This publication was supported by Award Number K22 CA218462-03 from the NIH National Cancer Institute. The HEK293-T cells expressing R-Spondin1 was a generous gift from Dr. Michael P. Verzi.

All the mouse experiments conducted, including the use of Tamoxifen and euthanasia by cervical dislocation, had the approval of the Institutional Animal Care and Use Committee at Stevens Institute of echnology.
1. Mice
NOTE: The generation of Smad4f/f; Catnblox(ex3)/+; Villin-CreERT2 mice have been previously described3. Adult female mice between eight to twelve weeks of agewere used.
<ol>
	<li>Induce the Smad4KO:&#946;-cateninGOF mutation in the Smad4f/f; Catnblox(ex3)/+; Villin-CreERT2 with intraperitoneal injection of Tamoxifen in corn oil for four consecutive days.</li>
	<li>Sacrifice the miceby cervical dislocation ten days after the first tamoxifen injection.</li>
	<li>Spray the abdomen with 70% ethanol to prevent mouse fur getting into the peritoneal cavity.</li>
	<li>Open the abdominal cavity with dissection scissors to expose the intestine. Isolate the intestine with the aid of the scissors and forceps. 
	&#8203;NOTE: Concomitant loss of Smad4 along with gain of function mutation of &#946;-catenin (Smad4KO;&#946;-cateninGOF) in intestinal epithelium was attained by injectingSmad4f/f; Catnblox(ex3)/+; Villin-CreERT2 mice every day for four consecutive days with 0.05 g Tamoxifen/kg body weight in corn oil. These mice are harvested ten days after the first tamoxifen injection to ensure the presence of cells expressing stem cell-associated markers in the dedifferentiating villi.</li>
</ol>
2. Duodenum isolation and preparation
<ol>
	<li>Dissect out the proximal half of the duodenum.</li>
	<li>Flush the duodenum with 5 mL of ice-cold PBS in a 10 mL syringe to clear the luminal content.</li>
	<li>Open the duodenum longitudinally with an angled scissor and lay the duodenum flat on a 15 cm Petri plate on ice with the lumen of the duodenum facing the operator.</li>
</ol>
3. Villi isolation by scraping
<ol>
	<li>Prior to beginning the scraping, place a 70 &#181;m mesh strainer in one of the wells of a 6-well tissue culture plate. Fill all the wells with 4 mL of 1x PBS and place the 6-well tissue culture plate on ice (Figure 1).</li>
	<li>Scrape the villi as follows using two microscopic glass slides: one to hold the duodenum down and the other to scrape (Figure 1B1).
	<ol>
		<li>Scrape the luminal side of the duodenum superficially to-and-fro twice to remove the mucus. Apply pressure such that this step removes the mucus layer, but not the villi.</li>
		<li>Scrape the duodenum again, to-and-fro twice applying the same pressure as in 3..1, during which villi can be seen collecting on the slides (Figure 1B2). This is the optimal pressure (which must be determined empirically by the operator) to yield the villi without the crypts tethered.</li>
	</ol>
	</li>
	<li>Use a 1 mL transfer pipet containing PBS to transfer the villi (that are collected on the slide from step 3.2.2.) to a 70 &#181;m mesh strainer placed in a 6-well dish. The villi are collected thus, after every scrape (Figure 1B2).</li>
	<li>Wash the villi collected in the 70 &#181;m strainer (from step 3.3) by transferring the strainer (with the villi) through a series of wells in a 6-well dish containing cold PBS (~ 4 mL/well). This is to remove loose crypts, if any.</li>
	<li>Using a p1000 pipet, transfer the villi suspension in PBS (~ 3 mL) from the 70 &#181;m strainer to a new 15 mL tube on ice.</li>
	<li>Use a 0.1% BSA coated blunt-ended p200 pipet tip to aspirate 50 &#181;L volume of the villi suspension onto a glass slide. Count the number of villi in the 50 &#181;L droplet under 4x magnification to determine the concentration of villi in the PBS suspension. For example, if there are 10 villi in the 50 &#181;L suspension examined, then the villi concentration is 0.2 villi/&#181;L. This is also the time to confirm the absence of tethered crypts.</li>
	<li>Calculate the volume of villi suspension required to plate at a concentration of 0.5 villi/&#181;L of BME-R1 matrix. Add an extra 100 &#181;L to account for pipetting error and the volume required for microscopic examination required to ensure the purity of the villi. 
	&#8203;CAUTION: The pressure used in the first two scrapes, that removes the mucus but not the villi, should be used in the subsequent scrapes during which villi will be released. Limit the number of to-and-fro scrapings to two after the villi release is first observed. This measure avoids release of villi with the crypts tethered. It is essential to microscopically evaluate the villi to ensure the absence of the tethered crypts.</li>
</ol>
4. Plating of villi on BME-R1 matrix
<ol>
	<li>Using a 0.1% BSA coated p200 blunt-ended pipet tip, transfer to a microcentrifuge tube the volume of villi (from step 3.6) required to plate at a density of ~ 6 villi per well in 12.5 &#181;L of BME-R1 matrix. The use of BSA-coated blunt-ended tip at this point ensures that the villi, which are too large for a pointed tip are aspirated without being blocked or lost from being stuck to the sides of the tip.</li>
	<li>Spin down the villi for 2 min at 200 x g in a refrigerated centrifuge (4 &#176;C) and remove the supernatant.</li>
	<li>Repeat step 4.2 to remove any residual PBS and proceed to the next step under a laminar flow-hood.</li>
	<li>Resuspend the villi pellet gently in the required amount of cold BME-R1 thawed on ice.</li>
	<li>Using a p20 pipet, plate 12.5 &#181;L/well of the villi in BME-R1 matrix in 3D in a prewarmed 96-well U-bottom plate.</li>
	<li>Incubate the plate in a tissue culture incubator at 37 &#176;C for 15 minutes to allow solidification of the BME-R1 matrix.</li>
	<li>Add 125 &#181;L/well of pre-warmed ENR (Epidermal growth factor/Noggin/R-spondin1) media: advanced DMEM F-12 media, supplemented with 1x Penicillin: Streptomycin, 10 mM HEPES, Glutamax, 50 ng/mL EGF, 100 ng/mL Noggin, 5% conditioned media from HEK293-T cells expressing R-Spondin1, 1x N2, 1x B27, 1 mM N-acetyl cysteine, and 0.05 mg/mL Primocin.</li>
	<li>Incubate the plated villi in a tissue culture incubator maintained at 37 &#176;C with 5% CO2, and change the media every other day.</li>
	<li>Discard any well in which organoids appear before two days of plating, since the earliest time point at which villi-derived organoids are expected is after two days. 
	NOTE: Any villi that has half or more than half the length of villi are counted as one villus. Unlike crypt-derived organoids, villi-derived organoids are not expected within 24 hours after plating. The organoid-initiating villi appear to be darkening and shrinking prior to formation of organoids. Hence, any wells with organoids developing within a day of plating should be discarded to avoid the plausibility of having derived from plausible crypt contamination. The methodology used to produce the conditioned media is available upon request.</li>
</ol>

