We are very grateful for the technical support from the Princess M&#225;xima Center for Pediatric Oncology and to the Hubrecht Institute and Zeiss for imaging support and collaborations. All the imaging was performed at the Princess M&#225;xima imaging center. This work was financially supported by the Princess M&#225;xima Center for Pediatric Oncology. JFD was supported by a Marie Curie Global Fellowship and a VENI grant from the Netherlands Organisation for Scientific Research (NWO). ACR was supported by a European Council (ERC) starting grant.

Use of mouse-derived organoids conformed to regulatory standards and was approved by the Walter and Eliza Hall Institute (WEHI) Animal Ethics Committee. All human organoid samples were retrieved from biobanks through the Hubrecht Organoid Technology (HUB, www.hub4organoids.nl). Authorizations were obtained by the Medical Ethical Committee of UMC Utrecht (METC UMCU) at request of the HUB in order to ensure compliance with the Dutch Medical Research Involving Human Subjects Act and informed consent was obtained from donors when appropriate.
1. Preparation of reagents
<ol>
	<li>To prepare 4% (w/v) paraformaldehyde (PFA), heat 400 mL of phosphate-buffered saline (PBS) to just under 60 &#176;C in a water bath. Add 20 g of PFA powder and dissolve using a stirrer.
	<ol>
		<li>Next, add a few drops of 10 M NaOH. Let cool on ice and add a few drops of 10 M HCl to adjust the pH to 7.4. Top up with PBS to 500 mL and aliquot (store at -20 &#176;C for up to 2 months). 
		NOTE: Do not heat above 60 &#176;C to avoid degradation of the PFA. Preparation time = 4 h.</li>
	</ol>
	</li>
	<li>To prepare PBS with Tween-20 (PBT) (0.1% v/v), add 1 mL of Tween-20 to 1 L of PBS (store at 4 &#176;C for up to 4 weeks). 
	NOTE: Preparation time = 10 min.</li>
	<li>To prepare 100 mL of 0.5 M ethylenediaminetetraacetic acid (EDTA), add 18.6 g of EDTA and 2.5 g of NaOH to 80 mL of dH2O. Adjust the pH to 8 with 1 M NaOH and fill to 100 mL with dH2O.</li>
	<li>To prepare 500 mL of 1 M Tris, dissolve 60.55 g of Tris with 42 mL of concentrated (36&#8722;38%) HCl in 300 mL of dH2O. Adjust the pH to 8 and fill to 500 mL.</li>
	<li>To prepare organoid washing buffer (OWB), add 1 mL of Triton X-100, 2 mL of 10% (w/v) SDS and 2 g of bovine serum albumin (BSA) to 1 L of PBS (store at 4 &#176;C up to 2 weeks). 
	NOTE: Preparation time = 10 min.</li>
	<li>To make 220 mL of FUnGI, mix 110 mL of glycerol with 20 mL of dH2O, 2.2 mL of Tris buffer (1 M, pH 8.0) and 440 &#181;L of EDTA (0.5 M). Add 50 g of fructose and mix at room temperature (RT) in the dark until dissolved. When clear, add 49 g of fructose and mix until dissolved. Then add 33.1 g of urea and mix until dissolved (store at 4 &#176;C in the dark). 
	NOTE: Do not heat as fructose caramelizes at higher temperatures. FUnGI consists of 50% (v/v) glycerol, 9.4% (v/v) dH2O, 10.6 mM tris base, 1.1 mM EDTA, 2.5 M fructose and 2.5 M urea. Preparation time = 1 day.</li>
	<li>To prepare PBS-BSA (1% w/v), dissolve 1 g of BSA in 100 mL of PBS (store at 4 &#176;C up to 2 weeks). 
	NOTE: Preparation time = 10 min.</li>
</ol>
2. Organoid recovery
NOTE: The following steps apply to organoids grown in basement membrane extract (BME) that were cultured in a 24 well plate with a size of 100&#8722;500 &#181;m .
<ol>
	<li>Aspirate the culture medium and wash 1x with ice-cold PBS. Avoid disrupting the 3D matrix.</li>
	<li>Put the plate on ice and add 1 mL of ice-cold cell recovery solution (Table of Materials) to each well. Incubate 30&#8722;60 min at 4 &#176;C on a horizontal shaker (40 rpm). 
	NOTE: The 3D matrix droplets should be completely dissolved.</li>
	<li>Coat a 1 mL pipette tip with BSA by dipping in 1% PBS-BSA and pipetting up and down 2x. This coating will prevent organoids from sticking to the tip. To coat the inner side of a 15 mL conical tube, fill with 5 mL of 1% PBS-BSA, invert 2&#8722;3x and discard the PBS-BSA. 
	NOTE: This coating is necessary for all plastic consumables until fixation (step 3.3).</li>
	<li>Using a coated tip, gently resuspend the content of the well 5&#8722;10x and transfer the organoids to coated 15 mL tubes. Organoids from different wells with the same identity can be pooled into the same tube.</li>
	<li>Add 1 mL of ice-cold 1% PBS-BSA to each culture well to rinse and collect all organoids.</li>
	<li>Add cold PBS to 10 mL and spin down for 3 min at 70 x g and 4 &#176;C to obtain a tight pellet without a visible layer of 3D matrix. Carefully remove the supernatant.</li>
</ol>
3. Fixation and blocking
<ol>
	<li>Carefully resuspend the organoids in 1 mL of ice-cold PFA using a coated 1 mL tip.</li>
	<li>Fix at 4 &#176;C for 45 min. Gently resuspend the organoids halfway through the fixation time using a coated 1 mL tip to ensure even fixation among all organoids.</li>
	<li>Add 10 mL of ice-cold PBT to the tube, gently mix by inverting the tube, incubate for 10 min and spin down at 70 x g, both at 4 &#176;C. 
	NOTE: From this step onwards coating of tips is generally not needed as most organoid types do not stick to the tip after fixation. However, some organoids may require coated plastics even after fixation.</li>
	<li>Block the organoids by resuspending the pellet in ice-cold OWB (at least 200 &#956;L of OWB per well) and transfer the organoids to a 24 well suspension plate. 
