Work in S.M.'s laboratory is funded by recruitment grants from CPRIT (R1220) and NIH (1R01GM113023-01).

1. Isolation of Neurospheres from the Adult Mouse Brain
<ol>
	<li>Euthanize an adult mouse (around 2 months old) by an overdose of isoflurane. Double-check that the mouse has stopped breathing and dissect immediately after death.</li>
	<li>Using scissors, make a midline incision to open the skull. Remove the brain.</li>
	<li>Place the brain in cold PBS in a 10 cm dish on ice. Follow the whole-mount dissection method to obtain the SVZ from the lateral ventricle48.</li>
	<li>Place the lateral ventricle into a 1.5 mL tube, add 500 &#181;L of 0.05% trypsin-EDTA in PBS, and incubate the tube for 15 min at 37 &#176;C in a water bath.</li>
	<li>After 15 min, add 500 &#181;L of stopping medium and gently pipet 20 - 30 times with a 1 mL tip. Avoid forming air bubbles during pipetting. 
	NOTE: This step is critical for cell survival.</li>
	<li>Spin down the cells at 500 x g for 8 min. Discard the supernatant, add 1 mL of PBS, and resuspend the cells by gently pipetting 5x with a 1 mL tip.</li>
	<li>Spin down at 500 x g for 8 min. Discard the supernatant using a 1 mL tip and add 1 mL of basal medium.</li>
	<li>(Optional) If cellular debris are observed, pass the cells through a 70 &#181;m cell-strainer.</li>
	<li>Count the number of cells with a hemocytometer; in general, about 30,000 - 60,000 cells/SVZ are obtained.</li>
	<li>Plate the cells from one SVZ into a 10 cm dish with 10 mL of NSC medium and culture at 37 &#176;C with 5% CO2.</li>
	<li>(Optional) To avoid fusion between spheres49, put 1,000 cells in a single well of an ultra-low-binding 6-well plate that is prefilled with 1.5 mL of NSC medium and culture at 37 &#176;C with 5% CO2. 
	NOTE: After 5-7 days, neurospheres can be observed (Figure 2A). The culturing period may differ with the age of mouse or the genetic background.</li>
	<li>Add 2 mL of NSC medium every 3-4 days to maintain the culture (do not remove the existing medium).</li>
</ol>
2. Analysis of the Differentiation Capacity of Neurospheres and Ciliogenesis Tests
<ol>
	<li>To analyze the differentiation capacity, analyze the neurospheres under adherent conditions in differentiation medium.</li>
	<li>Sterilize 12 mm round coverslips by autoclaving or with UV exposure prior to use. For an adherent cell culture, put a sterilized 12 mm round cover glass into a well of a 24-well plate under aseptic conditions.</li>
	<li>Coat the cover glass for 10 s with 500 &#181;L of 0.002% poly-L-Lysine (PLL). Aspirate the solution and dry it for 10-15 min.</li>
	<li>Add 500 &#181;L of laminin solution (5 &#181;g/&#181;L). Incubate the cover glass for 1 h at 37 &#176;C.</li>
	<li>Aspirate the laminin and add 500 &#181;L of differentiation medium or NSC medium (undifferentiated control).</li>
	<li>For the differentiation assay, pick up a 100 - 200 &#181;m sphere with a 200 &#181;L tip under the microscope. Add 5-10 neurospheres to each well of a 24-well plate and culture for 7-10 days in differentiation medium.</li>
	<li>To analyze undifferentiated neurospheres, add 5-10 neurospheres to each well of a 24-well plate and culture for 1-2 days in NSC medium. Attached neurospheres spread and grow as a monolayer (Figure 2B).</li>
	<li>After carefully removing the medium, fix the cells with 4% paraformaldehyde (PFA) in PBS for 15 min at RT, and then wash with PBS twice for 5 min at RT. The plate can be stored at 4 &#176;C for 1-2 months. 
	NOTE: To visualize neural stem/progenitor cells in NSC medium and differentiated cells in differentiation medium, perform immunostaining against Nestin (neural stem/progenitor cell marker), &#946;&#8211;tubulin III (TUJ1 monoclonal, neuronal marker), GFAP (Glial Fibrillary Acidic Protein, astrocyte marker), and O4 (oligodendrocyte marker) (Figure 2B-E). To analyze the cilia, perform immunostaining against Arl13b (primary cilia marker) and Gpr161 (ciliary GPCR) (Figure 3).</li>
	<li>Mount the cover glass with mounting solution onto a slide glass. Tilt the slide glass to remove excess solution.</li>
</ol>
3. Analysis of Ciliogenesis in Neurospheres
<ol>
	<li>To analyze cells in intact neurospheres by immunostaining, transfer 1 mL of culturing medium containing multiple neurospheres to a 1.5 mL tube and spin down at 500 x g for 8 min. Fix the spheres with 4% (PFA) after removing the medium for 15 min and wash with PBS.&#160;Spin down spheres at 500 x g for 8 min, remove supernatant, and incubate the spheres O/N with 30% sucrose at 4&#176;C.&#160;</li>
	<li>Discard the supernatant using a 1 mL tip and add 500 &#181;L of OCT solution using a cut 1 mL tip. Cut the edge of the tip to widen the opening, as OCT is viscous.</li>
	<li>Transfer the OCT solution containing the neurospheres into a disposable plastic freezing mold (10 mm x 10 mm x 5 mm).</li>
	<li>Freeze the mold on dry ice for at least 15 min.</li>
	<li>(Optional) Stop the experiment and store the mold in a -80 &#176;C freezer for up to 1 year.</li>
	<li>Cut the sections with a cryostat; the thickness of sections should be 15-30 &#181;m.</li>
	<li>To visualize the primary cilia in a neurosphere, perform immunostaining against Arl13b (Figure 4).</li>
</ol>
4. Culture and Passage of Neurospheres and Adherent NSCs
<ol>
	<li>Passage the neurospheres while the sphere size is between 100-200 &#181;m; when the neurospheres are too big (300 &#181;m or above), they are not ideal for experiments.</li>
	<li>Transfer the neurospheres into a 50 mL tube using a 1 mL tip and spin down at 500 x g for 8 min.</li>
	<li>Discard the supernatant using an aspirator, add 500 &#181;L of 0.05% trypsin-EDTA in PBS, and incubate for 5 min at 37 &#176;C. The amount of trypsin varies depending on the number of spheres.</li>
	<li>Add 500 &#181;L of serum medium and gently pipet 20 times with a 1 mL tip.</li>
	<li>Spin down at 500 x g for 8 min. Discard the supernatant using a 1 mL tip and add 1 mL of basal medium.</li>
	<li>(Optional) If cellular debris or undissociated neurospheres are seen, pass the cells through a 70 &#181;m cell-strainer.</li>
	<li>Passage the cells in a 10 cm dish at a density of 10,000 cells/cm2 in NSC medium; the cells will be ready for the next passage after a week.</li>
	<li>(Optional) To freeze the cells, add freezing medium to generate 500,000 to 1,000,000 cells/mL&#160;suspension. Freeze the cells using cryo-freezing containers; undissociated neurospheres can also be frozen.</li>
	<li>For the adherent culture, dillute the cells to 50,000 cells/cm2&#160;onto PLL- and Laminin-coated coverslips with NSC medium, and culture for 1-2 days.</li>
</ol>
5. Starvation and Analysis of Cilia
<ol>
	<li>Prepare starvation medium (Table 1).</li>
	<li>1-2 days after the initial plating of adherent cells after dissociation, change to NSC medium in one well (control) and starvation medium in another well (experiment).</li>
	<li>Culture the adherent cells for 24 h.</li>
	<li>Fix the cells with 4% PFA in PBS for 15 min at RT and wash twice with PBS&#160;for 5 min each at RT.</li>
	<li>(Optional) Stop the experiment and store the 24-well plates with coverslips in PBS at 4 &#176;C for 1-2 months.</li>
	<li>Perform an immunofluorescence protocol for staining7,50.</li>
	<li>Mount the coverslips using mounting solution. Dry the slide glass O/N&#160;at RT in the dark.</li>
	<li>Acquire images on a compound microscope at the necessary magnification. Use a microscope, a camera, and objectives (40X/1.3 oil and 63X/1.4 oil), controlled using the accompanied software. Take sufficient z-sections at 0.5-0.8 &#181;m intervals (Figure 5).</li>
	<li>For the quantitative analysis of ciliary localization in adherent cells, acquire stacks of images from 3-8 consecutive fields with confluent cells by looking into the DAPI channel. Quantify the number of primary cilia using ImageJ/Fiji. Typically, use the &#34;Cell counter&#34; tool in the ImageJ Plugins&#62;Analyze dialog box to count the cells with GPCR-positive cilia; maximal projections from stacks of images can also be exported from ImageJ/Fiji.Use similar image intensity and contrast parameters for all images from the same experiment for counting and exporting purposes.</li>
</ol>
6. Transfection of Neurospheres
<ol>
	<li>Adhere dissociated cells to coverslips for 24 h. Use a cell density typically between 75,000 and 150,000 cells in 500 &#181;L of NSC medium in a single well of a 24-well plate.</li>
	<li>Mix 25 &#181;L of reduced serum medium and 1.5 &#181;L of transfection reagent in a 0.5 mL microtube by vortexing for 5 s.</li>
	<li>Add 2.5 &#181;g of endotoxin-free plasmid DNA to a separate 0.5 mL microtube containing 25 &#181;L of reduced serum media and mix by vortexing for 5 s.</li>
	<li>Add 1 &#181;L of transfection reagent to the second microtube containing DNA and mix by vortexing for 5 s.</li>
	<li>Add the mixture from the DNA-containing tube to the first microtube and mix by pipetting.</li>
	<li>Incubate the mixture for 10-15 min at RT. After incubation, gently add the transfection mix dropwise to the wells on top of the NSC medium (500 &#181;L/well).</li>
	<li>Change the medium 24 h post-transfection to control medium (NSC medium) or starvation medium (500 &#181;L/well).</li>
	<li>Fix the cells by changing the medium to 4% PFA&#160;after 24 h and perform immunostaining; typically, up to a 10% transfection efficiency is obtained using this protocol.</li>
</ol>

