Triple-decker sandwich cultures of intestinal organoids for long-term live imaging, uniform perturbation and statistical sampling

Abstract

Three-dimensional organoid cultures enable the study of stem cell and tissue biology ex vivo providing improved access to cells for perturbation and live imaging. Typically, organoids are grown in hydrogel domes that are simple to prepare but lead to non-uniform tissue growth and viability. We recently developed a simple alternative culture method to embed intestinal organoids in multilayered hydrogels – “triple-decker sandwiches” – that align organoids in a common z-plane with uniform access to media. This culture configuration improves the growth and survival of organoids over a wide working area, and facilitates long-term confocal imaging and molecular perturbation. Here we present protocols for preparing organoids in triple-decker sandwich cultures and using them for live imaging, immunostaining, and single-cell RNA sequencing. We have tested our methods on mouse and human intestinal organoids and expect them to be useful for other highly proliferative three-dimensional cell cultures.

Basic Protocol 1: Pre-coating plates with PolyHEMA to prepare them for triple-decker sandwich culture

Support Protocol 1: Preparing PolyHEMA solution to coat glass-bottom dishes

Basic Protocol 2: Embedding intestinal organoids in triple-decker sandwiches

Alternate Protocol 2.1: Seeding single cells or organoids at low density in triple-decker sandwiches

Alternate Protocol 2.2: Embedding intestinal organoids in hydrogel domes

Support Protocol 2.1: Production of Wnt3a conditioned media

Support Protocol 2.2: Production of Rspo1 conditioned media

Basic Protocol 3: Live imaging of mouse intestinal organoids in triple-decker sandwich cultures

Alternate Protocol 3: Live imaging of vital-dye treated mouse intestinal organoids in triple-decker sandwich cultures

Basic Protocol 4: Immunofluorescence imaging of mouse organoids liberated from triple-decker sandwich cultures

Alternate Protocol 4.1: Liberating and fixing mouse intestinal organoids from dome cultures

Alternate Protocol 4.2: Measuring cell proliferation by EdU staining

Basic Protocol 5: Single-cell RNA sequencing and analysis of mouse intestinal organoids

INTRODUCTION:

Over the past decade organoid cultures have emerged as powerful tools for ex vivo studies of stem cell and tissue biology (Shamir and Ewald, 2014). Organoids facilitate long-term propagation of tissue stem cells in vitro and reproduce complex multicellular phenomena like patterning and morphogenesis. Compared to tissues in living animals, organoids are more accessible to microscopy – especially long-term time-lapse imaging – and to chemical, genetic, and biophysical perturbation (Murrow et al., 2017). As a result, organoids enable quantitative studies of tissue dynamics that would be difficult if not impractical to carry out in vivo (Paszek et al., 2005; Sato et al., 2011; Nguyen-Ngoc et al., 2012; Varner et al., 2014; Gracz et al., 2015; Farin et al., 2016; Langlands et al., 2016; Tallapragada, Cambra et al., 2021). At the same time, organoids can grow and behave more variably than tissues in vivo. It is therefore important to optimize organoid cultures for long-term imaging and scalability while minimizing heterogeneity in the dish.

Our work has required us to perturb mouse intestinal organoids and image them for long durations (Tallapragada, Cambra et al., 2021). To date, intestinal organoids have been cultured primarily in hemispherical hydrogel droplets – “domes” – that are easy to prepare but that lead to inhomogeneous access to media and variable distance from microscope objectives. One way to resolve these challenges is to grow organoids in planar hydrogel sandwiches that align them in space, but existing approaches to three-dimensional sandwich culture are incompatible with rapidly-growing organoids that may revert to two-dimensional monolayers upon contact with the cell culture dish (Tallapragada, Cambra et al., 2021).

Intestinal organoids proliferate rapidly in the absence of perturbation, and they expand even faster when carrying oncogenic mutations or when exposed to various small molecules and growth factors. We found that a “triple-decker” sandwich culture configuration makes these intestinal organoid cultures compatible with both high-quality imaging and long-term three-dimensional growth. A simple improvement over existing methods is the addition of a base layer of PolyHEMA – a hydrogel that organoids can neither penetrate nor digest – to prevent organoids from contacting the culture dish and reverting to cell monolayers.

Here we present protocols for establishing triple-decker sandwich cultures (Fig. 1) of intestinal organoids. For completeness, we also provide protocols describing their analysis by live and fixed imaging, and by single-cell RNA sequencing according to established practices. Basic Protocol 1 outlines how to coat glass-bottom plates and dishes with PolyHEMA in preparation for making triple-decker sandwich cultures, and Support Protocol 1 details how to make PolyHEMA solution required for coating. Basic Protocol 2 describes the process of seeding organoid cultures in “triple-decker sandwich” cultures for dissociated organoid fragments. Alternate Protocol 2.1 describes the same process for low density or single-cell seeding. For completeness and reproducibility, we describe our version of the standard “dome” culture protocol in Alternate Protocol 2.2. Support Protocols 2.1 and 2.2 are standard protocols for producing Wnt3a- and Rspo1-enriched conditioned media used to prepare complete organoid media (ENR and WENR; described in Reagents and Solutions). The remaining protocols primarily serve to consolidate relevant methods that have been described elsewhere and which we anticipate users of sandwich cultures will find useful. In several cases, modifications have been made to specifically adapt the methods to sandwich cultures. Basic Protocol 3 outlines steps for live imaging, as performed in Tallapragada, Cambra et al., 2021, focusing on endogenous fluorescent proteins. Alternate Protocol 3 describes how to use a vital dye to track dynamic processes in live organoids, with calcium staining as an example. Basic Protocol 4 details how to obtain high-quality antibody stains of organoids from triple-decker sandwich cultures; Alternate Protocol 4.1 provides the analogous steps for organoids in dome cultures. Both Basic Protocol 4 and Alternate Protocol 4.1 describe how to liberate organoids from Matrigel without disrupting organoid structure. Alternate Protocol 4.2 describes the protocol for obtaining high-quality images of fixed organoids in triple-decker sandwiches treated with dyes; we present the example of EdU staining, which is used to measure cell proliferation in organoids. Basic Protocol 5 describes how to prepare organoids for single cell RNA-sequencing, from dissociation to data processing after sequencing.

Figure 1. Schematic representation of triple-decker sandwich culture.

The three layers of the triple-decker sandwich culture are depicted as described in Basic Protocol 2; shown with the approximate dimensions of each layer for a glass-bottom 12-well plate with a #1.5 coverslip as the substrate for the PolyHEMA deposition and subsequent triple-decker sandwich culture.

STRATEGIC PLANNING

A flowchart for experimental planning is shown schematically in Fig. 2, starting with reagent preparation and ending with experiments on organoids in triple-decker sandwich cultures. It takes approximately 3 weeks to prepare reagents, materials, and cells to start culturing organoids in triple-decker sandwiches, for subsequent use in imaging, perturbation, and single-cell sequencing experiments (Fig. 2). This 3-week lead time includes generating conditioned media for culture media with and without Wnt3a (WENR and ENR media, respectively); preparing Poly(2-hydroxyethyl methacrylate) (PolyHEMA)-coated plates and establishing organoid cultures in ENR media from primary tissue or frozen stock.

Figure 2. Workflow for experiments using triple-decker sandwich cultures for long-term time-lapse imaging.

The media formulations used can be exchanged for the appropriate media for each type of organoid and their species of origin. In this example, the media used is formulated for the culture of mouse intestinal organoids. (A) Prepare batches of WENR media and ENR media using conditioned media from L-WRN and HEK-293-HA-Rspo1-Fc cells, respectively. In parallel, prepare PolyHEMA solution and coat glass-bottom plates. Plates can be stored for up to 3 weeks in 1x PBS at 37°C after coating. (B) Start organoid cultures, thawing and plating into dome cultures in WENR media. (C) Passage organoids in WENR media, and initiate ENR media dome cultures (D) Continue passaging the WENR organoids and initiating ENR cultures as needed. Use organoids cultured in ENR media for at least a week to initiate triple-decker sandwich cultures with PolyHEMA coated glass-bottom wells.

In this 3-week period, conditioned media should be prepared (requiring 1–1.5 weeks) before thawing organoids or generating them from primary tissue. For this, relevant cell lines must be obtained in advance (L-WRN and 293T-HA-RspoI-Fc cells, see Support Protocols 2.1 and 2.2). We advise producing enough conditioned media containing organoid growth factors in advance to use over the course of all planned experiments, to minimize experimental heterogeneity due to differences between batches of conditioned media. Alternatively, complete culture media (e.g., IntestiCult; Stem Cell Technologies) and recombinant growth factors (Wnt3a, Rspo1, and Noggin) are commercially available and can be used immediately at increased cost. The protocols for generating Wnt3a/Rspo1/Noggin conditioned media for WENR cultures, and for generating Rspo1 conditioned media for ENR culture media were reported in (Miyoshi and Stappenbeck, 2013) and (RandD Systems Protocol); for convenience, we reproduce them (with commentary) in Support Protocols 2.1 and 2.2.

Once media and PolyHEMA-coated plates are prepared, mouse intestinal organoid lines can be established as described (Sato and Clevers, 2013; Sato et al., 2009) and similarly, human intestinal organoids can be established as described in Sato et al., 2011. We recommend expanding organoid cultures soon after they are established from primary tissue and freezing organoid stocks in liquid nitrogen (−196°C) after 4 passages. One can maintain organoid cultures in Matrigel domes (Alternate Protocol 2.2), and establish all sandwich cultures from these organoids in domes. Over the period of an extended study, passaging of organoids in sandwich cultures may not be convenient, nor economical. Organoids can instead be maintained for passaging and expansion in dome cultures, using Wnt3a enriched media (WENR) that supports faster doubling times than in ENR media. If doing so, organoids should be transferred to ENR media for at least one week before being split and used for analysis (see flowchart, Fig. 2). Thus, users should plan for a lag time of two weeks between harvesting organoids from WENR media and analyzing them in sandwich cultures (one week of acclimation in dome culture in ENR media, followed by up to one week of growth in sandwich cultures).

Note: As is standard practice in tissue culture, mycoplasma tests (Lonza MycoAlert PLUS kit, cat. no. LT07–701) should be performed regularly, and all laboratory procedures related to organoid or cell culture passage and maintenance should be performed in a laminar flow hood to avoid contamination and to protect experimenters from biological risks.

BASIC PROTOCOL 1: PRE-COATING PLATES WITH POLYHEMA TO PREPARE THEM FOR TRIPLE-DECKER SANDWICH CULTURE

Triple-decker hydrogel sandwiches (Fig. 1) enable fast-growing organoids to expand in three dimensions close to a culture dish or coverslip. They prevent cells from spreading onto the culture dish by pre-coating the dish or coverslip with a non-adherent layer of PolyHEMA. In the case of mouse intestinal organoids, the base layer of PolyHEMA enables the growth of viable, multi-budded organoids with Lgr5+ stem cells located at the tips of the buds. This protocol describes how to coat tissue culture-grade glass-bottom dishes with PolyHEMA, the first of three hydrogel layers in a triple-decker sandwich. Support Protocol 1 describes how to prepare the PolyHEMA coating solution.

Materials:

• 70% (v/v) ethanol

• 1x tissue culture-grade PBS (Corning, cat. no. 21–040-CV)

• 0.02 g/mL tissue culture grade PolyHEMA (see recipe)

• 12-well glass-bottom plates (Mattek, P12G-1.5–14-F)

• Micropipet (P200 for 12 well plates)

• Pipet tips (1–200 μL for 12 well plates)

• Biosafety Class II tissue culture hood

• Vacuum desiccator (VWR; 24987–004)

• Parafilm (Millipore Sigma; P7793)

• 37°C incubator (aseptic) and 5% CO₂

Timing: 4–8 days

1. Add 187.5 μL of PolyHEMA solution to the center – i.e., atop the glass coverslip – of each well of a 12-well glass-bottomed tissue culture plate. Cover the plate with its lid.

   This volume is optimized for 12-well glass-bottom dishes, but the volume of plated PolyHEMA solution can be modified for other well sizes. As a rule of thumb, if X is the volume of cell culture media (in μL) normally added to a well, it takes X/8 μL PolyHEMA solution to coat that well.

