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

Summary

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

Abstract

The characterization of a large number of three-dimensional (3D) organotypic cultures (organoids) at different resolution scales is currently limited by standard imaging approaches. This protocol describes a way to prepare microfabricated organoid culture chips, which enable multiscale, 3D live imaging on a user-friendly instrument requiring minimal manipulations and capable of up to 300 organoids/h imaging throughput. These culture chips are compatible with both air and immersion objectives (air, water, oil, and silicone) and a wide range of common microscopes (e.g., spinning disk, point scanner confocal, wide field, and brightfield). Moreover, they can be used with light-sheet modalities such as the single-objective, single-plane illumination microscopy (SPIM) technology (soSPIM).

The protocol described here gives detailed steps for the preparation of the microfabricated culture chips and the culture and staining of organoids. Only a short length of time is required to become familiar with, and consumables and equipment can be easily found in normal biolabs. Here, the 3D imaging capabilities will be demonstrated only with commercial standard microscopes (e.g., spinning disk for 3D reconstruction and wide field microscopy for routine monitoring).

Introduction

In organotypic 3D cell cultures, hereafter referred to as organoids, stem cells differentiate and self-organize into spatial structures that share strong morphological and functional similarities with real organs. Organoids offer valuable models to study human biology and development outside the body^1,2,3. A growing number of models are being developed that mimic the liver, brain, kidney, lung, and many other organs^2,4,5. Differentiation in organoids is directed by the addition of soluble growth factors and an extracellular matrix in a precise time sequence. However, in marked contrast to organs, the development of organoids is quite heterogeneous.

Beyond numerous biological challenges^6,7, organoid cultures also pose technological challenges in terms of cell culture methods, characterization of transcriptomics, and imaging. In vivo organ development occurs in a biological environment that results in a highly stereotypical self-organization of cell arrangements. Any phenotypic alteration can be used as a proxy to diagnose a diseased state. In contrast, organoids develop in vitro in minimally controlled microenvironments compatible with cell culture conditions, resulting in large variability in the development path and shape formation for each individual organoid.

A recent study⁸ demonstrated that quantitative imaging of organoid shape (phenotype descriptors) coupled to the assessment of a few genetic markers allow the definition of phenotypic development landscapes. Arguably, the ability to relate the diversity of genomic expression in organoids with their phenotypic behavior is a major step toward unleashing the full potential of organotypic cultures. Thus, it begs for the development of dedicated, high-content imaging approaches allowing the characterization of organoid features at subcellular, multicellular, and whole-organoid scales in 3D^9,10.

We developed a versatile high-content screening (HCS) platform allowing streamlined organoid culture (from isolated human embryonic stem cells [hESCs], human induced pluripotent stem cells [hIPSCs], or primary cells to 3D, multicellular, differentiated organoids) and fast, non-invasive 3D imaging. It integrates a next-generation, miniaturized, 3D cell culture device, called the JeWells chip (the chip hereafter), that contains thousands of well-arrayed microwells flanked with 45° mirrors that allow fast, 3D, high-resolution imaging by single-objective light-sheet microscopy¹¹. Compatible with any standard, commercial, inverted microscope, this system enables the imaging of 300 organoids in 3D with subcellular resolution in <1 h.

The microfabrication of the cell culture device starts from an existing micro structured mold, which contains hundreds of micropyramids (Figure 1A) with a square base and sidewalls at 45° with respect to the base. Figure 1C shows electron microscope (EM) images of such structures. The mold itself is made of poly(dimethylsiloxane) (PDMS) and can be made as a replica cast of a primary mold (not shown here) with corresponding features (as cavities) using standard soft-lithographic procedures. The primary mold can be produced by different procedures. The one used for this protocol was made using silicon wet etching as reported in Galland et al.¹¹; the procedure for the fabrication of the primary mold is not critical for this protocol. The pyramids are arranged in a squared array, with the same pitch for the X and Y directions (in this case the pitch is 350 µm).

