This paper presents a fabrication protocol for a new type of culture substrate with hundreds of microcontainers per mm2, in which organoids can be cultured and observed using high-resolution microscopy. The cell seeding and immunostaining protocols are also detailed.
The characterization of a large number of three-dimensional (3D) organotypic cultures (organoids) at different resolution scales is currently limited by 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).
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 body1,2,3. A growing number of models are being developed that mimic the liver, brain, kidney, lung, and many other organs2,4,5. Differentiation in organoids is directed by the addition of soluble growth factors and an extracellular matrix in a precise time sequence. However, in marked contrast to organs, the development of organoids is quite heterogeneous.
Beyond numerous biological challenges6,7, organoid cultures also pose technological challenges in terms of cell culture methods, characterization of transcriptomics, and imaging. In vivo organ development occurs in a biological environment that results in a highly stereotypical self-organization of cell arrangements. Any phenotypic alteration can be used as a proxy to diagnose a diseased state. In contrast, organoids develop in vitro in minimally controlled microenvironments compatible with cell culture conditions, resulting in large variability in the development path and shape formation for each individual organoid.
A recent study8 demonstrated that quantitative imaging of organoid shape (phenotype descriptors) coupled to the assessment of a few genetic markers allow the definition of phenotypic development landscapes. Arguably, the ability to relate the diversity of genomic expression in organoids with their phenotypic behavior is a major step toward unleashing the full potential of organotypic cultures. Thus, it begs for the development of dedicated, high-content imaging approaches allowing the characterization of organoid features at subcellular, multicellular, and whole-organoid scales in 3D9,10.
We developed a versatile high-content screening (HCS) platform allowing streamlined organoid culture (from isolated human embryonic stem cells [hESCs], human induced pluripotent stem cells [hIPSCs], or primary cells to 3D, multicellular, differentiated organoids) and fast, non-invasive 3D imaging. It integrates a next-generation, miniaturized, 3D cell culture device, called the JeWells chip (the chip hereafter), that contains thousands of well-arrayed microwells flanked with 45° 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.
NOTE: The first part of this protocol details the microfabrication of the cell culture device. An original primary mold with pyramidal cavities can be produced in-house-if micro-fabrication facilities are available-or outsourced to external companies. The primary mold used in this work is produced in-house, with fabrication process steps described elsewhere11,13. A basic protocol for the microfabrication of the mold is available in Supplementary File 1. CRITICAL: The operations in steps 1 to 6 need to be conducted in a dust-free environment. A laminar flow hood or a clean room, if available, are preferred. All through these steps, personal protective equipment (PPE) needs to be used, such as gloves, a lab coat, and safety glasses.
1. Dicing of PDMS mold
2. Preparation of flat PDMS substrates
3. Production of the textured layer made of UV-curable adhesive
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.
5. Transfer of the textured film to the final substrate
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
7. Cell seeding
8. Immunostaining and imaging
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.
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 coverslip.
For the microwells to work, the textured film produced (as described in protocol step 3) needs to have both the top and the bottom sides of the pyramids open to ensure that cells during the seeding in step 7 can enter the cavities and remain contained once the organoids start forming. Figure 9 shows the increase in thickness of the coating by increasing the concentration of the biomimetic copolymer in the coating solution. Figure 10 shows the results of degassing the cell culture dishes to ensure no air is trapped in the pyramidal cavities. Figure 10A,B shows complete degassing, while Figure 10C,D shows air bubbles in the pyramidal cavities.
In Figure 11, cells are trapped inside the pyramids, and the corresponding organoids are formed after days 2 and 15 of culturing. If the top opening of the microwells is instead closed (as a consequence of incorrect step 3), no cells will be left after rinsing the sample after cell seeding. Figure 12 shows representative images of the organoids and a 3D reconstruction obtained using a spinning disk confocal microscope with a 40x air objective (numerical aperture 0.75). After release and collection, the organoids can be stored in a small vial and used for further analysis (e.g., RNA sequencing). Figure 13B shows an example of a sample of organoids collected as described.
Figure 1: Poly(dimethylsiloxane) mold. (A) The starting PDMS mold obtained as a cast replica of a 4" textured wafer (see Supplementary File 1). (B) 1 cm x 1 cm wide cut of the starting PDMS mold. (C) Scanning electron microscope images of the trapezoidal pillars arranged in a squared array, which populate the surface of the mold. Scale bars = 1 cm (A), 100 µm, and 200 µm (C top and bottom, respectively). Abbreviation: PDMS = poly(dimethysiloxane). Please click here to view a larger version of this figure.