2. Allow the plate to dry in a tissue culture hood for 2–4 days until all liquid has evaporated.

   This step may be shortened considerably depending on ambient temperature and humidity. Use a pipet tip to probe whether the coating is “sticky”; if it is, then it is still wet. This step reduces coating defects due to bubbles.

3. Repeat Steps 1–2; re-applying PolyHEMA ensures that the glass coverslip is coated evenly.

   After applying the second coat of PolyHEMA, wait to ensure that the PolyHEMA film is completely dry.

   PolyHEMA will seize up and detach from the coverslip if it comes in contact with an aqueous solution (e.g., PBS, media) before it is dry. (See Figure 3C)

4. Add 1.5 mL PBS to each coated well and place the plate in a tissue culture incubator at 37°C for at least 3 hours.

   This step is simultaneously a wash step (to remove precipitated PolyHEMA, debris, etc.) and a way to bring to the surface any bubbles still trapped in the PolyHEMA film. It is better that these bubbles are allowed to come to the surface prior to embedding organoids in the hydrogel sandwich rather than afterward. A coated plate with PBS-filled wells can sit in a 37°C incubator for as little as 3 hours or as long as one month awaiting further use (with periodic replacement of 1x PBS). When using a PolyHEMA-coated plate that is >1 week old, confirm using a tissue culture microscope that the PolyHEMA layer is still intact and free of major defects.

Figure 3. Examples of coating dishes with PolyHEMA for imaging.

(A) A usable single well of a glass-bottom 12-well plate coated with PolyHEMA as in Basic Protocol 1. (B) A poor-quality single well of a glass-bottom 12-well plate that may be prone to delamination and detachment from the coverslip. Such wells may be usable as assessed after quality control testing by incubation with 1x PBS as instructed in Basic Protocol 1. (C) A defective single well of a glass-bottom 12-well plate coated with PolyHEMA. (D) A tiled micrograph of a usable single well of a glass-bottom 12 well-plate after triple-decker sandwich culture plating. (E) A tiled image of a defective single well of a glass-bottom 12-well plate coated with PolyHEMA showing signs of excessive delamination. (F) A tiled image of a defective single well of a glass-bottom 12-well plate coated with PolyHEMA showing signs of excessive bubbling. Scale bars = 1 mm. Panels D-F acquired at 10X magnification.

SUPPORT PROTOCOL 1: PREPARING POLYHEMA SOLUTION TO COAT GLASS-BOTTOM DISHES

This protocol describes how to make the PolyHEMA solution used to coat glass-bottom dishes in Basic Protocol 1. PolyHEMA solution should be kept under tight seal at room temperature when not in use, and after it is sterile filtered it should only be opened in a Biosafety Class II tissue culture hood using aseptic technique. We suggest using a hot plate with a feedback system or probe to monitor the actual temperature of this solution as it is being mixed. Otherwise, take great care to monitor and adjust the hot plate temperature so that the PolyHEMA solution reaches 65°C for 3 hours while stirring, or until all PolyHEMA crystals are dissolved. Undissolved PolyHEMA interferes with organoid imaging. Once this reagent is made, it can be stored for up to 3 months at room temperature. We caution against leaving coated plates with 1x PBS in the incubator for more than 3 weeks. The coatings may degrade by this time. Always visually check coating integrity before using it for culture.

Materials:

• Poly(2-hydroxyethyl methacrylate) (PolyHEMA) (2g) (Sigma, P3932)

• 95% (v/v) ethanol

• Vacuum desiccator (VWR, cat. no. 24987–004)

• 0.22 μm cellulose acetate membrane with bottle top vacuum filter system (Corning, cat. no. 430756)

• Hot plate and magnetic stirrer

• Magnetic stir bar

• 200 mL beaker

• Parafilm (Millipore Sigma, cat. no. P7793)

• Glass thermometer (Thomas Scientific, cat. no. 9313A27)

• Aluminum foil

• Biosafety Class II tissue culture hood

Timing: 4 hours

1. Weigh and add 2g PolyHEMA to 100 mL 95% ethanol in a glass beaker.

2. Add a magnetic stir bar to the beaker and place the beaker on a hot plate with a magnetic stirrer and stir at 65°C until fully dissolved (typically ≥3 hours). Cover first with aluminum foil, followed by generous amounts of Parafilm to prevent evaporation.

   Poke a glass thermometer through the top of the aluminum foil and Parafilm to ensure that the temperature of the PolyHEMA solution is as close to 65°C as possible.

3. Allow the PolyHEMA solution to cool to room temperature.

4. In a tissue culture hood, filter the PolyHEMA solution through a 0.22 μm filter using the vacuum filter system.

5. In a tissue culture hood, place the bottle of filtered PolyHEMA solution (with cap off) in a vacuum desiccator for 30 minutes to degas the solution.

6. In a tissue culture hood, after the solution is degassed, replace the screw cap and use Parafilm to seal the cap-bottle junction on the outside of the bottle, as an extra measure to prevent unwanted gas exchange.

BASIC PROTOCOL 2: EMBEDDING INTESTINAL ORGANOIDS IN TRIPLE-DECKER SANDWICHES

This protocol describes how to embed mouse and human organoids in triple-decker sandwiches (Fig. 1) after tissue culture plates have been coated with PolyHEMA following the steps in Basic Protocol 1. Organoids are sandwiched between two thin layers of Matrigel that are deposited on top of the base layer of PolyHEMA. By effectively confining organoids to a single, common z-plane, this configuration enables both long-term and high-throughput organoid imaging. It also improves organoid viability, and the reproducibility of responses to molecular perturbations, by homogenizing access to diffusible molecules in media (Tallapragada, Cambra et al., 2021). This culture system is still compatible with functional studies of luminal swelling, such as forskolin treatment (Tallapragada, Cambra, et al., 2021).

In this protocol we discuss how to re-plate organoids as crypt-sized multicellular fragments rather than single cells. When single cells and dilute multicellular fragments are seeded, the topcoat and basecoat Matrigel layers tend to separate, and so the topcoat must be applied differently. In Alternate Protocol 2.1 we address how to deposit single cells, or organoid fragments at very low concentrations. We did not observe that either organoid growth or viability correlated with organoid density, provided that organoids were not over-fragmented during passaging. Lower density facilitates imaging because it reduces the likelihood of organoids growing into each other. This protocol has also been used to culture human intestinal organoids alongside dome cultures of human intestinal organoids (see Fig. 5). Human intestinal organoids are similarly less variable in their distance from the coverslip (z-distances are 3-fold narrower in their average range; 1480 μm for dome and 491 μm for sandwich cultures, n=2) when cultured in sandwich culture as compared to culturing in dome culture (Fig. 5E).

Figure 5. Comparison of Human intestinal organoids grown in sandwich culture and dome culture.

Human organoids were generated from biopsy samples collected from a healthy, de-identified pediatric donor undergoing endoscopy for gastrointestinal complaints following Informed Consent from the donor’s guardian, as previously described (PMID: 29440725), and with approval from the Institutional Review Board of Boston Children’s Hospital (Protocol number IRB-P00000529). Organoids shown are on passage 19 and grown in Human duodenum growth media (see Reagents and Solutions). (A) Organoids grown in sandwich culture for 3 days post-plating (dpp). (B) Same well of organoids as in (A), 8 dpp. (C) Organoids grown in dome culture, 3 dpp. (D) Same well of dome cultured organoids as in (C) 8dpp. (E) Median centered distance of organoids in the Z-plane (from objective) in each culture condition, for 2 wells per culture condition. Scale bar = 1 mm.

Materials:

• Matrigel, growth factor reduced, phenol red-free (Corning, cat. no. 356231)

  Thaw aliquots of Matrigel overnight in the 4°C.

  Aliquots can be kept at 4°C for about a week, and should not go through multiple freeze-thaw cycles.

• DMEM/F12 (1:1) with L-glutamine and phenol red (ThermoFisher, cat. no. 11320033)

• Cell Recovery Solution (Corning, cat. no. 354253)

• Complete crypt culture media containing EGF, Noggin, and R-Spondin-1 (ENR) media (see recipe) OR media of choice for culture (e.g., human duodenum growth media as in Sato et al., 2011; see Reagents and Solutions for recipe)

• 70 μm cell strainer (pluriSelect, cat. no. 43–50070-51)

• 1000 μL pipet tips

• Glass aspirator tips

• 5, 10 mL serological pipets

• 15, 50 mL Falcon tubes

• PolyHEMA coated glass-bottom 12 well plates (see Basic Protocol 1)

  Plates should be equilibrated in the 37°C (aseptic) incubator prior to use, aiding in the solidification of the Matrigel.

• Swinging bucket rotor centrifuge (set to 4°C)

• 37°C incubator (aseptic) and 5% CO₂

Timing: 3–4 hours

• 1Before beginning, calculate the amount by which to dilute cells to achieve a desired split ratio.

  Whether they are dissociated into multicellular fragments or into single cells, organoids must be resuspended in media before deposition in a triple-decker sandwich. The split ratio is the ratio between the initial volume of organoids during dissociation and the final volume of organoids in media during deposition. The following example illustrates this concept, starting with organoids in a 24-well plate and ending with organoids in triple-decker sandwiches in a 12-well plate:

  Initial: 3 wells x 500 μL/well = 1.5 mL organoids in Cell Recovery Solution

  Final: 5 wells x 1.25 mL organoids in media/well = 6.25 mL organoids in media

  The split ratio in this scenario is 1.5 mL : 6.25 mL, or ≈ 1:4.

Dissociate organoids and deposit Matrigel basecoat

• 2Start dissociating organoids by incubating them in Cell Recovery Solution in a 50 mL Falcon tube on ice for 45–60 minutes.
• 3In parallel, in a 15 or 50 mL Falcon tube on ice (depending on the number of wells being prepared), prepare a solution of 5% (v/v) Matrigel in DMEM/F12.

  As an example, coating 1 well of a 12-well plate requires 1–1.5 mL of this 5% Matrigel solution. Invert the tube 10–15 times and swirl the contents gently to ensure that Matrigel is well mixed in DMEM.

• 4Add 5% Matrigel solution to each PolyHEMA-coated well. Incubate the plate at 37°C for 30–60 minutes.

  If you plan to image organoids, do not incubate the plate for longer than 1 hour at this step. The deposited layer of Matrigel will become too thick, and organoids will end up too far from the coverslip for high-resolution imaging with low laser exposure.

• 5In the last 10–15 minutes of incubation, pre-warm ENR media in anticipation of depositing dissociated cell clusters.

Seed organoids on Matrigel basecoat

• 2Once dissociation is complete, pellet cells by centrifugation at 250g and 4°C for 5 minutes.
• 3Aspirate supernatant.
• 4Re-suspend pellet in 1–2 mL of ENR media (or growth media of choice), pipetting up and down vigorously 40–50 times.
• 5Add the remaining volume of media required to achieve the desired split ratio (See Step 1 above for guidance) and gently mix the contents of the tube (e.g., by pipetting up and down 5 times with a serological pipet and inverting the tube 5 times)

  Using complete media (containing all growth factors and supplements necessary for long-term culture) minimizes the stress on cells during deposition. To reduce cost, it may be possible to substitute DMEM/F12 for complete media in Steps 7 and 8.

• 6Filter the solution into a new Falcon tube through a cell strainer with a 70 μm filter.
• 7Add 1.25 mL of cells in media to each well being prepared. To ensure that fragments of roughly consistent size are deposited in each well, draw from the middle of the liquid in the Falcon tube when pipetting, not the bottom or the air-media interface.

  In order to ensure that each well gets a consistent density of material, we pipet 612.5 μL into every well before topping up with the remaining 612.5 μL, and we deposit the solution in reverse order when topping up. For example, when adding cells to every well of a 12-well plate, moving from the upper left to the lower right, then on the second pass move from the lower right to the upper left.

• 8Shake plate briefly left and right (in “x”) and back and forth (in “y”), but not up and down (in “z”), to ensure that organoid fragments are evenly dispersed in each well.

  Do not swirl the plate, as this will cause organoid fragments to accumulate in the center of well rather than spreading evenly across the coverslip. Use a standard tissue culture microscope to confirm that fragments have settled and have spread out across the well.

Allow organoids to adhere to Matrigel basecoat, deposit Matrigel topcoat, and add media

• 2Transfer the plate to a tissue culture incubator at 37°C and incubate for 30 minutes.
• 3Remove the plate from the incubator and aspirate the media in each well.

