applying pressure at the sides with forceps.</li>
		<li>Remove the cerebellum, keep only the forebrain and remove the meninges with forceps.</li>
		<li>Place the isolated tissue in the non-enzymatic dissociation buffer and repeat the dissection steps with the other embryos. Insert the tissue from 2&#8211;3 animals into each tube containing the buffer.</li>
		<li>Incubate at 37 &#176;C for 15 min under continuous shaking.</li>
		<li>After incubation, add 850 &#181;L of standard medium and mix by pipetting until the suspension is free of clumps.</li>
		<li>If non-dissociated tissue is still visible, wait for 2 min at RT until it deposits at the bottom of the tube.</li>
		<li>When dissociation is complete, count the cells and plate them in suspension at a density of 10&#8211;50 cells/&#181;L in a T-25 or T-45 flask containing 10&#8211;30 mL of neurosphere medium, kept in a vertical position to avoid cell adhesion.</li>
	</ol>
	</li>
	<li>Adult NSC cultures
	<ol>
		<li>Sacrifice animals by cervical dislocation.</li>
		<li>Collect brains from 4&#8211;5 mice in a 50 mL tube containing ice cold HBSS.</li>
		<li>Place the brain on a cold sterile surface. For this purpose, use a T-25 flask filled with water and placed at -20 &#176;C overnight. At the time of the experiment, cover the flask with sterile aluminum foil.</li>
		<li>Place the brain ventral side downwards, in rostro-caudal direction, and remove the olfactory bulbs using a razor blade.</li>
		<li>Using a razor blade, cut 2&#8211;3 coronal slices of 1 mm thickness, from the cortex to the optical chiasma.</li>
		<li>Place the slices on the cold surface in a ventro-dorsal position and identify the corpus callosum and the two lateral ventricles.</li>
		<li>Using magnifying glasses or a stereoscope, isolate the walls of the lateral ventricles, taking care not to carry pieces of the corpus callosum.</li>
		<li>Put the isolated tissue in the enzymatic dissociation buffer (5&#8211;10 mL) and incubate at 37 &#176;C for 15 min.</li>
		<li>Mix the solution, pipetting several times (at least 50), and incubate again at 37 &#176;C for 10 min.</li>
		<li>Neutralize the trypsin by adding 5 mL of standard culture medium and filter the solution using a 70 &#181;m filter.</li>
		<li>Centrifuge the filtered solution for 5 min at 400 x g.</li>
		<li>Resuspend the pellet in the sucrose solution and centrifuge for 10 min at 500 x g.</li>
		<li>Resuspend the pellet in BSA washing solution and centrifuge for 7 min at 400 x g.</li>
		<li>Resuspend the pellet in the standard culture medium, count the cells, and perform plating as described above (in step 2.1.10).</li>
	</ol>
	</li>
</ol>
3. Primary neurospheres
<ol>
	<li>Add the growth factors (bFGF/EGF) every 2 days.</li>
	<li>Every 4&#8211;6 days (depending on cell density), change half of the medium as follows:
	<ol>
		<li>Transfer the entire cell suspension to a 15 or 50 mL tube.</li>
		<li>Centrifuge for 5 min at 400 x g.</li>
		<li>Remove half of the volume.</li>
		<li>Add the same amount of fresh medium, gently mix by pipetting, and add growth factors.</li>
	</ol>
	</li>
</ol>
4. Oligospheres
NOTE: Oligodendrocyte differentiation is performed following the Chen protocol10 with modifications.
<ol>
	<li>When the neurospheres reach a diameter of 100&#8211;150 &#181;m, they are ready to be passed. To do so, transfer the entire cell suspension to a 15 or 50 mL tube, and centrifuge for 5 min at 400 x g.
	<ol>
		<li>Rapidly evaluate the diameter by taking pictures of the spheres using an inverted transmitted light microscope and opening them by ImageJ software.</li>
		<li>Click on the Analyze menu and from the Tools window, select Scale bar.</li>
		<li>Set 150 &#181;m as Width in microns and compare the scale bar with the spheres.</li>
	</ol>
	</li>
	<li>Remove the entire volume by inversion and resuspend the pellet in 180 &#181;L of fresh standard culture medium. Pipette 50 times to allow disaggregation of the spheres.</li>
	<li>Add 810 &#181;L of fresh standard culture medium, count the cells, and re-plate them as described for the neurospheres.</li>
	<li>Add bFGF/PDGF-AA 10 ng/mL every 2 days.</li>
	<li>Every 4&#8211;6 days (depending on cell density), change half of the medium as follows:</li>
	<li>Transfer the entire cell suspension to a 15 or 50 mL tube.</li>
	<li>Centrifuge for 5 min at 400 x g.</li>
	<li>Remove half of the volume.</li>
	<li>Add the same amount of fresh medium, gently mix by pipetting, and add growth factors.</li>
</ol>
5. Plate coating
<ol>
	<li>Poly-D,L-ornithine/laminin coating: at least 2 days before plating the OPCs, add 50 &#181;g/mL poly-D,L-ornithine solution, diluted in PBS, to each well (40 &#181;L/well for 96-well plates) and incubate at RT overnight.</li>
	<li>The following day, remove the liquid and wash three times with distilled sterile water.</li>
	<li>Let the plates dry at RT overnight. The following day, add a laminin solution diluted in PBS (5 &#181;g/mL; 40 &#181;L/well for 96-well plates) and incubate for 2 h at 37 &#176;C.</li>
</ol>
6. Cell seeding
<ol>
	<li>When the oligospheres reach a diameter of 100&#8211;150 &#181;m, they are ready to be dissociated and seeded on the poly-D,L-ornithine/laminine coated plates. To do so, transfer the entire cell suspension to a 15 or 50 mL tube, and centrifuge for 5 min at 400 x g (as indicated in step 4.1)</li>
	<li>Remove the entire volume by inversion and resuspend the pellet in 180 &#181;L of fresh standard culture medium. Pipette 50 times to allow disaggregation of the spheres.</li>
	<li>Add 810 &#181;L of fresh standard culture medium and count the cells.</li>
	<li>Remove the laminin solution from the wells and plate the cells at 3,000 cell/cm2 density (100 &#181;L/well for 96-well plates).</li>
</ol>
7. OPC differentiation induction
<ol>
	<li>After 3 days, remove the entire medium and add the same volume of oligodendrocyte differentiation medium.</li>
	<li>Change half of the medium every 4 days and add fresh differentiation mix (T3/CNTF/NAC) every 2 days.</li>
</ol>
8. Induction of inflammation-mediated differentiation block
<ol>
	<li>After neurosphere dissociation and oligosphere production (section 4), add the cytokine mix to the culture medium and keep oligospheres exposed to cytokines for the whole spheres formation step. 
	NOTE: The volume depends on the number of cells, since for the spheres forming cells are seeded at 10&#8211;50 cells/&#181;L.</li>
	<li>If the medium needs to be changed, change the entire volume and add the cytokine mix once more.</li>
</ol>
9. Induction of oxygen-glucose deprivation cell death
<ol>
	<li>At -1 DIV (2 days after cell seeding in multiwell plates), remove the medium and conserve it in a new multiwell plate.</li>
	<li>Add half the volume (50 &#181;L for 96-well plates) of OGD-medium (OGD group) or fresh medium (control group). The half amount of volume is used to reduce the exchange of oxygen between the liquid and the air.</li>
	<li>Place the OGD group cultures in an airtight hypoxia chamber saturated with 95% N2 and 5% CO2. To achieve saturation of the chamber, let the gas mixture flow for 6 min at 25 l/min before closing the chamber pipes.</li>