The authors declare no competing financial interests.

Here, we describe a method to generate and maintain neurosphere cultures from adult mouse SVZ. A few pertinent points regarding the cultures are as follows. First, the sizes of the spheres are typically between 50 - 200 &#181;m. In our experience, when a neurosphere gets larger than 300 &#181;m in diameter, the optimal time for passaging has been missed. These larger spheres contain dead cells in the core. Second, as neurospheres are commonly used to study neural stem/progenitor cells, it is important to use EGF and FGF-basic (bFGF) to maintain the stemness of these cells. Therefore, factors that induce differentiation, such as FBS, must be avoided during the maintenance of the neurospheres. It has been shown that 10% FBS can differentiate neurospheres into astrocytes47. Third, the degree of neuronal differentiation might depend on the age of mouse. If one desires more neuronal differentiation from neurospheres, younger mice (postnatal day 0) can be used. Fourth, if starting new cultures, it is always good to examine the differentiation capacity of the neurospheres. Fifth, our method also allows freezing and thawing from established neurosphere cultures for future experimental use. After thawing, cells typically grow as adherent spindle-shaped cells without poly-L-lysine or laminin. This may be caused by freeze-thaw damage to the cells. It is important to keep the cells growing until they become 60 - 80% confluent before passage. After the passage, cells typically grow as neurospheres and can generally be passaged up to 5 - 10 times.
Neurosphere-based culture methods allow us to study ciliary trafficking and signaling pathways in neural stem/progenitor cells and differentiated neurons. We examined primary cilia in floating/adherent and differentiated neurospheres. It is important to acquire several z-sections to visualize cilia throughout the full extent of the neurospheres. Furthermore, after generating adherent cultures of these neurospheres, and starving them for 24 h, most of the cells are ciliated (Figure 5) and enriched in the orphan GPCR, Gpr161. However, prolonged starvation of the adherent cultures results in subcellular vacuolar structures. Thus, it is important to carefully optimize starvation periods for particular experimental purposes.
Currently, studies on ciliogenesis are predominantly performed in a few ciliated, immortalized cell lines57. These cell lines do not always faithfully recapitulate neurons and neural stem/progenitor cells in expressing signaling components, such as GPCRs or Shh pathway machinery, making it imperative to develop newer methods to study the role of cilia in biological processes. The neurosphere-based methods described here allow us to study cilia-regulated cellular pathways, including GPCR and Shh signaling in the context of neural stem/progenitor cells and differentiated neurons.