  It is critical to completely remove media at this step. If a layer of media remains in the well, the topcoat of Matrigel may not adhere properly to the cells and basecoat of Matrigel below. In order to remove all media in a well, use a standard aspirator in a tissue culture hood, as follows. First, tilt the plate towards you and remove media by placing the aspirator tip along the wall of the well. Then place the aspirator tip close to the edge of the glass coverslip in the middle of the well to remove residual media. Be careful not to aspirate cells, Matrigel, or PolyHEMA while doing this.

• 4Add 70 μL ice-cold Matrigel to the center of each well (above the glass coverslip and PolyHEMA).

  For maximally even coverage, we recommend depositing this volume in three parts (i.e., by partially emptying the pipet tip) at different positions across the coverslip to ensure an even coating.

• 5Incubate the plate at 37°C for 45–60 minutes to ensure that the Matrigel sets completely.
• 6Add 1.5 mL warm media to each well and return the plate to the incubator. Triple-decker sandwich preparation is complete.

ALTERNATE PROTOCOL 2.1: SEEDING SINGLE CELLS OR ORGANOIDS AT LOW DENSITY IN TRIPLE-DECKER SANDWICHES

This protocol has the same endpoint as Basic Protocol 2, with modifications to allow single cells or organoid fragments at low density to be seeded in triple-decker sandwich cultures. In a triple-decker sandwich, organoids support the attachment of the Matrigel topcoat to the basecoat. If plated organoid fragments are not dense enough, or if single cells are deposited, the Matrigel topcoat is less likely to strongly attach to the basecoat. To circumvent this issue, low-density fragments or single cells can be mixed and deposited together with the Matrigel topcoat.

Additional Materials:

• Instead of a 70 μm cell strainer, use a 40 μm cell strainer (pluriSelect, cat. no. 43–10040-60)

Timing: ≥ 18 hours

• 1Before beginning, calculate how much to dilute cells to achieve a desired split ratio.

  Whether they are dissociated into multicellular fragments or into single cells, organoids must be resuspended in media before deposition in a hydrogel triple-decker sandwich. The split ratio is the ratio between the initial volume of organoids during dissociation and the final volume of organoids in media during deposition. The following example illustrates this concept, starting with organoids in a 24-well plate and ending with organoids in triple-decker sandwiches in a 12-well plate:

  Initial: 3 wells x 500 μL/well = 1.5 mL organoids in Cell Recovery Solution

  Final: 5 wells x (1.25 mL organoids in media+10% Matrigel)/well = 6.25 mL organoids in media+10% Matrigel

  The split ratio in this scenario is 1.5 mL : 6.25 mL ≈ 1:4.

Dissociate organoids and deposit Matrigel basecoat

• 2Start dissociating organoids by incubating them in Cell Recovery Solution in a 50 mL Falcon tube on ice for 45–60 minutes.
• 3In parallel, in a 15 or 50 mL Falcon tube on ice (depending on the number of wells being prepared), prepare a solution of 5% (v/v) Matrigel in DMEM/F12.

  As an example, coating 1 well of a 12-well plate requires 1–1.5 mL of this 5% Matrigel solution. You must invert the tube 10–15 times and swirl the contents gently to ensure that Matrigel is well mixed in DMEM.

• 4Add 5% Matrigel solution to each PolyHEMA-coated well. Incubate the plate at 37°C for 30–60 minutes.

  If you plan to image organoids, do not incubate the plate for longer than 1 hour at this step. The deposited layer of Matrigel will become too thick, and organoids will end up too far from the coverslip for high-resolution imaging with low laser exposure.

• 5Once dissociation is complete, pellet cells by centrifugation at 250g and 4°C for 5 minutes.

Deposit organoid-Matrigel-media mixture on Matrigel basecoat

• 2Aspirate supernatant.
• 3Resuspend pellet in 1 mL of ENR media, pipetting up and down vigorously 40–50 times.
• 4Add the remaining amount of media required to achieve the desired split ratio (See Step 1 above for guidance) and gently mix the contents of the tube (e.g., by pipetting up and down 5 times with a serological pipet and inverting the tube 5 times)

  Using complete media (containing all growth factors and supplements necessary for long-term culture) minimizes the stress on cells during deposition. To reduce cost, it may be possible to substitute DMEM/F12 for complete media in Steps 8 and 9. This media should be cold, as Matrigel is being diluted into it along with the cells being deposited.

• 5Filter the solution into a new Falcon tube through a cell strainer with a 40 μm filter.
• 6Add Matrigel to the cell solution at a final concentration of 10% (v/v) Matrigel and mix well immediately after straining. (Revisit your calculations from Step 1 for guidance)
• 7Add 1.25 mL of cold cell solution (Matrigel + cells + ENR) to each well being prepared. Be sure to draw from the middle of the liquid in the Falcon tube when pipetting, rather than the bottom or the air-media interface.

  In order to ensure that each well gets a consistent density of material, we pipet 612.5 μL into every well before topping up with the remaining 612.5 μL, and we deposit solution in reverse order when topping up. For example, when adding cells to every well of a 12-well plate, moving from the upper left to the lower right, then on the second pass move from the lower right to the upper left.

• 8Shake plate briefly left and right (in the “x” plane) and back and forth (in the “y” plane), but not up and down (in the “z” plane), to ensure that organoid fragments or single cells are evenly dispersed in each well. Use a standard tissue culture microscope to confirm that cells are settling and have spread out.

  Do not swirl the plate, or else cells will accumulate in the center of well, rather than evenly across the coverslip.

Incubate and feed organoids to complete deposition in a triple-decker sandwich

• 2Transfer the plate to a tissue culture incubator at 37°C.
• 3Leave the plate in the incubator for at least 12 hours.
• 4After prolonged incubation, remove the plate from the incubator and aspirate the media in each well.
• 5Add 1.5 mL warm ENR media to each well and return the plate to the incubator; sample preparation is complete.

ALTERNATE PROTOCOL 2.2: EMBEDDING INTESTINAL ORGANOIDS IN HYDROGEL DOMES

This protocol describes how to culture intestinal organoids in Matrigel “domes”, reproducing the protocol from Sato and Clevers, 2013 with minor modifications. Use this protocol to maintain organoid cultures from week to week after thawing (e.g., in WENR media), and to prepare wells of organoids (e.g., ENR to induce differentiation) with material to seed in triple-decker sandwich cultures. Note, if adapting this protocol to human intestinal organoids (e.g., Figure 5), organoids can be similarly grown in a pro-growth media (e.g., Human duodenum growth media), and then transferred to differentiation media at least one week before use in experiments (see Figure 2).

Additional Materials:

• Complete crypt culture media containing Wnt3a, EGF, Noggin, and R-Spondin-1 (WENR) media (see recipe)

• Instead of a 12-well glass-bottom plate coated with PolyHEMA, use a 24-well plastic tissue culture plate, or an uncoated 12-well glass-bottom plate; similarly, place plates in the 37°C (sterile) incubator to equilibrate them to the temperature before use.

Timing: Approximately 1 hour

1. Check organoids using a standard tissue culture microscope to confirm size and confluence and place tissue culture plates to be used in the incubator to equilibrate them to 37°C. Equilibrating the plates helps Matrigel set upon contact with the bottom of the well, preventing the droplet from spreading.

2. Aspirate old media.

3. Add 500 μL ice cold Cell Recovery Solution to each well.

4. Scrape each well with a P1000 tip several times to detach Matrigel domes from the culture dish.

   With the tissue culture plate tilted towards you, collect and dispense the contents of each well using a pipet, using the liquid to wash the well from top to bottom. This process facilitates the mechanical dissociation of organoids and increases the recovery of organoids attached to the cell culture dish. Tilt the entire plate by 45° to aspirate organoid solution into pipet tip from the bottom of the well and release at the top of the well to make sure the Matrigel is completely removed from the plate.

5. Pipet solution into a 50 mL Falcon Tube.

6. Incubate the Falcon Tube on ice for 30–40 minutes.

   Do not keep on ice for longer than 1 hour. Extended exposure to Cell Recovery Solution reduces organoid viability.

7. Spin at 300g at 4°C for 5 minutes.

   If a pellet does not obviously form, remove some supernatant, incubate on ice for 5–10 minutes, and re-spin.

8. Keep the Falcon Tube on ice and aspirate the supernatant.

9. With the Falcon Tube still on ice, add enough Matrigel to replate a desired number of wells – generally, 50 μL per well.

   Organoids should generally be plated at a 1:3 ratio, adjusted up or down depending on organoid size and confluence. For example, starting with 2 wells’ worth of organoids, add enough Matrigel to replate 6 wells, with 10% extra volume as buffer in anticipation of bubble formation when pipetting viscous Matrigel (see Step 10) – 1.1 x (6 × 50 μL) = 330 μL, in this case

10. Pipet the Matrigel and organoids vigorously for approximately 1 minute or until you can see the organoids breaking up. Keep the Falcon tube on ice during this step. Take great care not to introduce bubbles.

    1. Count to one minute while breaking up organoids via trituration in Matrigel.

    2. If you are working with a large volume of Matrigel (e.g., 750 μL), bending the pipet tip to make a smaller, more elliptical inlet/outlet will help break apart the organoids better; this technique is also routinely used especially for passaging human intestinal organoid, regardless of volumes.

    3. If you are working with a small volume of Matrigel, do not bend the pipet tip. It increases the likelihood that you will introduce bubbles into the Matrigel, which is worse for organoid growth than incomplete fragmentation into smaller clusters.

11. Add a 50 μL drop of the Matrigel-organoid mixture to the center of a well in a 24 well plate.

    Depending on the protein concentration, to prevent the domes from collapsing after deposition, you can keep the tube out of the ice for this step. Take the Matrigel off ice while you label the plate to let the Matrigel and organoid mixture warm up slightly. However, do not spend more than 5 minutes plating (off ice). After you finish adding the Matrigel to the plate, you can incubate the plate upside down so that the deposited Matrigel forms better domes.

    See Troubleshooting Guide for optimization.

12. Incubate the plate with the newly plated dome cultures of mouse intestinal organoids in a 37°C incubator for 10–15 minutes.

13. Warm WENR and/or ENR media while Matrigel with embedded organoids sets in the 37°C incubator.

14. For a 24-well plate, add 500 μL pre-warmed media per well with organoids, then return plate to incubator.

    Adjust this volume per well as necessary for different cell culture plates.

SUPPORT PROTOCOL 2.1: PRODUCTION OF WNT3A CONDITIONED MEDIA

This protocol details how to produce Wnt3a conditioned media used to prepare WENR media as described in Miyoshi and Stappenbeck, 2013. L-WRN conditioned media prepared according to the protocol reproduced here has been shown to be replicable across batches and labs (VanDussen et al., 2019).

Materials:

• L-WRN cells (ATCC, CRL-3276)

  These cells are now commercially available; we obtained cells from Dr. Stappenbeck at Washington University (Miyoshi and Stappenbeck, 2013)

• Primary culture media (see recipe)

• G418 (50mg/mL) (Sigma, cat. no. D6429)

• Hygromycin (100mg/mL) (InvivoGen, cat. no. ant-hg-1)

• L-Cell media (see recipe)

• 0.25% Trypsin-EDTA (see recipe)

• PBS-EDTA (0.5 mM) (see recipe)

• 150 cm² culture flasks (Corning, cat. no. CLS430825)

Timing: Approximately 10 days

1. Thaw L-WRN Cells and incubate for 1 day at 37°C.

2. Change media and add G418 (500μg/mL) and Hygromycin (500μg/mL)

3. Wait until the 150 cm² flask is confluent (2–3 days).

4. Wash cells with 20 mL PBS-EDTA and aspirate.

5. Add 1 mL Trypsin-EDTA and tap to coat the flask surface.

6. Incubate the flask for 3–5 minutes at 37°C.

7. Suspend the cells in 12 mL L-cell media.

8. Add 120 mL L-cell media into the flask and aliquot 25 mL into five (5) 150 cm² culture flasks.

9. Incubate the flasks in a cell culture incubator (without G418 and Hygromycin, to prevent carryover of drugs in the conditioned media) until cells become over-confluent and many cell aggregates detach from the plate (3–4 days).

10. Wash cells with 10 mL Primary Culture Media per flask and aspirate.

11. Add 25 mL (per flask) primary culture media.

12. Incubate the flasks in the cell culture incubator for 24 hours.

13. Recover the conditioned media into 50 mL conical tubes and add new primary culture media to the flasks. Return flasks to the incubator.