	<li>Incubate the hypoxic chamber in the incubator for 3 h. The control group and plates containing the medium removed and conserved at step 9.1 should also be left in the incubator.</li>
	<li>Remove the glucose-free (OGD group) or the new medium (control group) and add the medium removed and conserved at step 9.1.</li>
</ol>
10. Immunocytochemistry
<ol>
	<li>At the desired time point, fix the cells with cold 4% paraformaldehyde for 20 min at RT.</li>
	<li>Wash twice with PBS (10 min of incubation for each wash at RT).</li>
	<li>Incubate 1 h at RT with the blocking solution&#160;(PBS Triton 0.3% containing 1% BSA and 1% donkey/goat normal serum).</li>
	<li>Incubate with primary antibody mix (Table 1), diluted in PBS triton 0.3%, overnight at 4 &#176;C.</li>
	<li>Wash twice with PBS (10 min of incubation for each wash at RT).</li>
	<li>Incubate with secondary antibody (Table 1) solution diluted in PBS triton 0.3% adding Hoechst 33258 for 30 min at 37 &#176;C.</li>
	<li>Wash twice with PBS (10 min of incubation for each wash at RT).</li>
</ol>
11. HCS analysis of cell viability, lineage composition, and lineage-specific cell death
NOTE: The HCS representative images and workflow are shown in Figure 2A,B.
<ol>
	<li>Select the Compartmental Analysis algorithm from the main menu of the software (HCS Studio v 6.6.0) and select Scan from the main menu Develop Assay/Scan Plate.</li>
	<li>In the iDev window, select New and then select General intensity Measurement Tool from the Develop Assay template.</li>
	<li>Click on Create on the right side of the menu, selecting the 10x objective.</li>
	<li>This will open the Configure Acquisition menu. In this window, select the following parameters: (a) number of channels: the first one for the Hoechst nuclear staining (BGRFR_386) and one for each lineage-specific marker used in the reaction (b) select software focus on channel 1 and autofocus interval as 1 (c) select the plate model from the list.</li>
	<li>From the acquisition menu, look at the quality of the staining in different wells and different fields and manually select the exposure time by selecting Fixed Exposure Time in the menu.</li>
	<li>Once the acquisitions parameters are set, select Mini Scan on the top of the menu and select ten fields per well in two wells per experimental condition. This will allow the set-up of all the analysis parameter in a subset of fields for the entire plate.</li>
	<li>When the mini scan is finished, click on the Configure Assay Parameter to configure the algorithm of the analysis.</li>
	<li>Click on Configure Groups in the right side of the window and drag-and-drop the wells of the miniscan. Click on the Add button in the Groups sub-section to configure the different groups.</li>
	<li>Follow the workflow on the left side of the window step by step to develop the whole algorithm. First select Process Image for each channel and click on Background Removal and on the desired level.</li>
	<li>First identify and select the nuclei by nuclear staining. Click on Identify Primary Object &#8211; Channel 1 to select the real nuclei and avoid analyzing artifact and debris. For this purpose, zoom in a representative picture of nuclear staining and check whether the nuclei are well surrounded by the perimeter built by the software. It is possible to change the thresholding value and to apply segmentation algorithms to better identify single nuclei.</li>
	<li>Once the nuclei are defined correctly, click on the following step: Validate Primary Object. Select Object.BorderObject.Ch1 to avoid the analysis of nuclei at the border of each field image. Select Object.Area.Ch1 and, by moving the &#8220;low&#8221; and &#8220;high&#8221; bars on the histograms, remove all the identified debris or big-objects corresponding to aggregates or artifacts.</li>
	<li>Check all the mini scan representative images of all the experimental conditions to be sure that the selected parameters fit with all of them.</li>
	<li>Click on Identify Spots for each channel corresponding to the specific lineage markers, and select the Ring values: Width = 3 and Distance = 0. This will allow the identification of the cytoplasmatic fluorescence. According to the cell density, these values can be adapted. The software will automatically avoid the overlapping between adjacent rings.</li>
	<li>Select Reference Levels in the workflow to build the analysis. The setting of the reference levels will allow the automatic counting of condensed nuclei, based on the nuclear size and nuclear staining intensity, and of specific marker-positive cells, based on the cytoplasmatic fluorescence identified by the Ring.</li>
	<li>First click on Object.Area.Ch1. In the mini scan images, select a condensed nucleus and move the &#8220;LOW&#8221; bar on the histograms in order to select as &#8220;condensed&#8221; all the nuclei under this size.</li>
	<li>Click on Object.AvgIntensity.Ch1. In the mini scan images, select a condensed nucleus and move the &#8220;HIGH&#8221; bar on the histograms in order to select as &#8220;condensed&#8221; all the nuclei above this fluorescence intensity.</li>
	<li>Click on Object.RingAvgIntensity for each channel of lineage specific markers. Select in your mini scan images a positive cell and move the &#8220;HIGH&#8221; bar on the histograms in order to select as &#8220;positive&#8221; all the cells above this fluorescence intensity.</li>
	<li>Check all the mini scan representative images of all the experimental conditions to be sure that the selected parameters fit with all of them.</li>
	<li>On the top menu, select Population Characterization and select Event Subpopulation.</li>
	<li>As Type 1 Event, select ObjectAreaCh1 on the left list, then click on the AND &#62; button and finally select ObjectAvgIntensityCh1. This will allow the identification of condensed nuclei, as a combination of low area and high intensity.</li>
	<li>In the same window, deselect all the Scan Limits.</li>
	<li>Click on Select Features to Store in the top menu, to choose the parameters to keep in the analysis.</li>
	<li>Select Well Features and move from the left list to the right only the desired parameters: (a) SelectedObjectCountPErValidField (b) %EventType1ObjectCount (c) %High_RingAvgIntensity (For each channel of the specific lineage markers). 
	NOTE: This analysis will give as readout the total number of cells, the percentage of condensed nuclei, and the percentage of lineage-specific positive cells for each analyzed marker on the total cell number. If the percentage of the different lineages is needed only on live cells, is it possible either to keep the value &#8220;High_RingAvgIntensity&#8221; for the channel (absolute number of positive cells) and recalculate the percentage on total cell numbers after the subtraction of the percentage of dead cells.
	<ol>
		<li>Alternatively, it is possible to remove the dead cells from the analysis setting the same parameters used to identify condensed nuclei (steps 11.14&#8211;11.15) on the nuclei validation (step 11.11).</li>
	</ol>
	</li>
	<li>Select Scan Plate from the main top menu and click on the plate symbol on Scan Setting sub-menu on the top section to identify the well to analyze.</li>
	<li>Write the name of the experiment and the description and once all the settings are completed, press the play symbol.</li>
</ol>