The primary cilium is a microtubule-based dynamic subcellular compartment that functions as a sensory antenna in coordinating cellular signaling pathways, including the Sonic Hedgehog (Shh) pathway during embryonic neuronal development1,2, and compartmentalized subcellular signaling in adult neuronal function3,4. Signaling components of these pathways, such as the Shh receptor Patched5; the pathway activator Smoothened (Smo)6; and Gpr1617, an orphan G Protein-coupled Receptor (GPCR) that negatively regulates the Shh pathway, localize to cilia in a dynamic fashion. Multiple GPCRs have been reported to localize to the cilia in neurons in the brain7,8,9,10,11,12,13,14,15,16. Defects in cilia and cilia-generated signaling pathways affect multiple tissues and are collectively known as ciliopathies17,18,19. The ciliopathy disease spectrum frequently includes neurodevelopmental defects, such as craniofacial abnormalities20,21,22. In addition, primary cilia in hypothalamic neurons regulate central satiety pathways, and defects result in central obesity23, mirroring obesity in syndromic ciliopathies such as Bardet Biedel syndrome24. In addition, neuropeptide receptor signaling in cilia regulates central satiety pathways11,14. Ciliary localization of Adenylyl Cyclase III (ACIII) and GPCRs such as somatostatin receptor 3 in hippocampal neurons result in novel object recognition defects and memory deficits25,26 and parallels a lack of ciliary integrity27. The developmental aspects of cilia-generated signaling are closely tied to tissue homeostasis; in particular, cilia are important to the progression of Shh-subtype medulloblastomas arising from granule progenitors in the cerebellum28,29. Thus, primary cilia play important roles during embryonic, postnatal, and adult neuronal development and function.
Neural Stem Cells (NSCs) reside in the subventricular zone (SVZ) of the lateral ventricle, the subgranular zone of the dentate gyrus of the hippocampus, and the ventricular zone of the third ventricle in the hypothalamus in mammals30,31,32. NSCs are multipotent, possess the capacity for self-renewal, and are important for brain development and regenerative medicine30. Most NSCs in the SVZ are quiescent and possess a solitary primary cilium that, in many cases, extends out to the lateral ventricle33. The primary cilium signals via the localization of various receptors, inducing downstream cellular responses, particularly in relation to Shh, TGF&#946;, and receptor tyrosine kinase pathways2,34,35,36. Since primary cilia extend into the lateral ventricle, it is hypothesized that primary cilia detect cytokines in the cerebrospinal fluid (CSF) to activate NSCs37. Recent studies suggest that the Shh signaling pathway and primary cilia are critical for the activation of stem cells in the repair and regeneration of multiple tissues, including the olfactory epithelium, lung, and kidney38,39,40,41. However, the mechanisms by which CSF communicates with NSCs and whether primary cilia are involved are not known. Adherent NSCs in culture are ciliated; localize Shh pathway components, such as Smo and Gpr161 in cilia; and are Shh responsive42. Thus, NSCs can serve as an important model system to study the Shh pathway, ciliary trafficking, and neuronal differentiation pathways. In addition, neurons differentiated from NSCs can also be used for ciliary trafficking assays.
Neurospheres are constituted of clusters of free-floating cells arising from the proliferation of neural stem/progenitor cells that grow in the presence of specific growth factors and nonadhesive surfaces43,44. Neurospheres serve as important in vitro culture models to study neural stem/progenitor cells in normal development and disease31,45,46,47. Here, we describe a neurosphere-based assay for culturing neural stem/progenitor cells and for differentiation into neurons/glia. We particularly emphasize the trafficking of signaling components to cilia of neural stem/progenitor cells and differentiated neurons (Figure 1). As opposed to culturing primary neurons, primary neurospheres are relatively easy to culture, are amenable to multiple passages and freeze-thaw cycles, and can undergo differentiation into neurons/glia. Importantly, we determined that neurosphere-derived neural stem/progenitor cells and differentiated neurons are ciliated in culture and localize signaling molecules relevant to ciliary function in these compartments. Neurosphere-based culturing methods can serve as an ideal model system for studying ciliogenesis and ciliary trafficking in NSCs and differentiated neurons.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>12 mm round cover glass</td>
			<td>Fisherbrand</td>
			<td>12-545-80&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>24 well plate</td>
			<td>Falcon</td>
			<td>353047</td>
			<td></td>
		</tr>
		<tr>
			<td>4 % paraformaldehyde</td>
			<td>Affymetrix</td>
			<td>19943</td>
			<td></td>
		</tr>
		<tr>
			<td>50 ml tube</td>
			<td>Falcon</td>
			<td>352098</td>
			<td></td>
		</tr>
		<tr>
			<td>95 mm X 15 mm petri dish, slippable lid</td>
			<td>Fisherbrand</td>
			<td>FB0875714G</td>
			<td>10 cm dish</td>
		</tr>
		<tr>
			<td>70 &#181;m cell strainer</td>
			<td>Falcon</td>
			<td>352350</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 Affinipure Donkey Anti-Rabbit IgG (H+L)</td>
			<td>Jackson Immunoresearch</td>
			<td>711-545-152</td>
			<td>Donkey anti Rabbit, Alexa 488 secondary antibody</td>
		</tr>
		<tr>
			<td>Arl13B, Clone N295B/66</td>
			<td>Neuromab</td>
			<td>AB_11000053</td>
			<td></td>
		</tr>
		<tr>
			<td>B-27 Supplement (50X), serum free</td>
			<td>ThermoFisher Scientific</td>
			<td>17504001</td>
			<td>B27</td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>Thermo scientific</td>
			<td>ST 40R</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryogenic vial</td>
			<td>Corning</td>
			<td>430488</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma</td>
			<td>D9542-10MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Deoxyribonuclease I from bovine pancrease</td>
			<td>Sigma</td>
			<td>D5025-15KU</td>
			<td>Dnase I</td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide</td>
			<td>Sigma</td>
			<td>D8418-100ML</td>
			<td>DMSO</td>
		</tr>
		<tr>
			<td>Disposable Vinyl Specimen Molds</td>
			<td>Sakura Tissue-Tek Cryomold</td>
			<td>4565</td>
			<td>10 mm X 10 mm X 5 mm</td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Phosphate Buffered Saline 10&#215;, Modified, without calcium chloride and magnesium chloride, liquid, sterile-filtered, suitable for cell culture</td>
			<td>Sigma</td>
			<td>D1408-500ML</td>
			<td>PBS</td>
		</tr>
		<tr>
			<td>Dumont #5 Forceps</td>
			<td>Fine science tools</td>
			<td>11254-20</td>
			<td></td>
		</tr>
		<tr>
			<td>FBS&#160;</td>
			<td>Sigma</td>
			<td>F9026-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoromount-G solution</td>
			<td>Southern Biotech</td>
			<td>0100-01</td>
			<td>mounting solution</td>
		</tr>
		<tr>
			<td>GFAP</td>
			<td>DAKO</td>
			<td>Z0334</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti Mouse IgG1 Secondary Antibody, Alexa Fluor 555 conjugate</td>
			<td>ThermoFisher Scientific</td>
			<td>A-21127</td>
			<td>Goat anti Mouse IgG1, Alexa 555 secondary antibody</td>
		</tr>
		<tr>
			<td>Goat anti Mouse IgG2a Secondary Antibody, Alexa Fluor 555 conjugate</td>
			<td>ThermoFisher Scientific</td>
			<td>A-21137</td>
			<td>Goat anti Mouse IgG2a, Alexa 555 secondary antibody</td>
		</tr>
		<tr>
			<td>Gpr161</td>
			<td>home made</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>human bFGF</td>
			<td>Sigma</td>
			<td>F0291</td>
			<td>FGF</td>
		</tr>
		<tr>
			<td>hemocytometer</td>
			<td>Hausser Scientific</td>
			<td>0.100 mm deep</td>
			<td>improved neubauer</td>
		</tr>
		<tr>
			<td>Isothesia</td>
			<td>Henry Schein</td>
			<td>NDC 11695-0500-2</td>
			<td>Isofluorane</td>
		</tr>
		<tr>
			<td>Laminin from Engelbreth-Holm-Swarm Sarcoma basement membrane</td>
			<td>Sigma</td>
			<td>L2020</td>
			<td>Laminin</td>
		</tr>
		<tr>
			<td>L-Glutamine (200mM)</td>
			<td>Sigma</td>
			<td>G7513</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipofectamine 3000 Transfection Reagent</td>
			<td>ThermoFisher Scientific</td>
			<td>L3000</td>
			<td></td>
		</tr>
		<tr>
			<td>Mr. Frosty</td>
			<td>Nalgene&#160;</td>
			<td>5100-0036</td>
			<td></td>
		</tr>
		<tr>
			<td>N-2 supplement (100X)</td>
			<td>ThermoFisher Scientific</td>
			<td>17502001</td>
			<td>N2</td>
		</tr>
		<tr>
			<td>Neurobasal medium</td>
			<td>Gibco</td>
			<td>21103-049</td>
			<td></td>
		</tr>
		<tr>
			<td>Normal Donkey Serum</td>
			<td>Jackson ImmunoResearch</td>
			<td>017-000-121</td>
			<td></td>
		</tr>
		<tr>
			<td>OCT compound</td>
			<td>Sakura Tissue-Tek</td>
			<td>4583</td>
			<td>OCT</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Sigma</td>
			<td>P4333-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-L-lysine</td>
			<td>Sigma</td>
			<td>P4707</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant human EGF protein, CF</td>
			<td>R and D systems</td>
			<td>236-EG-200</td>
			<td>EGF</td>
		</tr>
		<tr>
			<td>Scissor</td>
			<td>Fine science tools</td>
			<td>14060-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Superfrost plus microscope slide</td>
			<td>Fisher scientific</td>
			<td>12-550-15</td>
			<td>slides</td>
		</tr>
		<tr>
			<td>Triton X</td>
			<td>Bio-Rad</td>
			<td>161-0407</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin-EDTA solution (10X)</td>
			<td>Sigma</td>
			<td>T4174-100</td>
			<td>Trypsin</td>
		</tr>
		<tr>
			<td>COSTAR 6 Well Plate, With Lid Flat Bottom Ultra-Low Attachment Surface Polystyrene, Sterile</td>
			<td>Corning</td>
			<td>3471</td>
			<td>ultra-low binding 6 well plate</td>
		</tr>
		<tr>
			<td>&#946;-tubulin III</td>
			<td>Covance</td>
			<td>MMS-435P</td>
			<td>TUJ1</td>
		</tr>
	</tbody>
</table>