	NOTE: Organoids from one large pellet can be split over multiple wells to perform different stainings.</li>
	<li>Incubate at 4 &#176;C for at least 15 min.</li>
</ol>
4. Immunolabeling
<ol>
	<li>Pipette 200 &#956;L of OWB in an empty well to serve as a reference well. 
	NOTE: The immunolabeling can also be performed in 48- or 96-wells plates to reduce antibody usage. However, the user should be aware that both staining and washing performance could be reduced due to the smaller volume.</li>
	<li>Allow the organoids to settle at the bottom of the plate. 
	NOTE: This can be checked using a stereomicroscope and is made easier by using a dark background.</li>
	<li>Tilt the plate 45&#176; and remove OWB leaving the organoids in 200 &#956;L of OWB (use the reference well to estimate 200 &#956;L).</li>
	<li>Add 200 &#956;L of OWB with primary antibodies 2x concentrated (e.g., E-cadherin [1:400] and Ki67 [1:200] for results in Figure 1; keratin 5 [1:500], keratin 8/18 [1:200], MRP2 [1:50], and Ki67 [1:200] for results in Figure 2) and incubate overnight at 4 &#176;C while mildly rocking/shaking (40 rpm on horizontal shaker).</li>
	<li>The next day, add 1 mL of OWB.</li>
	<li>Allow the organoids to settle at the bottom of the plate for 3 min. Remove OWB leaving 200 &#956;L in the plate. Add 1 mL of OWB and wash 2 h with mild rocking/shaking.</li>
	<li>Repeat step 4.6 two more times.</li>
	<li>Allow the organoids to settle at the bottom of the plate for 3 min. Remove OWB leaving 200 &#956;L in the well.</li>
	<li>Add 200 &#956;L of OWB with secondary antibodies, conjugated antibodies and dyes 2x concentrated (e.g., DAPI [1:1000], rat-AF488 [1:500], mouse-AF555 [1:500], phalloidin-AF647 [1:100] for results in Figure 1; DAPI [1:1000], rat-AF488 [1:500], rabbit-AF555 [1:500], mouse-AF555 [1:500], phalloidin-AF647 [1:100] for results in Figure 2; DAPI [1:1000], phalloidin-AF647 [1:100] for results in Figure 3) and incubate overnight at 4 &#176;C while mildly rocking/shaking.</li>
	<li>The next day, repeat steps 4.5&#8722;4.7.</li>
	<li>Transfer the organoids to a 1.5 mL tube and spin down at 70 x g for 3 min.</li>
</ol>
5. Optical clearing of organoids
<ol>
	<li>Remove as much as possible the OWB by pipetting without disrupting the organoids.</li>
	<li>Add FUnGI (at least 50 &#956;L, RT) using a 200 &#956;L tip with the end cut off and resuspend gently to prevent bubble formation. Incubate at RT for 20 min. 
	NOTE: Optical clearing by FUnGI may cause minor tissue shrinkage. This will not affect the general morphology of monolayered and multilayered organoids; however, spherical shaped monolayered organoids with large lumens may collapse. The protocol can be paused here and samples can be stored at 4 &#176;C (for at least 1 week) or at -20 &#176;C (for at least 6 months).</li>
</ol>
6. Slide preparation for confocal imaging
<ol>
	<li>Prepare a 10 mL syringe with a silicone sealant (Table of Materials). Attach a 200 &#956;L tip and cut off the end to allow a gentle flow of the viscous silicone after pressing the syringe. Use the syringe to draw a rectangle of 1 cm x 2 cm in the middle of a slide.</li>
	<li>Cut off the end of a 200 &#956;L tip and transfer the organoids in FUnGI to the middle of the rectangle.</li>
	<li>Place a coverslip on top. To minimize trapped air bubbles, place the left side of the coverslip first, then slowly lower the coverslip from left to right until there is no trapped air and then release the coverslip. 
	NOTE: Spacers that are simililar in size to the organoids can be used to prevent them from being damaged.</li>
	<li>Gently apply pressure on all edges of the coverslip to firmly attach it to the silicone sealant. The slide is now ready for imaging. 
	NOTE: The protocol can be paused here and the sample can be stored at 4 &#176;C (for at least 1 week) or -20 &#176;C (for at least 6 months).</li>
</ol>
7. Image acquisition and processing
<ol>
	<li>Using a confocal laser scanning microscope (Table of Materials), image the slide with a multi- immersion 25x or oil immersion 40x objective for confocal imaging.
	<ol>
		<li>Use the following acquisition settings for the 25x objective: scan mode frame, frame size 1024 x 1024, voxel size 332 nm x 332 nm x 1.2 &#181;m, pixel dwell time &#60;2 &#181;s, bidirectional scanning, averaging number 1, bit depth 8. To reduce photobleaching, use low laser power (&#60;5% in general, &#60;10% for weak staining).</li>
		<li>Use the Z-stack mode to define the lower and upper bounds and set the Z-step size to optimal. When imaging large organoid structures or multiple organoids together, use the tiling mode with 10% overlap and indicate the area of interest. 
		NOTE: With these settings, the data size for a typical organoid with a diameter of &#60;300 &#181;m is &#60;1 GB.</li>
	</ol>
	</li>
	<li>For tile scan datasets, stitch the imaging files in the software accompanied with the microscope (Table of Materials). In the processing section, select Stitching as the method, choose New Output under parameters and select the file to stitch. Press Apply to start stitching.</li>
	<li>Obtain a 3D rendered representation of the imaging under the 3D View tab in the imaging software (Table of Materials) and subsequently optimize brightness, contrasting and 3D rendering properties. Export RGB snapshots of the results as TIFF files.</li>
</ol>

