The research is supported by the CALIPSO project supported by the National Research Foundation, Prime Minister&#39;s Office, Singapore, under its Campus for Research Excellence and Technological Enterprise (CREATE) programme. V.V. acknowledges the support of NRF investigator NRF-NRFI2018-07, MOE Tier 3 MOE2016-T3-1-005, MBI seed funding, and ANR ADGastrulo. A.B. and G.G. acknowledge the support from MBI core funding. A.B. acknowledges Andor Technologies for the loan of the BC43 microscope.

NOTE: The first part of this protocol details the microfabrication of the cell culture device. An original primary mold with pyramidal cavities can be produced in-house-if micro-fabrication facilities are available-or outsourced to external companies. The primary mold used in this work is produced in-house, with fabrication process steps described elsewhere11,13. A basic protocol for the microfabrication of the mold is available in Supplementary File 1. CRITICAL: The operations in steps 1 to 6 need to be conducted in a dust-free environment. A laminar flow hood or a clean room, if available, are preferred. All through these steps, personal protective equipment (PPE) needs to be used, such as gloves, a lab coat, and safety glasses.
1. Dicing of PDMS mold
<ol>
	<li>Cut small portions of the PDMS mold to the dimension required for the final device (e.g., 1 cm x 1 cm [Figure 1B]). Cut them parallel to the XY directions of the array. 
	&#8203;CRITICAL: When cutting the PDMS mold in 1 cm x 1 cm dices, execute the cuts in single steps; using a one-sided razorblade, apply pressure and cut through the PDMS in one single step. This is to prevent the formation of small particles of PDMS, which can deposit on the surface of the mold and affect the quality of the replica steps (step 3).</li>
</ol>
2. Preparation of flat PDMS substrates
<ol>
	<li>Weigh ~15-20 g of PDMS base resin and 1.5-2 g of its reticulation agent (i.e., a ratio of 10:1), mix them carefully, and degas in a vacuum jar for ~20 min.</li>
	<li>Slowly pour the degassed PDMS onto a plastic, 15 cm wide (diameter) Petri dish.</li>
	<li>Cure the PDMS by keeping the Petri dish in an oven set at 65&#176; C for 2 h. After waiting for the curing to complete, a flat PDMS film of ~1.5 mm thickness (contained in the Petri dish) is ready for use (Figure 2A).</li>
	<li>Using a sharp blade, cut 2 cm x 2 cm pieces out of the PDMS (Figure 2B-D). 
	&#8203;NOTE: The exact thickness of this PDMS sheet is not critical; ~1-2 mm is a suitable range. The dimension for the cut should be bigger than the textured mold but not much larger, which would result in wasting of material.</li>
</ol>
3. Production of the textured layer made of UV-curable adhesive
<ol>
	<li>Place one PDMS mold dice (as prepared in step 1) face-down on top of the flat PDMS cut (as prepared in step 2) (Figure 3A,B). 
	NOTE: Make sure the pyramidal protrusions in the mold are oriented toward the flat PDMS substrate and that only the top surface of the truncated pyramids is in contact. Assess the correct placement with an optical microscope (Figure 3C,D).</li>
	<li>To one side of the mold, using a pipette, drop a small amount (two drops, approximately 0.1-0.2 mL) of UV-curable adhesive (Figure 4A). 
	NOTE: As the liquid gets in contact with the edge of the mold, capillarity will drive the liquid to fill the cavity between the mold itself and the flat cut of PDMS used as substrate.
	<ol>
		<li>Follow the progression of the liquid inside the cavity, for example, using an inverted optical microscope with a 10x magnification objective (Figure 4B,C).</li>
	</ol>
	</li>
	<li>Expose the UV-curable adhesive to UV light to cure it. Adjust the time of exposure depending on the power density of the UV source used (e.g., a UV-LED box with a power density of 35 mW/cm2; 2 min at 50% power in this protocol to cure the adhesive completely).</li>
	<li>Using the excess amount of adhesive at one edge, hold the cured adhesive on the flat PDMS substrate by gently pressing with a finger (Figure 5A,B). Meanwhile, use tweezers to pinch one corner of the mold next to the same edge being held down and slowly peel it off while making sure that the textured film is not lifted up as well. 
	NOTE: Figure 5C shows the result of a proper mold removal procedure; Figure 5E-G shows the wrong procedure.</li>
	<li>Trim the excess adhesive and the excess PDMS substrate using a razor blade to leave the cured, textured, adhesive layer flat on the PDMS with excess PDMS on one edge only (Figure 5D), which will be needed in step 5.</li>
</ol>
4. Coverslip substrate preparation
NOTE: As a substrate for the final device, standard rounded 1.5H coverslips with 25 mm diameter are used. Before they can be used, they need to be cleaned to remove dust and/or organic residual from their surface.
<ol>
	<li>Coverslip cleaning
	<ol>
		<li>Immerse the coverslips in soapy water for 5 min while the ultrasound is applied (40 kHz, 110 W sonic power).</li>
		<li>Wash the coverslips in clean deionized (DI) water, first by immersion and finally with running DI water from a tap. Dry the coverslips with an N2 gas blow-gun.</li>
		<li>Immerse the coverslips in an acetone bath for 5 min; immediately move to a 2-propanol (IPA) bath for an additional 5 min.</li>
		<li>Rinse the coverslips with clean IPA using a squeeze bottle. Dry the coverslips with the N2 gas blow-gun. 
		NOTE: Coverslips cleaned using this procedure can be stored in a closed container until needed. Keep them in a dry cabinet to avoid humidity depositing on their surface.</li>
	</ol>
	</li>
	<li>When ready to be used, treat a clean coverslip with a short O2 plasma process to improve its hydrophilicity: O2 20 sccm, 3 mbar pressure, 60 W at the RF power generator, and 60 s duration.</li>
	<li>Immediately after the plasma activation, proceed with spin-coating the thin layer of UV-curable adhesive by placing the coverslip on the vacuum chuck of a standard spin-coater and pouring a small drop of adhesive at the center of the coverslip (Figure 6). Run the spin-coating process: spreading for 5 s at 500 rpm, coating at 3,000 rpm for 45 s (with the acceleration and deceleration set at 100 rpm/s).
	<ol>
		<li>If spin-coating is not available, use the following alternative way to produce a thin film of UV adhesive on the coverslips:
		<ol>
			<li>On a clean coverslip, drop ~0.1 mL of UV-curable adhesive using a pipette (Figure 7A).</li>
			<li>Take a second coverslip and place it on top of the first one to make the liquid adhesive spread evenly between the two coverslips (Figure 7B-D).</li>
			<li>Once the spreading adhesive has reached the edges of the coverslips, gently separate them by sliding one over the other. Once separated, both coverslips are fully coated with a thin layer of liquid adhesive (Figure 7E). 
			NOTE: The coating might not be uniform and smooth only if the separation is not done with a smooth and continuous movement.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Precuring of UV-curable adhesive
	<ol>
		<li>After spin-coating, precure the adhesive by exposure to UV. Adjust the time of exposure depending on the power density of the UV source used (here, a UV-LED box with a power density of 35 mW/cm2 was used for 1 min at 50% power). 
		CRITICAL: The adhesive used here is an optical glue. See the discussion for key points related to the energy doses for its curing.</li>
	</ol>
	</li>
</ol>
5. Transfer of the textured film to the final substrate
<ol>
	<li>Take one of the textured films (prepared in step 3) and place it in contact with the adhesive-coated coverslip (prepared in step 4). Make sure the contact between the partially cured adhesive on the coverslip and the textured film is as uniform as possible (Figure 8A-C). 
	CRITICAL: At this stage, the adhesive on the coverslip should be solid enough to avoid reflowing, which would fill the pyramidal cavities of the textured film when placed in contact but also be plastic and adhesive enough that contact can be optimized by gently pressing on the textured film.</li>
	<li>Expose the coverslips to UV light until the adhesive layer coated on the coverslip is fully cured. This will seal the textured film on the coverslip and provide leak-proof isolation between the pyramidal cavities.</li>