using primary neurosphere cultures to study primary cilia

The primary cilium is fundamentally important for the proliferation of neural stem/progenitor cells and for neuronal differentiation during embryonic, postnatal, and adult life. In addition, most differentiated neurons possess primary cilia that house signaling receptors, such as G-protein-coupled receptors, and signaling molecules, such as adenylyl cyclases. The primary cilium determines the activity of multiple developmental pathways, including the sonic hedgehog pathway during embryonic neuronal development, and also functions in promoting compartmentalized subcellular signaling during adult neuronal function. Unsurprisingly, defects in primary cilium biogenesis and function have been linked to developmental anomalies of the brain, central obesity, and learning and memory deficits. Thus, it is imperative to study primary cilium biogenesis and ciliary trafficking in the context of neural stem/progenitor cells and differentiated neurons. However, culturing methods for primary neurons require considerable expertise and are not amenable to freeze-thaw cycles. In this protocol, we discuss culturing methods for mixed populations of neural stem/progenitor cells using primary neurospheres. The neurosphere-based culturing methods provide the combined benefits of studying primary neural stem/progenitor cells: amenability to multiple passages and freeze-thaw cycles, differentiation potential into neurons/glia, and transfectability. Importantly, we determined that neurosphere-derived neural stem/progenitor cells and differentiated neurons are ciliated in culture and localize signaling molecules relevant to ciliary function in these compartments. Utilizing these cultures, we further describe methods to study ciliogenesis and ciliary trafficking in neural stem/progenitor cells and differentiated neurons. These neurosphere-based methods allow us to study cilia-regulated cellular pathways, including G-protein-coupled receptor and sonic hedgehog signaling, in the context of neural stem/progenitor cells and differentiated neurons.