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O., 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=SMAD4+Suppresses+WNT-Driven+Dedifferentiation+and+Oncogenesis+in+the+Differentiated+Gut+Epithelium.">SMAD4 Suppresses WNT-Driven Dedifferentiation and Oncogenesis in the Differentiated Gut Epithelium.</a> Cancer Research. 78 (17), 4878-4890 (2018).</li><li>Roche, J. 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H., 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=Dll1++secretory+progenitor+cells+revert+to+stem+cells+upon+crypt+damage.">Dll1+ secretory progenitor cells revert to stem cells upon crypt damage.</a> Nature Cell Biology. 14 (10), 1099-1104 (2012).</li><li>Tetteh, P. 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The authors declare no conflicts of interest.

This method can confirm the self-renewal capacity of dedifferentiating villi epithelium that acquire stem cell markers in vivo. The normal intestinal epithelium can give rise to organoids from the crypt but not the villi compartment, when cultured in 3D because of the presence of stem cells in the crypts1. Thus, organoid formation from the dedifferentiated villi epithelium&#160;cultured in 3D cultures confirms stem cell formation from cell fate reversal. Reports on cell fate reversal in t...

The intestinal crypts but not villi, form organoids when cultured in Matrigel or BME-R1 matrix. These organoids are self-organizing structures, often referred to as the &#34;mini-gut&#34; owing to the presence of the various differentiated lineages, progenitor, and stem cells present in the intestinal epithelium in vivo. The potential to form organoids from crypts is attributed to the presence of stem cells1. The intestinal villi on the other hand consist only of differentiated cells, and hence cannot form organoids. However, mutations2 or conditions that permit dedifferentiation of the villus epithelium may lead to stem cells in the villi2,3. This fate change resulting in stemness in the villi epithelium can be confirmed by plating the dedifferentiating villus epithelium in 3D matrix to determine their organoid-forming potential as an indicator of de-novo stemness in the villus epithelium. Hence, the critical aspect of this procedure is to ensure absence of crypt contamination.
The Smad4KO:&#946;-cateninGOF conditional mutation causes dedifferentiation in the intestinal epithelium marked by the expression of proliferation and stem cell markers in the villi, and eventually the formation of crypt-like structures in the villi referred to as ectopic crypts The presence of stem cells these dedifferentiated villi was determined by the expression of stem cell markers in the ectopic crypts (in vivo) and the ability of the mutant villi to form organoids when plated Matrigel3. The below mentioned procedure elaborates the methodology used to confirm the stemness of the dedifferentiating intestinal epithelium in the Smad4KO:&#946;-cateninGOF mutant mice. A key feature of this methodology for isolating villi was the use of scraping of the intestinal lumen, as opposed to the EDTA chelation method4. Unlike in the EDTA chelation method, villi isolation by scraping retains majority of the underlying mesenchyme and allows adjust the pressure of scraping to yield villi without tethered crypts. The pressure of scraping is subjective to the operator, hence, the optimal pressure to yield villi without crypts tethered must be determined empirically by the operator. The critical aspect of this procedure is to ensure the absence of crypt contamination by microscopic examination of the villi both before and after plating in the BME-R1 matrix.