	<li>Finally, peel off the PDMS flat substrate (Figure 8D-F). Using tweezers, pinch the PDMS on one corner at the edge where excess material was left after trimming (step 3.5). This way, the textured film layer is left adhesive to the coverslip with open access at the top for cell seeding.CRITICAL: When peeling off the flat PDMS, the textured film should remain well attached to the coverslip. Adhesion failures are easily confirmed if the textured film can be peeled off from the coverslip after final exposure to UV without any effort.</li>
</ol>
6. Long-term passivation of the cell culture coverslip for cell culture
NOTE: Passivation is achieved by generating a conformal and continuous coating of a biomimetic copolymer with a structure similar to the polar group of phospholipids in the cell membrane. This conformal coating prevents cells adhesion to the cell culture device
<ol>
	<li>Prepare a solution with 0.5% (w/v) of the biomimetic copolymer dissolved in pure ethanol. Store the solution at 4 &#176;C for future use.</li>
	<li>Place the cell culture coverslip in a 35 mm Petri dish and fully cover it with the biomimetic copolymer solution.</li>
	<li>After 5 min, remove the cell culture coverslip from the container with the biomimetic copolymer solution and leave it to dry at room temperature inside the final dish in a biosafety hood (&#62;1 h). 
	NOTE: A thicker coating can be produced by increasing the concentration of the biomimetic copolymer in the coating solution; the results of a thicker coating are visible under a brightfield microscope (Figure 9A).</li>
</ol>
7. Cell seeding
<ol>
	<li>Degassing and sterilization
	<ol>
		<li>Just before cell seeding, dispense sterile phosphate-buffered saline (PBS) into the cell culture dishes (typically 1 mL for a 35 mm Petri dish). Degas the dish with the sterile PBS using an ultrasonic device for ~10 min, followed by several rounds of pipetting to remove all the bubbles. 
		CRITICAL: If air is trapped inside the pyramidal cavities, it will prevent the cells from entering them. To make sure there is no air trapped in the pyramid before cell seeding, it is recommended to visually ensure (under a benchtop brightfield microscope at 10x or 20x magnification) the absence of air in these cavities (Figure 10).</li>
		<li>Replace the PBS with sterile culture medium and sterilize the plate with UV light for 30 min under a cell culture hood. 
		NOTE: From this step onward, the dish should be considered as sterile and manipulated using sterile techniques. An alternative way to remove trapped air from the cell culture device is to use a vacuum jar with a vacuum pump.</li>
	</ol>
	</li>
	<li>Cell suspension preparation 
	NOTE: Cells can be seeded as single or small cell aggregates and enter the sample wells through the top aperture. Over time, the inserted cells aggregate and grow inside the sample wells into spheroids of a size greater than the size of the aperture. As validated cell line models, use HCT116 (CCL-247 ATCC) or MCF7 (HTB-22 ATCC) cancer cells maintained in recommended culture medium (ATCC guidelines).
	<ol>
		<li>Prepare a cell suspension (e.g., using a trypsinization process following ATCC guidelines). Follow trypsinization/cell suspension preparation recommendations for the cells of interests.</li>
		<li>Count and adjust the cell concentration to 0.5 &#215; 106 cells/mL in the recommended culture medium.</li>
	</ol>
	</li>
	<li>Cell dispensing
	<ol>
		<li>Remove the PBS buffer from the 35 mm cell culture dish and then dispense 1 mL of the adjusted cell suspension. See Figure 11A for an optical microscope image of a cell seeding procedure with adequate cell density and homogeneity.</li>
		<li>Place the cell culture dish back into the cell incubator (37 &#176;C, 5% CO2, and 100% humidity) for 10 min. Approximately 80-100 cells will enter each pyramidal cavity. 
		NOTE: It is possible to increase the number of cells per cell culture dish by increasing either the cell concentration or the time spent before cell suspension removal. Typically, spheroid formation takes several hours (depending on cell type) after cell seeding and can be followed with a brightfield microscope (4x to 40x objectives; Figure 11). From here, culture medium, extracellular matrices, and differentiation growth factors can be changed or added to the cell culture dish containing the spheroids in accordance with typical differentiation protocols that could last a few days, weeks, or months.</li>
		<li>After the 10 min incubation, recover the cell culture dish from the incubator and gently aspirate the cellular suspension to remove untrapped cells. Add 1 mL of culture medium to a 35 mm dish and place it back into the cell incubator. 
		CRITICAL: At this stage, because spheroids have not formed yet, it is very important to avoid strong aspiration or dispensing that will result in loss of the cells. Visual control using a benchtop brightfield microscope is highly recommended at this step.</li>
	</ol>
	</li>
</ol>
8. Immunostaining and imaging
<ol>
	<li>Fixation and staining 
	NOTE: Different classical procedures of fixation and immunostaining are completely compatible with the cell culture dish. One typical protocol is described here.
	<ol>
		<li>Fix the organoids/spheroids in the cell culture dish for 20 min in 4% paraformaldehyde at room temperature.</li>
		<li>Permeabilize the organoids for 24 h in 1% surfactant solution in sterile PBS at 4 &#176;C on an orbital shaker and incubate for 24 h in blocking buffer (2% bovine serum albumin [BSA] and 1% surfactant in sterile PBS) at 4 &#176;C on an orbital shaker.</li>
		<li>Incubate the samples with primary antibodies of interest at a dilution between 1/50 and 1/200 (or according to the manufacturer&#39;s recommendations) in antibody dilution buffer (2% BSA and 0.2% surfactant in sterile PBS) at 4 &#176;C for 48 h.</li>
		<li>Rinse the samples 3x with washing buffer on an orbital shaker (3% NaCl and 0.2% surfactant in sterile PBS) and incubate with corresponding secondary antibodies in antibody dilution buffer (dilution between 1/100 and 1/300 or according to the manufacturer&#39;s recommendations), 0.5 &#181;g/mL 4&#39;,6-diamidino-2-phenylindole (DAPI), and 0.2 &#181;g/mL Alexa Fluor 647 or 488 Phalloidin at 4 &#176;C for 24 h on an orbital shaker, followed by five rinsing steps with PBS. Optionally, mount the samples using a water-soluble clearing agent prewarmed to 37 &#730;C.</li>
	</ol>
	</li>
	<li>Imaging 
	NOTE: At this stage, the organoids in the microwell plate can be considered as a normal culture dish containing fixed and stained samples for imaging: any standard imaging procedure can be used with no adaptation or modification required. Figure 12 illustrates a representative result of images and 3D reconstruction obtained using a spinning disk confocal microscope, with a 40x air objective (numerical aperture 0.75).
	<ol>
		<li>Use constructor software for automatic image acquisition process with the following settings: exposure time = 50 ms, with a z motorized stage to acquire a z-stack (1 &#181;m Z-step, for a total height of 70 &#181;m).</li>
		<li>Perform 3D reconstruction using image analysis software.</li>
	</ol>
	</li>
</ol>
9. Release and collection of the organoids
NOTE: The textured adhesive layer of the cell culture dish can be detached from the coverslip to release the living spheroids/organoids (before fixation) contained inside the pyramidal cavities for analysis of the cells with other procedures such as RNA sequencing, -omic approaches, in vitro experiments, and in vivo transplantation.
<ol>
	<li>With the sample still in the Petri dish and in a biosafety hood, use a blade such as a scalpel to cut a corner of the textured adhesive layer.</li>
	<li>With tweezers, pinch the textured adhesive layer on the cut edge and gently peel it off from the glass coverslip but keep it immersed in the medium (some organoids might be remaining with the adhesive layer, Figure 13).</li>
	<li>Rinse 3x with culture medium and collect the organoids by pipetting.</li>
</ol>