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The authors have nothing to disclose.

Here, we put forward a detailed protocol for 3D imaging of intact organoids with single-cell resolution. To successfully perform this protocol, some critical steps have to be taken. In this section we highlight these steps and provide troubleshooting.
The first critical step is the removal of the 3D matrix. Most organoids are propagated in vitro with the use of matrices that mimic the in vivo extracellular environment to enhance the formation of well-polarized 3D structures. Fixating and subsequent staining within the 3D matrix is possible, but can be disadvantageous for the penetration of antibodies or can generate high background signal (data not shown). Efficient removal of matrices can be influenced by the type of matrix, the amount and size of the organoids and prolonged culturing. Therefore, optimization may be required for different culture conditions. For organoids cultured in Matrigel or BME, a 30&#8722;60 min step in ice-cold cell recovery solution is sufficient to dissolve the matrix without damaging the organoids. In addition, removal of the supporting 3D matrix could result in loss of native structures and disruption of organoid contacts with other cell types, for instance when organoids are co-cultured with fibroblasts or immune cells. Furthermore, optimal organoid fixation is crucial in preserving 3D tissue architecture, protein antigenicity and minimizing autofluorescence. Fixing for 45 min with 4% PFA at 4 &#176;C is normally sufficient for labeling of a wide range of organoids and antigens. However, a longer fixation step, up to 4 h, is typically more appropriate for organoids expressing fluorescent reporter proteins, but will require optimization for different fluorophores. Fixation times shorter than 20 min are insufficient to properly label F-actin using phalloidin probes. Another common issue is the loss of organoids during the protocol. It is therefore important to (i) carefully coat pipet tips and tubes with 1% BSA-PBS as described when handling unfixed organoids to prevent them from sticking to plastics, (ii) use low-adherence or suspension plates to avert the sample from sticking to the plate, and (iii) allow enough time for the organoids to settle at the bottom of the plate before carefully removing buffers. Pipetting viscous FUnGI may introduce bubbles. Handling the cleared sample at RT decreases viscosity and improves ease-of-use, thereby minimizing loss of organoids. While most organoids are easy to handle, cystic organoids with an enlarged lumen have a high tendency to collapse when fixing with 4% PFA or when cleared with FUnGI. This effect can be reduced, but not completely prevented, by using a different fixative (e.g., formalin or PFA-glutaraldehyde). However, this could potentially impact autofluorescence, antibody penetration and epitope availability. When cystic organoids appear folded after clearing, it is advised to skip the clearing step and image by multi-photon microscopy, which is less hampered by light scattering. Lastly, obtaining the entire 3D structure of organoids can be challenging and requires minimal distance between coverslip and organoid. In addition, when organoids have room to move in their mounting agent, this can result in X- and Y-shifts while recording data in Z-depth. Using less silicone sealant during slide preparation can solve suboptimal mounting between coverslip and microscope slide. However, too little silicone may lead to organoid compression and loss of their inherent 3D structure. FUnGI improves handling for slide mounting and stability of organoids while imaging, due to its higher viscosity.
While this protocol can be used for a broad range of applications to study in depth cellular content and 3D architecture of intact organoids, certain limitations should be considered. This methodology is rather low-throughput and time-consuming. Indeed, users should bear in mind that imaging large intact organoids in 3D requires both tiling and sample acquisition in Z, leading to prolonged acquisition times. Faster imaging could be achieved by using microscope assets, including resonant or spinning disk scanner, or by light sheet microscope technology26. Another consideration is that markers can be heterogeneously expressed between different organoids from the same sample. Therefore, multiple organoids should be acquired to better capture this organoid heterogeneity in culture. Lastly, while the complete wet lab procedure is straightforward, postprocessing of data requires skills in image analysis software for 3D visualization and quantification, as well as statistics for mining all the information present in the dataset.
In the last decade, the field of volume imaging has greatly advanced, due to both the development of a wide range of optical clearing agents and improvements in microscopy and computational technologies27,30,31. While in the past most studies focused on large volume imaging of organs or associated tumors, more recently methods for smaller and more fragile tissues, including organoid structures, have been developed36,37,38. We recently published a simple and fast method for imaging whole-mount organoids of various origin, size and shape at the single cell level for subsequent 3D rendering and image analysis26, which we presented here with some improvements (e.g., FUnGI, silicone mounting) and accompanied by a video protocol. This method is superior to conventional 2D section-based imaging in deciphering complex cell morphology and tissue architecture (Figure 2) and easy to implement in laboratories with a confocal microscope. With slight adaptations to the protocol, the samples can be made compatible with, super-resolution confocal, multi-photon as well as light sheet imaging, which makes this protocol widely applicable and provides users with a powerful tool to better comprehend the multidimensional complexity that can be modeled with organoids.