After plating the cells from the SVZ in NSC medium for a week, floating neurospheres were observed (Figure 2A). The sizes of the spheres varied between 50 and 200 &#181;m. To examine if the spheres were derived from neural stem/progenitor cells, the neurospheres were plated onto PLL- and Laminin-coated coverslips in NSC medium for 2 days. They were then immunostained against the neural stem/progenitor cell marker, Nestin. Two days were needed to allow the spheres to attach to the coverslips and to grow as a monolayer of cells. The monolayer of cells was positive for Nestin (Figure 2B). To examine the differentiation capacity of the neurospheres, we plated them following steps outlined in section 2 onto PLL- and Laminin-coated coverslips in differentiation medium without dissociation for 7-10 days. It takes at least a week for differentiation. The neurospheres attached to the coverslips and expanded to grow as a monolayer of cells. We fixed the cells and immunostained against neuron, astrocyte, and oligodendrocyte markers. We observed adherent neurospheres that differentiated into &#946;-Tubulin III-positive neurons, GFAP-positive astrocytes, and O4-positive oligodendrocytes (Figure 2C-E). Thus, multipotent neurospheres can be derived from adult mouse SVZ.
To determine if adherent neurospheres and differentiated neurons have primary cilia, we immunostained against Arl13b (primary cilium marker)51 and Gpr161 (ciliary-localized GPCR)7. In our experience, in available mouse models expressing ARL13B-mCherry, and in studies with developing mouse cortex, most (if not all) neuronal cilia in the brain express Arl13b, while all neurons are not ACIII-positive52,53,54. Thus, we mainly rely on Arl13b to detect neuronal cilia. We detected primary cilia and the localization of Gpr161 to the cilia of progenitor cells in adhered neurospheres (Figure 3A) and in differentiated neurons (Figure 3B). Thus, adherent neurospheres and differentiated lineages are ciliated in culture and localize signaling molecules.
To determine if floating neurospheres have primary cilia, we cryosectioned floating neurospheres after fixation, as outlined in section 3. We observed primary cilia in neural stem/progenitor cells in the sectioned neurospheres (Figure 4A-B). It is important to acquire z-sections across the neurospheres to fully visualize primary cilia, because it is difficult to examine the primary cilia in a single z-plane. In summary, undifferentiated neurospheres have heterogeneous populations of ciliated and non-ciliated cells.
To analyze the ciliogenesis of neural stem/progenitor cells, we dissociated neurospheres and plated the cells onto PLL- and Laminin-coated coverslips. After culturing them in starvation medium, we observed an increased percentage of cilia, as detected by Arl13b, as compared to those cultured in NSC medium (% ciliation) (Figure 5A-B). In addition, we noted an increased number of Gpr161-positive cilia upon starvation (% Gpr161-positive cilia/Arl13b-positive cilia) (Figure 5A-B). Interestingly, a previous study using undifferentiated human embryonic stem cell lines demonstrated that human embryonic stem cells in culture also possess primary cilia55. The number and length of cilia increase upon serum starvation in these cell lines, and these cilia also possess components of the Shh machinery.
We also transfected adherent neural stem/progenitor cells, as outlined in section 6. We typically detected a transfection efficiency of up to 10%, similar to primary neuron cultures in the lab (Figure 6).
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Solution</td>
			<td>Component</td>
		</tr>
		<tr>
			<td>1x PBS</td>
			<td>Dilute 10x PBS with autoclaved water</td>
		</tr>
		<tr>
			<td>NSC medium</td>
			<td>2 mL 50x B27</td>
		</tr>
		<tr>
			<td></td>
			<td>1 mL 100x N2</td>
		</tr>
		<tr>
			<td></td>
			<td>1 mL 200 mM L-glutamine</td>
		</tr>
		<tr>
			<td></td>
			<td>1 mL 100x Pen/Strep</td>
		</tr>
		<tr>
			<td></td>
			<td>FGF-basic (human bFGF) (Stock solution 25 ng/mL in Neurobasal medium): 20 ng/mL or 2 mg total for 100 mL media</td>
		</tr>
		<tr>
			<td></td>
			<td>EGF (Stock solution 25 ng/mL in Neurobasal medium): 20 ng/mL or 2 mg total for 100 mL media&#160;</td>
		</tr>
		<tr>
			<td></td>
			<td>95 mL basal medium</td>
		</tr>
		<tr>
			<td>Human FGF-basic (human bFGF)</td>
			<td>1 mL basal medium to commercial human bFGF 25 &#956;g bottle</td>
		</tr>
		<tr>
			<td></td>
			<td>Store in sterile microtubes in -20 &#176;C or lower. Use a single 80 &#956;L aliquot or 2 &#956;g/100 mL NSC media.</td>
		</tr>
		<tr>
			<td>EGF</td>
			<td>Add 8 mL basal medium to commercial EGF 20 &#956;g bottle</td>
		</tr>
		<tr>
			<td></td>
			<td>Store as sterile microtubes in -20 &#176;C or lower. Use a single 80 &#956;g aliquot/100 mL media.</td>
		</tr>
		<tr>
			<td>Stopping medium</td>
			<td>1 mL FBS</td>
		</tr>
		<tr>
			<td></td>
			<td>75 U Dnase I (75 U/uL in PBS)</td>
		</tr>
		<tr>
			<td></td>
			<td>9 mL PBS</td>
		</tr>
		<tr>
			<td>Serum medium</td>
			<td>1 mL FBS</td>
		</tr>
		<tr>
			<td></td>
			<td>9 mL Basal medium</td>
		</tr>
		<tr>
			<td>Freezing medium</td>
			<td>4 mL DMSO</td>
		</tr>
		<tr>
			<td></td>
			<td>24 mL basal medium</td>
		</tr>
		<tr>
			<td></td>
			<td>12 mL 30% FBS</td>
		</tr>
		<tr>
			<td>Differentiation medium</td>
			<td>90 mL basal medium</td>
		</tr>
		<tr>
			<td></td>
			<td>5 mL 5% FBS</td>
		</tr>
		<tr>
			<td></td>
			<td>1 mL 100x Pen/Strep</td>
		</tr>
		<tr>
			<td></td>
			<td>1 mL 200 mM L-glutamine</td>
		</tr>
		<tr>
			<td></td>
			<td>1 mL 100x N2</td>
		</tr>
		<tr>
			<td></td>
			<td>2 mL 50x B27</td>
		</tr>
		<tr>
			<td></td>
			<td>FGF-basic (human bFGF) (Stock solution of 25 ng/mL in Neurobasal medium): 10 ng/mL or 1 mg total for 100 mL</td>
		</tr>
		<tr>
			<td>Starvation medium</td>
			<td>95 mL basal medium</td>
		</tr>
		<tr>
			<td></td>
			<td>1 mL 100x Pen/Strep</td>
		</tr>
		<tr>
			<td></td>
			<td>1 mL 200 mM L-glutamine</td>
		</tr>
		<tr>
			<td></td>
			<td>1 mL 100x N2</td>
		</tr>
		<tr>
			<td></td>
			<td>2 mL 50x B27</td>
		</tr>
	</tbody>
</table>
Table 1: Recipe of Solutions.
<img alt="Figure 1" src="/files/ftp_upload/55315/55315fig1.jpg" /> 
Figure 1: Schematic Diagram of Neurosphere Culture and Differentiation Protocols. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55315/55315fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55315/55315fig2.jpg" /> 
Figure 2: Neurospheres Exhibit Differentiation Capacity. (A) Representative neurospheres are shown. (B) Neurospheres were plated onto PLL- and Laminin-coated coverslips in NSC medium for 2 days and were immunostained against Nestin (neural stem/progenitor cell marker). (C-E) Neurospheres were plated following the steps outlined in section 2 onto PLL- and Laminin-coated coverslips in differentiation medium for 7 days without dissociation; fixed; and immunostained against &#946;-Tubulin III (neuronal marker), GFAP (astrocyte marker), and O4 (oligodendrocyte marker). The nuclei were stained by DAPI. Scale bar = 100 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55315/55315fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a> 
&#160;
<img alt="Figure 3" src="/files/ftp_upload/55315/55315fig3.jpg" /> 
Figure 3: Neurospheres and Neurons in Adherent Differentiated Neurospheres are Ciliated. (A) Undissociated neurospheres were plated onto coated coverslips in NSC medium for 2 d; fixed; and immunostained against Gpr161 (green), Arl13b (red), and DNA (blue). (B) Differentiated neurons were immunostained against Gpr161 (green), Arl13b (red, A) or &#946;-Tubulin III (red, B), and DNA (blue). White arrows indicate Gpr161 localization in cilia. The right panels in (A) are enlarged views of the primary cilium indicated by the white arrow. The Gpr161-positive cilium in the white-boxed neuron is shown to the right in (B). Scale = 10 &#181;m (A); 50 &#181;m (B). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55315/55315fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/55315/55315fig4.jpg" /> 
Figure 4: Floating, Undifferentiated Neurospheres Possess Ciliated Cells. Floating neurospheres were fixed, embedded in OCT, cryosectioned, and processed for immunofluorescence against Arl13b (red) and DNA. Arl13b-positive primary cilia in a neurosphere are shown. The panel in (A) is a maximum-intensity projection of 14 z-sections at 0.8 &#181;m intervals from a neurosphere. The panels in (B) show the enlarged view of ciliated cells in the white-boxed region in (A). The maximal projection image from all z-sections is shown as a stack, while numbers 1 to 14 indicate individual z-sections. The white arrows indicate primary cilia. Scale bar = 50 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55315/55315fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/55315/55315fig5.jpg" /> 
Figure 5: Starvation Induces Ciliogenesis in Dissociated Adherent Neural Stem/Progenitor Cells. (A) Dissociated cells were plated onto coated coverslips in NSC medium or starvation medium for 24 h; fixed; and immunostained against Gpr161 (green), Arl13b (red), and DNA (blue). Gpr161-positive cilia are observed in control (left) and starved (right) cells. The lower panels are enlarged views of the indicated white boxes in the respective upper panels. The white arrows and arrowheads indicate Gpr161-positive and -negative cilia, respectively. Scale bar = 100 &#181;m. (B) Quantification of Arl13b-positive ciliated cells (left graph) and Gpr161 localizing in Arl13b-positive cilia (right graph) in control and starved cells. The percentages of both Arl13b-positive ciliated cells and Gpr161 localizing in Arl13b-positive cilia increased upon starvation. The quantification was performed from two fields of view each from two technical replicates of a single experiment. The data are represented as the mean from all 4 fields of view &#177;the standard deviation. *, P &#60;0.05; **, P &#60;0.01. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55315/55315fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" src="/files/ftp_upload/55315/55315fig6.jpg" /> 
Figure 6: Transfection in Dissociated Adherent Neural Stem/Progenitor Cells. Dissociated cells plated on coated coverslips were transfected with a LAP-TULP3 N-terminus (1-183 aa) mut12 construct that does not bind to the core IFT-A complex56. The medium was changed to starvation medium 24 h post-transfection. The cells were fixed after 24 h and immunostained against GFP (green), Gpr161 (red), Arl13b (magenta), and DNA (blue). A transfected cell with Gpr161- and Arl13b-positive cilium is marked by a white arrow, while a transfected cell without cilia is shown with a white arrowhead. Yellow arrows and arrowheads point to untransfected cells with or without Gpr161/Arl13b, respectively. Scale bar = 10 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55315/55315fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Using Primary Neurosphere Cultures to Study Primary Cilia at JoVE.com