<ol>
	<li>Kim, J., Koo, B. -. K., Knoblich, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+organoids:+model+systems+for+human+biology+and+medicine.">Human organoids: model systems for human biology and medicine.</a> Nature Reviews Molecular Cell Biology. 21 (10), 571-584 (2020).</li><li>Takebe, T., Wells James, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organoids+by+design.">Organoids by design.</a> Science. 364 (6444), 956-959 (2019).</li><li>Kratochvil, M. J., 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=Engineered+materials+for+organoid+systems.">Engineered materials for organoid systems.</a> Nature Reviews Materials. 4 (9), 606-622 (2019).</li><li>Rossi, G., Manfrin, A., Lutolf, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Progress+and+potential+in+organoid+research.">Progress and potential in organoid research.</a> Nature Reviews Genetics. 19 (11), 671-687 (2018).</li><li>O'Connell, L., Winter, D. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organoids:+past+learning+and+future+directions.">Organoids: past learning and future directions.</a> Stem Cells and Development. 29 (5), 281-289 (2020).</li><li>Vives, J., Batlle-Morera, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+challenge+of+developing+human+3D+organoids+into+medicines.">The challenge of developing human 3D organoids into medicines.</a> Stem Cell Research &amp; Therapy. 11 (1), 72 (2020).</li><li>Busslinger, G. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+potential+and+challenges+of+patient-derived+organoids+in+guiding+the+multimodality+treatment+of+upper+gastrointestinal+malignancies.">The potential and challenges of patient-derived organoids in guiding the multimodality treatment of upper gastrointestinal malignancies.</a> Open Biology. 10 (4), 190274 (2020).</li><li>Lukonin, I., 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=Phenotypic+landscape+of+intestinal+organoid+regeneration.">Phenotypic landscape of intestinal organoid regeneration.</a> Nature. 586 (7828), 275-280 (2020).</li><li>Rios, A. C., Clevers, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+organoids:+a+bright+future+ahead.">Imaging organoids: a bright future ahead.</a> Nature Methods. 15 (1), 24-26 (2018).</li><li>Dekkers, J. F., 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=High-resolution+3D+imaging+of+fixed+and+cleared+organoids.">High-resolution 3D imaging of fixed and cleared organoids.</a> Nature Protocols. 14 (6), 1756-1771 (2019).</li><li>Galland, R., 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=3D+high-+and+super-resolution+imaging+using+single-objective+SPIM.">3D high- and super-resolution imaging using single-objective SPIM.</a> Nature Methods. 12 (7), 641-644 (2015).</li><li>Beghin, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High+content+3D+imaging+method+for+quantitative+characterization+of+organoid+development+and+phenotype.">High content 3D imaging method for quantitative characterization of organoid development and phenotype.</a> bioRxiv. , (2021).</li><li>Beghin, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+high-speed+3D+imaging+of+organoid+cultures+with+multi-scale+phenotypic+quantification.">Automated high-speed 3D imaging of organoid cultures with multi-scale phenotypic quantification.</a> Nature Methods. 19 (7), 881-892 (2022).</li></ol>