The advancement of novel culture methods, such as organoid technology, has enabled the culture of organs in a dish1. Organoids grow into three-dimensional (3D) structures that ressemble their tissue of origin as they preserve phenotypic and functional traits. Organoids are now instrumental for addressing fundamental biological questions2, modelling diseases including cancer3, and developing personalized treatment strategies4,5,6,7. Since the first protocol for generating organoids derived from intestinal adult stem cells8, organoid technology has extended to include a wide range of healthy and cancerous tissues derived from organs including prostate9, brain10, liver11,12, stomach13, breast14,15, endometrium16, salivary gland17, taste bud18, pancreas19, and kidney20.
The development of organoids has concurred with the rise of new volumetric microscopy techniques that can visualize the architecture of whole mount tissue in 3D21,22,23,24. 3D imaging is superior to traditional 2D tissue section imaging in visualizing the complex organization of biological specimens. 3D information proves to be essential for understanding cellular composition, cell shape, cell-fate decisions and cell-cell interactions of intact biological samples. Nondestructive optical sectioning techniques, such as confocal or multi-photon laser scanning microscopy (CLSM and MLSM) and light sheet fluorescence microscopy (LSFM), now enable the combined visualization of both fine details, as well as general tissue architecture, within a single biological specimen. This powerful imaging approach provides the opportunity to study the structural complexity that can be modeled with organoids25 and map the spatial distribution, phenotypic identity and cellular state of all individual cells composing these 3D structures.
Recently we published a detailed protocol for high-resolution 3D imaging of fixed and cleared organoids26. This protocol is specifically designed and optimized for processing delicate organoid structures, as opposed to methodologies for large intact tissues such as DISCO27,28, CUBIC29,30,31, and CLARITY32,33. As such, this method is generally applicable to a wide variety of organoids differing in origin, size and shape and cellular content. Furthermore, as compared to other volumetric imaging protocols that often require considerable time and effort, our protocol is undemanding and can be completed within 3 days. We have applied our 3D imaging protocol to visualize the architecture and cellular composition of newly developed organoid systems derived from various tissues, including human airways34, kidney20, liver11, and human breast cancer organoids15. In combination with multicolored fluorescent lineage tracing, this method has also been used to reveal the biopotency of basal cells in mouse mammary organoids14.
Here, we refine the protocol by introducing the nontoxic clearing agent FUnGI35. FUnGI clearing is achieved in a single incubation step, is easier to mount due to its viscosity, and better preserves fluorescence during storage. In addition, we introduce sodium dodecyl sulfate (SDS) to the wash buffer to enhance nuclear stainings as well as a silicone based-mounting method for easy slide preparation prior to microscopy. Figure 1 provides the graphical overview of the protocol (Figure 1A) and examples of 3D-imaged organoids (Figure 1B&#8722;D). In short, organoids are recovered from their 3D matrix, fixed and immunolabeled, optically cleared, imaged using confocal microscopy and then 3D rendered with visualization software.&#160;