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Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oligodendrocyte+Progenitor+Cells+(OPCs)+from+adult+human+brain+expressed+distinct+microRNAs+compared+to+OPCs+in+development.">Oligodendrocyte Progenitor Cells (OPCs) from adult human brain expressed distinct microRNAs compared to OPCs in development.</a> Neurology. 82, (2014).</li><li>Bribi&#225;n, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+heterogeneity+of+mouse+and+human+brain+OPCs:+relevance+for+preclinical+studies+in+multiple+sclerosis.">Functional heterogeneity of mouse and human brain OPCs: relevance for preclinical studies in multiple sclerosis.</a> Journal of Clinical Medicine. 9, 1681 (2020).</li><li>J&#228;kel, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Altered+Human+Oligodendrocyte+Heterogeneity+In+Multiple+Sclerosis.">Altered Human Oligodendrocyte Heterogeneity In Multiple Sclerosis.</a> Nature. 566, 543-547 (2019).</li><li>Medina-Rodr&#237;guez, E. M., Arenzana, F. J., Bribi&#225;n, A., de Castro, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protocol+to+isolate+a+large+amount+of+functional+oligodendrocyte+precursor+cells+from+the+cerebral+cortex+of+adult+mice+and+humans.">Protocol to isolate a large amount of functional oligodendrocyte precursor cells from the cerebral cortex of adult mice and humans.</a> PloS One. 8, 81620 (2013).</li><li>Baldassarro, V. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+death+in+pure-neuronal+and+neuron-astrocyte+mixed+primary+culture+subjected+to+oxygen-glucose+deprivation:+the+contribution+of+poly(ADP-Ribose)+polymerases+and+caspases.">Cell death in pure-neuronal and neuron-astrocyte mixed primary culture subjected to oxygen-glucose deprivation: the contribution of poly(ADP-Ribose) polymerases and caspases.</a> Microchemical Journal. 136, 215-222 (2018).</li></ol>