14. Centrifuge the tubes at 2,000g for 5 minutes and decant the supernatant carefully in a 1 L bottle. Store the bottle at 4°C (1^st conditioned media).

15. Every 24 hours for the next 3 days (2^nd, 3^rd and 4^th collection), collect conditioned media in the same bottle, following Steps 13 and 14.

16. After the 4^th collection, sterile-filter, aliquot, and store conditioned media at −20°C.

SUPPORT PROTOCOL 2.2: PRODUCTION OF RSPO1 CONDITIONED MEDIA

This protocol details the steps to produce Rspo1 conditioned media used to prepare ENR media as described in Sato et al., 2011. For quality control assays see the RandD Systems R-spondin1 conditioned media Protocol, which provides protocols for both a Western blot assay and a TOPFlash assay to assess R-spondin concentration. Assays for quality control are described therein. We have observed differences in organoid growth, inflationary dynamics, and Lgr5+ stem cell zone fission rates across experiments, but these have correlated with differences in thawed cell aliquots and passage numbers. Standard experimental design practices should control for these sources of experimental variation by matching control and perturbation experiments.

Additional Materials:

• 293T-HA-RspoI-Fc Cells (Trevigen, cat. no. 3710–001-01)

  We used an R-spondin1-secreting cell line generously provided by the Kuo Lab at Stanford University (Li et al., 2014)

• Growing media (see recipe)

• Conditioned Media AD-DF +++ (see recipe)

• PBS-EDTA (0.5 mM) (see recipe)

• 0.25% Trypsin-EDTA (see recipe)

• Zeocin (100 mg/mL) (Life Technologies, cat. no. R25001)

• 150 cm² culture flasks (Corning, cat. no. CLS430825)

Timing: Approximately 8 days

1. Thaw 293T-HA-RspoI-Fc cells and incubate at 37°C for 1 day.

2. Change media and add 3 μl per mL of Zeocin so that it is 300 μg/mL. Wait until cells are confluent (2–3 days).

3. Aspirate old media.

4. Wash cells with an appropriate amount of warm 0.5 mM PBS-EDTA and aspirate.

5. Add an appropriate amount of warm 0.25% Trypsin-EDTA. Make sure the dish/flask is evenly coated. (2 mL for 100 mm, 3 mL for 75 cm², 6 mL for 150 cm²).

6. Incubate the dish/flask for 3–5 minutes at 37°C.

7. Suspend the cells in 12 mL warm Growing Media. Gently pipet cell solution up and down to release cells from the bottom of the flask.

8. Add 140 mL warm Growing Media into flask and aliquot 25 mL into six 150 cm² culture flasks.

9. Add 75 μL Zeocin (300 μg/ml) to 1 flask. This flask will be used to continue the cell line, freeze cells, or make more conditioned media.

10. Leave 5 flasks with no Zeocin.

11. After the flasks growing without Zeocin are confluent (3–4 days). Aspirate old media and replace with 50 mL AD-DF+++ media per flask.

12. Incubate for 7 days.

13. Remove media from flasks and spin down for 5 minutes at 1500 rpm to remove floating cells.

14. Sterile filter through a 0.22 μM filter and store aliquots in −20°C.

15. Cells can be used to harvest conditioned medium for 10–12 passages.

BASIC PROTOCOL 3: LIVE IMAGING OF MOUSE INTESTINAL ORGANOIDS IN TRIPLE-DECKER SANDWICH CULTURES

This protocol describes how to set-up a long-term time-lapse imaging experiment using mouse intestinal organoids embedded in triple-decker sandwich cultures. We used a point-scanning confocal microscope (see Table 1) and objectives from Nikon, but similar equipment from any manufacturer should work. The basic requirements are as follows: (1) a confocal microscope (point-scanning or spinning disk); (2) imaging objectives, especially an index-matched water-immersion lens, if available; and (3) an incubator (stage-top or live box) compatible with your microscope. Time-lapse imaging of multiple organoids requires a motorized z-stage.

Table 1.

Relevant parameters and tradeoffs for long-term time-lapse imaging with a confocal microscope.

Option	Recommended choice or value	Trade-off
Microscope	Point-scanning confocal	Image quality vs. imaging speed and phototoxicity
Culture configuration	Hydrogel sandwich	Resolution and throughput vs. preparation time
Objective	10x dry (whole well imaging) 20x dry (imaging multiple organoids in multiple wells) 40x water immersion (high-resolution imaging)	Throughput vs. spatial resolution
Laser power	High enough that pixels are barely saturated but with < 10 mW cumulative exposure per pixel per z-stack (“< 2%” cumulative laser power across all channels on a typical point-scanning confocal)	SNR vs. tissue health
Number of z-slices imaged (inversely related to z-step)	≤ 40 z-slices (organoid mid-plane ± 20 z-slices)	Depth of imaging vs. tissue health
Parameters specific to a point-scanning confocal microscope
Scan area	High resolution: 1024 px x 1024 px (0.31 μm/px) High throughput: 512 px x 512 px (+ scan area zoom, if desired)	Resolution vs. tissue health
Photomultiplier tube (PMT) gain	High enough that pixels are barely saturated	Detectability vs. noise
Pixel dwell time	~2 μs per pixel	SNR vs. tissue health
Pinhole size	1.2–1.8 Airy units (AU)	Signal vs. confocality
Averaging	None (live), 2x-8x (fixed)	SNR vs. tissue health and throughput

Note: In most labs the confocal microscope is in a non-sterile environment. Plates placed on the microscope ideally should not be brought back into a lab’s primary tissue culture room, or at least should not be placed in the same incubators and hoods as cell cultures that have not left the tissue culture room. In our tissue culture room, we use a designated hood to feed organoids that have been taken to the microscope room (e.g., during multi-day time-lapse imaging), and an alternate incubator to store plates after imaging (e.g., after time-lapse imaging, but before fixation and immunostaining).

Materials:

• Mouse intestinal organoids cultured in triple-decker sandwiches (see Basic Protocol 2)

• Immersol W 2010 (Zeiss, cat. no. 444969–0000-000)

• Complete crypt culture media containing EGF, Noggin, and R-Spondin-1 (ENR) media (see recipe) (or WENR)

• Nikon inverted A1R point-scanning confocal microscope with the following features:

  1. 10X dry objective
  2. 20X dry objective (NA 0.75)
  3. 40X water immersion objective (NA 1.15)
  4. Motorized z-stage
  5. Stage-top incubator allowing CO₂ concentration and heat control (Tokai Hit)
  6. Solid-state lasers in:
     • Blue (405 nm)
     • Green (488 nm)
     • Red (555 nm)
     • Far-red (640 nm)

• Imaris 8.4 (Oxford Instruments)

  Any image analysis software capable of handling 3+ dimensional images can be used (e.g., Fiji).

1. Plate organoids in only the 2 central wells of a 12-well plate to minimize variation in temperature and humidity between wells during imaging. Add 1x PBS to the empty wells surrounding these central wells to minimize evaporation.

   See (Shamir and Ewald, 2014) for a more in-depth discussion of temperature and humidity considerations during live imaging.

2. Turn on the confocal microscope, and allow the stage-top incubator to equilibrate at 37°C and 5% CO₂ before continuing with setup.

3. Prepare a suitable objective for use. For high-resolution imaging with a 40X water immersion objective, follow Steps 3–6. Otherwise, proceed to Step 7.

4. For high resolution imaging we recommend using a 40X (NA 1.15) water immersion objective and Immersol W as the immersion fluid.

5. Apply 2–3 drops of Immersol W to the objective and ensure that it spreads evenly over the objective. Use the Immersol applicator to dislodge any bubbles visible in the immersion fluid.

6. After applying immersion fluid, place your sample in a glass-bottom dish on the stage top and slowly bring it into focus. Looking through the microscope eyepiece, you will notice the image sharpen suddenly when immersion fluid is in contact with both the objective and the glass coverslip.

7. Find a z-plane with a reference object — e.g., a Paneth cell, or a speck of debris — and focus on it. Record this initial z-position. Wait 5 min and refocus on that reference object. Record the z-position and note whether it has changed. Repeat this process every 5 min until the z-position of the reference object no longer changes. Proceed to Step 8.

   This procedure allows immersion fluid to equilibrate before time-lapse imaging. Significant z-drift is common when samples are first placed on the microscope, as the drop of immersion fluid spreads out between the coverslip and the objective.

8. For imaging multiple organoids across multiple wells, we recommend using the 10X magnification dry objective (e.g., to capture whole-well images and tiled movies of large fields of view), or a 20X dry objective (NA 0.75) (e.g., to capture images of tens of organoids at intermediate resolution for segmenting stem cell zone sizes).

   You can use a low-magnification (e.g., 10X dry) objective to take tiled images of large fields of view (> 1mm x 1mm) and record the position of every organoid in a whole well or large field of view. Pre-recording the positions of organoids facilitates selecting a random subset for imaging after perturbation.

   Tiled images can be captured using the Multipoints feature in NIS Elements; to cover an entire triple-decker sandwich in a 12-well glass-bottom plate, set the microscope to scan an 14.5 mm x 14.5 mm square centered at the middle of the sandwich. By right-clicking on organoids in the tiled image and selecting “Add point to ND Acquisition” from the dropdown menu, you can record the positions of all organoids in the field of view and randomly select a subset for imaging. We recommend this step for perturbation experiments, which require unbiased comparisons of organoid growth dynamics and morphology in different conditions. If you decide to pre-select multiple organoids following this process, be sure to switch to the objective being used for data acquisition before proceeding.

9. Adjust laser power, pinhole size, and camera settings, ensuring – by referring to the image histogram –that you are not saturating the signal, and that you are using the lowest possible laser power to achieve adequate signal. These considerations enable quantitative comparisons between images, and they help reduce phototoxicity due to repeated laser exposure.

10. Start imaging.

    If you find that organoids undergo phototoxic lysis (Fig. 6) within 12–24 hours, consider the following changes for the next imaging run: reducing laser power; increasing photomultiplier tube (PMT) gain; increasing pinhole size; reducing pixel dwell time; increasing time between frames; reducing the number of z-slices imaged per z-stack. Table 1 summarizes these suggestions and relevant trade-offs.

    We have successfully captured time-lapse movies for >1 week with a 15-minute interval between frames, and for 72 hours with a 6-minute interval between frames. In all cases we capture z-stacks with no more than 41 z-slices (organoid mid-plane +/− 20 slices), adjusting z-step size (in μm) accordingly to cover the z-range in no more than that number of steps.

11. Render 3D volumetric images and movies in image analysis software of your choice. We use Imaris 8.4 (Oxford Instruments).

Figure 6. 3D fluorescent time-lapse imaging of organoids (reproduced from Tallapragada, Cambra et al., 2021).

Using a standard confocal microscope imaging of organoids derived from LGR-DTR-EGFP/H2B-mCherry transgenic mice at 15 minute time intervals is phototoxic when imaging at 200 μm from the coverslip, but not at 75 μm distance typical of sandwich cultures, because organoids close to the coverslip require less laser power (laser exposure per voxel) to produce signal than organoids farther away. Scale bars: 100 μm.

ALTERNATE PROTOCOL 3: LIVE IMAGING OF VITAL DYE-TREATED INTESTINAL ORGANOIDS IN TRIPLE-DECKER SANDWICH CULTURES

This protocol details how to image multiple organoids in a triple-decker sandwich culture over time after treatment with a vital dye, using live calcium imaging as an illustrative example. We followed this protocol to measure intracellular calcium activity in organoids in response to luminal inflation and cell stretch (Tallapragada, Cambra et al., 2021) using a live calcium stain (Cal-630 AM). This protocol can be modified for use with other vital dyes. The key parameters to change will be the duration to incubate organoids in dye, and additional reagents (e.g., Probenecid and Pluronic F-127) that may facilitate dye uptake or prevent dye efflux.

Additional Materials:

• 10 mM Cal-630™ AM (see recipe)

• 10% Pluronic® F-127 (see recipe)

• 25 mM Probenecid (see recipe)

• HHBS (AAT Bioquest, cat. no. 20011)

Timing: ≥ 4 hours

1. Prepare fresh reagents immediately prior to use. Prepare the confocal microscope for live imaging (turn on CO₂ and stage-top incubator; set to 37°C and 5% CO₂), and allow the system to equilibrate for 15 minutes prior to use.