As an illustration, proof-of-concept experiments¹² were published to demonstrate that the chip allows long-term culture (months) and differentiation protocols while precisely defining the number of initial cells in the wells. Individual development of a large number of organoids can be automatically monitored live using standard brightfield and 3D light-sheet fluorescence microscopes. Moreover, organoids can be retrieved to perform further biological investigations (e.g., transcriptomic analysis). This paper outlines detailed protocols for the fabrication of the cell culture coverslips, the seeding and staining procedure for fluorescence microscopy, as well as the retrieval of the organoids.

Protocol

NOTE: The first part of this protocol details the microfabrication of the cell culture device. An original primary mold with pyramidal cavities can be produced in-house-if micro-fabrication facilities are available-or outsourced to external companies. The primary mold used in this work is produced in-house, with fabrication process steps described elsewhere^11,13. A basic protocol for the microfabrication of the mold is available in Supplementary File 1. CRITICAL: The operations in steps 1 to 6 need to be conducted in a dust-free environment. A laminar flow hood or a clean room, if available, are preferred. All through these steps, personal protective equipment (PPE) needs to be used, such as gloves, a lab coat, and safety glasses.

1. Dicing of PDMS mold

1. Cut small portions of the PDMS mold to the dimension required for the final device (e.g., 1 cm x 1 cm [Figure 1B]). Cut them parallel to the XY directions of the array.
   ​CRITICAL: When cutting the PDMS mold in 1 cm x 1 cm dices, execute the cuts in single steps; using a one-sided razorblade, apply pressure and cut through the PDMS in one single step. This is to prevent the formation of small particles of PDMS, which can deposit on the surface of the mold and affect the quality of the replica steps (step 3).

2. Preparation of flat PDMS substrates

1. Weigh ~15-20 g of PDMS base resin and 1.5-2 g of its reticulation agent (i.e., a ratio of 10:1), mix them carefully, and degas in a vacuum jar for ~20 min.
2. Slowly pour the degassed PDMS onto a plastic, 15 cm wide (diameter) Petri dish.
3. Cure the PDMS by keeping the Petri dish in an oven set at 65° C for 2 h. After waiting for the curing to complete, a flat PDMS film of ~1.5 mm thickness (contained in the Petri dish) is ready for use (Figure 2A).
4. Using a sharp blade, cut 2 cm x 2 cm pieces out of the PDMS (Figure 2B-D).
   ​NOTE: The exact thickness of this PDMS sheet is not critical; ~1-2 mm is a suitable range. The dimension for the cut should be bigger than the textured mold but not much larger, which would result in wasting of material.

3. Production of the textured layer made of UV-curable adhesive

1. Place one PDMS mold dice (as prepared in step 1) face-down on top of the flat PDMS cut (as prepared in step 2) (Figure 3A,B).
   NOTE: Make sure the pyramidal protrusions in the mold are oriented toward the flat PDMS substrate and that only the top surface of the truncated pyramids is in contact. Assess the correct placement with an optical microscope (Figure 3C,D).
2. To one side of the mold, using a pipette, drop a small amount (two drops, approximately 0.1-0.2 mL) of UV-curable adhesive (Figure 4A).
   NOTE: As the liquid gets in contact with the edge of the mold, capillarity will drive the liquid to fill the cavity between the mold itself and the flat cut of PDMS used as substrate.
   1. Follow the progression of the liquid inside the cavity, for example, using an inverted optical microscope with a 10x magnification objective (Figure 4B,C).
3. Expose the UV-curable adhesive to UV light to cure it. Adjust the time of exposure depending on the power density of the UV source used (e.g., a UV-LED box with a power density of 35 mW/cm²; 2 min at 50% power in this protocol to cure the adhesive completely).
4. Using the excess amount of adhesive at one edge, hold the cured adhesive on the flat PDMS substrate by gently pressing with a finger (Figure 5A,B). Meanwhile, use tweezers to pinch one corner of the mold next to the same edge being held down and slowly peel it off while making sure that the textured film is not lifted up as well.
   NOTE: Figure 5C shows the result of a proper mold removal procedure; Figure 5E-G shows the wrong procedure.
5. Trim the excess adhesive and the excess PDMS substrate using a razor blade to leave the cured, textured, adhesive layer flat on the PDMS with excess PDMS on one edge only (Figure 5D), which will be needed in step 5.

4. Coverslip substrate preparation

NOTE: As a substrate for the final device, standard rounded 1.5H coverslips with 25 mm diameter are used. Before they can be used, they need to be cleaned to remove dust and/or organic residual from their surface.