Intestinal villi are scraped off the intestinal lumen with glass slides and placed in a 70 &#181;m filter and washed with PBS to get rid of loose cells or crypts, if any, prior to plating in BME-R1 matrix. The method stresses on the following criteria to avoid crypt contamination: a) confining the villi harvest the proximal half of the duodenum where the villi are the longest, b) minimizing the number of villi-yielding scrapes, c) washing the filter containing the villi through a series of PBS in a six-well dish, and d) confirming the absence of crypt contamination by microscopic examination prior to and after plating in BME-R1 matrix. Villi isolation by scraping, rather than by EDTA chelation, prevents the complete loss of the underlying mesenchyme that may provide the niche signals5, 6, 7, 8, if required, for organoid initiation from the villus epithelium.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Advanced DMEM F-12 media</td>
			<td>Gibco</td>
			<td>12634010</td>
			<td></td>
		</tr>
		<tr>
			<td>3,3-diaminobenzidine</td>
			<td>Vector Labs</td>
			<td>SK-4105</td>
			<td></td>
		</tr>
		<tr>
			<td>96 well U-bottom plate</td>
			<td>Fisher Scientific</td>
			<td>FB012932</td>
			<td></td>
		</tr>
		<tr>
			<td>ABC kit</td>
			<td>Vector Labs</td>
			<td>&#160;PK4001</td>
			<td></td>
		</tr>
		<tr>
			<td>Angled scissor</td>
			<td>Fisher Scientific</td>
			<td>11-999</td>
			<td></td>
		</tr>
		<tr>
			<td>Animal-Free Recombinant Human EGF</td>
			<td>Peprotech</td>
			<td>AF-100-15</td>
			<td></td>
		</tr>
		<tr>
			<td>B-27 Supplement (50X), minus vitamin A</td>
			<td>Gibco</td>
			<td>12587010</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA) Protease-free Powder</td>
			<td>Fisher Scientific</td>
			<td>BP9703100</td>
			<td></td>
		</tr>
		<tr>
			<td>CD44 antibody</td>
			<td>BioLegend</td>
			<td>1030001</td>
			<td></td>
		</tr>
		<tr>
			<td>Cdx2 antibody</td>
			<td>Cell Signaling</td>
			<td>12306</td>
			<td></td>
		</tr>
		<tr>
			<td>Corn oil</td>
			<td>Sigma-Aldrich</td>
			<td>C8267-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning 70-micron cell strainer</td>
			<td>Life Sciences</td>
			<td>431751</td>
			<td></td>
		</tr>
		<tr>
			<td>Cultrex Reduced Growth Factor Basement Membrane Extract, Type R1</td>
			<td>R&#38;D</td>
			<td>3433-005-R1</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection scissors</td>
			<td>Fisher Scientific</td>
			<td>22-079-747</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Fisher Scientific</td>
			<td>17-456-209</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutamax (100X)</td>
			<td>Gibco</td>
			<td>35050-061</td>
			<td></td>
		</tr>
		<tr>
			<td>HEK 293-T cells expressing RSPO-1</td>
			<td>Gift from Dr. Michael Verzi</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES (1M)</td>
			<td>Gibco</td>
			<td>15630-080</td>
			<td></td>
		</tr>
		<tr>
			<td>Histogel</td>
			<td>Thermoscientific</td>
			<td>HG-4000-012</td>
			<td></td>
		</tr>
		<tr>
			<td>Mesh filter</td>
			<td>Fisher Scientific</td>
			<td>07-201-431</td>
			<td></td>
		</tr>
		<tr>
			<td>Micrscope glass slide</td>
			<td>VWR</td>
			<td>89218-844</td>
			<td></td>
		</tr>
		<tr>
			<td>N-2 Supplement (100X)</td>
			<td>Gibco</td>
			<td>17502048</td>
			<td></td>
		</tr>
		<tr>
			<td>N-acetyl cysteine</td>
			<td>Sigma-Aldrich</td>
			<td>A9165</td>
			<td></td>
		</tr>
		<tr>
			<td>p200 Blunt tips</td>
			<td>VWR</td>
			<td>46620-642</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin (10,000 U/mL)</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>Primocin (50mg/mL)</td>
			<td>Invivogen</td>
			<td>ant-pm-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Quality Biological Inc PBS (10X)</td>
			<td>Fisher Scientific</td>
			<td>50-146-770</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant Murine Noggin</td>
			<td>Peprotech</td>
			<td>250-38</td>
			<td></td>
		</tr>
		<tr>
			<td>Signal diluent</td>
			<td>Cell Signaling</td>
			<td>8112L</td>
			<td></td>
		</tr>
		<tr>
			<td>Tamoxifen</td>
			<td>Sigma-Aldrich</td>
			<td>T5648-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>6-well tissue culture plate</td>
			<td>Fisher Scientific</td>
			<td>50-146-770</td>
			<td></td>
		</tr>
	</tbody>
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3d culturing of organoids from the intestinal villi epithelium undergoing dedifferentiation