2. To estimate background fluorescence in the absence of calcium dye, image organoids before adding calcium working solution, using the same laser channel and the same settings (laser power, gain, camera settings). Take multiple images for multiple settings if you are unsure – this is a critical control for quantitative imaging.

   Be careful about overexposure to laser light and the effects of prolonged imaging! We recommend testing imaging parameters on an organoid that will not be used subsequently for live imaging.

   We recommend keeping the plate on the stage for the entire duration of the experiment to minimize xyz-drift in organoid positions. Recentering organoids in the field of view can be time-consuming. Be sure to adjust volumes and concentrations accordingly to reach final target volumes, when adding multiple reagents over time.

3. Prepare dye working solution:

   Example for 1 well in a 12-well glass-bottom dish for triple-decker sandwich cultured organoids, for a total of 500 μL of 2X working solution:

   Final in-well concentration of Cal-630™ AM: 5 uM

   Final in-well concentration of Pluronic® F-127: 0.04%

   Final in-well concentration of Probenecid: 1 mM

   In a suitable container mix 1 μL of Cal-630 AM, 4 μL of 10% Pluronic F-127, and 40 μL of 25 mM Probenecid. Next, add HHBS until the volume is 0.5 mL. (455 uL)

4. Mix 500 uL ENR and 500 uL 2X dye working solution. Incubate on the stage for 1–2 hours. Use incubation time to select xyz-coordinates of organoids for imaging (see Step 4 below).

   You can add 2X dye working solution to a multi-well plate while it is on the microscope stage, if you do not wish to remove the plate from the stage. To do this, ensure that wells contain 500 μL ENR (or half of the target volume). It may be possible to incubate organoids with dye for less than 1 hour, but we did not test this possibility.

5. If you have not done so already, choose coordinates of organoids for time-lapse imaging, and choose parameters for recording time-lapse images.

6. Begin post-dye time-lapse imaging.

BASIC PROTOCOL 4: IMMUNOFLUORESCENCE IMAGING OF ORGANOIDS LIBERATED FROM TRIPLE-DECKER SANDWICH CULTURES

This protocol describes how to immunostain mouse intestinal organoids from triple-decker sandwiches and image them with low background. Like many other organoids cultured in hydrogels, intestinal organoids should be released, or “liberated”, from the hydrogel to reduce background fluorescence and non-specific staining. Organoids grown in a triple-decker sandwich culture must be carefully extracted from the sandwich through gentle digestion of the Matrigel layers, in order to preserve three-dimensional structure and to avoid fragmenting organoids. Here we present our protocol for the multi-day process of liberating, fixing, blocking, washing, and immunostaining organoids. The protocol is adapted from a MilliporeSigma protocol to which we have linked in the Internet Resources section, and is expected to also work for human intestinal organoids. See Fig. 7 for an example of the application of this protocol to mouse intestinal organoids and Table 2 for antibody information (e.g., manufacturer and dilution recommendations).

Figure 7. Immunofluorescence of mouse intestinal organoids using Basic Protocol 4.

(A) Lysozyme [1:100] staining for Paneth cells. (B) Chromogranin A [1:100] staining for enteroendocrine cells. (C) Villin [1:100] staining for enteroendocrine cells. (D) CLCA1 [1:200] staining for goblet cells. Blue: Nuclei (DAPI/H2B-mCherry); Gray: Signal for respective antigen. Scale bars: 50 μm.

Table 2.

Antibodies for cell types immunostained using Basic Protocol 4 (see Figure 7)

Antibody Target	Cell type	Manufacturer	Catalog No.	Recommended Concentration(s)
Chromogranin A	Enteroendocrine	Abcam	ab15160	[1:100], [1:200]
CLCA1	Goblet	Abcam	ab180851	[1:100], [1:200]
Lysozyme	Paneth	Agilent (DAKO)	F037201–1	[1:100], [1:200]
Villin	Enterocyte	Santa Cruz Biotechnology	sc-58897 AF647	[1:100], [1:200]
BASP1	Stretch-responsive	Abcam	ab214322	[1:100]
ANXA1	Stretch-responsive	Abcam	ab214486	[1:100]
Rabbit IgG (secondary, 647)	-	Life Technologies	A21245	(1:1000)
Rabbit IgG (secondary, 555)	-	Thermo Fisher Scientific	A21428	(1:1000)

Materials:

• Mouse intestinal organoids cultured in triple-decker sandwiches (see Basic Protocol 2)

• Cell Recovery Solution (Corning, cat. no. 354253)

• 1x PBS (Corning, cat. no. 21–040-CV)

• 4% PFA in 1x PBS

  Prepared from 16% paraformaldehyde (Fisher Scientific, cat. no. 50–980-487), and diluted with 10X PBS (Thermo Fisher Scientific, cat. no. 14200075) in sterile water.

• Wash Buffer (see recipe)

• Permeabilization buffer (see recipe)

• Antibody Wash Buffer (see recipe)

• Blocking Buffer (see recipe)

• Antibodies and stains

  Consult Table 2 for primary and secondary antibodies used with this protocol

  Optional: DAPI (1:1000)

• 1000 μL pipet tip, 2–3 cm cut off the tip (10% BSA washed)

  Prepare at least one tip per pooled condition. You can use a razor blade or scalpel to enlarge the 1000 μL pipet tip inlet/outlet; this allows for the whole organoids to be transferred without shearing the tissue.

• Glass-bottom 96 well plate (Mattek, cat. no. P96G-1.5–5-F)

  Final containment vessel of mouse intestinal organoids for imaging.

• 50-mL Falcon tubes (10% BSA washed)

• Rocking platform

Timing: 4 days

Day 1: Fixation, permeabilization, and blocking

• 1Prepare all reagents you will need for fixation and staining.

  Prepare buffers, if necessary. Wash falcon tubes with 10% BSA. Prepare several cut 1000 μL pipet tips, and wash with 10% BSA.

• 2Image all wells prior to removal from the glass-bottom dish and/or fixation in domes to be able to confirm the number of viable organoids for imaging at the start and end of the protocol to document success.

  This step becomes optional with experience.

• 3Begin liberating organoids from Matrigel by aspirating media from wells and adding 0.5 mL ice-cold Cell Recovery Solution. Incubate plate on ice for 10 minutes.
• 4Add 4% PFA at room temperature, and incubate for 15 minutes, with gentle rocking.
• 5Transfer liberated organoids from sandwiches into BSA-washed Falcon tubes using a BSA-washed cut 1-mL tip. Allow fixed and liberated organoids to settle to the bottom of the Falcon collection tube for 15 minutes.
• 6Meanwhile, wash and incubate each sandwich well with Wash Buffer for 15 minutes at room temperature, with gentle rotation.

  This step both binds free aldehydes and releases organoids from the dish. Check the wells/dishes under the microscope to see if organoids left on the dish (failed to completely liberate) are being liberated after washing.

• 7Aspirate PFA (Wash Buffer during repeated steps) from Falcon tubes containing settled organoids, and transfer freshly-liberated organoids now in the Wash Buffer remaining in sandwich wells using a cut, BSA-washed 1000 μL pipet tip.

  Be sure to get the pipet tip as close as possible when depositing freshly liberated organoids onto the settled organoids to avoid disturbing them too much.

• 8Repeat steps 4–6 two more times, for a total of three washes.

  Ensure that few organoids remain in the washed sandwich wells. If there are still organoids remaining in the thrice washed culture dish, you may try another round of washing or it is possible you may need to optimize incubation times with Cell Recovery Solution and PFA fixation.

• 9Add 1 mL of fresh Wash Buffer to settled organoids, resuspend, and transfer organoids from the Falcon tube to a 96-well glass-bottom plate using a cut, BSA-washed 1000 μL pipet tip, filling approximately 10 wells (100 μL each) from each collection tube.
• 10Verify that organoids are present in the 96-well plate using a standard tissue culture microscope.
• 11Remove excess Wash Buffer from each of the 96-well plate wells with organoids by careful aspiration with a 1000 μL pipet tip, and incubate all organoids for 45 minutes in 200 μL Permeabilization Buffer at room temperature, gently rocking.
• 12Aspirate Permeabilization Buffer and incubate wells in 200 μL Blocking Buffer overnight, or up to 24 hours.

Day 2: Primary Staining

• 13Prepare fresh blocking buffer for staining, and master mixes for primary antibodies.
• 14Add 200 μL of blocking buffer and 200 μL of primary antibody solution to each well of organoids in the 96-well plate. If staining organoids with different antibodies in parallel, ensure that the right primary antibodies end up in the right wells.
• 15Incubate overnight, or up to 24 hours.

Day 3: Washing, Secondary Staining

• 13Wash organoids three times with Antibody Wash Buffer, for 20 minutes each time.

  In between washes, carefully aspirate supernatant in each well using a 1 mL pipet. Do not use vacuum suction.

• 14If you are using primary antibodies directly conjugated to fluorophores, samples are now ready to be imaged on a confocal microscope. However, you can leave them in 1x PBS overnight and wait to image until the next day.

  If you would like to stain nuclei, add 1:1000 DAPI to the sample well and proceed to Day 4 (Step 20). If not, you can skip to Day 4 (Step 22).

• 15If you are using unconjugated primary antibodies, prepare secondary antibodies (1:1000) in Blocking Buffer and add to the appropriate wells of organoids.
• 16Stain plate with secondary antibody at (1:1000) overnight with gentle rocking at 4°C.

Day 4: Washing and imaging secondary-stained samples

• 13Wash secondary-stained samples once for 20 minutes with Wash Buffer at room temperature, gently rotating.

  Optionally, add 200 μL of 1:1000 DAPI to each of the samples requiring nuclear staining.

• 14Incubate each sample for 20 minutes at room temperature with gentle rocking.
• 15Wash twice for 20 minutes each with Antibody Wash Buffer.
• 16Wash once with 1x PBS for 15 minutes.
• 17Image organoids using a 20X dry objective (NA 0.75) or 40X water immersion objective (NA 1.15) on a confocal microscope (e.g., Nikon A1R point-scanning)

  We use Immersol W 2010 as the immersion fluid for our 40X water immersion objective, because it is index-matched to water but evaporates far slower.

  If you have a motorized stage, we recommend taking tiled images of entire wells with a 10X dry objective, in order to identify every organoid in a well and facilitate high-throughput imaging for statistically robust measurements using the higher magnification objectives.

ALTERNATE PROTOCOL 4.1: LIBERATING AND FIXING MOUSE INTESTINAL ORGANOIDS FROM DOME CULTURES

This protocol also enables immunostaining of organoids, but starting from dome cultures instead of triple-decker sandwich cultures. Liberating organoids from Matrigel domes requires slight modifications to the procedure in Steps 0–7 of Basic Protocol 4; but Steps 8–23 of Basic Protocol 4 can be applied as is.

Additional Materials:

• Instead of triple-decker sandwich cultured organoids, use dome cultured mouse intestinal organoids (see Alternate Protocol 2)

Timing: 90 minutes

1. Prepare all reagents you will need for fixation and staining.

   Prepare buffers, if necessary. Wash falcon tubes with 10% BSA. Prepare several cut 1000 μL pipet tips, and wash with 10% BSA.

2. Image all wells prior to removal from the glass-bottom dish and/or fixation in domes to be able to confirm the number of viable organoids for imaging at the start and end of the protocol to document success.

   This step becomes obsolete with experience.

3. Transfer whole domes to 10% BSA washed 6 mm petri dishes.

   This can be done by gently disrupting the adherence of the Matrigel dome to the glass or plastic bottom of the culture dish, followed by suction with the cut 1000 μL pipet tip.

4. Incubate domes/sandwiches for liberation in ice cold Cell Recovery Solution on ice for 30 minutes.

   This step initiates the digestion process of the Matrigel hydrogel.

5. Add 4% room temperature PFA for 30 minutes, gently rotating.

   This step serves both to fix and to further liberate the organoids.

6. Transfer liberated organoids from petri dishes into BSA washed 50-mL falcon tubes by pouring the loose organoids from dishes.

7. Wash each dish with Wash Buffer, gently rotating on a blot mixer or gentle nutator at room temperature for 15 minutes. Allow organoids to settle to the bottom of the 50-mL Falcon tube during this time.