Using Primary Neurosphere Cultures to Study Primary Cilia

The primary cilium is fundamentally important in neural progenitor cell proliferation, neuronal differentiation, and adult neuronal function. Here, we describe a method to study ciliogenesis and the trafficking of signaling proteins to cilia in neural stem/progenitor cells and differentiated neurons using primary neurosphere cultures.

The primary cilium is fundamentally important for the proliferation of neural stem/progenitor cells and for neuronal differentiation during embryonic, ...

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Nanyang Technological University

Research

JoVE Journal

Developmental Biology

9.3K Views.  University of Texas Southwestern Medical Center. The primary cilium is fundamentally important in neural progenitor cell proliferation, neuronal differentiation, and adult neuronal function. Here, we describe a method to study ciliogenesis and the trafficking of signaling proteins to cilia in neural stem/progenitor cells and differentiated neurons using primary neurosphere cultures.

Video: Using Primary Neurosphere Cultures to Study Primary Cilia

Production and Use of Lentivirus to Selectively Transduce Primary Oligodendrocyte Precursor Cells for In Vitro Myelination Assays

Myelination is a complex process that involves both neurons and the myelin forming glial cells, oligodendrocytes in the central nervous system (CNS) and Schwann cells in the peripheral nervous system (PNS). We use an in vitro myelination assay, an established model for studying CNS myelination in vitro. To do this, oligodendrocyte precursor cells (OPCs) are added to the purified primary rodent dorsal root ganglion (DRG) neurons to form myelinating co-cultures. In order to specifically interrogate the roles that particular proteins expressed by oligodendrocytes exert upon myelination we have developed protocols that selectively transduce OPCs using the lentivirus overexpressing wild type, constitutively active or dominant negative proteins before being seeded onto the DRG neurons. This allows us to specifically interrogate the roles of these oligodendroglial proteins in regulating myelination. The protocols can also be applied in the study of other cell types, thus providing an approach that allows selective manipulation of proteins expressed by a desired cell type, such as oligodendrocytes for the targeted study of signaling and compensation mechanisms. In conclusion, combining the in vitro myelination assay with lentiviral infected OPCs provides a strategic tool for the analysis of molecular mechanisms involved in myelination.

Production and Use of Lentivirus to Selectively Transduce Primary Oligodendrocyte Precursor Cells for In Vitro Myelination Assays

Here we present protocols that offer a flexible and strategic foundation for virally manipulating oligodendrocyte precursor cells to overexpress proteins of interest in order to specifically interrogate their role in oligodendrocytes via the in vitro model of central nervous system myelination.