Clonogenicity of organoids from the intestinal epithelium is attributed to the presence of stem cells therein. The mouse small intestinal epithelium is compartmentalized into crypts and villi: the stem and proliferating cells are confined to the crypts, whereas the villi epithelium contains only differentiated cells. Hence, the normal intestinal crypts, but not the villi, can give rise to organoids in 3D cultures. The procedure described here is applicable only to villus epithelium undergoing dedifferentiation leading to stemness. The method described uses the Smad4-loss-of-function:&#946;-catenin gain-of-function (Smad4KO:&#946;-cateninGOF) conditional mutant mouse. The mutation causes the intestinal villi to dedifferentiate and generate stem cells in the villi. Intestinal villi undergoing dedifferentiation are scraped off the intestine using glass slides, placed in a 70 &#181;m strainer and washed several times to filter out any loose cells or crypts prior to plating in BME-R1 matrix to determine their organoid-forming potential. Two main criteria were used to ensure that the resulting organoids were developed from the dedifferentiating villus compartment and not from the crypts: 1) microscopically evaluating&#160;the isolated villi to ensure absence of any tethered crypts, both before and after plating in the 3D matrix, and 2) monitoring the time course of organoid development from the villi. Organoid initiation from the villi occurs only two to five days after plating and appears irregularly shaped, whereas the crypt-derived organoids from the same intestinal epithelium are apparent within sixteen hours of plating and appear spherical. The limitation of the method, however, is that the number of organoids formed, and the time required for organoid initiation from the villi vary depending on the degree of dedifferentiation. Hence, depending upon the specificity of the mutation or the insult causing the dedifferentiation, the optimal stage at which villi can be harvested to assay their organoid forming potential, must be determined empirically.