The authors have nothing to disclose.

The complex nature of myelination/remyelination processes and demyelinating events makes the development of predictive in vitro systems extremely challenging. The most widely used in vitro drug screening systems are mostly human cell lines or primary pure OL cultures, with increasing use of more complex co-cultures or organotypic systems15. Even if such systems are coupled with high content technologies, pure OL cultures remain the method of choice when developing screening platforms<sup class="xr...

In the central nervous system (CNS), myelin forming cells (oligodendrocytes, OLs) and their precursors (oligodendrocyte precursor cells, OPCs) are responsible for developmental myelination, a process which occurs during the peri- and post-natal periods, and for myelin turnover and repair (remyelination) in adulthood1. These cells are highly specialized, interacting anatomically and functionally with all the other glial and neuronal components, making them a fundamental part of CNS structure and function.
Demyelinating events are involved in different CNS injuries and diseases2, and mainly act on OPCs and OLs by way of multifactorial mechanisms, both during development and adulthood. The undifferentiated precursors are driven by differentiating factors, mainly thyroid hormone (TH), in a synchronized process3 which leads the OPC to recognize and respond to specific stimuli which induce proliferation, migration to the non-myelinated axon, and differentiation into mature OLs which in turn develop the myelin sheath4. All these processes are finely controlled and occur in a complex environment.
Due to the complex nature of myelination, remyelination and demyelination events, there is a great need for a simplified and reliable in vitro method to study the underlying mechanisms and to develop new therapeutic strategies, focusing on the main cellular player: the OPC5.
For an in vitro system to be reliable, a number of factors need to be taken into account: the complexity of the cellular environment, age-related cell-intrinsic differences, physiological TH-mediated differentiation, pathological mechanisms, and the robustness of the data6. Indeed, the unmet need in the field is a model which mimics the complexity of the in vivo condition, not successfully achieved through the use of isolated pure OPC cultures. In addition, the two main components of demyelinating events, inflammation and hypoxia/ischemia (HI), directly involve other cell components that may indirectly affect the physiological differentiation and maturation of OPCs, an aspect which cannot be studied in over-simplified in vitro models.
Starting from a highly predictive culture system, the subsequent and more general challenge is the production of robust and reliable data. In this context, cell-based high-content screening (HCS) is the most suitable technique7, since our aim is firstly to analyze the entire culture in an automatic workflow, avoiding the bias of choosing representative fields, and secondly to obtain the automatic and simultaneous generation of imaging-based high-content data8.
Given that the main need is to achieve the best balance between in vitro simplification and in vivo-mimicking complexity, here we present a highly reproducible method for obtaining OPCs derived from neural stem cells (NSCs) isolated from the fetal forebrain and the adult sub-ventricular zone (SVZ). This in vitro model encompasses the entire OPC differentiation process, from multipotent NSC to mature/myelinating OL, in a physiological TH-dependent manner. The resulting culture is a dynamically differentiating/maturating system which results in a spontaneous co-culture consisting mainly of differentiating OPCs and astrocytes, with a low percentage of neurons. This primary culture better mimics the complex in vivo environment, while its stem cell derivation allows simple manipulations to be performed to obtain the cell lineage enrichment desired.
On the contrary to other drug screening strategies using cell lines or pure cultures of primary OPCs, the method described here allows the study of the effect of pathological interferents or therapeutic molecules in a complex environment, without losing the focus on the desired cell type. The HCS workflow described permits an analysis of cell viability and lineage specification, as well as lineage-specific cell death and morphological parameters.

<table>
	<tbody>
		<tr>
			<td>96-well plates - untreated</td>
			<td>NUNC</td>
			<td>267313</td>
			<td></td>
		</tr>
		<tr>
			<td>B27 supplement (100x)</td>
			<td>GIBCO</td>
			<td>17504-044</td>
			<td></td>
		</tr>
		<tr>
			<td>basic Fibroblast Growth Factor (bFGF)</td>
			<td>GIBCO</td>
			<td>PHG0024</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma-Aldrich</td>
			<td>A2153</td>
			<td></td>
		</tr>
		<tr>
			<td>Ciliary Neurotropic Factor (CNTF)</td>
			<td>GIBCO</td>
			<td>PHC7015</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM w/o glucose</td>
			<td>GIBCO</td>
			<td>A14430-01</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM/F12 GlutaMAX</td>
			<td>GIBCO</td>
			<td>31331-028</td>
			<td></td>
		</tr>
		<tr>
			<td>DNase</td>
			<td>Sigma-Aldrich</td>
			<td>D5025-150KU</td>
			<td></td>
		</tr>
		<tr>
			<td>EBSS</td>
			<td>GIBCO</td>
			<td>14155-048</td>
			<td></td>
		</tr>
		<tr>
			<td>Epidermal Growth Factor (EGF)</td>
			<td>GIBCO</td>
			<td>PHG6045</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>GIBCO</td>
			<td>14170-088</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>GIBCO</td>
			<td>15630-056</td>
			<td></td>
		</tr>
		<tr>
			<td>Hyaluronidase</td>
			<td>Sigma-Aldrich</td>
			<td>H3884</td>
			<td></td>
		</tr>
		<tr>
			<td>IFN-&#947;</td>
			<td>Origene</td>
			<td>TP721239</td>
			<td></td>
		</tr>
		<tr>
			<td>IL-17A</td>
			<td>Origene</td>
			<td>TP723199</td>
			<td></td>
		</tr>
		<tr>
			<td>IL-1&#946;</td>
			<td>Origene</td>
			<td>TP723210</td>
			<td></td>
		</tr>
		<tr>
			<td>IL-6</td>
			<td>Origene</td>
			<td>TP723240</td>
			<td></td>
		</tr>
		<tr>
			<td>laminin</td>
			<td>GIBCO</td>
			<td>23017-051</td>
			<td></td>
		</tr>
		<tr>
			<td>N-acetyl-L-cysteine</td>
			<td>Sigma-Aldrich</td>
			<td>A9165</td>
			<td></td>
		</tr>
		<tr>
			<td>N2 supplement (50x)</td>
			<td>GIBCO</td>
			<td>17502-048</td>
			<td></td>
		</tr>
		<tr>
			<td>Non-enzymatic dissociation buffer</td>
			<td>GIBCO</td>
			<td>13150-016</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>GIBCO</td>
			<td>70011-036</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin / Streptomycin</td>
			<td>Sigma-Aldrich</td>
			<td>P4333</td>
			<td></td>
		</tr>
		<tr>
			<td>Platelet Derived Growth Factor (PDGF-AA)</td>
			<td>GIBCO</td>
			<td>PHG0035</td>
			<td></td>
		</tr>
		<tr>
			<td>poly-D,L-ornitine</td>
			<td>Sigma-Aldrich</td>
			<td>P4957</td>
			<td></td>
		</tr>
		<tr>
			<td>TGF-&#946;1</td>
			<td>Origene</td>
			<td>TP720760</td>
			<td></td>
		</tr>
		<tr>
			<td>TNF-&#945;</td>
			<td>Origene</td>
			<td>TP723451</td>
			<td></td>
		</tr>
		<tr>
			<td>Triiodothyronine</td>
			<td>Sigma-Aldrich</td>
			<td>T2752-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>Sigma-Aldrich</td>
			<td>T1426</td>
			<td></td>
		</tr>
	</tbody>
</table>

high-content screening differentiation and maturation analysis of fetal and adult neural stem cell-derived oligodendrocyte precursor cell cultures