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Using Ex Vivo Upright Droplet Cultures of Whole Fetal Organs to Study Developmental Processes during Mouse Organogenesis

Investigating organogenesis in utero is a technically challenging process in placental mammals due to inaccessibility of reagents to embryos that develop within the uterus. A newly developed ex vivo upright droplet culture method provides an attractive alternative to studies performed in utero. The ex vivo droplet culture provides the ability to examine and manipulate cellular interactions and diverse signaling pathways through use of various blocking and activating compounds; additionally, the effects of various pharmacological reagents on the development of specific organs can be studied without unwanted side effects of systemic drug delivery in utero. As compared to other in vitro systems, the droplet culture not only allows for the ability to study three-dimensional morphogenesis and cell-cell interactions, which cannot be reproduced in mammalian cell lines, but also requires significantly less reagents than other ex vivo and in vitro protocols. This paper demonstrates proper mouse fetal organ dissection and upright droplet culture techniques, followed by whole organ immunofluorescence to demonstrate the effectiveness of the method. The ex vivo droplet culture method allows the formation of organ architecture comparable to what is observed in vivo and can be utilized to study otherwise difficult-to-study processes due to embryonic lethality in in vivo models. As a model application system, a small-molecule inhibitor will be utilized to probe the role of vascularization in testicular morphogenesis. This ex vivo droplet culture method is expandable to other fetal organ systems, such as lung and potentially others, although each organ must be extensively studied to determine any organ-specific modifications to the protocol. This organ culture system provides flexibility in experimentation with fetal organs, and results obtained using this technique will help researchers gain insights into fetal development.

Using Ex Vivo Upright Droplet Cultures of Whole Fetal Organs to Study Developmental Processes during Mouse Organogenesis

The&#160;ex vivo&#160;upright droplet culture is an alternative to current&#160;in vitro&#160;and&#160;in vivo&#160;experimental techniques. This protocol is easy to perform and requires smaller amounts of reagent, while permitting the ability to manipulate and study fetal vascularization, morphogenesis, and organogenesis.

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Generating Primary Fibroblast Cultures from Mouse Ear and Tail Tissues

Primary cells are derived directly from tissue and are thought to be more representative of the physiological state of cells in vivo than established cell lines. However, primary cell cultures usually have a finite life span and need to be frequently re-established. Fibroblasts are an easily accessible source of primary cells. Here, we discuss a simple and quick experimental procedure to establish primary fibroblast cultures from ears and tails of mice. The protocol can be used to establish primary fibroblast cultures from ears stored at RT for up to 10 days. When the protocol is carefully followed, contaminations are unlikely to occur despite the use of non-sterile tissue stored for extended time in some cases. Fibroblasts proliferate rapidly in culture and can be expanded to substantial numbers before undergoing replicative senescence.

We describe a simple and quick experimental procedure for generating primary fibroblasts from the ears and tails of mice. The procedure does not require special animal training and can be used for the generation of fibroblast cultures from ears stored at RT for up to 10 days.

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Human Primary Trophoblast Cell Culture Model to Study the Protective Effects of Melatonin Against Hypoxia/reoxygenation-induced Disruption

This protocol describes how villous cytotrophoblast cells are isolated from placentas at term by successive enzymatic digestions, followed by density centrifugation, media gradient isolation and immunomagnetic purification. As observed in vivo, mononucleated villous cytotrophoblast cells in primary culture differentiate into multinucleated syncytiotrophoblast cells after 72 hr. Compared to normoxia (8% O2), villous cytotrophoblast cells that undergo hypoxia/reoxygenation (0.5% / 8% O2) undergo increased oxidative stress and intrinsic apoptosis, similar to that observed in vivo in pregnancy complications such as preeclampsia, preterm birth, and intrauterine growth restriction. In this context, primary villous trophoblasts cultured under hypoxia/reoxygenation conditions represent a unique experimental system to better understand the mechanisms and signalling pathways that are altered in human placenta and facilitate the search for effective drugs that protect against certain pregnancy disorders. Human villous trophoblasts produce melatonin and express its synthesizing enzymes and receptors. Melatonin has been suggested as a treatment for preeclampsia and intrauterine growth restriction because of its protective antioxidant effects. In the primary villous cytotrophoblast cell model described in this paper, melatonin has no effect on trophoblast cells in normoxic state but restores the redox balance of syncytiotrophoblast cells disrupted by hypoxia/reoxygenation. Thus, human villous trophoblast cells in primary culture are an excellent approach to study the mechanisms behind the protective effects of melatonin on placental function during hypoxia/reoxygenation.

This manuscript presents a unique in vitro model of immunopurified human villous cytotrophoblast cells cultured under hypoxia/reoxygenation. This model is suitable to study the protective effects of promising treatments, such as melatonin, on pregnancy complications associated with increased oxidative stress and altered placental function.

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Reprogramming Primary Amniotic Fluid and Membrane Cells to Pluripotency in Xeno-free Conditions

Autologous cell-based therapies got a step closer to reality with the introduction of induced pluripotent stem cells. Fetal stem cells, such as amniotic fluid and membrane mesenchymal stem cells, represent a unique type of undifferentiated cells with promise in tissue engineering and for reprogramming into iPSC for future pediatric interventions and stem cell banking. The protocol presented here describes an optimized procedure for extracting and culturing primary amniotic fluid and membrane mesenchymal stem cells and generating episomal induced pluripotent stem cells from these cells in fully chemically defined culture conditions utilizing human recombinant vitronectin and the E8 medium. Characterization of the new lines by applying stringent methods &#8211; flow cytometry, confocal imaging, teratoma formation and transcriptional profiling &#8211; is also described. The newly generated lines express markers of embryonic stem cells &#8211; Oct3/4A, Nanog, Sox2, TRA-1-60, TRA-1-81, SSEA-4 &#8211; while being negative for the SSEA-1 marker. The stem cell lines form teratomas in scid-beige mice in 6-8 weeks and the teratomas contain tissues representative of all three germ layers. Transcriptional profiling of the lines by submitting global expression microarray data to a bioinformatic pluripotency assessment algorithm deemed all lines pluripotent and therefore, this approach is an attractive alternative to animal testing. The new iPSC lines can readily be used in downstream experiments involving the optimization of differentiation and tissue engineering.

This protocol describes the reprogramming of primary amniotic fluid and membrane mesenchymal stem cells into induced pluripotent stem cells using a non-integrating episomal approach in fully chemically defined conditions. Procedures of extraction, culture, reprogramming, and characterization of the resulting induced pluripotent stem cells by stringent methods are detailed.