The determining factor for the success of the procedure is preventing crypt contamination. Organoid development from the villi (and not from any contaminating crypts) is ensured by confirming four major criteria: 1) ensuring the purity of the harvested villi by microscopic examination before and after plating the villi in BME-R1, 2) plating limited number of villi per well to allow visualization of all the plated villi individually, 3) onitoring the development of organoid daily; images of the time course show developmen...

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3D Culturing of Organoids from the Intestinal Villi Epithelium Undergoing Dedifferentiation

The procedure describes isolation of the villi from the mouse intestinal epithelium undergoing dedifferentiation to determine their organoid forming potential.

Clonogenicity of organoids from the intestinal epithelium is attributed to the presence of stem cells therein. The mouse small intestinal epithelium is ...

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4.8K Views.  Stevens Institute of Technology. The procedure describes isolation of the villi from the mouse intestinal epithelium undergoing dedifferentiation to determine their organoid forming potential.

Video: 3D Culturing of Organoids from the Intestinal Villi Epithelium Undergoing Dedifferentiation

A 3D System for Culturing Human Articular Chondrocytes in Synovial Fluid

Cartilage destruction is a central pathological feature of osteoarthritis, a leading cause of disability in the US. Cartilage in the adult does not regenerate very efficiently in vivo; and as a result, osteoarthritis leads to irreversible cartilage loss and is accompanied by chronic pain and immobility 1,2. Cartilage tissue engineering offers promising potential to regenerate and restore tissue function. This technology typically involves seeding chondrocytes into natural or synthetic scaffolds and culturing the resulting 3D construct in a balanced medium over a period of time with a goal of engineering a biochemically and biomechanically mature tissue that can be transplanted into a defect site in vivo 3-6. Achieving an optimal condition for chondrocyte growth and matrix deposition is essential for the success of cartilage tissue engineering. 
 In the native joint cavity, cartilage at the articular surface of the bone is bathed in synovial fluid. This clear and viscous fluid provides nutrients to the avascular articular cartilage and contains growth factors, cytokines and enzymes that are important for chondrocyte metabolism 7,8. Furthermore, synovial fluid facilitates low-friction movement between cartilaginous surfaces mainly through secreting two key components, hyaluronan and lubricin 9 10. In contrast, tissue engineered cartilage is most often cultured in artificial media. While these media are likely able to provide more defined conditions for studying chondrocyte metabolism, synovial fluid most accurately reflects the natural environment of which articular chondrocytes reside in.
 Indeed, synovial fluid has the advantage of being easy to obtain and store, and can often be regularly replenished by the body. Several groups have supplemented the culture medium with synovial fluid in growing human, bovine, rabbit and dog chondrocytes, but mostly used only low levels of synovial fluid (below 20&#37;) 11-25. While chicken, horse and human chondrocytes have been cultured in the medium with higher percentage of synovial fluid, these culture systems were two-dimensional 26-28. Here we present our method of culturing human articular chondrocytes in a 3D system with a high percentage of synovial fluid (up to 100&#37;) over a period of 21 days. In doing so, we overcame a major hurdle presented by the high viscosity of the synovial fluid. This system provides the possibility of studying human chondrocytes in synovial fluid in a 3D setting, which can be further combined with two other important factors (oxygen tension and mechanical loading) 29,30 that constitute the natural environment for cartilage to mimic the natural milieu for cartilage growth. Furthermore, This system may also be used for assaying synovial fluid activity on chondrocytes and provide a platform for developing cartilage regeneration technologies and therapeutic options for arthritis.

A 3D system of culturing human articular chondrocytes in high levels of synovial fluid is described. Synovial fluid reflects the most natural microenvironment for articular cartilage, and can be easily obtained and stored. This system thus can be used for studying cartilage regeneration and for screening therapeutics for treating arthritis.

Cartilage destruction is a central pathological feature of osteoarthritis, a leading cause of disability in the US. Cartilage in the adult does not ...