The main hurdle in developing drug screening techniques for assessing the efficacy of therapeutic strategies in complex diseases is striking a balance between in vitro simplification and recreating the complex in vivo environment, along with the main aim, shared by all screening strategies, of obtaining robust and reliable data, highly predictive for in vivo translation.
In the field of demyelinating diseases, the majority of drug screening strategies are based on immortalized cell lines or pure cultures of isolated primary oligodendrocyte precursor cells (OPCs) from newborn animals, leading to strong biases due to the lack of age-related differences and of any real pathological condition or complexity.
Here we show the setup of an in vitro system aimed at modeling the physiological differentiation/maturation of neural stem cell (NSC)-derived OPCs, easily manipulated to mimic pathological conditions typical of demyelinating diseases. Moreover, the method includes isolation from fetal and adult brains, giving a system which dynamically differentiates from OPCs to mature oligodendrocytes (OLs) in a spontaneous co-culture which also includes astrocytes. This model physiologically resembles the thyroid hormone-mediated myelination and myelin repair process, allowing the addition of pathological interferents which model disease mechanisms. We show how to mimic the two main components of demyelinating diseases (i.e., hypoxia/ischemia and inflammation), recreating their effect on developmental myelination and adult myelin repair and taking all the cell components of the system into account throughout, while focusing on differentiating OPCs.
This spontaneous mixed model, coupled with cell-based high-content screening technologies, allows the development of a robust and reliable drug screening system for therapeutic strategies aimed at combating the pathological processes involved in demyelination and at inducing remyelination.

The first phase of the culture may vary in duration, depending on seeding density and on whether the spheres are of fetal or adult origin. Moreover, oligospheres display a reduced population doubling compared to neurospheres (Figure 1B). Moreover, spheres production from adult tissue is slower and it may take 2&#8211;3 weeks to generate oligospheres compared to fetal that may take 1&#8211;2 weeks, depending on the seeding density.
Once seeded, the entire different...

Watch this Scientific Journal Video about High-Content Screening Differentiation and Maturation Analysis of Fetal and Adult Neural Stem Cell-Derived Oligodendrocyte Precursor Cell Cultures at JoVE.com

High-Content Screening Differentiation and Maturation Analysis of Fetal and Adult Neural Stem Cell-Derived Oligodendrocyte Precursor Cell Cultures

We describe the production of mixed cultures of astrocytes and oligodendrocyte precursor cells derived from fetal or adult neural stem cells differentiating into mature oligodendrocytes, and in vitro modeling of noxious stimuli. The coupling with a cell-based high-content screening technique builds a reliable and robust drug screening system.

The main hurdle in developing drug screening techniques for assessing the efficacy of therapeutic strategies in complex diseases is striking a balance ...

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2.7K Views.  University of Bologna. We describe the production of mixed cultures of astrocytes and oligodendrocyte precursor cells derived from fetal or adult neural stem cells differentiating into mature oligodendrocytes, and in vitro modeling of noxious stimuli. The coupling with a cell-based high-content screening technique builds a reliable and robust drug screening system.

Video: High-Content Screening Differentiation and Maturation Analysis of Fetal and Adult Neural Stem Cell-Derived Oligodendrocyte Precursor Cell Cultures

Growth and Differentiation of Adult Hippocampal Arctic Ground Squirrel Neural Stem Cells

Arctic ground squirrels (Urocitellus parryii, AGS) are unique in their ability to hibernate with a core body temperature near or below freezing 1. These animals also resist ischemic injury to the brain in vivo 2,3 and oxygen-glucose deprivation in vitro 4,5. These unique qualities provided the impetus to isolate AGS neurons to examine inherent neuronal characteristics that could account for the capacity of AGS neurons to resist injury and cell death caused by ischemia and extremely cold temperatures. Identifying proteins or gene targets that allow for the distinctive properties of these cells could aid in the discovery of effective therapies for a number of ischemic indications and for the study of cold tolerance. Adult AGS hippocampus contains neural stem cells that continue to proliferate, allowing for easy expansion of these stem cells in culture. We describe here methods by which researchers can utilize these stem cells and differentiated neurons for any number of purposes. By closely following these steps the AGS neural stem cells can be expanded through two passages or more and then differentiated to a culture high in TUJ1-positive neurons (~50%) without utilizing toxic chemicals to minimize the number of dividing cells. Ischemia induces neurogenesis 6 and neurogenesis which proceeds via MEK/ERK and PI3K/Akt survival signaling pathways contributes to ischemia resistance in vivo7 and in vitro 8 (Kelleher-Anderson, Drew et al., in preparation). Further characterization of these unique neural cells can advance on many fronts, using some or all of these methods.

Neural stem cells were prepared from the hippocampus of adult non-hibernating yearling Arctic ground squirrels (AGS). These neural stem cells can be expanded through numerous passages, differentiated and maintained as a nearly 50:50 neuron to glial culture.

Arctic ground squirrels (Urocitellus parryii, AGS) are unique in their ability to hibernate with a core body temperature near or below freezing 1. These ...