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Calcific aortic valve disease (CAVD) is characterized by the progressive thickening of the aortic valve leaflets. It is a condition frequently found in the elderly and end-stage renal disease (ESRD) patients, who commonly suffer from hyperphosphatemia and hypercalcemia. At present, there are no medication therapies that can stop its progression. The mechanisms that underlie this pathological process remain unclear. The aortic valve leaflet is composed of a thin layer of valve endothelial cells (VECs) on the outer surfaces of the aortic cusps, with valve interstitial cells (VICs) sandwiched between the VECs. The use of a rat model enables the in vitro study of ectopic calcification based on the in vivo physiopathological serum phosphate (Pi) and calcium (Ca) levels of patients who suffer from hyperphosphatemia and hypercalcemia. The described protocol details the isolation of a pure rat VIC population as shown by the expression of VIC markers: alpha-smooth muscle actin (&#945;-SMA) vimentin and tissue growth factor beta (TGF&#946;) 1 and 2, and the absence of cluster of differentiation (CD) 31, a VEC marker. By expanding these VICs, biochemical, genetic, and imaging studies can be performed to study and unravel the key mediators underpinning CAVD.

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The main function of keratinocytes is to provide the structural integrity of the epidermis, thereby maintaining a mechanical barrier to the outside world. In addition, keratinocytes play an essential role in the initiation, maintenance, and regulation of epidermal immune responses by being part of the innate immune system responding to antigenic stimuli in a fast, nonspecific manner. Here, we describe a protocol for isolation of primary human keratinocytes from adult skin, and demonstrate that these cells respond to calcium-induced terminal differentiation, as measured by an increased expression of the differentiation marker involucrin. In addition, we show that the isolated keratinocytes are responsive to IL-1&#946;-induced activation of intracellular signaling pathways as measured by the activation of the p38 MAPK pathway. Taken together, we describe a method for isolation and culturing of primary human keratinocytes from adult skin. Because the keratinocytes are the predominant cell type in the epidermis, this method is useful to study molecular mechanisms in cutaneous biology in vitro.

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Intestinal organoid cultures provide a unique opportunity to investigate intestinal stem cell and crypt biology in vitro, although efficient approaches to manipulate gene expression in organoids have made limited progress in this arena. While CRISPR/Cas9 technology allows for precise genome editing of cells for organoid generation, this strategy requires extensive selection and screening by sequence analysis, which is both time-consuming and costly. Here, we provide a detailed protocol for efficient viral transduction of intestinal organoids. This approach is rapid and highly efficient, thus decreasing the time and expense inherent in CRISPR/Cas9 technology. We also present a protocol to generate frozen sections from intact organoid cultures for further analysis with immunohistochemical or immunofluorescent staining, which can be used to confirm gene expression or silencing. After successful transduction of viral vectors for gene expression or silencing is achieved, intestinal stem cell and crypt function can be rapidly assessed. Although most organoid studies employ in vitro assays, organoids can also be delivered to mice for in vivo functional analyses. Moreover, our approaches are advantageous for predicting therapeutic responses to drugs because currently available therapies generally function by modulating gene expression or protein function rather than altering the genome.

We describe step-by-step instructions to: 1) efficiently engineer intestinal organoids using magnetic nanoparticles for lenti- or retroviral transduction, and, 2) generate frozen sections from engineered organoids. This approach provides a powerful tool to efficiently alter gene expression in organoids for investigation of downstream effects.

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Induction and Validation of Cellular Senescence in Primary Human Cells

Cellular senescence is a state of permanent cell cycle arrest activated in response to different damaging stimuli. Activation of cellular senescence is a hallmark of various pathophysiological conditions including tumor suppression, tissue remodeling and aging. The inducers of cellular senescence in vivo are still poorly characterized. However, a number of stimuli can be used to promote cellular senescence ex vivo. Among them, most common senescence-inducers are replicative exhaustion, ionizing and non-ionizing radiation, genotoxic drugs, oxidative stress, and demethylating and acetylating agents. Here, we will provide detailed instructions on how to use these stimuli to induce fibroblasts into senescence. This protocol can easily be adapted for different types of primary cells and cell lines, including cancer cells. We also describe different methods for the validation of senescence induction. In particular, we focus on measuring the activity of the lysosomal enzyme Senescence-Associated &#946;-galactosidase (SA-&#946;-gal), the rate of DNA synthesis using 5-ethynyl-2&#39;-deoxyuridine (EdU) incorporation assay, the levels of expression of the cell cycle inhibitors p16 and p21, and the expression and secretion of members of the Senescence-Associated Secretory Phenotype (SASP). Finally, we provide example results and discuss further applications of these protocols.

Here, we discuss a series of protocols for induction and validation of cellular senescence in cultured cells. We focus on different senescence-inducing stimuli and describe the quantification of common senescence-associated markers. We provide technical details using fibroblasts as a model, but the protocols can be adapted to various cellular models.

Cellular senescence is a state of permanent cell cycle arrest activated in response to different damaging stimuli. Activation of cellular senescence is a ...

A Simplified and Efficient Method to Isolate Primary Human Keratinocytes from Adult Skin Tissue

Primary human keratinocytes isolated from fresh skin tissues and their expansion in vitro have been widely used for laboratory research and for clinical applications. The conventional isolation method of human keratinocytes involves a two-step sequential enzymatic digestion procedure, which has been proven to be inefficient in generating primary cells from adult tissues due to the low cell recovery rate and reduced cell viability. We recently reported an advanced method to isolate human primary epidermal progenitor cells from skin tissues that utilizes the Rho kinase inhibitor Y-27632 in the medium. Compared with the traditional protocol, this new method is simpler, easier, and less time-consuming, and increases epithelial stem cell yield and enhances their stem cell characteristics. Moreover, the new methodology does not require the separation of the epidermis from the dermis, and, therefore, is suitable for isolating cells from different types of adult tissues. This new isolation method overcomes the major shortcomings of conventional methods and is more suitable for producing large numbers of epidermal cells with high potency both for laboratory and for clinical applications. Here, we describe the new method in detail.

Here we present a protocol to efficiently isolate primary human keratinocytes from adult skin tissues. This method simplifies the conventional procedure by using the ROCK Inhibitor Y-27632 in the inoculation medium to spontaneously separate epidermal cells from dermal cells.

Primary human keratinocytes isolated from fresh skin tissues and their expansion in vitro have been widely used for laboratory research and for clinical ...