MicroRNA Expression Profiles of Human iPS Cells, Retinal Pigment Epithelium Derived From iPS, and Fetal Retinal Pigment Epithelium

The objective of this report is to describe the protocols for comparing the microRNA (miRNA) profiles of human induced-pluripotent stem (iPS) cells, retinal pigment epithelium (RPE) derived from human iPS cells (iPS-RPE), and fetal RPE. The protocols include collection of RNA for analysis by microarray, and the analysis of microarray data to identify miRNAs that are differentially expressed among three cell types. The methods for culture of iPS cells and fetal RPE are explained. The protocol used for differentiation of RPE from human iPS is also described. The RNA extraction technique we describe was selected to allow maximal recovery of very small RNA for use in a miRNA microarray. Finally, cellular pathway and network analysis of microarray data is explained. These techniques will facilitate the comparison of the miRNA profiles of three different cell types.

The microRNA (miRNA) profiles of human induced-pluripotent stem (iPS) cells, retinal pigment epithelium (RPE) derived from human induced-pluripotent stem (iPS) cells (iPS-RPE), and fetal RPE, were compared.

The objective of this report is to describe the protocols for comparing the microRNA (miRNA) profiles of human induced-pluripotent stem (iPS) cells, ...

A Protocol for Lentiviral Transduction and Downstream Analysis of Intestinal Organoids

Intestinal crypt-villus structures termed organoids, can be kept in sustained culture three dimensionally when supplemented with the appropriate growth factors. Since organoids are highly similar to the original tissue in terms of homeostatic stem cell differentiation, cell polarity and presence of all terminally differentiated cell types known to the adult intestinal epithelium, they serve as an essential resource in experimental research on the epithelium. The possibility to express transgenes or interfering RNA using lentiviral or retroviral vectors in organoids has increased opportunities for functional analysis of the intestinal epithelium and intestinal stem cells, surpassing traditional mouse transgenics in speed and cost. In the current video protocol we show how to utilize transduction of small intestinal organoids with lentiviral vectors illustrated by use of doxycylin inducible transgenes, or IPTG inducible short hairpin RNA for overexpression or gene knockdown. Furthermore, considering organoid culture yields minute cell counts that may even be reduced by experimental treatment, we explain how to process organoids for downstream analysis aimed at quantitative RT-PCR, RNA-microarray and immunohistochemistry. Techniques that enable transgene expression and gene knock down in intestinal organoids contribute to the research potential that these intestinal epithelial structures hold, establishing organoid culture as a new standard in cell culture.

In this video protocol we give a step by step explanation of lentiviral transduction in organoids of primary intestinal epithelium and of processing and downstream analysis of these cultures by quantitative RT-PCR, RNA-microarray and immunohistochemistry.

Intestinal crypt-villus structures termed organoids, can be kept in sustained culture three dimensionally when supplemented with the appropriate growth ...

Primary Cell Cultures from the Mouse Retinal Pigment Epithelium

The retinal pigment epithelium (RPE) is a highly polarized multi-functional epithelium that is located between the neural retina and the choroid of the eye. It is a single sheet of pigmented cells that are hexagonally packed and connected by tight junctions. The main functions of the RPE include absorption of light, phagocytosis of the shed photoreceptor outer segments, spatial buffering of ions, transport of nutrients, ions and water as well as active involvement in the visual cycle. With such important and diverse functions, it is critically important to study the biology of RPE cells. A number of RPE cell lines have been established; however, passaged and immortalized cells are known to quickly lose some of the morphological and physiological characteristics of natural RPE cells. Thus, primary cells are more suitable for studying different aspects of RPE cell biology and function. Mouse primary RPE cell culture is very useful to researchers since mouse models are widely used in biological studies, however collecting RPE cells from mouse is also very challenging due to their small size. Here, we present a protocol for establishing primary mouse RPE cell cultures which includes enucleation and dissection of the eyes and isolation of the RPE sheets to yield the cells for culturing. This method enables efficient cell recovery. The RPE cells obtained from two mice&#160;can reach confluency on one 12 mm polyester membrane insert pre-loaded in culture plate after one week of culture and display some of the original properties of bona fide RPE cells such as hexagonal shape and pigmentation after two weeks of culture.

The retinal pigment epithelium (RPE) is a multi-functional epithelium of the eye. Here we present a protocol to establish primary cell cultures derived from the murine RPE.

The retinal pigment epithelium (RPE) is a highly polarized multi-functional epithelium that is located between the neural retina and the choroid of the ...