Generation of Neural Stem Cells from Discarded Human Fetal Cortical Tissue

Neural stem cells (NSCs) reside along the ventricular zone neuroepithelium during the development of the cortical plate. These early progenitors ultimately give rise to intermediate progenitors and later, the various neuronal and glial cell subtypes that form the cerebral cortex. The capacity to generate and expand human NSCs (so called neurospheres) from discarded normal fetal tissue provides a means with which to directly study the functional aspects of normal human NSC development 1-5. This approach can also be directed toward the generation of NSCs from known neurological disorders, thereby affording the opportunity to identify disease processes that alter progenitor proliferation, migration and differentiation 6-9. We have focused on identifying pathological mechanisms in human Down syndrome NSCs that might contribute to the accelerated Alzheimer's disease phenotype 10,11. Neither in vivo nor in vitro mouse models can replicate the identical repertoire of genes located on human chromosome 21. 
Here we use a simple and reliable method to isolate Down syndrome NSCs from aborted human fetal cortices and grow them in culture. The methodology provides specific aspects of harvesting the tissue, dissection with limited anatomical landmarks, cell sorting, plating and passaging of human NSCs. We also provide some basic protocols for inducing differentiation of human NSCs into more selective cell subtypes.

A simple and reliable method on isolation and culture of neural stem cells from discarded human fetal cortical tissue is described. Cultures derived from known human neurological disorders can be used for characterization of pathological cellular and molecular processes, as well as provide a platform to assess pharmacological efficacy.

Neural stem cells (NSCs) reside along the ventricular zone neuroepithelium during the development of the cortical plate. These early progenitors ...

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 ...

Analysis of Neural Crest Migration and Differentiation by Cross-species Transplantation

Avian embryos provide a unique platform for studying many vertebrate developmental processes, due to the easy access of the embryos within the egg. Chimeric avian embryos, in which quail donor tissue is transplanted into a chick embryo in ovo, combine the power of indelible genetic labeling of cell populations with the ease of manipulation presented by the avian embryo.
Quail-chick chimeras are a classical tool for tracing migratory neural crest cells (NCCs)1-3. NCCs are a transient migratory population of cells in the embryo, which originate in the dorsal region of the developing neural tube4. They undergo an epithelial to mesenchymal transition and subsequently migrate to other regions of the embryo, where they differentiate into various cell types including cartilage5-13, melanocytes11,14-20, neurons and glia21-32. NCCs are multipotent, and their ultimate fate is influenced by 1) the region of the neural tube in which they originate along the rostro-caudal axis of the embryo11,33-37, 2) signals from neighboring cells as they migrate38-44, and 3) the microenvironment of their ultimate destination within the embryo45,46. Tracing these cells from their point of origin at the neural tube, to their final position and fate within the embryo, provides important insight into the developmental processes that regulate patterning and organogenesis.
Transplantation of complementary regions of donor neural tube (homotopic grafting) or different regions of donor neural tube (heterotopic grafting) can reveal differences in pre-specification of NCCs along the rostro-caudal axis2,47. This technique can be further adapted to transplant a unilateral compartment of the neural tube, such that one side is derived from donor tissue, and the contralateral side remains unperturbed in the host embryo, yielding an internal control within the same sample2,47. It can also be adapted for transplantation of brain segments in later embryos, after HH10, when the anterior neural tube has closed47.
Here we report techniques for generating quail-chick chimeras via neural tube transplantation, which allow for tracing of migratory NCCs derived from a discrete segment of the neural tube. Species-specific labeling of the donor-derived cells with the quail-specific QCPN antibody48-56 allows the researcher to distinguish donor and host cells at the experimental end point. This technique is straightforward, inexpensive, and has many applications, including fate-mapping, cell lineage tracing, and identifying pre-patterning events along the rostro-caudal axis45. Because of the ease of access to the avian embryo, the quail-chick graft technique may be combined with other manipulations, including but not limited to lens ablation40, injection of inhibitory molecules57,58, or genetic manipulation via electroporation of expression plasmids59-61, to identify the response of particular migratory streams of NCCs to perturbations in the embryo's developmental program. Furthermore, this grafting technique may also be used to generate other interspecific chimeric embryos such as quail-duck chimeras to study NCC contribution to craniofacial morphogenesis, or mouse-chick chimeras to combine the power of mouse genetics with the ease of manipulation of the avian embryo.62

An approach for analyzing migration and eventual fate of avian neural crest cells in quail-chick chimeric embryos is described. This method is a simple and straightforward technique for tracing neural crest cells during migration and differentiation that are otherwise difficult to distinguish within an unmanipulated chick embryo.

Avian embryos provide a unique platform for studying many vertebrate developmental processes, due to the easy access of the embryos within the egg. ...