   This step serves to wash and bind free aldehydes (block) and further releases organoids from the dish. Check the wells/dishes under a bright field scope to see if they are being liberated after washing.

8. Aspirate supernatant from the Falcon tubes and add newly released organoids in Wash Buffer to the Falcon tube with settled organoids. Repeat steps 6–7 for a total of 3 washes.

9. Follow Basic Protocol 4 from Step 8 to Step 23 to use liberated organoids in an immunostaining experiment.

ALTERNATE PROTOCOL 4.2: MEASURING CELL PROLIFERATION BY EDU STAINING MOUSE INTESTINAL ORGANOIDS

This protocol outlines an assay for using the Click-IT EdU Kit with mouse intestinal organoids grown in sandwich culture. Organoids do not need to be liberated from the Matrigel for this assay if no antibodies are used jointly with EdU staining. Thus, this protocol can be completed within a day, from fixation to imaging. For example, results of applying this protocol to mouse intestinal organoids, see Tallapragada, Cambra et al., 2021. This protocol is also expected to work with human intestinal organoids.

Materials:

• Click-IT™ EdU Plus staining kit with (Alexa Fluor® 647) (Thermo Fisher Scientific, C10640) (see recipe for staining cocktail)

Timing: Approximately 4 hours

Incubate organoids in EdU

• 1Begin 10 μM EdU incubation in ENR media for 2 hours at 37°C in the tissue culture incubator.

  Optional: Add perturbation reagent (i.e., small molecule drug) prior to incubation and incubate with perturbation reagent.

  1. Remove 0.5X of the media in each treated well

  2. Add fresh ENR, 2X EdU, and 1x small molecule drug treatment to desired concentration from stock.

  3. Add 1 mL of mixture to 1 mL of media and small molecule drug

• 2Fix mouse intestinal organoids in triple-decker sandwiches for 15 minutes with 4% PFA, gently rotating the plate at room temperature.
• 3Wash twice with Wash Buffer for 5 minutes each, gently rotating at room temperature.
• 4Incubate for 20 minutes in Permeabilization Buffer, gently rotating at room temperature.
• 5Wash twice with Wash Buffer for 5 minutes each, gently rotating at room temperature.

Prepare and incubate in EdU staining cocktail

• 6While samples are being washed, prepare 1 mL of Click-It EdU Kit cocktail (see recipe) and immediately after making it, add 125 μL to each well (12-well glass-bottom plate), directly on top of the sandwich culture.

  Adjust volume as necessary for different surface areas to ensure coverage of the surface area of the triple-decker sandwich culture.

• 7Incubate the plate for 30 minutes at room temperature, gently rotating and covered from light.

  Keep the plate protected from light for the remainder of the protocol.

• 8Aspirate each well to remove the EdU reaction cocktail.

Wash and Image

• 6Wash each well once with Wash Buffer.

  Optionally, if you need to label nuclei, add DAPI in 1x PBS and stain for 20 minutes, then wash once more post-staining with Wash Buffer before moving on.

• 7Remove Wash Buffer and add 1x PBS.
• 8Image using a 20X dry objective (NA 0.75) or 40X water immersion objective (NA 1.15) on a confocal microscope (e.g., Nikon A1R point-scanning). See Basic Protocol 3 for additional guidance on image capture.

BASIC PROTOCOL 5: SINGLE-CELL RNA SEQUENCING AND ANALYSIS OF MOUSE INTESTINAL ORGANOIDS

This protocol describes the process of single-cell RNA sequencing mouse intestinal organoids. We specifically cover steps for single cell dissociation prior to analysis. Expectations from single cell data and recommended pipelines are discussed in “Understanding Results” later in this paper, and we provide example Python notebooks to facilitate data analysis. This protocol is also expected to work for human intestinal organoids.

Dissociation of mouse intestinal organoids into single cells

Materials:

• Triple-decker sandwich cultured mouse intestinal organoids (see Basic Protocol 2)

  Or dome cultured mouse intestinal organoids (see Alternate Protocol 2)

  Enough material to obtain approximately 5×10⁵ cells/mL, if subjecting to single cell RNA sequencing.

• Trypan Blue (Bio-Rad, cat. no. 1450021)

• 1x PBS (Corning, cat. no. 21–040-CV)

• TrypLE Express (Thermo Fisher Scientific, cat. no. 12605010)

• 1% (w/v) BSA (Millipore Sigma, cat. no. A9418) in 1x PBS

  For resuspension of cells prior to single cell RNA-sequencing

• 10% (w/v) BSA in 1x PBS

  For coating pipet tips and tubes that come into contact with the cell suspension. This prevents cellular material from sticking to the walls of the containment and transfer vessels.

• 15 mL falcon tube (washed with 10% BSA)

• 1000 μL pipet tips (washed with 10% BSA)

• 1 mL test tube (washed with 10% BSA)

• Swinging bucket centrifuge set to 4°C

• 40 μm filter (pluriSelect, cat. no. 43–10040-60)

• Bright field microscope

• Hemocytometer

Timing: 45 minutes

1. Aspirate media from wells.

2. Wash wells twice with 1x PBS (1.5 mL for one well of a 12-well plate).

3. Add 0.5- to 1-mL TrypLE Express to each well and mechanically detach the Matrigel from the bottom of each well with a pipet tip.

   Add 1 mL for 12-well glass-bottom dishes. When comparing dome v. sandwich cultures, plate them both on the glass-bottom covered dishes. Take care to break up large chunks of Matrigel first, as clumps of cells will dissociate into single cells largely enzymatically. Note that when applying this to triple-decker sandwich cultures, you may lift the PolyHEMA coating when you scrape. Try not to clog the pipet tip in the process, by being mindful of these sticky clumps. Also be careful not to detach the glass coverslip from the bottom of the well with too much force.

4. Incubate wells at 37°C for approximately 25 minutes (no less than 20 minutes and no longer than 30 minutes), triturating solution gently with a P1000 pipet every 5 minutes. Avoid causing the solution to bubble.

   Monitor the dissociation process by periodically examining wells using a standard tissue culture microscope. Stop dissociation after 30 minutes, or when you see many single cells and no clumps larger than 2–3 cells. You should observe decreasing amounts of material stuck to the bottom of the well.

5. Collect the total volume of organoid solution in TrypLE in a washed 15 mL Falcon tube and dilute it 1:1 with cold DMEM/F12 media.

6. Strain the solution through a 40 μm strainer. Keep this solution on ice.

   Note that PolyHEMA fragments may affect the ability of the solution to pass through the filter. Lift the serological pipet off of the filter occasionally to relieve pressure build-up.

7. Spin the remainder at 250g for 1 minute at 4°C.

8. Aspirate TrypLE and media. Resuspend cells in 200 μL 1x PBS, 1% BSA at 4°C.

9. Immediately check cells for viability and dissociation quality.

   Confirm single cell dissociation and viability with a cell counter and/or a hemocytometer, using trypan blue stain for viability. To do so, transfer 10 μL from the cell solution into an Eppendorf tube, add 10 μL of trypan blue (1:1), and sample 10 μL of that for the cell counter or hemocytometer.

10. Place cells on ice and proceed to single-cell RNA sequencing, if cells are at required density.

11. Load the cells for single cell RNA-sequencing, for example using inDrops (Zilionis et al., 2017), Seq-Well (Gierahn et al., 2017), a 10X Chromium system (Zheng et al., 2017), or another single cell RNA-sequencing method of your choosing.

    If you choose to use inDrops, we recommend generating per-cell gene expression counts from raw sequencing reads using the v3 iteration of the publicly available (github.com/indrops) data processing pipeline as described (Klein et al., 2015). Parameters used with the indrop.py pipeline for data provided in the publicly accessible submission of data from Tallpragada et al. 2021 on GEO:GSE164638 are specified in yaml files as provided on GitHub (github.com/AllonKleinLab/paper-data/tree/master/Tallapragada_Cambra_2021).

12. Use a standard pipeline for analyzing single cell RNA-seq data, such as Seurat (Satija et al., 2015) or Scanpy (Wolf et al., 2018). See Understanding Results to gauge expectations for computational analysis results, and Supplementary Materials for a set of example notebooks to process data.

REAGENTS AND SOLUTIONS:

Antibody Wash Buffer

• 1x PBS (Corning, cat. no. 21–040-CV)

• 0.2% Triton

• 0.04% Tween-20

  Store at 4°C for up to 1 month.

Blocking Buffer

• 0.5% (v/v) Triton X-100

  Permeabilizes – in this protocol the permeabilization is combined with blocking.

• 5% (w/v) BSA (Millipore Sigma, cat. no. A9418)

  Competes with all weakly bound antibodies to prevent non-specific binding of antibodies.

• 10% (v/v) goat serum (abcam, cat. no. ab7481)

  Quenches non-specific secondary signal.

• 0.3M Glycine

  Binds remaining free aldehydes.

• 1x PBS (Corning, cat. no. 21–040-CV)

Cal-630 AM (AATBioquest, cat. no. 20530) (5 mM)

Prepared according to manufacturer (AATBioquest) provided instructions:

1. In a suitable container (e.g., 500 μL test tube), mix 50 μg of Cal-630™ AM with 7.79 μL of anhydrous DMSO.

   The exact concentration of the indicator required for loading must be determined empirically.

2. Dye should be incubated with mouse intestinal organoids for at least 1 hour.

   As indicated in the kit instructions: It is recommended to prepare and use the Cal-630 AM stock solution on the same day it is used. However, if stock solutions need to be prepared in advance, store the Cal-630 AM stock solution as aliquots in tightly sealed vials at −20°C, in a bag or box with a desiccant and protected from light. Under these conditions, AM esters should be stable for 3 months.

Click-IT EdU Kit cocktail (Thermo Fisher Scientific, cat. no. C10640)

Make the cocktail in the order of the components listed below and prepare fresh during wash steps following permeabilization. Be sure to adjust the total volume of cocktail for each well according to the (you can use as little as 125 μL on a 12-well glass-bottom plate, so long as you coat the surface area of the triple-decker sandwich)

1. 86% (v/v) 1x Click-iT® reaction buffer

   Store in 860 μL aliquots at 4°C after reconstitution

2. 4% (v/v) CuSO₄ (Component E)

   Store in 1.2 μL aliquots at 4°C after reconstitution, protected from light

3. 0.24% (v/v) Alexa Fluor® azide

   Store in 1.2 μL aliquots at −20°C after reconstitution, protected from light

4. 10% (v/v) 1x Reaction buffer additive

   Store 5 μL aliquots of 10X Reaction buffer additive at −20°C after reconstitution

   Prepare a 1x solution with the 10X solution as indicated in the manufacturer’s protocol with DI water.