Analyzing Oxidative Stress in Murine Intestinal Organoids using Reactive Oxygen Species-Sensitive Fluorogenic Probe

Reactive oxygen species (ROS) play essential roles in intestinal homeostasis. ROS are natural by-products of cell metabolism. They are produced in response to infection or injury at the mucosal level as they are involved in antimicrobial responses and wound healing. They are also critical secondary messengers, regulating several pathways, including cell growth and differentiation. On the other hand, excessive ROS levels lead to oxidative stress, which can be deleterious for cells and favor intestinal diseases like chronic inflammation or cancer. This work provides a straightforward method to detect ROS in the intestinal murine organoids by live imaging and flow cytometry, using a commercially available fluorogenic probe. Here the protocol describes assaying the effect of compounds that modulate the redox balance in intestinal organoids and detect ROS levels in specific intestinal cell types, exemplified here by the analysis of the intestinal stem cells genetically labeled with GFP. This protocol may be used with other fluorescent probes.

The present protocol describes a method to detect reactive oxygen species (ROS) in the intestinal murine organoids using qualitative imaging and quantitative cytometry assays. This work can be potentially extended to other fluorescent probes to test the effect of selected compounds on ROS.

Reactive oxygen species (ROS) play essential roles in intestinal homeostasis. ROS are natural by-products of cell metabolism. They are produced in ...

A High-Throughput Platform for Culture and 3D Imaging of Organoids

The characterization of a large number of three-dimensional (3D) organotypic cultures (organoids) at different resolution scales is currently limited by standard imaging approaches. This protocol describes a way to prepare microfabricated organoid culture chips, which enable multiscale, 3D live imaging on a user-friendly instrument requiring minimal manipulations and capable of up to 300 organoids/h imaging throughput. These culture chips are compatible with both air and immersion objectives (air, water, oil, and silicone) and a wide range of common microscopes (e.g., spinning disk, point scanner confocal, wide field, and brightfield). Moreover, they can be used with light-sheet modalities such as the single-objective, single-plane illumination microscopy (SPIM) technology (soSPIM). 
The protocol described here gives detailed steps for the preparation of the microfabricated culture chips and the culture and staining of organoids. Only a short length of time is required to become familiar with, and consumables and equipment can be easily found in normal biolabs. Here, the 3D imaging capabilities will be demonstrated only with commercial standard microscopes (e.g., spinning disk for 3D reconstruction and wide field microscopy for routine monitoring).

This paper presents a fabrication protocol for a new type of culture substrate with hundreds of microcontainers per mm2, in which organoids can be cultured and observed using high-resolution microscopy. The cell seeding and immunostaining protocols are also detailed.

The characterization of a large number of three-dimensional (3D) organotypic cultures (organoids) at different resolution scales is currently limited by ...

Culturing, Freezing, Processing, and Imaging of Entire Organoids and Spheroids While Still in a Hydrogel

Organoids and spheroids, three-dimensional growing structures in cell culture labs, are becoming increasingly recognized as superior models compared to two-dimensional culture models, since they mimic the human body better and have advantages over animal studies. However, these studies commonly face problems with reproducibility and consistency. During the long experimental processes - with transfers of organoids and spheroids between different cell culture vessels, pipetting, and centrifuging - these susceptible and fragile 3D growing structures are often damaged or lost. Ultimately, the results are significantly affected, since the 3D structures cannot maintain the same characteristics and quality. The methods described here minimize these stressful steps and ensure a safe and consistent environment for organoids and spheroids throughout the processing sequence while they are still in a hydrogel in a multipurpose device. The researchers can grow, freeze, thaw, process, stain, label, and then examine the structure of organoids or spheroids under various high-tech instruments, from confocal to electron microscopes, using a single multipurpose device. This technology improves the studies&#39; reproducibility, reliability, and validity, while maintaining a stable and protective environment for the 3D growing structures during processing. In addition, eliminating stressful steps minimizes handling errors, reduces time taken, and decreases the risk of contamination.

The present study describes methodologies for culturing, freezing, thawing, processing, staining, labeling, and examining entire spheroids and organoids under various microscopes, while they remain intact in a hydrogel within a multipurpose device.

Organoids and spheroids, three-dimensional growing structures in cell culture labs, are becoming increasingly recognized as superior models compared to ...