<table><tbody><tr><td>1.5 ml safe-lock centrifuge tubes</td><td>Eppendorf</td><td>EP0030 120.094</td><td></td></tr><tr><td>2 ml safe-lock centrifuge tubes</td><td>Eppendorf</td><td>EP0030 120.094</td><td></td></tr><tr><td>Bovine Serum Albumin (BSA)</td><td>Sigma Aldrich</td><td>A3059</td><td></td></tr><tr><td>Cell recovery solution</td><td>Corning</td><td>354253</td><td></td></tr><tr><td>Confocal microscope</td><td>Zeiss</td><td>LSM880</td><td></td></tr><tr><td>Confocal microscope software ZEN</td><td>Zeiss</td><td>ZEN black/blue</td><td></td></tr><tr><td>Conical tubes 15 ml</td><td>Greiner Bio-One</td><td>5618-8271</td><td></td></tr><tr><td>Coverglass #1.5 24x60mm</td><td>Menzel-Glazer</td><td>G418-15</td><td></td></tr><tr><td>Coverglass #1.5 48x60mm</td><td>ProSciTech</td><td>G425-4860</td><td></td></tr><tr><td>DAPI</td><td>ThermoFisher</td><td>D3571</td><td>dilution 1:1000</td></tr><tr><td>Dissection Stereomicroscope</td><td>Leica</td><td>M205 FA</td><td></td></tr><tr><td>Double sided sticky tape 12,7 mm 6,35 m</td><td>Scotch 3M</td><td></td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate-bufferd Saline (DPBS)</td><td>Gibco</td><td>14190144</td><td>1x</td></tr><tr><td>EDTA</td><td>Invitrogen</td><td>15576-028</td><td></td></tr><tr><td>Focus Clear</td><td>CelExplorer</td><td>FC-101</td><td></td></tr><tr><td>Fructose</td><td>Sigma Aldrich</td><td>F0127</td><td></td></tr><tr><td>Glycerol</td><td>Boom</td><td>76050771.0500</td><td></td></tr><tr><td>Graduated Transfer Pipets</td><td>Samco</td><td>222-15</td><td></td></tr><tr><td>Horizontal shaker</td><td>VWR</td><td>444-2900</td><td></td></tr><tr><td>Hydrochloric acid (HCl)</td><td>Ajax Firechem.</td><td>265.2.5L-PL</td><td>10M stock solution, corrosive</td></tr><tr><td>Imaris software</td><td>Bitplane</td><td></td><td></td></tr><tr><td>Microscope slides Superfrost</td><td>ThermoFisher</td><td>10143352</td><td>ground edge, 90 degrees 26 mm~76 mm</td></tr><tr><td>Paraformaldehyde</td><td>Sigma Aldrich</td><td>P6148-500g</td><td>Hazardous</td></tr><tr><td>Phalloidin Alexa Fluor 647</td><td>ThermoFisher</td><td>A22287</td><td>dilution 1:100-200</td></tr><tr><td>E-cadherin</td><td>ThermoFisher</td><td>13-1900</td><td>dilution 1:400</td></tr><tr><td>Ki-67</td><td>BD Biosciences</td><td>B56</td><td>dilution 1:200</td></tr><tr><td>Keratin 5</td><td>BioLegend</td><td>905501</td><td>dilution 1:500</td></tr><tr><td>Keratin 8/18</td><td>DSHB (University of Iowa)</td><td></td><td>dilution 1:200</td></tr><tr><td>MRP2</td><td>Abcam</td><td>ab3373</td><td>dilution 1:50</td></tr><tr><td>Secondary Rat IgG (H+L) Alexa Fluor 488</td><td>ThermoFisher</td><td>A-21208</td><td>dilution 1:500</td></tr><tr><td>Secondary Mouse IgG (H+L) Alexa Fluor 555</td><td>ThermoFisher</td><td>A-31570</td><td>dilution 1:500</td></tr><tr><td>Secondary Rabbit IgG (H+L) Alexa Fluor 555</td><td>ThermoFisher</td><td>A-31572</td><td>dilution 1:500</td></tr><tr><td>Silicone Sealant</td><td>Griffon</td><td>S-200</td><td></td></tr><tr><td>Sodium Dodecyl Sulfate</td><td>ThermoFisher</td><td>28312</td><td></td></tr><tr><td>Sodium Hydroxide (NaOH) pellets</td><td>Merck Millipore</td><td>567530</td><td>10 M stock solution, corrosive</td></tr><tr><td>Suspension cell culture plates</td><td>Greiner Bio-One</td><td>662102</td><td>24-well</td></tr><tr><td>Tris</td><td>Fisher Scientific</td><td>11486631</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>T8532</td><td>Hazardous</td></tr><tr><td>Tween-20</td><td>Sigma Aldrich</td><td>P1379</td><td></td></tr><tr><td>UltraPure Low Melting Point (LMP) Agarose</td><td>ThermoFisher</td><td>16520050</td><td></td></tr><tr><td>Urea</td><td>Sigma Aldrich</td><td>51456</td><td></td></tr><tr><td>Workstation</td><td>Dell</td><td></td><td></td></tr></tbody></table>