Isolation and Culture of Neural Crest Cells from Embryonic Murine Neural Tube

The embryonic neural crest (NC) is a multipotent progenitor population that originates at the dorsal aspect of the neural tube, undergoes an epithelial to mesenchymal transition (EMT) and migrates throughout the embryo, giving rise to diverse cell types 1-3. NC also has the unique ability to influence the differentiation and maturation of target organs4-6. When explanted in vitro, NC progenitors undergo self-renewal, migrate and differentiate into a variety of tissue types including neurons, glia, smooth muscle cells, cartilage and bone. 
NC multipotency was first described from explants of the avian neural tube7-9. In vitro isolation of NC cells facilitates the study of NC dynamics including proliferation, migration, and multipotency. Further work in the avian and rat systems demonstrated that explanted NC cells retain their NC potential when transplanted back into the embryo10-13. Because these inherent cellular properties are preserved in explanted NC progenitors, the neural tube explant assay provides an attractive option for studying the NC in vitro. 
To attain a better understanding of the mammalian NC, many methods have been employed to isolate NC populations. NC-derived progenitors can be cultured from post-migratory locations in both the embryo and adult to study the dynamics of post-migratory NC progenitors11,14-20, however isolation of NC progenitors as they emigrate from the neural tube provides optimal preservation of NC cell potential and migratory properties13,21,22. Some protocols employ fluorescence activated cell sorting (FACS) to isolate a NC population enriched for particular progenitors11,13,14,17. However, when starting with early stage embryos, cell numbers adequate for analyses are difficult to obtain with FACS, complicating the isolation of early NC populations from individual embryos. Here, we describe an approach that does not rely on FACS and results in an approximately 96% pure NC population based on a Wnt1-Cre activated lineage reporter23. 
The method presented here is adapted from protocols optimized for the culture of rat NC11,13. The advantages of this protocol compared to previous methods are that 1) the cells are not grown on a feeder layer, 2) FACS is not required to obtain a relatively pure NC population, 3) premigratory NC cells are isolated and 4) results are easily quantified. Furthermore, this protocol can be used for isolation of NC from any mutant mouse model, facilitating the study of NC characteristics with different genetic manipulations. The limitation of this approach is that the NC is removed from the context of the embryo, which is known to influence the survival, migration and differentiation of the NC2,24-28.

Isolation of embryonic neural crest from the neural tube facilitates the use of in vitro methods for studying migration, self-renewal, and multipotency of neural crest.

The embryonic neural crest (NC) is a multipotent progenitor population that originates at the dorsal aspect of the neural tube, undergoes an epithelial to ...

Progenitor-derived Oligodendrocyte Culture System from Human Fetal Brain

Differentiation of human neural progenitors into neuronal and glial cell types offers a model to study and compare molecular regulation of neural cell lineage development. In vitro expansion of neural progenitors from fetal CNS tissue has been well characterized. Despite the identification and isolation of glial progenitors from adult human sub-cortical white matter and development of various culture conditions to direct differentiation of fetal neural progenitors into myelin producing oligodendrocytes, acquiring sufficient human oligodendrocytes for in vitro experimentation remains difficult. Differentiation of galactocerebroside+ (GalC) and O4+ oligodendrocyte precursor or progenitor cells (OPC) from neural precursor cells has been reported using second trimester fetal brain. However, these cells do not proliferate in the absence of support cells including astrocytes and neurons, and are lost quickly over time in culture. The need remains for a culture system to produce cells of the oligodendrocyte lineage suitable for in vitro experimentation.
Culture of primary human oligodendrocytes could, for example, be a useful model to study the pathogenesis of neurotropic infectious agents like the human polyomavirus, JCV, that in vivo infects those cells. These cultured cells could also provide models of other demyelinating diseases of the central nervous system (CNS). Primary, human fetal brain-derived, multipotential neural progenitor cells proliferate in vitro while maintaining the capacity to differentiate into neurons (progenitor-derived neurons, PDN) and astrocytes (progenitor-derived astrocytes, PDA) This study shows that neural progenitors can be induced to differentiate through many of the stages of oligodendrocytic lineage development (progenitor-derived oligodendrocytes, PDO). We culture neural progenitor cells in DMEM-F12 serum-free media supplemented with basic fibroblast growth factor (bFGF), platelet derived growth factor (PDGF-AA), Sonic hedgehog (Shh), neurotrophic factor 3 (NT-3), N-2 and triiodothyronine (T3). The cultured cells are passaged at 2.5e6 cells per 75cm flasks approximately every seven days. Using these conditions, the majority of the cells in culture maintain a morphology characterized by few processes and express markers of pre-oligodendrocyte cells, such as A2B5 and O-4. When we remove the four growth factors (GF) (bFGF, PDGF-AA, Shh, NT-3) and add conditioned media from PDN, the cells start to acquire more processes and express markers specific of oligodendrocyte differentiation, such as GalC and myelin basic protein (MBP). We performed phenotypic characterization using multicolor flow cytometry to identify unique markers of oligodendrocyte.

Primary, human fetal brain-derived, multipotential progenitor cells proliferate in vitro while maintaining the capacity to differentiate into neurons and astrocytes. This work shows that neural progenitors can be induced to differentiate through stages of the oligodendrocytic lineage by conditioning with select growth factors.

Differentiation of human neural progenitors into neuronal and glial cell types offers a model to study and compare molecular regulation of neural cell ...

One Mouse, Two Cultures: Isolation and Culture of Adult Neural Stem Cells from the Two Neurogenic Zones of Individual Mice

The neurosphere assay and the adherent monolayer culture system are valuable tools to determine the potential (proliferation or differentiation) of adult neural stem cells in vitro. These assays can be used to compare the precursor potential of cells isolated from genetically different or differentially treated animals&#160;to determine the effects of exogenous factors on neural precursor cell proliferation and differentiation and to generate neural precursor cell lines that can be assayed over continuous passages. The neurosphere assay is traditionally used for the post-hoc identification of stem cells, primarily due to the lack of definitive markers with which they can be isolated from primary tissue&#160;and has the major advantage of giving a quick estimate of precursor cell numbers in brain tissue derived from individual animals. Adherent monolayer cultures, in contrast, are not traditionally used to compare proliferation between individual animals, as each culture is generally initiated from the combined tissue of between 5-8 animals. However, they have the major advantage that, unlike neurospheres, they consist of a mostly homogeneous population of precursor cells and are useful for following the differentiation process in single cells. Here, we describe, in detail, the generation of neurosphere cultures and, for the first time, adherent cultures from individual animals. This has many important implications including paired analysis of proliferation and/or differentiation potential in both the subventricular zone (SVZ) and dentate gyrus (DG) of treated or genetically different mouse lines, as well as a significant reduction in animal usage.

Here we describe a detailed protocol for the simultaneous generation of neural precursor cell cultures, as either adherent monolayers or neurospheres, from the subventricular zone and dentate gyrus of individual adult mice.