Establishing 3D Endometrial Organoids from the Mouse Uterus

Endometrial tissue lines the inner cavity of the uterus and is under the cyclical control of estrogen and progesterone. It is a tissue that is composed of luminal and glandular epithelium, a stromal compartment, a vascular network, and a complex immune cell population. Mouse models have been a powerful tool to study the endometrium, revealing critical mechanisms that control implantation, placentation, and cancer. The recent development of 3D endometrial organoid cultures presents a state-of-the-art model to dissect the signaling pathways that underlie endometrial biology. Establishing endometrial organoids from genetically engineered mouse models, analyzing their transcriptomes, and visualizing their morphology at a single-cell resolution are crucial tools for the study of endometrial diseases. This paper outlines methods to establish 3D cultures of endometrial epithelium from mice and describes techniques to quantify gene expression and analyze the histology of the organoids. The goal is to provide a resource that can be used to establish, culture, and study the gene expression and morphological characteristics of endometrial epithelial organoids.

This protocol describes methodologies to establish mouse endometrial epithelial organoids for gene expression and histological analyses.

Endometrial tissue lines the inner cavity of the uterus and is under the cyclical control of estrogen and progesterone. It is a tissue that is composed of ...

Culture of Piglet Intestinal 3D Organoids from Cryopreserved Epithelial Crypts and Establishment of Cell Monolayers

Intestinal organoids are increasingly being used to study the gut epithelium for digestive disease modeling, or to investigate interactions with drugs, nutrients, metabolites, pathogens, and the microbiota. Methods to culture intestinal organoids are now available for multiple species, including pigs, which is a species of major interest both as a farm animal and as a translational model for humans, for example, to study zoonotic diseases. Here, we give an in-depth description of a procedure used to culture pig intestinal 3D organoids from frozen epithelial crypts. The protocol describes how to cryopreserve epithelial crypts from the pig intestine and the subsequent procedures to culture 3D intestinal organoids. The main advantages of this method are (i) the temporal dissociation of the isolation of crypts from the culture of 3D organoids, (ii) the preparation of large stocks of cryopreserved crypts derived from multiple intestinal segments and from several animals at once, and thus (iii) the reduction in the need to sample fresh tissues from living animals. We also detail a protocol to establish cell monolayers derived from 3D organoids to allow access to the apical side of epithelial cells, which is the site of interactions with nutrients, microbes, or drugs. Overall, the protocols described here is a useful resource for studying the pig intestinal epithelium in veterinary and biomedical research.

Here, we describe a protocol to culture pig intestinal 3D organoids from cryopreserved epithelial crypts. We also describe a method to establish cell monolayers derived from 3D organoids, allowing access to the apical side of epithelial cells.

Intestinal organoids are increasingly being used to study the gut epithelium for digestive disease modeling, or to investigate interactions with drugs, ...

3D Culture of Organoids from Murine Intestinal Crypts and Single Stem Cell

The 3D Culturing of Organoids from Murine Intestinal Crypts and a Single Stem Cell for Organoid Research

At present, organoid culture represents an important tool for in vitro studies of different biological aspects and diseases in different organs. Murine small intestinal crypts can form organoids that mimic the intestinal epithelium when cultured in a 3D extracellular matrix. The organoids are composed of all cell types that fulfill various intestinal homeostatic functions. These include Paneth cells, enteroendocrine cells, enterocytes, goblet cells, and tuft cells. Well-characterized molecules are added into the culture medium to enrich the intestinal stem cells (ISCs) labeled with leucine-rich repeats containing G protein-coupled receptor 5 and are used to drive differentiation down specific lineages; these molecules include epidermal growth factor, Noggin (a bone morphogenetic protein), and R-spondin 1. Additionally, a protocol to generate organoids from a single erythropoietin-producing hepatocellular receptor B2 (EphB2)-positive ISC is also detailed. In this methods article, techniques to isolate small intestinal crypts and a single ISC from tissues and ensure the efficient establishment of organoids are described.

Author Spotlight: The 3D Culturing of Organoids from Murine Intestinal Crypts and a Single Stem Cell for Organoid Research  

We describe a protocol to isolate murine small intestinal crypts and culture intestinal 3D organoids from the crypts. Additionally, we describe a method to generate organoids from a single intestinal stem cell in the absence of a sub-epithelial cellular niche.

At present, organoid culture represents an important tool for in vitro studies of different biological aspects and diseases in different organs. Murine ...