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.

Imaging organoids in 3D enables visualization of architecture, cellular composition as well intracellular processes in great detail. The presented technique is undemanding and can presumably be applied to a wide range of organoid systems that are derived from various organs or host species.
The strength of 3D imaging compared to 2D imaging is illustrated by images of mouse mammary gland organoids that were generated using recently published methods14. The central layer of these organoids consists of columnar-shaped K8/K18-positive luminal cells and the outer layer contains elongated K5-postive basal cells (Figure 2A), which recapitulates the morphology of the mammary gland in vivo. This polarized organization is challenging to appreciate from a 2D optical section of that same organoid (Figure 2B, middle panel). Another example of a complex structure that is impossible to interpret without 3D information is the network of MRP2-positive canaliculi that facilitate the collection of the bile fluid of human liver organoids11 (Figure 2B). This exemplifies how our method allows visualization of essential structural features of organoids. Moreover, the obtained quality and resolution allows for semi-automated segmentation and image analysis. Thus, total cell numbers and presence of markers can be quantified in specific cellular subtypes in whole organoids. We illustrate this by segmenting the nuclei of an entire organoid containing 140 cells, of which 3 cells display high positivity for the Ki67 cell cycle marker (Figure 2C). The DAPI channel is selected as source channel, and segments are generated based on an intensity thresholding step and a sphere diameter of 10 &#181;m. Touching objects are split by region growing from seed points. Lastly, a size filter of 10 voxels is applied to remove small noise induced segments. For every segment representing a nucleus, the mean intensity of the Ki67 channel is then exported for plotting.
We recently developed the optical clearing agent FUnGI35, which we now integrated into this protocol to refine the transparency of the organoids. FUnGI is easy to use, as clearing is readily achieved by a single incubation step after immunofluorescent staining. An added advantage of the agent is its viscosity, which makes it easier for sample handling during slide mounting. Fluorescent samples in FUnGI preserve their fluorescence even when stored for multiple months at -20 &#176;C. We demonstrate that FUnGI outperforms uncleared and fructose-glycerol in fluorescent signal quality deep in the organoid (Figure 3A,B), and that FunGI-cleared organoids have overall enhanced fluorescence intensity compared to uncleared organoids (Figure 3C).
In summary, we describe an undemanding, reproducible 3D imaging technique for acquiring volumetric data of immunolabeled organoids. This protocol can be readily used to image a variety of organoids including those of both mouse and human origin, from healthy and disease models. The straightforward sample preparation can be adapted to facilitate confocal, multi-photon and light sheet fluorescent microscopes to obtain cellular to subcellular resolution of entire organoids.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60709/60709fig1.jpg" /> 
Figure 1: Schematic overview of the high-resolution 3D imaging protocol. Organoids are recovered from their 3D matrix. Fixation and blocking is performed prior to immunolabeling with antibodies and dyes. Optical clearing is achieved in a single step using the FUnGI clearing agent. 3D rendering of images can be performed by using imaging software. (A) Schematic overview of the procedure. (B) Cleared whole-mount 3D confocal image of a human colonic organoid immunolabeled for F-actin and E-cadherin (E-cad) (25x oil objective). Scale bar = 40 &#956;m. (C) Cleared whole-mount 3D confocal image. Scale bar = 20 &#956;m. (D) Enlarged optical section of a human colonic organoid immunolabeled for F-actin, E-cadherin (E-cad) and Ki67 (25x oil objective). Scale bar = 5 &#956;m. This figure has been modified from Dekkers et al.26. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60709/60709fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60709/60709fig2.jpg" /> 
Figure 2: Volumetric imaging visualizes complex 3D architecture. Confocal images representing whole-mount 3D datasets (left panel), 2D optical sections (middle panel) and 3D areas of an enlarged region (right panel). (A) An organoid derived from a single basal cell of the mouse mammary gland illustrating the 3D organization of elongated mammary basal cells that surround luminal cells or labeled for K8/18, K5 and F-actin (fructose-glycerol clearing; 25x oil objective). Scale bars represent 55 &#956;m (left panel) and 40 &#956;m (middle and right panels). (B) A human fetal liver organoid with a complex 3D network of MRP2-positive canaliculi, labeled for DAPI, MRP2 and F-actin (fructose-glycerol clearing; 40x oil objective). Scale bars represent 25 &#956;m (left panel) and 8 &#956;m (middle and right panels). (C) Confocal 3D whole-mount image of a human fetal liver organoid labeled with DAPI and Ki67 (left panel) and a segmented image on the DAPI channel using imaging software (middle panel). Scale bar = 15 &#956;m. Graph plotted representing the Ki67 mean intensity in all the cells (DAPI-segmented) of the entire organoid (140 cells) (right panel). This figure has been modified from Dekkers et al.26. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60709/60709fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60709/60709fig3.jpg" /> 
Figure 3: Optical clearing of organoids with FUnGI.&#160;(A)&#160;Representative images of human colonic organoids labeled with F-actin (green) and DAPI (grey) and imaged with no clearing, cleared with fructose-glycerol or cleared with FUnGI (25x oil objective). Left panel: 3D rendering of the organoid. Right panel: optical-section of the organoid at 150 &#956;m depth. For the &#8220;no clearing&#8221; condition the brightness of the image had to be increased in comparison to the &#8220;fructose-glycerol&#8221; and &#8220;FUnGI&#8221; conditions to visualize the organoid. Scale bar = 50 &#956;m. (B) Nonlinear regression fit showing the decrease of DAPI intensity with increasing Z-depth for different optical clearing methods. Values represent intensities of individual cells detected by DAPI segmentation and are normalized to the average DAPI intensity of the first 50 &#956;m of the organoid. To avoid underestimation of the attenuation caused by brighter cells on the deeper edges and budding structures, only the center regions of the organoids were analyzed. (C) Three organoids per condition of similar size and depth towards the coverslip were imaged using identical microscope settings. The full 3D datasets were single cell segmented on DAPI signal for comparison. Bar graph showing average DAPI intensity with different clearing methods on full segmented datasets. Data are depicted as mean &#177; SD. Values are intensities of &#62;3800 individual cells detected by DAPI segmentation. **** = p &#60; 0.0001, Kruskal-Wallis test with two-sided Dunn&#8217;s multiple comparison post-hoc testing. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60709/60709fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Single-Cell Resolution Three-Dimensional Imaging of Intact Organoids at JoVE.com