1. Coverslip cleaning
   1. Immerse the coverslips in soapy water for 5 min while the ultrasound is applied (40 kHz, 110 W sonic power).
   2. Wash the coverslips in clean deionized (DI) water, first by immersion and finally with running DI water from a tap. Dry the coverslips with an N₂ gas blow-gun.
   3. Immerse the coverslips in an acetone bath for 5 min; immediately move to a 2-propanol (IPA) bath for an additional 5 min.
   4. Rinse the coverslips with clean IPA using a squeeze bottle. Dry the coverslips with the N₂ gas blow-gun.
      NOTE: Coverslips cleaned using this procedure can be stored in a closed container until needed. Keep them in a dry cabinet to avoid humidity depositing on their surface.
2. When ready to be used, treat a clean coverslip with a short O₂ plasma process to improve its hydrophilicity: O₂ 20 sccm, 3 mbar pressure, 60 W at the RF power generator, and 60 s duration.
3. Immediately after the plasma activation, proceed with spin-coating the thin layer of UV-curable adhesive by placing the coverslip on the vacuum chuck of a standard spin-coater and pouring a small drop of adhesive at the center of the coverslip (Figure 6). Run the spin-coating process: spreading for 5 s at 500 rpm, coating at 3,000 rpm for 45 s (with the acceleration and deceleration set at 100 rpm/s).
   1. If spin-coating is not available, use the following alternative way to produce a thin film of UV adhesive on the coverslips:
      1. On a clean coverslip, drop ~0.1 mL of UV-curable adhesive using a pipette (Figure 7A).
      2. Take a second coverslip and place it on top of the first one to make the liquid adhesive spread evenly between the two coverslips (Figure 7B-D).
      3. Once the spreading adhesive has reached the edges of the coverslips, gently separate them by sliding one over the other. Once separated, both coverslips are fully coated with a thin layer of liquid adhesive (Figure 7E).
         NOTE: The coating might not be uniform and smooth only if the separation is not done with a smooth and continuous movement.
4. Precuring of UV-curable adhesive
   1. After spin-coating, precure the adhesive by exposure to UV. Adjust the time of exposure depending on the power density of the UV source used (here, a UV-LED box with a power density of 35 mW/cm² was used for 1 min at 50% power).
      CRITICAL: The adhesive used here is an optical glue. See the discussion for key points related to the energy doses for its curing.

5. Transfer of the textured film to the final substrate

1. Take one of the textured films (prepared in step 3) and place it in contact with the adhesive-coated coverslip (prepared in step 4). Make sure the contact between the partially cured adhesive on the coverslip and the textured film is as uniform as possible (Figure 8A-C).
   CRITICAL: At this stage, the adhesive on the coverslip should be solid enough to avoid reflowing, which would fill the pyramidal cavities of the textured film when placed in contact but also be plastic and adhesive enough that contact can be optimized by gently pressing on the textured film.
2. Expose the coverslips to UV light until the adhesive layer coated on the coverslip is fully cured. This will seal the textured film on the coverslip and provide leak-proof isolation between the pyramidal cavities.
3. Finally, peel off the PDMS flat substrate (Figure 8D-F). Using tweezers, pinch the PDMS on one corner at the edge where excess material was left after trimming (step 3.5). This way, the textured film layer is left adhesive to the coverslip with open access at the top for cell seeding.CRITICAL: When peeling off the flat PDMS, the textured film should remain well attached to the coverslip. Adhesion failures are easily confirmed if the textured film can be peeled off from the coverslip after final exposure to UV without any effort.

6. Long-term passivation of the cell culture coverslip for cell culture

NOTE: Passivation is achieved by generating a conformal and continuous coating of a biomimetic copolymer with a structure similar to the polar group of phospholipids in the cell membrane. This conformal coating prevents cells adhesion to the cell culture device

1. Prepare a solution with 0.5% (w/v) of the biomimetic copolymer dissolved in pure ethanol. Store the solution at 4 °C for future use.
2. Place the cell culture coverslip in a 35 mm Petri dish and fully cover it with the biomimetic copolymer solution.
3. After 5 min, remove the cell culture coverslip from the container with the biomimetic copolymer solution and leave it to dry at room temperature inside the final dish in a biosafety hood (>1 h).
   NOTE: A thicker coating can be produced by increasing the concentration of the biomimetic copolymer in the coating solution; the results of a thicker coating are visible under a brightfield microscope (Figure 9A).