Complete crypt culture media containing EGF, Noggin, and R-Spondin-1 (ENR) media

• 86% Advanced DMEM/F12 (Thermo, 11320–033)

• 10% Rspo1 conditioned media (see Support Protocol 2.2)

• 1% Glutamax (Thermo, cat. no. 35050–079)

• 1% 1M HEPES (Thermo, cat. no. 15630–080)

• 1% B-27 Supplement (Thermo, cat. no. 17504–044)

• 0.5% N2 (Thermo, cat. no. 17502–048)

• 0.25% N-acetyl cysteine (500mM) (Sigma, A7250–10G)

• 0.2% Primocin (50 mg/mL) (Invivogen, cat. no. ant-pm-1)

• 0.1% Noggin (100 ng/mL ) (Millipore Sigma, cat. no. SRP3227–20UG)

• 0.01% EGF (500 μg/mL) (Thermo Fisher, cat. no. PMG8041)

Complete crypt culture media containing Wnt3a, EGF, Noggin, and R-Spondin-1 (WENR) media

• 50% Wnt3a/Rspo1/Noggin conditioned media (see Support Protocol 2.1)

• 46% Advanced DMEM/F12 (Thermo, 11320–033)

• 1% Glutamax (Thermo, cat. no. 35050–079)

• 1% 1M HEPES (Thermo, cat. no. 15630–080)

• 1% B-27 Supplement (Thermo, cat. no. 17504–044)

• 0.5% N2 (Thermo, cat. no. 17502–048))

• 0.25% N-acetyl cysteine (500mM) (Sigma, A7250–10G)

• 0.2% Primocin (50 mg/mL) (Invivogen, cat. no. ant-pm-1)

• 0.01% EGF (500 μg/mL) (Thermo Fisher, cat. no. PMG8041)

Conditioned Media AD-DF +++ (500 mL)

• 5 mL 100x Penicillin/Streptomycin (Life Technologies, cat. no. 15140122)

• 5 mL 1M HEPES (Life Technologies, cat. no. 15630)

• 5 mL Glutamax (Life Technologies, cat. no. 35050)

• 485 mL Advanced DMEM/F12 (Life Technologies, cat. no. 12634)

Growing Media (500 mL)

• 5 mL 100x Penicillin/Streptomycin

• 60 mL FBS Life Technologies

• 435 mL DMEM, high glucose, GlutaMAX

Human duodenum growth media (100 mL)

• 50% Wnt3a/RSpo1/Noggin conditioned media (see Support Protocol 2.1)

• 45% Advanced DMEM/F12 (Life Technologies, cat no. 12634–028)

• 1% Glutamax (Life Technologies, cat no. 35050–061)

• 1% 1M HEPES (Life Technologies, cat no. 15630080)

• 0.2% Primocin (50 mg/mL) (Invivogen, cat. no. ant-pm-1)

• 0.2% Normocin (50mg/mL, Invivogen, cat. no. ant-nr-1)

• 1% B-27 Supplement (50x) (Life Technologies, cat no. 12587–010)

• 1% ​​Nicotinamide (1M) (Sigma, N0636)

• 0.5% N2 (100x) (Life Technologies, cat no. 17502–001)

• 0.20% N-acetyl cysteine (500mM) (Sigma, A7250–10G)

• 0.1% A-83–01 (500μM) (Sigma SML0788)

• 0.0332% SB202190 (30mM) (Sigma, S7067)

• 0.01% EGF (500 μg/mL) (Thermo Fisher, cat. no. PMG8041)

• 0.01% Gastrin (500 μM) (Sigma, G9145)

L-Cell Media (500 mL)

• 5 mL 100x Penicillin/Streptomycin (Invitrogen, cat. no. 15140122)

• 50 mL FBS (Gibco, cat. no. 26140)

• 445 mL DMEM, high glucose (Invitrogen, cat. no. 11965–092)

  Store at 4°C

PBS-EDTA (0.5 mM)

• 500 μL 0.5M EDTA (Invitrogen, cat. no. 15575020)

• 495.5 mL Dulbecco’s PBS (Sigma, cat. no. D8537)

  Store at room temperature

Permeabilization buffer

• 0.5% (v/v) Triton X-100

• 1x PBS (Corning, cat. no. 21–040-CV)

  Store at 4°C for up to 1 month.

Pluronic F-127 (AAT Bioquest, cat. no. 20050) (10%)

Pluronic® F-127 (PF-127) is a nonionic surfactant and relatively non-toxic to cells. PF-127 is commonly used with dye AM esters to improve their aqueous solubility. Prepared according to AAT Bioquest instructions.

1. Dissolve 1 g of Pluronic F-127 in 10 mL of distilled water to make a 10% (w/v) stock solution.

2. Heat 10% (w/v) Pluronic F-127 stock solution for about 30 minutes at a temperature ranging from 40 to 50°C.

3. Store excess 10% Pluronic F-127 at room temperature for no more than 6 months.

Primary Culture Media

1. 5 mL 100x Penicillin/Streptomycin (Invitrogen, cat. no. 15140122)

2. 5 mL 200mM L-Glutamine (Sigma, cat. no. G7513)

3. 100 mL FBS (Gibco, cat. no. 26140)

4. 390 mL Advanced DMEM/F12 (Invitrogen, cat. no. 12634010)

   Store at 4°C

Probenecid (25 mM)

Probenecid (0.5–1.0 mM) is added to the dye working solution to reduce the leakage of the de-esterified indicators via organic anion-transporters. Prepared according to AAT Bioquest instructions. Avoid repeated freeze-thaw cycles.

1. In a suitable container, dissolve 1 vial (72 mg) of Probenecid (AATBioquest, cat. no. 20060) in 0.3 mL of 1 M NaOH.

2. Add HHBS or a buffer of your choice until the volume is 10 mL.

3. Aliquot and store at −20°C and protected from light for up to 6 months.

0.25% Trypsin-EDTA

1. 2.5% Trypsin (Invitrogen, cat. no.15090046)

2. PBS-EDTA (0.5 mM)

   For a total volume of X 0.25% Trypsin-EDTA, combine 0.1× 2.5% Trypsin and 0.9X PBS-EDTA (0.5 mM). Handle reagents in a tissue culture hood (BSL2) and filter sterilize.

   Aliquots can be stored at −20°C for 1 month.

Wash solution

• 0.75% Glycine

  Binds unreacted aldehydes after fixation with 4% PFA

• 10% BSA (w/v) (Millipore Sigma, cat. no. A9418)

  Prevents organoids from adhering to containment and transfer vessels

• 1x PBS (Corning, cat. no. 21–040-CV)

  It is recommended to make this solution fresh, but it can be stored at 4°C for 1–2 weeks if handled appropriately (always check for turbidity; discard if at all opaque). Use serological and autoclaved pipets and equipment to prevent contamination during use. Aliquot and use only what is needed per day from stock.

COMMENTARY

BACKGROUND INFORMATION:

Over the past decade, several protocols have been developed to grow organoids in uniform environments close to coverslips for imaging. One approach cultures organoids between a Transwell insert on top and a glass coverslip on the bottom, aligning organoids in a single z-plane relative to a microscope objective (Saarela et al., 2017). Another approach confines organoids between two hydrogel layers with a thin base layer of hydrogel between organoids and the glass coverslip to facilitate systematic microscopy of multiple organoids. This technique has been applied to culture mammary organoids (Debnath et al., 2003; Lee et al., 2007; Mroue and Bissell, 2013; Gajewska and McNally, 2017) and bronchospheres (Hild and Jaffe, 2016). This current protocol represents a simple extension of these approaches by adding a third passivation layer to culture. An alternative to these methods is the use of modern structured illumination to collect full-thickness, frequent images of organoids (Pampaloni et al., 2015; McKinley et al., 2018). Compared to such approaches, our method requires simpler sample preparation, a standard microscope, and it very significantly increases the number of organoids that can be imaged at once.

CRITICAL PARAMETERS:

When preparing PolyHEMA-coated plates following Basic Protocol 1, imperfections in the coating often arise. Small defects such as the bubbles seen in Fig. 4B rarely interfere with organoid growth and imaging away from the defects. However, if bubbles are too widespread or large (Fig. 3C,F), the PolyHEMA coating may detach from the plate as you prepare sandwich cultures, especially when depositing organoids in aqueous media. One purpose of incubating the PolyHEMA-coated plates in 1x PBS at 37°C for hours to days before plating is to see which coatings detach from the plate and which coatings do not. If the application is successful, we have found that 70–100% of prepared plates pass this QC step. We recommend preparing several plates in advance so that you can choose the best ones for coating.

Figure 4. Comparison of densely and sparsely plated sandwich cultures.

(A) Tiled image of the entire well of a 12 well glass-bottom dish showing an evenly spread and dense triple-decker sandwich culture. (B) Tiled image of the same size well with a sparsely plated triple-decker sandwich culture. Scale bars = 1 mm.

When preparing triple-decker sandwiches following Basic Protocol 2, you should expect a relatively uniform distribution of organoids across the surface area of the well (Fig. 4A). If organoids are not distributed evenly (Fig. 4B), pay attention to the way you disperse organoids when plating. As discussed in the protocol, swirling motions concentrate organoids at the center of a well and should be avoided. Organoids should be healthy for at least one week after plating.

During routine passaging in dome cultures, (Alternate Protocol 2.2), when preparing Matrigel domes we have noticed that Matrigel can become runny and fail to form a hemispherical drop at the center of a well, pooling instead along the edges. In our experience, organoids in these “failed” domes end up less healthy and give rise to less healthy sandwich cultures. If you notice that Matrigel is not forming domes, ensure that you have not thawed the Matrigel stock too many times; that the original 10 mL vial is mixed well before aliquoting; and that you allow Matrigel to warm slightly before attempting to plate it. One way to test if the problem is your Matrigel or your technique is to attempt to make domes on glass-bottom plates, as Matrigel forms domes more easily on glass. If it is still runny, check the Certificate of Analysis for your lot of Matrigel to see if the total protein concentration is low (e.g., ~<9 mg/mL). If all else fails, order and try a different batch of Matrigel.

Live imaging organoids requires that you have healthy organoids at the outset and that you can maintain culture conditions while you are imaging. It is important to use the lowest laser power settings possible to maximize the duration you can image organoids without inducing phototoxicity. If you notice you need to use high laser powers, consider whether your assay uses any reagents that affect the fluors you are using, and check that the distance between the coverslip and your sample is <100μm. If most organoids are found beyond this distance, you should check that you are coating wells with the appropriate volume of PolyHEMA; too much will make the layer thicker than it needs to be. It is also important to maintain 5% CO₂ and 37°C for the duration of organoid imaging. Finally, long-term imaging experiments will require you either to add extra media when plating organoids (overfeeding wells) or to add fresh media during the experiment to compensate for evaporation as well to feed cells. Plan accordingly if you need to add particular concentrations of a reagent for your assay.

Basic Protocol 4 describes stringent staining techniques to minimize background fluorescence and non-specific staining. We have found it difficult to specifically antibody stain organoids in intact triple-decker sandwiches or Matrigel domes. When staining organoids in triple-decker sandwiches in situ, in our hands we have seen delamination of the sandwich culture and PolyHEMA turning cloudy, preventing signal acquisition. However, we have successfully stained organoids with chemical dyes in triple-decker sandwiches in situ, without liberation from Matrigel. See Alternate Protocol 4.2 for more details. When liberating organoids from Matrigel for staining, they will stick to any surface not coated with 10% BSA during and after liberation; major loss of material will occur if vessels and transfer tools are not washed with BSA prior to coming into contact with the organoids. It is important to be gentle when transferring and washing organoids, again to prevent material loss.

Basic Protocol 5 may require coordination with a single-cell sequencing core, and it assumes know-how or support with bioinformatic analysis of scRNA-Seq data. When designing experiments, each of your samples should have enough cells to represent expected cell states within the data. A typical number of thousands of cells per replicate is suitable to capture all of the major cell types. Once data is obtained, you may adapt the provided python notebooks for analysis (see Github Repository for Tallapragada, Cambra et al., 2021 in Internet Resources). These notebooks make use of the scanPy environment for single cell RNA-Seq data analysis to produce the results in Figure 7. These notebooks cover: data quality checks and background filtering; variable gene selection; dimensionality reduction by principal component analysis; embedding and visualization using UMAP (McInnes et al., 2018); unsupervised clustering; cell type classification; classification refinement using unsupervised clustering; and plotting steps to produce figures such as in (Tallapragada, Cambra et al., 2021). When using classification methods that assign a cell type identity to each observed cell transcriptome, one should check the classified data by examining the expression of canonical markers of enterocytes, Paneth cells, goblet cells, enteroendocrine cells, and intestinal stem cells to gain confidence that the data is consistent with prior observations. Lists of marker genes, and Python code to carry out visualization can be found in the Github Repository for Tallapragada, Cambra et al., 2021 (See Internet Resources). Note also that in organoids, unlike in primary tissues, Paneth cells and goblet cells often do not appear as distinct transcriptomic clusters after embedding; they may be annotated confidently as secretory cells together.

Troubleshooting Guide

Understanding Results:

Basic Protocol 1 will yield results that look like the images in Figs. 3A,D if you take care to completely dry the coatings prior to adding a second coating of PolyHEMA or 1x PBS. If you still have issues, consider making a fresh batch of PolyHEMA to see if this is the source of excessive defects, as PolyHEMA will form precipitates over time. Minor defects are acceptable if they leave the majority of the surface area imageable (free of bubbles/defects) and they do not cause the PolyHEMA coating to lift in the process of making triple-decker sandwiches.

Organoids plated in the triple-decker sandwich (Basic Protocol 2) should be uniformly distributed; if they are not, ensure that you are not swirling the dish when plating organoids, and that you have plated cells with a 1:2 or 1:3 split ratio.

Live imaging results (Basic Protocol 3) are sensitive to environmental conditions and camera settings. If your organoids are not surviving very long after imaging, look for sources of phototoxicity (high laser power, long laser dwell times) and check that the temperature of the plate, media volume, and carbon dioxide concentrations are appropriate for culturing while imaging.