The neurosphere assay and the adherent monolayer culture system are valuable tools to determine the potential (proliferation or differentiation) of adult ...

Organotypic Slice Cultures to Study Oligodendrocyte Dynamics and Myelination

NG2 expressing cells (polydendrocytes, oligodendrocyte precursor cells) are the fourth major glial cell population in the central nervous system. During embryonic and postnatal development they actively proliferate and generate myelinating oligodendrocytes. These cells have commonly been studied in primary dissociated cultures, neuron cocultures, and in fixed tissue. Using newly available transgenic mouse lines slice culture systems can be used to investigate proliferation and differentiation of oligodendrocyte lineage cells in both gray and white matter regions of the forebrain and cerebellum. Slice cultures are prepared from early postnatal mice and are kept in culture for up to 1 month. These slices can be imaged multiple times over the culture period to investigate cellular behavior and interactions. This method allows visualization of NG2 cell division and the steps leading to oligodendrocyte differentiation while enabling detailed analysis of region-dependent NG2 cell and oligodendrocyte functional heterogeneity. This is a powerful technique that can be used to investigate the intrinsic and extrinsic signals influencing these cells over time in a cellular environment that closely resembles that found in vivo.

A technique to study NG2 cells and oligodendrocytes using a slice culture system of the forebrain and cerebellum is described. This method allows examination of the dynamics of proliferation and differentiation of cells within the oligodendrocyte lineage where the extracellular environment can be easily manipulated while maintaining tissue cytoarchitecture.

NG2 expressing cells (polydendrocytes, oligodendrocyte precursor cells) are the fourth major glial cell population in the central nervous system. During ...

Functional Evaluation of Biological Neurotoxins in Networked Cultures of Stem Cell-derived Central Nervous System Neurons

Therapeutic and mechanistic studies of the presynaptically targeted clostridial neurotoxins (CNTs) have been limited by the need for a scalable, cell-based model that produces functioning synapses and undergoes physiological responses to intoxication. Here we describe a simple and robust method to efficiently differentiate murine embryonic stem cells (ESCs) into defined lineages of synaptically active, networked neurons. Following an 8 day differentiation protocol, mouse embryonic stem cell-derived neurons (ESNs) rapidly express and compartmentalize neurotypic proteins, form neuronal morphologies and develop intrinsic electrical responses. By 18 days after differentiation (DIV 18), ESNs exhibit active glutamatergic and &#947;-aminobutyric acid (GABA)ergic synapses and emergent network behaviors characterized by an excitatory:inhibitory balance. To determine whether intoxication with CNTs functionally antagonizes synaptic neurotransmission, thereby replicating the in vivo pathophysiology that is responsible for clinical manifestations of botulism or tetanus, whole-cell patch clamp electrophysiology was used to quantify spontaneous miniature excitatory post-synaptic currents (mEPSCs) in ESNs exposed to tetanus neurotoxin (TeNT) or botulinum neurotoxin (BoNT) serotypes /A-/G. In all cases, ESNs exhibited near-complete loss of synaptic activity within 20 hr. Intoxicated neurons remained viable, as demonstrated by unchanged resting membrane potentials and intrinsic electrical responses. To further characterize the sensitivity of this approach, dose-dependent effects of intoxication on synaptic activity were measured 20 hr after addition of BoNT/A. Intoxication with 0.005 pM BoNT/A resulted in a significant decrement in mEPSCs, with a median inhibitory concentration (IC50) of 0.013 pM. Comparisons of median doses indicate that functional measurements of synaptic inhibition are faster, more specific and more sensitive than SNARE cleavage assays or the mouse lethality assay. These data validate the use of synaptically coupled, stem cell-derived neurons for the highly specific and sensitive detection of CNTs.

A custom protocol is described to differentiate mouse ES cells into defined populations of highly pure neurons exhibiting functioning synapses and emergent network behavior. Electrophysiological analysis demonstrates the loss of synaptic transmission following exposure to botulinum neurotoxin serotypes /A-/G and tetanus neurotoxin.

Therapeutic and mechanistic studies of the presynaptically targeted clostridial neurotoxins (CNTs) have been limited by the need for a scalable, ...

Modelling Zika Virus Infection of the Developing Human Brain In Vitro Using Stem Cell Derived Cerebral Organoids

The recent emergence of Zika virus (ZIKV) in susceptible populations has led to an abrupt increase in microcephaly and other neurodevelopmental conditions in newborn infants. While mosquitos are the main route of viral transmission, it has also been shown to spread via sexual contact and vertical mother-to-fetus transmission. In this latter case of transmission, due to the unique viral tropism of ZIKV, the virus is believed to predominantly target the neural progenitor cells (NPCs) of the developing brain.
Here a method for modeling ZIKV infection, and the resulting microcephaly, that occur when human cerebral organoids are exposed to live ZIKV is described. The organoids display high levels of virus within their neural progenitor population, and exhibit severe cell death and microcephaly over time. This three-dimensional cerebral organoid model allows researchers to conduct species-matched experiments to observe and potentially intervene with ZIKV infection of the developing human brain. The model provides improved relevance over standard two-dimensional methods, and contains human-specific cellular architecture and protein expression that are not possible in animal models.

Modelling Zika Virus Infection of the Developing Human Brain In Vitro Using Stem Cell Derived Cerebral Organoids

This protocol describes a technique used to model Zika virus infection of the developing human brain. Using wildtype or engineered stem cell lines, researchers may use this technique to uncover the various mechanisms or treatments that may affect early brain infection and resulting microcephaly in Zika virus-infected embryos.

The recent emergence of Zika virus (ZIKV) in susceptible populations has led to an abrupt increase in microcephaly and other neurodevelopmental conditions ...