7. Cell seeding

1. Degassing and sterilization
   1. Just before cell seeding, dispense sterile phosphate-buffered saline (PBS) into the cell culture dishes (typically 1 mL for a 35 mm Petri dish). Degas the dish with the sterile PBS using an ultrasonic device for ~10 min, followed by several rounds of pipetting to remove all the bubbles.
      CRITICAL: If air is trapped inside the pyramidal cavities, it will prevent the cells from entering them. To make sure there is no air trapped in the pyramid before cell seeding, it is recommended to visually ensure (under a benchtop brightfield microscope at 10x or 20x magnification) the absence of air in these cavities (Figure 10).
   2. Replace the PBS with sterile culture medium and sterilize the plate with UV light for 30 min under a cell culture hood.
      NOTE: From this step onward, the dish should be considered as sterile and manipulated using sterile techniques. An alternative way to remove trapped air from the cell culture device is to use a vacuum jar with a vacuum pump.
2. Cell suspension preparation
   NOTE: Cells can be seeded as single or small cell aggregates and enter the sample wells through the top aperture. Over time, the inserted cells aggregate and grow inside the sample wells into spheroids of a size greater than the size of the aperture. As validated cell line models, use HCT116 (CCL-247 ATCC) or MCF7 (HTB-22 ATCC) cancer cells maintained in recommended culture medium (ATCC guidelines).
   1. Prepare a cell suspension (e.g., using a trypsinization process following ATCC guidelines). Follow trypsinization/cell suspension preparation recommendations for the cells of interests.
   2. Count and adjust the cell concentration to 0.5 × 10⁶ cells/mL in the recommended culture medium.
3. Cell dispensing
   1. Remove the PBS buffer from the 35 mm cell culture dish and then dispense 1 mL of the adjusted cell suspension. See Figure 11A for an optical microscope image of a cell seeding procedure with adequate cell density and homogeneity.
   2. Place the cell culture dish back into the cell incubator (37 °C, 5% CO₂, and 100% humidity) for 10 min. Approximately 80-100 cells will enter each pyramidal cavity.
      NOTE: It is possible to increase the number of cells per cell culture dish by increasing either the cell concentration or the time spent before cell suspension removal. Typically, spheroid formation takes several hours (depending on cell type) after cell seeding and can be followed with a brightfield microscope (4x to 40x objectives; Figure 11). From here, culture medium, extracellular matrices, and differentiation growth factors can be changed or added to the cell culture dish containing the spheroids in accordance with typical differentiation protocols that could last a few days, weeks, or months.
   3. After the 10 min incubation, recover the cell culture dish from the incubator and gently aspirate the cellular suspension to remove untrapped cells. Add 1 mL of culture medium to a 35 mm dish and place it back into the cell incubator.
      CRITICAL: At this stage, because spheroids have not formed yet, it is very important to avoid strong aspiration or dispensing that will result in loss of the cells. Visual control using a benchtop brightfield microscope is highly recommended at this step.