Liberation and immunostaining (Basic Protocol 4) requires the reagents provided in order to ensure the results presented in Figure 7. Omission of any of these ingredients may result in poor signal quality or high background. If you are having issues executing this protocol and reproducing results, make sure that all of your reagents are fresh and appropriately stored (for example, store 4% PFA in PBS at −20°C and only thaw when you are ready to use); be aware of temperature requirements at each step, and when troubleshooting, take images of your plate after each major step and pay attention to material loss and sources of material loss as you go along.

Before using Basic Protocol 5 for a full scRNA-Seq experiment, we strongly recommend that you first practice it by dissociating organoids and then monitoring the dissociated cells. This can be done by periodically sampling dissociated cells and scoring their number and viability on a tissue culture microscope using Trypan Blue staining. The cells should remain viable on ice for an hour or more after dissociation. Viability should remain above 75%. Adjust the total number of organoids to ensure that you have enough cells for your experiment, as per the requirement of the platform you will be using. We recommend doing this with the exact experimental set-up (e.g. including perturbations) to anticipate how much material you will need and to optimize your technique so that you have the highest number of viable cells possible. Remember to be gentle in dissociation, to monitor the organoids every five minutes as they dissociate, and to prevent over-incubation in TrypLE (or else cells are more likely to rupture during encapsulation). Figure 8A is representative of cells dissociated and subsequently used for downstream analysis.

Figure 8. Single cell RNA-sequencing of mouse intestinal organoids (adapted from Tallapragada, Cambra et al., 2021).

(A) Dissociated cells prior to sequencing. This represents a sample of approximately 96 pooled organoids from 4 wells of a 12-well (glass-bottom) triple-decker sandwich culture plate. Scale bar: 100 μm. (B) Histograms of reads, aligned reads, and UMI-filtered counts obtained during preprocessing of scRNA-Seq data from the same sample as cells shown in (A). Grey line shows a filtering threshold for the UMI-filtered counts. (C) UMAP visualization of scRNA-Seq data obtained from both sandwich and dome cultures, merged, with cell states annotated following preprocessing and annotation (adapted from Tallapragada, Cambra et al, 2021).

During processing of the single cell RNA-seq data, we recommend following current best practices (Luecken and Theis, 2019), including filtering out background cell barcodes (see Figure 8B), annotating doublets (Wolock et al., 2018), variable gene selection, dimensionality reduction, visualization by two-dimensional embedding (e.g. UMAP), clustering, and classifying cell types. We provide Python notebooks that carry out these key steps and can be used as templates for analysis (Github Repository for Tallapragada, Cambra et al., 2021). Note that when working with murine intestinal organoids, goblet and Paneth cells may not separate neatly into two distinct clusters, as is the case for the provided example data. In this case, group the states together into a secretory state.

Time Considerations:

PolyHEMA coating in Basic Protocol 1 will take approximately 4–8 days depending on the humidity and temperature of the ambient air in the tissue culture hood. Drier, hotter ambient air likely accelerates the drying process, but this has not been systematically tested in our hands. Support Protocol 1 should take approximately 4 hours to complete, including cool down and filtration of the PolyHEMA solution.

The triple-decker sandwich plating protocol (Basic Protocol 2) takes approximately 3–4 hours to complete, depending on the incubation times the user chooses within the acceptable ranges given. Seeding triple-decker cultures from low-density or single cells requires an overnight step in order to ensure proper cross-linking of the Matrigel layers, and should take at least 18 hours to complete. Note that organoids should first be plated using Alternate Protocol 2.2 for 2 weeks following thaw: one week in WENR and the second week in ENR (see Reagents and Solutions), prior to the third week’s plating in triple-decker sandwich cultures for experimentation.

Support Protocols 2.1 and 2.2 detail the production of Wnt3a conditioned media used in WENR media and Rspo-1 conditioned media (used in both ENR and WENR media recipes), and require between one to one and a half weeks to generate from L-WRN and 293T-HA-RspoI-Fc cells to culture, respectively. This time consideration includes thawing to the final collection of conditioned media.

Basic Protocol 3 lasts as long as the duration of a live imaging experiment. In our hands, we have not imaged organoids for more than 170 hours (~7 days) after plating, but this is not a fundamental limit of the protocol or organoid culture.

Basic Protocol 4 takes approximately 4 days. Alternate Protocol 4.1 is an adaptation of the liberation steps outlined in Basic Protocol 4, for dome cultures instead of sandwich cultures. Alternate Protocol 4.2 outlines a proliferation assay for organoids and takes approximately 4 hours, including EdU incubation time before fixation. This protocol does not require antibody staining, and thus does not require the more stringent application of blocking, wash, and incubation steps as they are applied in Basic Protocol 4, although some reagents for these protocols are shared.

Basic Protocol 5 takes at least 45 minutes for dissociation, and the time to encapsulate and generate a library for a single-cell RNA sequencing experiment will vary by platform (typically <1 hour for cell encapsulation, and 1–2 days for library preparation). Final sequencing library QC and sequencing, which have not been described in this protocol, are typically carried out by sequencing facilities and take <1 week, although lag times can add 2–3 weeks delay. Bioinformatic analysis of scRNA-Seq data can require as little as a few days using the Jupyter Notebooks provided.

Table 3.

Common troubleshooting problems and solutions

Problem	Possible Cause(s)	Solution(s)
Basic Protocol 1
Excessive bubbles or delamination after the first round of coating or addition of 1x PBS (see Figure 3C).	PolyHEMA did not dry completely prior to addition of 1x PBS (most likely) OR prior to the second coat of PolyHEMA.	Use a pipet tip to check that the PolyHEMA coating is completely dry before adding a second coat or 1x PBS; check the vacuum desiccator to ensure there is a tight seal.
Excessive delamination; opacity (see Figure 3E).	Old PolyHEMA solution.	PolyHEMA solution has a shelf-life of about 3 months. If your solution is over 3 months old, do not use it for coating and prepare fresh solution instead.
Excessive bubbles (see Figure 3F).	Bubbles were introduced during the coating step.	If you have any bubbles in your PolyHEMA aliquot (in your pipet tip) or on the plate, pop or aspirate the bubbles if possible. Avoid introducing bubbles at any step of coating.
Support Protocol 1
PolyHEMA solution has particulates in it.	If this issue occurs after 3 hours at 65°C, but before filtration, the PolyHEMA may not be entirely dissolved yet. Also, Parafilm particulates may get into the solution if you are not careful; the melting temperature of Parafilm is 60°C, so it is important to cover the beaker (or other glass vessel for mixing) with aluminum foil before adding layers of Parafilm on top, to keep Parafilm from melting and coming in contact with PolyHEMA solution. If this issue occurs after cooling but before filtering, it may be that the PolyHEMA isn’t fully dissolved, or PolyHEMA has precipitated out as a result of evaporation of ethanol.	Check to make sure the solution is actually at 65°C and has been for 3 hours; use a hot plate with temperature feedback. Use a thick layer of Parafilm to seal the top of the beaker; only fill half of the total volume of the beaker when making the solution. Filter solution a second time; if the particulates are precipitated PolyHEMA due to evaporated ethanol, the concentration of PolyHEMA may not have changed significantly. As long as PolyHEMA evenly coats glass coverslips, any concentration of PolyHEMA solution is acceptable; assess the quality of the solution with a trial run of coating a plate. Ensure that glassware and containment vessels you are using are all autoclaved and free of particulates at every step.
Alternate Protocol 2.2
When plating dome cultures, Matrigel isn’t viscous enough and pools around the edges of a well.	Matrigel has low protein content; be sure to thoroughly mix the Matrigel before aliquoting. Matrigel and cell suspension is slightly too cold for plating.	Always thoroughly mix Matrigel before aliquoting. Hold the tube of Matrigel and cell suspension in your hands for a few seconds to help it firm up before plating. Plate on glass instead of on plastic, if you can; Matrigel domes resist running, even when less viscous, when plated on glass coverslips. You can opt to quickly deposit the droplets of Matrigel and flip the plate so that the dome cultures dry upside down, forcing the formation of a dome shape through gravity.
Basic Protocol 2; Alternate Protocols 2.1, 2.2
Plating is too sparse.	You are diluting too much during passaging/seeding. Organoids are unhealthy.	Try to stay at a 1:2 or 1:3 ratio when splitting. Organoids tend to be healthiest in relatively dense cultures. Do not passage past ~P16.
Matrigel is too viscous.	Matrigel aliquot has higher than average protein content. Matrigel was subjected to too many freeze-thaw cycles. Matrigel is old. Matrigel has already reached room temperature.	Always thoroughly mix Matrigel before aliquoting. Thaw only once to aliquot and once for use. Keep at 4°C for a maximum of one week. Always keep Matrigel on ice until you are ready to plate it.
Basic Protocol 2
Organoids cluster at the center of the dish in sandwich culture.	Swirling after plating. Otherwise failing to distribute organoids evenly when plating (e.g., no movement).	Do not swirl the dish or plate. This will cause organoids to accumulate at the centers of wells. Do gently distribute organoids by moving the plate forward and backward (“y”) and side to side (“x”), as indicated in the protocol (NOT up and down, in “z”).
Entire top layer appears to be loose (“jiggly”).	5% Matrigel layer was left to incubate for too long. 100% Matrigel was too viscous when used.	Do not exceed the maximum incubation periods indicated in Basic Protocol 2. Do not subject Matrigel to multiple freeze-thaw cycles, storage at 4°C for longer than one week, and keep Matrigel ice cold to prevent premature hardening.
Center of well is detaching.	Matrigel is low in protein concentration.	Incubate Matrigel layers for longer durations.
Basic Protocol 3
Organoids do not survive more than a few hours during long-term time-lapse imaging.	If you are using a vital dye, the concentration may be too high. Phototoxicity as a result of camera/time-lapse settings. Incubator set-up has malfunctioned. Organoids were unhealthy at the start of the experiment.	Try reducing the concentration of vital dye, or otherwise conduct a dose response experiment. Laser power may be too high; for long-term time-lapse imaging, we recommend keeping laser power to 1.5% of its maximum value or less. Frequency of imaging may be too high for the organoids under the conditions you are imaging them (e.g., drug perturbation). Double check that the incubator set-up you are using is actually supplying carbon dioxide and heating the sample plate/dish. Adjust set-up as necessary. Use only healthy organoids (e.g., no visible signs of cell death beyond the normal luminal cellular debris; organoids have not been passaged beyond P16).
Basic Protocol 4
Organoids do not remain intact post-liberation.	Incubation in Cell Recovery Solution was too long. Containment vessels and pipet tips were not properly coated with 10% BSA. Pipetting was too vigorous.	Incubation duration may vary when liberating organoids from Matrigel, depending on the batch of Matrigel and the integrity of the organoid cultures. Observe the effects of dissociation every 5 minutes to gauge how long to incubate in order to liberate organoids without rupturing them. Ensure that ALL surfaces coming into contact with organoids except the 96-well glass-bottom dish are washed with 10% BSA prior to use. Once liberated from Matrigel, organoids will stick to surfaces that are not coated in 10% BSA. At each step, organoids should be handled with care, avoiding excessive shear stress and pressure during transfer and wash steps.
Basic Protocol 5
Single cells show low viability.	Incubation in TrypLE was too long. Cells were left on ice for too long. Organoids were unhealthy and undergoing apoptosis prior to dissociation.	Be sure to monitor incubation in TrypLE and cease incubation when organoids are dissociated into single cells.
Low numbers of overall cells (less than 5.5 ×106 cells/mL).	Too few organoids dissociated per sample.	Increase the number of organoids pooled for the run – you should have at least ~100 organoids per sample, or at least one densely cultured well of sandwich or dome cultured organoids.
Doublet rate is high/clumps of cells are visible after dissociation.	Organoids were not sufficiently dissociated into single cells.	Be sure to filter with a 40 μm pore size. Incubation in TrypLE was too short.
Cells fail to pass quality control testing during computational preprocessing.	Too few cells were sequenced (a lot of empty droplets). Cells were too unhealthy (e.g., high mitochondrial genes). Read depth was too low (too few genes).	Collect approximately 3000 cells per sample, anticipating that 50–70% may be lost during filtering steps. Keep cells on ice at all times after dissociation in 1% BSA in 1x PBS until they are suspended for single cell RNA-seq. Aim for at least 1×106 aligned reads per sample.