Single-Cell Resolution Three-Dimensional Imaging of Intact Organoids

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 ...

single-cell-resolution-three-dimensional-imaging-of-intact-organoids

Universidad San Sebastian

Research

JoVE Journal

Biology

15.9K Views.  Princess Máxima Center for Pediatric Oncology. 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.

Video: Single-Cell Resolution Three-Dimensional Imaging of Intact Organoids

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

Three-Dimensional Culture of Murine Colonic Crypts to Study Intestinal Stem Cell Function Ex Vivo

The intestinal epithelium regenerates every 5-7 days, and is controlled by the intestinal epithelial stem cell (IESC) population located at the bottom of the crypt region. IESCs include active stem cells, which self-renew and differentiate into various epithelial cell types, and quiescent stem cells, which serve as the reserve stem cells in the case of injury. Regeneration of the intestinal epithelium is controlled by the self-renewing and differentiating capabilities of these active IESCs. In addition, the balance of the crypt stem cell population and maintenance of the stem cell niche are essential for intestinal regeneration. Organoid culture is an important and attractive approach to studying proteins, signaling molecules, and environmental cues that regulate stem cell survival and functions. This model is less expensive, less time-consuming, and more manipulatable than animal models. Organoids also mimic the tissue microenvironment, providing in vivo relevance. The present protocol describes the isolation of colonic crypts, embedding these isolated crypt cells into a three-dimensional gel matrix system and culturing crypt cells to form colonic organoids capable of self-organization, proliferation, self-renewal, and differentiation. This model allows one to manipulate the environment-knocking out specific proteins such as claudin-7, activating/deactivating signaling pathways, etc.-to study how these effects influence the functioning of colonic stem cells. Specifically, the role of tight junction protein claudin-7 in colonic stem cell function was examined. Claudin-7 is vital for maintaining intestinal homeostasis and barrier function and integrity. Knockout of claudin-7 in mice induces an inflammatory bowel disease-like phenotype exhibiting intestinal inflammation, epithelial hyperplasia, weight loss, mucosal ulcerations, epithelial cell sloughing, and adenomas. Previously, it was reported that claudin-7 is required for intestinal epithelial stem cell functions in the small intestine. In this protocol, a colonic organoid culture system is established to study the role of claudin-7 in the large intestine.