An international patent application has been published with the Publication Number WO 2021/167535 A1.

The procedure for the fabrication of the microwell culture dish, which allows high-density organoid culture and differentiation, has been described in this paper. Owing to the geometry and arrangement of the microcavities, thousands of spheroids can be cultured and stained in a single plate for long periods of time (several weeks or more) with nearly no loss of material. As a comparison, an area of 4 mm x 2 mm on the cell culture plate can contain as many spheroids as a single 384-well plate with an area of 12 cm x 8 cm....

In organotypic 3D cell cultures, hereafter referred&#160;to as organoids, stem cells differentiate and self-organize into spatial structures that share strong morphological and functional similarities with real organs. Organoids offer valuable models to study human biology and development outside the body1,2,3. A growing number of models are being developed that mimic the liver, brain, kidney, lung, and many other organs2,4,5. Differentiation in organoids is directed by the addition of soluble growth factors and an extracellular matrix in a precise time sequence. However, in marked contrast to organs, the development of organoids is quite heterogeneous.
Beyond numerous biological challenges6,7, organoid cultures also pose technological challenges in terms of cell culture methods, characterization of transcriptomics, and imaging. In vivo organ development occurs in a biological environment that results in a highly stereotypical self-organization of cell arrangements. Any phenotypic alteration can be used as a proxy to diagnose a diseased state. In contrast, organoids develop in vitro in minimally controlled microenvironments compatible with cell culture conditions, resulting in large variability in the development path and shape formation for each individual organoid.
A recent study8 demonstrated that quantitative imaging of organoid shape (phenotype descriptors) coupled to the assessment of a few genetic markers allow the definition of phenotypic development landscapes. Arguably, the ability to relate the diversity of genomic expression in organoids with their phenotypic behavior is a major step toward unleashing the full potential of organotypic cultures. Thus, it begs for the development of dedicated, high-content imaging approaches allowing the characterization of organoid features at subcellular, multicellular, and whole-organoid scales in 3D9,10.
We developed a versatile high-content screening (HCS) platform allowing streamlined organoid culture (from isolated human embryonic stem cells [hESCs], human induced pluripotent stem cells [hIPSCs], or primary cells to 3D, multicellular, differentiated organoids) and fast, non-invasive 3D imaging. It integrates a next-generation, miniaturized, 3D cell culture device, called the JeWells chip (the chip hereafter), that contains thousands of well-arrayed microwells flanked with 45&#176; mirrors that allow fast, 3D, high-resolution imaging by single-objective light-sheet microscopy11. Compatible with any standard, commercial, inverted microscope, this system enables the imaging of 300 organoids in 3D with subcellular resolution in &#60;1 h.
The microfabrication of the cell culture device starts from an existing micro structured mold, which contains hundreds of micropyramids (Figure 1A) with a square base and sidewalls at 45&#176; with respect to the base. Figure 1C shows electron microscope (EM) images of such structures. The mold itself is made of poly(dimethylsiloxane) (PDMS) and can be made as a replica cast of a primary mold (not shown here) with corresponding features (as cavities) using standard soft-lithographic procedures. The primary mold can be produced by different procedures. The one used for this protocol was made using silicon wet etching as reported in Galland et al.11; the procedure for the fabrication of the primary mold is not critical for this protocol. The pyramids are arranged in a squared array, with the same pitch for the X and Y directions (in this case the pitch is 350 &#181;m).
As an illustration, proof-of-concept experiments12 were published to demonstrate that the chip allows long-term culture (months) and differentiation protocols while precisely defining the number of initial cells in the wells. Individual development of a large number of organoids can be automatically monitored live using standard brightfield and 3D light-sheet fluorescence microscopes. Moreover, organoids can be retrieved to perform further biological investigations (e.g., transcriptomic analysis). This paper outlines detailed protocols for the fabrication of the cell culture coverslips, the seeding and staining procedure for fluorescence microscopy, as well as the retrieval of the organoids.