Figure 2: Flat PDMS substrate. (A) A layer of PDMS, ~1 mm thick, is prepared in a standard 15 cm wide Petri dish. (B) Peeling out of a fresh cut of PDMS. (C) The remaining PDMS on the Petri dish can be cut into more pieces for further usage. (D) Flat PDMS cut and PDMS mold side by side; the flat PDMS is bigger than the mold. Abbreviation: PDMS = poly(dimethysiloxane). Please click here to view a larger version of this figure.
Figure 3: Placing the mold on the substrate. (A) The mold is placed on the flat substrate with the pyramids facing down. (B) Different appearance of poor contact (top) and good contact (bottom) between the mold and PDMS substrate. (C) Optical microscope image of an area with poor contact; the red circles highlight the pillars not in contact with the substrate-they appear of a brighter color than the ones in contact in the same image. (D) Optical image of an area with all pillars in good contact. Scale bars = 200 µm (C,D). Abbreviation: PDMS = poly(dimethysiloxane). Please click here to view a larger version of this figure.
Figure 4: Capillary filling of UV-curable adhesive. (A) This sequence of images shows (from left to right) the pouring of a small quantity of adhesive on one edge and the progression of the liquid adhesive in the cavity between the mold and substrate. The yellow arrows point at the direction of the liquid flow. (B) Sequence of optical images (40x magnification) of the liquid front moving; when in good contact, the front of the liquid moves around the edges of the pyramids without leaks. The yellow arrows point at the flow direction and the red arrows point at the leading edge of the liquid front moving around the contact edges. (C) Sequence of optical images (40x) of the liquid front moving when the contact is poor; the red arrows point at the leading edge of the liquid infiltrating between the mold and the substrate. Please click here to view a larger version of this figure.
Figure 5: Removal of the mold after adhesive curing. (A–C) Correct procedure for peeling off the mold: while gently pressing on the excess of adhesive at the left edges with tweezers, the mold is pinched near the same edge and slowly bent to peel off. (D) The result of the correct procedure; the excess is trimmed out on three edges, while a small excess of PDMS is left on the fourth side (yellow arrow). (E–G) Example of an incorrect procedure for removing the mold: the mold is pinched with tweezers from the opposite edge, resulting in peeling off from the flat substrate of the adhesive film together with the mold. Please click here to view a larger version of this figure.
Figure 6: Coating of the glass coverslip. (A,B) Example of a good coating procedure: the glass coverslip is placed on the vacuum chuck of a spin-coater, and enough adhesive is dispensed at the center, giving a homogeneous coating of the coverslip. (C,D) Example of a bad coating procedure: not enough adhesive is dispensed, resulting in non-uniform coating of the coverslip. (E) From left to right, examples of good, acceptable, and bad coating. Please click here to view a larger version of this figure.
Figure 7: Alternative coating process. (A) A small drop of liquid adhesive is placed at the center of the coverslip. (B) A second coverslip is gently placed on top of the first one. (C,D) The liquid spread between the two coverslips and evenly distributed. (E) Examples of results after correct separation of the coverslips (left) or incorrect separation (right). Please click here to view a larger version of this figure.
Figure 8: Transfer to the coverslip. (A) The cured and trimmed adhesive film (Figure 5D) is placed on the coverslip with partially cured adhesive. (B) Example of good contact between the textured film and the coverslip. The inset is an optical image showing uniform contact of the areas between the trapezoidal cavities (dark uniform color). (C) Example of poor contact between the textured film and the coverslip. The brighter areas are not in contact, the red arrows point at those areas. The inset is an optical image showing the different appearance of good contact (two on the left) and poor contact (two on the right). Scale bars (yellow, B,C) = 200 µm. (D,E) Correct procedure for removing the PDMS flat substrate: tweezers are used to pinch the PDMS at the edge with excess of PDMS and bend to gently peel off. (F) The result with the UV adhesive-textured film successfully transferred to the coverslip. (G,H) Example of incorrect peeling procedure: tweezers are used to pinch the flat PDMS on one of the edges where the excess material was trimmed-off; thus, the film is removed together with the flat PDMS. Abbreviation: PDMS = poly(dimethysiloxane). Please click here to view a larger version of this figure.
Figure 9: Coating with biomimetic copolymer. (A) Cell culture device coated with biomimetic copolymer using a solution of 0.5% (left) or (B) 1% (right) (w/v) in pure ethanol. Scale bar = 100 µm. Please click here to view a larger version of this figure.
Figure 10: Degassing of cell culture device. (A) Cell culture device before complete degassing. (B) Cell culture device after complete degassing; (C,D) cell culture device with only partial degassing where air bubbles are trapped in the pyramids. Scale bars = 200 µm (A–D). Please click here to view a larger version of this figure.
Figure 11: Representative brightfield images of cell seeding and growth of organoids. (A) Cells are visible floating on the top. (B) Cells trapped in the pyramidal cavities after rinsing away the cell suspension. Only trapped cells will be retained as shown. (C) Cells aggregate to form spheroids in each cavity (day 2 after cell seeding). (D) Image at day 15 after cell seeding of an organoid that has grown in a pyramidal cavity. Scale bars = 200 µm (A B), 120 µm (C), and 150 µm (D). Please click here to view a larger version of this figure.