Three-Dimensional Culture of Murine Colonic Crypts to Study Intestinal Stem Cell Function Ex Vivo

The present protocol describes establishing a murine colonic organoid system to study the activity and functioning of colonic stem cells in a claudin-7 knockout model.

The intestinal epithelium regenerates every 5-7 days, and is controlled by the intestinal epithelial stem cell (IESC) population located at the bottom of ...

Cancer Research

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 ...

Adapting Gastrointestinal Organoids for Pathogen Infection and Single Cell Sequencing under Biosafety Level 3 (BSL-3) Conditions

Human intestinal organoids constitute the best cellular model to study pathogen infections of the gastrointestinal tract. These organoids can be derived from all sections of the GI tract (gastric, jejunum, duodenum, ileum, colon, rectum) and, upon differentiation, contain most of the cell types that are naturally found in each individual section. For example, intestinal organoids contain nutrient-absorbing enterocytes, secretory cells (Goblet, Paneth, and enteroendocrine), stem cells, as well as all lineage-specific differentiation intermediates (e.g.,&#160;early or immature cell types). The greatest advantage in using gastrointestinal tract-derived organoids to study infectious diseases is the possibility of precisely identifying which cell type is targeted by the enteric pathogen and to address whether the different sections of the gastrointestinal tract and their specific cell types similarly respond to pathogen challenges. Over the past years, gastrointestinal models, as well as organoids from other tissues, have been employed to study viral tropism and the mechanisms of pathogenesis. However, utilizing all the advantages of using organoids when employing highly pathogenic viruses represents a technical challenge and requires strict biosafety considerations. Additionally, as organoids are often grown in three dimensions, the basolateral side of the cells is facing the outside of the organoid while their apical side is facing the inside (lumen) of the organoids. This organization poses a challenge for enteric pathogens as many enteric infections initiate from the apical/luminal side of the cells following ingestion. The following manuscript will provide a comprehensive protocol to prepare human intestinal organoids for infection with enteric pathogens by considering the infection side (apical vs. basolateral) to perform single-cell RNA sequencing to characterize cell-type-specific host/pathogen interactions. This method details the preparation of the organoids as well as the considerations needed to perform this work under biosafety level 3 (BSL-3) containment conditions.

Adapting Gastrointestinal Organoids for Pathogen Infection and Single Cell Sequencing under Biosafety Level 3 BSL-3 Conditions

This protocol describes how to infect human intestinal organoids from either their apical or basolateral side to characterize host/pathogen interactions at the single-cell level using single-cell RNA sequencing (scRNAseq) technology.

Human intestinal organoids constitute the best cellular model to study pathogen infections of the gastrointestinal tract. These organoids can be derived ...

Immunology and Infection

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

Mucociliary Epithelial Organoids from Xenopus Embryonic Cells: Generation, Culture and High-Resolution Live Imaging

Mucociliary epithelium provides the first line of defense by removing foreign particles through the action of mucus production and cilia-mediated clearance. Many clinically relevant defects in the mucociliary epithelium are inferred as they occur deep within the body. Here, we introduce a tractable 3D model for mucociliary epithelium generated from multipotent progenitors that were microsurgically isolated from Xenopus laevis embryos. The mucociliary epithelial organoids are covered with newly generated epithelium from deep ectoderm cells and later decorated with distinct patterned multiciliated cells, secretory cells, and mucus-producing goblet cells that are indistinguishable from the native epidermis within 24 h. The full sequences of dynamic cell transitions from mesenchymal to epithelial that emerge on the apical surface of organoids can be tracked by high-resolution live imaging. These in vitro cultured, self-organizing mucociliary epithelial organoids offer distinct advantages in studying the biology of mucociliary epithelium with high-efficiency in generation, defined culture conditions, control over number and size, and direct access for live imaging during the regeneration of the differentiated epithelium.

We describe a simple protocol to develop mucociliary epithelial organoids from deep ectoderm cells isolated from Xenopus laevis embryos. The multipotent progenitors regenerate epithelial goblet cell precursors and allow live tracking of the initiation and progression of the cell transitions on the surface of organoids.

Mucociliary epithelium provides the first line of defense by removing foreign particles through the action of mucus production and cilia-mediated ...

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

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 ...

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, ...

Single-Cell Resolution Three-Dimensional Imaging of Intact Organoids

Summary

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Chapters in this video

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

Three-Dimensional Culture of Murine Colonic Crypts to Study Intestinal Stem Cell Function Ex Vivo

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

Adapting Gastrointestinal Organoids for Pathogen Infection and Single Cell Sequencing under Biosafety Level 3 (BSL-3) Conditions

Culture and Imaging of Human Nasal Epithelial Organoids

Mucociliary Epithelial Organoids from Xenopus Embryonic Cells: Generation, Culture and High-Resolution Live Imaging

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

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

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

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