<table><tbody><tr><td>2-Propanol</td><td>Thermofisher scientific</td><td>AA19397K7</td><td></td></tr><tr><td>Acetone</td><td>Thermofisher scientific</td><td>AA19392K7</td><td></td></tr><tr><td>BC43 Benchtop Confocal Microscope</td><td>Andor Technology</td><td></td><td>spinning disk confocal microscope</td></tr><tr><td>bovine serum albumin&amp;nbsp;</td><td>Thermofisher scientific</td><td>37525</td><td></td></tr><tr><td>Buffered oxide etching solution</td><td>Merck</td><td>901621-1L</td><td></td></tr><tr><td>CEE Spin Coater</td><td>Brewer Science</td><td>200X</td><td></td></tr><tr><td>DAPI</td><td>Thermofisher scientific</td><td>62248</td><td></td></tr><tr><td>Developer AZ400K</td><td>Merck</td><td>18441223164</td><td></td></tr><tr><td>DI Milliq water</td><td>Millipore</td><td></td><td></td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>Invitrogen</td><td>10082147</td><td></td></tr><tr><td>Glass coverslips</td><td>Marienfled</td><td>117650</td><td>1.5H, round 25 mm diameter</td></tr><tr><td>Hepes</td><td>Invitrogen</td><td>15630080</td><td></td></tr><tr><td>Imaris software</td><td>BitPlane</td><td></td><td>image analysis software</td></tr><tr><td>Inverted Transmission optical microscope</td><td>Nikon</td><td>TSF100-F</td><td></td></tr><tr><td>Labsonic M</td><td>Sartorius Stedium Biotech</td><td></td><td>Ultrasonic homogenizer</td></tr><tr><td>Lipidure</td><td>NOF America</td><td>CM5206</td><td>bio-mimetic copolymer</td></tr><tr><td>NOA73</td><td>Norland Products</td><td>17-345</td><td>UV curable adhesive</td></tr><tr><td>Penicillin-Streptomycin</td><td>Invitrogen</td><td>15070063</td><td></td></tr><tr><td>Phalloidin</td><td>Thermofisher scientific&amp;nbsp;</td><td>A12379</td><td>Alexa Fluor</td></tr><tr><td>Phosphate Buffer Solution</td><td>Thermofisher scientific</td><td>10010023</td><td></td></tr><tr><td>Photo Resist AZ5214E</td><td>Merck</td><td>14744719710</td><td></td></tr><tr><td>Pico Plasma tool</td><td>Diener Electronic GmbH + Co. KG</td><td>Pico Plasma</td><td>For O&lt;sub&gt;2&lt;/sub&gt; plasma treatment</td></tr><tr><td>RapiClear 1.52</td><td>Sunjin lab</td><td>RC 152001</td><td>water-soluble clearing agent</td></tr><tr><td>RCT Hot Plate/Stirrer</td><td>IKA (MY)</td><td></td><td></td></tr><tr><td>Reactive Ion Etching tool</td><td>Samco Inc. (JPN)</td><td>RIE-10NR</td><td></td></tr><tr><td>RPMI 1640</td><td>Invitrogen</td><td>11875093</td><td>culture medium for HCT116 cells</td></tr><tr><td>Sylgard 184 Silicone Elastomer Kit</td><td>Dow Corning</td><td>4019862</td><td>Polydimethylsiloxane or in short, PDMS</td></tr><tr><td>Trichloro(1H,1H,2H,2H-perfluorooctyl)silane</td><td>Sigma Aldrich</td><td>448931-10G</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>T9284</td><td>surfactant</td></tr><tr><td>Trypsin EDTA</td><td>Thermofisher scientific</td><td>15400054</td><td></td></tr><tr><td>Ultrasonic Cleaner</td><td>Bransonic</td><td>CPX2800</td><td></td></tr><tr><td>UV-KUB 2</td><td>KLOE</td><td></td><td>UV-LED light source, 365 nm wavelength, 35 mW/cm&lt;sup&gt;2&lt;/sup&gt; power density</td></tr><tr><td>UV mask aligner</td><td>SUSS Microtec Semiconductor (DE)</td><td>MJB4</td><td></td></tr></tbody></table>

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).

Figure 8F shows the typical aspect of a cell culture coverslip after successful fabrication. The UV-curable adhesive layer appears flat and adheres well to the coverslip. The transfer of the adhesive layer on the coverslip might fail if the layer on the coverslip is overcured, or if the removal of the flat PDMS substrate is done incorrectly (as shown in Figure 8G,H). In both cases, the failure is evident as no textured film is transferred to the...

Watch this Scientific Journal Video about A High-Throughput Platform for Culture and 3D Imaging of Organoids at JoVE.com

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

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 ...

a-high-throughput-platform-for-culture-and-3d-imaging-of-organoids

Research

JoVE Journal

Biology

2.4K Views.  National University of Singapore. 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.