8. Immunostaining and imaging

1. Fixation and staining
   NOTE: Different classical procedures of fixation and immunostaining are completely compatible with the cell culture dish. One typical protocol is described here.
   1. Fix the organoids/spheroids in the cell culture dish for 20 min in 4% paraformaldehyde at room temperature.
   2. Permeabilize the organoids for 24 h in 1% surfactant solution in sterile PBS at 4 °C on an orbital shaker and incubate for 24 h in blocking buffer (2% bovine serum albumin [BSA] and 1% surfactant in sterile PBS) at 4 °C on an orbital shaker.
   3. Incubate the samples with primary antibodies of interest at a dilution between 1/50 and 1/200 (or according to the manufacturer's recommendations) in antibody dilution buffer (2% BSA and 0.2% surfactant in sterile PBS) at 4 °C for 48 h.
   4. Rinse the samples 3x with washing buffer on an orbital shaker (3% NaCl and 0.2% surfactant in sterile PBS) and incubate with corresponding secondary antibodies in antibody dilution buffer (dilution between 1/100 and 1/300 or according to the manufacturer's recommendations), 0.5 µg/mL 4',6-diamidino-2-phenylindole (DAPI), and 0.2 µg/mL Alexa Fluor 647 or 488 Phalloidin at 4 °C for 24 h on an orbital shaker, followed by five rinsing steps with PBS. Optionally, mount the samples using a water-soluble clearing agent prewarmed to 37 ˚C.
2. Imaging
   NOTE: At this stage, the organoids in the microwell plate can be considered as a normal culture dish containing fixed and stained samples for imaging: any standard imaging procedure can be used with no adaptation or modification required. Figure 12 illustrates a representative result of images and 3D reconstruction obtained using a spinning disk confocal microscope, with a 40x air objective (numerical aperture 0.75).
   1. Use constructor software for automatic image acquisition process with the following settings: exposure time = 50 ms, with a z motorized stage to acquire a z-stack (1 µm Z-step, for a total height of 70 µm).
   2. Perform 3D reconstruction using image analysis software.

9. Release and collection of the organoids

NOTE: The textured adhesive layer of the cell culture dish can be detached from the coverslip to release the living spheroids/organoids (before fixation) contained inside the pyramidal cavities for analysis of the cells with other procedures such as RNA sequencing, -omic approaches, in vitro experiments, and in vivo transplantation.

1. With the sample still in the Petri dish and in a biosafety hood, use a blade such as a scalpel to cut a corner of the textured adhesive layer.
2. With tweezers, pinch the textured adhesive layer on the cut edge and gently peel it off from the glass coverslip but keep it immersed in the medium (some organoids might be remaining with the adhesive layer, Figure 13).
3. Rinse 3x with culture medium and collect the organoids by pipetting.

Results

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

Discussion

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

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
2-Propanol	Thermofisher scientific	AA19397K7
Acetone	Thermofisher scientific	AA19392K7
BC43 Benchtop Confocal Microscope	Andor Technology		spinning disk confocal microscope
bovine serum albumin	Thermofisher scientific	37525
Buffered oxide etching solution	Merck	901621-1L
CEE Spin Coater	Brewer Science	200X
DAPI	Thermofisher scientific	62248
Developer AZ400K	Merck	18441223164
DI Milliq water	Millipore
Fetal Bovine Serum (FBS)	Invitrogen	10082147
Glass coverslips	Marienfled	117650	1.5H, round 25 mm diameter
Hepes	Invitrogen	15630080
Imaris software	BitPlane		image analysis software
Inverted Transmission optical microscope	Nikon	TSF100-F
Labsonic M	Sartorius Stedium Biotech		Ultrasonic homogenizer
Lipidure	NOF America	CM5206	bio-mimetic copolymer
NOA73	Norland Products	17-345	UV curable adhesive
Penicillin-Streptomycin	Invitrogen	15070063
Phalloidin	Thermofisher scientific	A12379	Alexa Fluor
Phosphate Buffer Solution	Thermofisher scientific	10010023
Photo Resist AZ5214E	Merck	14744719710
Pico Plasma tool	Diener Electronic GmbH + Co. KG	Pico Plasma	For O2 plasma treatment
RapiClear 1.52	Sunjin lab	RC 152001	water-soluble clearing agent
RCT Hot Plate/Stirrer	IKA (MY)
Reactive Ion Etching tool	Samco Inc. (JPN)	RIE-10NR
RPMI 1640	Invitrogen	11875093	culture medium for HCT116 cells
Sylgard 184 Silicone Elastomer Kit	Dow Corning	4019862	Polydimethylsiloxane or in short, PDMS
Trichloro(1H,1H,2H,2H-perfluorooctyl)silane	Sigma Aldrich	448931-10G
Triton X-100	Sigma Aldrich	T9284	surfactant
Trypsin EDTA	Thermofisher scientific	15400054
Ultrasonic Cleaner	Bransonic	CPX2800
UV-KUB 2	KLOE		UV-LED light source, 365 nm wavelength, 35 mW/cm2 power density
UV mask aligner	SUSS Microtec Semiconductor (DE)	MJB4