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

Enhanced Viability for Ex vivo 3D Hydrogel Cultures of Patient-Derived Xenografts in a Perfused Microfluidic Platform

Patient-derived xenografts (PDX), generated when resected patient tumor tissue is engrafted directly into immunocompromised mice, remain biologically stable, thereby preserving molecular, genetic, and histological features, as well as heterogeneity of the original tumor. However, using these models to perform a multitude of experiments, including drug screening, is prohibitive both in terms of cost and time. Three-dimensional (3D) culture systems are widely viewed as platforms in which cancer cells retain their biological integrity through biochemical interactions, morphology, and architecture. Our team has extensive experience culturing PDX cells in vitro using 3D matrices composed of hyaluronic acid (HA). In order to separate mouse fibroblast stromal cells associated with PDXs, we use rotation culture, where stromal cells adhere to the surface of tissue culture-treated plates while dissociated PDX tumor cells float and self-associate into multicellular clusters. Also floating in the supernatant are single, often dead cells, which present a challenge in collecting viable PDX clusters for downstream encapsulation into hydrogels for 3D cell culture. In order to separate these single cells from live cell clusters, we have employed density step gradient centrifugation. The protocol described here allows for the depletion of non-viable single cells from the healthy population of cell clusters that will be used for further in vitro experimentation. In our studies, we incorporate the 3D cultures in microfluidic plates which allow for media perfusion during culture. After assessing the resultant cultures using a fluorescent image-based viability assay of purified versus non-purified cells, our results show that this additional separation step substantially reduced the number of non-viable cells from our cultures.

This protocol demonstrates methods to enable extended in vitro culture of patient-derived xenografts (PDX). One step enhances overall viability of multicellular cluster cultures in 3D hydrogels, through straightforward removal of non-viable single cells. A secondary step demonstrates best practices for PDX culture in a perfused microfluidic platform.

Patient-derived xenografts (PDX), generated when resected patient tumor tissue is engrafted directly into immunocompromised mice, remain biologically ...

Cancer Research

Three-Dimensional Imaging of Organoids to Study Primary Ciliogenesis During ex vivo Organogenesis

Organoids are stem cell-derived three-dimensional structures that reproduce ex vivo the complex architecture and physiology of organs. Thus, organoids represent useful models to study the mechanisms that control stem cell self-renewal and differentiation in mammals, including primary ciliogenesis and ciliary signaling. Primary ciliogenesis is the dynamic process of assembling the primary cilium, a key cell signaling center that controls stem cell self-renewal and/or differentiation in various tissues. Here we present a comprehensive protocol for the immunofluorescence staining of cell lineage and primary cilia markers, in whole-mount mouse mammary organoids, for light sheet microscopy. We describe the microscopy imaging method and an image processing technique for the quantitative analysis of primary cilium assembly and length in organoids. This protocol enables a precise analysis of primary cilia in complex three-dimensional structures at the single cell level. This method is applicable for immunofluorescence staining and imaging of primary cilia and ciliary signaling in mammary organoids derived from normal and genetically modified stem cells, from healthy and pathological tissues, to study the biology of the primary cilium in health and disease.

Three-Dimensional Imaging of Organoids to Study Primary Ciliogenesis During ex vivo Organogenesis

Stem cell-derived organoids facilitate the analysis of molecular and cellular processes that regulate stem cell self-renewal and differentiation during organogenesis in mammalian tissues. Here we present a protocol for the analysis of the biology of the primary cilium in mouse mammary organoids.

Organoids are stem cell-derived three-dimensional structures that reproduce ex vivo the complex architecture and physiology of organs. Thus, organoids ...

Genetics

Medium-Throughput Drug- and Radiotherapy Screening Assay using Patient-Derived Organoids

Patient-derived organoid (PDO) models allow for long-term expansion and maintenance of primary epithelial cells grown in three dimensions and a near-native state. When derived from resected or biopsied tumor tissue, organoids closely recapitulate in vivo tumor morphology and can be used to study therapy response in vitro. Biobanks of tumor organoids reflect the vast variety of clinical tumors and patients and therefore hold great promise for preclinical and clinical applications. This paper presents a method for medium-throughput drug screening using head and neck squamous cell carcinoma and colorectal adenocarcinoma organoids. This approach can easily be adopted for use with any tissue-derived organoid model, both normal and diseased. Methods are described for in vitro exposure of organoids to chemo- and radiotherapy (either as single-treatment modality or in combination). Cell survival after 5 days of drug exposure is assessed by measuring adenosine triphosphate (ATP) levels. Drug sensitivity is measured by the half-maximal inhibitory concentration (IC50), area under the curve (AUC), and growth rate (GR) metrics. These parameters can provide insight into whether an organoid culture is deemed sensitive or resistant to a particular treatment.

We describe detailed protocols to use patient-derived organoids for medium-throughput therapy sensitivity screenings. Therapies tested include chemotherapy, radiotherapy, and chemo-radiotherapy. Adenosine triphosphate levels are used as a functional readout.

Patient-derived organoid (PDO) models allow for long-term expansion and maintenance of primary epithelial cells grown in three dimensions and a ...

High-Throughput In Vitro Assay using Patient-Derived Tumor Organoids

Patient-derived tumor organoids (PDOs) are expected to be a preclinical cancer model with better reproducibility of disease than traditional cell culture models. PDOs have been successfully generated from a variety of human tumors to recapitulate the architecture and function of tumor tissue accurately and efficiently. However, PDOs are unsuitable for an in vitro high-throughput assay system (HTS) or cell analysis using 96-well or 384-well plates when evaluating anticancer drugs because they are heterogeneous in size and form large clusters in culture. These cultures and assays use extracellular matrices, such as Matrigel, to create tumor tissue scaffolds. Therefore, PDOs have a low throughput and high cost, and it has been difficult to develop a suitable assay system. To address this issue, a simpler and more accurate HTS was established using PDOs to evaluate the potency of anticancer drugs and immunotherapy. An in vitro HTS was created that uses PDOs established from solid tumors cultured in 384-well plates. An HTS was also developed for assessment of antibody-dependent cellular cytotoxicity activity to represent the immune response using PDOs cultured in 96-well plates.

High-Throughput In Vitro Assay using Patient-Derived Tumor Organoids

A highly accurate in vitro high-throughput assay system was developed to evaluate anticancer drugs using patient-derived tumor organoids (PDOs), similar to cancer tissues but are unsuitable for in vitro high-throughput assay systems with 96-well and 384-well plates.

Patient-derived tumor organoids (PDOs) are expected to be a preclinical cancer model with better reproducibility of disease than traditional cell culture ...

Multiparametric Tumor Organoid Drug Screening Using Widefield Live-Cell Imaging for Bulk and Single-Organoid Analysis

Patient-derived tumor organoids (PDTOs) hold great promise for preclinical and translational research and predicting the patient therapy response from ex vivo drug screenings. However, current adenosine triphosphate (ATP)-based drug screening assays do not capture the complexity of a drug response (cytostatic or cytotoxic) and intratumor heterogeneity that has been shown to be retained in PDTOs due to a bulk readout. Live-cell imaging is a powerful tool to overcome this issue and visualize drug responses more in-depth. However, image analysis software is often not adapted to the three-dimensionality of PDTOs, requires fluorescent viability dyes, or is not compatible with a 384-well microplate format. This paper describes a semi-automated methodology to seed, treat, and image PDTOs in a high-throughput, 384-well format using conventional, widefield, live-cell imaging systems. In addition, we developed viability marker-free image analysis software to quantify growth rate-based drug response metrics that improve reproducibility and correct growth rate variations between different PDTO lines. Using the normalized drug response metric, which scores drug response based on the growth rate normalized to a positive and negative control condition, and a fluorescent cell death dye, cytotoxic and cytostatic drug responses can be easily distinguished, profoundly improving the classification of responders and non-responders. In addition, drug-response heterogeneity can by quantified from single-organoid drug response analysis to identify potential, resistant clones. Ultimately, this method aims to improve the prediction of clinical therapy response by capturing a multiparametric drug response signature, which includes kinetic growth arrest and cell death quantification.

This protocol describes a semi-automated method for medium- to high-throughput organoid drug screenings and microscope-agnostic, automated image analysis software to quantify and visualize multiparametric, single-organoid drug responses to capture intratumor heterogeneity.

Patient-derived tumor organoids (PDTOs) hold great promise for preclinical and translational research and predicting the patient therapy response from ex ...

3D Analysis of Multi-cellular Responses to Chemoattractant Gradients

Various limitations of 2D cell culture systems have sparked interest in 3D cell culture and analysis platforms, which would better mimic the spatial and chemical complexity of living tissues and mimic in vivo tissue functions. Recent advances in microfabrication technologies have facilitated the development of 3D in vitro environments in which cells can be integrated into a well-defined extracellular matrix (ECM) and a defined set of soluble or matrix associated biomolecules. However, technological barriers have limited their widespread use in research laboratories. Here, we describe a method to construct simple devices for 3D culture and experimentation with cells and multicellular organoids in 3D microenvironments with a defined chemoattractant gradient. We illustrate the use of this platform for analysis of the response of epithelial cells and organoids to gradients of growth factors, such as epidermal growth factor (EGF). EGF gradients were stable in the devices for several days leading to directed branch formation in breast organoids. This analysis allowed us to conclude that collective gradient sensing by groups of cells is more sensitive vs. single cells. We also describe the fabrication method, which does not require photolithography facilities nor advanced soft lithography techniques. This method will be helpful to study 3D cellular behaviors in the context of the analysis of development and pathological states, including cancer.

We describe a method to construct devices for 3D culture and experimentation with cells and multicellular organoids. This device allows analysis of cellular responses to soluble signals in 3D microenvironments with defined chemoattractant gradients. Organoids are better than single cells at detection of weak noisy inputs.

Various limitations of 2D cell culture systems have sparked interest in 3D cell culture and analysis platforms, which would better mimic the spatial and ...

Bioengineering

Facilitating Cerebral Organoid Culture via Lateral Soft Light Illumination

At present, cerebral organoid culture technology is still complicated to operate and difficult to apply on a large scale. It is necessary to find a simple and practical solution. Therefore, a more feasible cerebral organoid protocol is proposed in the present study. To solve the unavoidable inconvenience in medium change and organoid transfer in the early stage, the current research optimizes the operation technology by applying the engineering principle. A soft light lamp was adopted to laterally illuminate the embryoid body (EB) samples, allowing the EBs to be seen by the naked eye through the enhanced diffuse reflection effect. Using the principle of secondary flow generated by rotation, the organoids gather toward the center of the well, which facilitates the operation of medium change or organoid transfer. Compared to the dispersed cell, the embryoid body settles faster in the pipette. Using this phenomenon, most of the free cells and dead cell fragments can be effectively removed in a simple way, preventing EBs from incurring damage from centrifugation. This study facilitates the operation of cerebral organoid culture and helps to promote the application of brain organoids.

Facilitating Cerebral Organoid Culture via Lateral Soft Light Illumination

Cerebral organoids provide unprecedented opportunities for studying organ development and human disease pathology. Although great success has been achieved with cerebral organoid culture systems, there are still operational difficulties in applying this technology. The present protocol describes a cerebral organoid procedure that facilitates medium change and organoid transfer.

At present, cerebral organoid culture technology is still complicated to operate and difficult to apply on a large scale. It is necessary to find a simple ...

Culture and Imaging of Human Nasal Epithelial Organoids

Individualized therapy for cystic fibrosis (CF) patients can be achieved with an in vitro disease model to understand baseline Cystic Fibrosis Transmembrane conductance Regulator (CFTR) activity and restoration from small molecule compounds. Our group recently focused on establishing a well-differentiated organoid model directly derived from primary human nasal epithelial cells (HNE). Histology of sectioned organoids, whole-mount immunofluorescent staining, and imaging (using confocal microscopy, immunofluorescent microscopy, and bright field) are essential to characterize organoids and confirm epithelial differentiation in preparation for functional assays. Furthermore, HNE organoids produce lumens of varying sizes that correlate with CFTR activity, distinguishing between CF and non-CF organoids. In this manuscript, the methodology for culturing HNE organoids are described in detail, focusing on the assessment of differentiation using the imaging modalities, including the measurement of baseline lumen area (a method of CFTR activity measurement in organoids that any laboratory with a microscope can employ) as well as the developed automated approach to a functional assay (which requires more specialized equipment).

A detailed protocol is presented here to describe an in vitro organoid model from human nasal epithelial cells. The protocol has options for measurements requiring standard laboratory equipment, with additional possibilities for specialized equipment and software.

Individualized therapy for cystic fibrosis (CF) patients can be achieved with an in vitro disease model to understand baseline Cystic Fibrosis ...

Medicine

Single-Cell Resolution Three-Dimensional Imaging of Intact Organoids

Organoid technology, in vitro 3D culturing of miniature tissue, has opened a new experimental window for cellular processes that govern organ development and function as well as disease. Fluorescence microscopy has played a major role in characterizing their cellular composition in detail and demonstrating their similarity to the tissue they originate from. In this article, we present a comprehensive protocol for high-resolution 3D imaging of whole organoids upon immunofluorescent labeling. This method is widely applicable for imaging of organoids differing in origin, size and shape. Thus far we have applied the method to airway, colon, kidney, and liver organoids derived from healthy human tissue, as well as human breast tumor organoids and mouse mammary gland organoids. We use an optical clearing agent, FUnGI, which enables the acquisition of whole 3D organoids with the opportunity for single-cell quantification of markers. This three-day protocol from organoid harvesting to image analysis is optimized for 3D imaging using confocal microscopy.

The entire 3D structure and cellular content of organoids, as well as their phenotypic resemblance to the original tissue can be captured using the single-cell resolution 3D imaging protocol described here. This protocol can be applied to a wide range of organoids varying in origin, size and shape.

Organoid technology, in vitro 3D culturing of miniature tissue, has opened a new experimental window for cellular processes that govern organ development ...

Staining and High-Resolution Imaging of Three-Dimensional Organoid and Spheroid Models

In vitro three-dimensional (3D) cell culture models, such as organoids and spheroids, are valuable tools for many applications including development and disease modeling, drug discovery, and regenerative medicine. To fully exploit these models, it is crucial to study them at cellular and subcellular levels. However, characterizing such in vitro 3D cell culture models can be technically challenging and requires specific expertise to perform effective analyses. Here, this paper provides detailed, robust, and complementary protocols to perform staining and subcellular resolution imaging of fixed in vitro 3D cell culture models ranging from 100 &#181;m to several millimeters. These protocols are applicable to a wide variety of organoids and spheroids that differ in their cell-of-origin, morphology, and culture conditions. From 3D structure harvesting to image analysis, these protocols can be completed within 4-5 days. Briefly, 3D structures are collected, fixed, and can then be processed either through paraffin-embedding and histological/immunohistochemical staining, or directly immunolabeled and prepared for optical clearing and 3D reconstruction (200 &#181;m depth) by confocal microscopy.

Here, we provide detailed, robust, and complementary protocols to perform staining and subcellular resolution imaging of fixed three-dimensional cell culture models ranging from 100 &#181;m to several millimeters, thus enabling the visualization of their morphology, cell-type composition, and interactions.

In vitro three-dimensional (3D) cell culture models, such as organoids and spheroids, are valuable tools for many applications including development and ...

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

Summary

Explore More Videos

Chapters in this video

Enhanced Viability for Ex vivo 3D Hydrogel Cultures of Patient-Derived Xenografts in a Perfused Microfluidic Platform

Three-Dimensional Imaging of Organoids to Study Primary Ciliogenesis During ex vivo Organogenesis

Medium-Throughput Drug- and Radiotherapy Screening Assay using Patient-Derived Organoids

High-Throughput In Vitro Assay using Patient-Derived Tumor Organoids

Multiparametric Tumor Organoid Drug Screening Using Widefield Live-Cell Imaging for Bulk and Single-Organoid Analysis

3D Analysis of Multi-cellular Responses to Chemoattractant Gradients

Facilitating Cerebral Organoid Culture via Lateral Soft Light Illumination

Culture and Imaging of Human Nasal Epithelial Organoids

Single-Cell Resolution Three-Dimensional Imaging of Intact Organoids

Staining and High-Resolution Imaging of Three-Dimensional Organoid and Spheroid Models