Figure 12: 3D imaging of organoids. (A) Three different Z plans acquired using a spinning disk confocal (lens 40x, air, numerical aperture: 0.75) of an organoid under neuroectoderm differentiation (day 7after cell seeding). Actin staining using phalloidin (Alexa Fluor 647) in orange and N-cadherin immunostaining (Alexa Fluor 561) in blue. (B) A 3D reconstruction using image analysis software of the corresponding organoids (80 Z plans, total height of 100 µm). White arrows: clusters of cells around a streak with high level of actin expression. Blue arrows: cell clusters positive for N-cadherin protein expression. Please click here to view a larger version of this figure.
Figure 13: Release of organoids. (A) Removing the textured adhesive layer with tweezers; (B) brightfield image of an organoid suspension obtained after the removal. Scale bar = 200 µm (B). Please click here to view a larger version of this figure.
Supplementary File 1: A step-by-step protocol for the microfabrication of a Si mold with pyramidal microcavities is presented. Please click here to download this File.
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.
The regular distribution of the microcavities and the flat standard coverslip used as the substrate enable the monitoring of thousands of live or fixed organoids in 3D, without the need for complex and time-consuming sample manipulation between culturing vessels and imaging supports. Due to the pyramid shape with a smaller top than the base, there is no loss of organoids due to pipetting steps during the medium exchange, dispensing of extracellular matrices, or staining procedures. All these steps could be performed directly, as for a 2D cell monolayer without any extra precautions, unlike classical organoid generation techniques (e.g., hanging droplets, low attachment plates, Aggrewell).
Moreover, compared to these existing techniques, the microwells have been designed for high-resolution 3D imaging compatible with all types of immersion lenses (air, water, oil, and silicon). Long-term maintenance of organoids closed to a flat glass coverslip makes the cell culture coverslip compatible with high-resolution light microscopes, which is impossible with the existing methods without complex handling and fastidious organoid transfer. Further, an optimized passivation procedure avoids any adhesion of cells/organoids for long-term culture, thus controlling the differentiation of the 3D culture Finally, since the textured adhesive layer is removable, living organoids can be retrieved to perform other types of experiments such as -omics assays or in vivo transplantation.
It should be noted that the pyramidal shape and array arrangement of the microcavities used for this protocol are derived from the primary mold, which has been fabricated by means of silicon wet-etching to provide surfaces with mirror-like finishing and 45° inclination to allow for single-objective light-sheet microscopy (soSPIM11). While a discussion of these requirements and a full description on how to produce such molds can be found elsewhere12,13, a simple protocol is also given in Supplementary File 1. Different ways of producing a primary mold are available, which are fully compatible with the fabrication of the chips in this protocol. Laser etching of glass and high-resolution 3D printing, for example, could be used to produce a primary mold with suitable cavities for the containment of the organoids, but are not suitable for soSPIM imaging.
The fabrication protocol disclosed here can be adapted for use in any standard biological laboratory, as no dedicated, advanced microfabrication tools are required. Some critical steps of the fabrication of the microwells are the production through capillary filling of the textured adhesive film and its transfer to the glass substrate. However, in our experience, they can be mastered with high reproducibility in a short period of time. The distance between the pyramids should be as small as possible to increase the density of cavities for the growth of organoids, but it also needs to be large enough so that the cavities can be sealed on the substrate without any leak or lack of sealing between them. We found that a distance of 50 µm is a good compromise for this case.
During cell seeding, a critical aspect is the removal of any air bubbles trapped inside the cavities to ensure cell entry. We describe here a validated solution, which uses only classical laboratory equipment (e.g., an ultrasonic homogenizer or a vacuum jar). To allow better homogenization of the number of cells per cell culture dish, it is recommended to dispense the cell suspension from the side of the dish and not directly above the coverslip. Gentle manual agitation could also help to homogenize the cellular solution.
After the seeding of cells, the cell culture dish with organoids can be treated like any other classical culture support; no specific manipulation or critical steps are required. All the protocols that have been used with the microwell devices are practically the same as those used in low-attachment dishes such as U-bottom plates. Hence, organoid culture in these microwells does not require any change in culture conditions compared to other standards.
Another critical factor is that the adhesive used here is an optical glue. In its suggested best usage (also refer to application notes from the vendor), two optical elements to be glued are aligned and UV-curable adhesive is applied at the interface. A first exposure to UV of an energy insufficient to cause full polymerization is used to solidify the glue while maintaining adhesive properties. The optical components can be moved to achieve optimal alignment, and then the glue is fully cured to its final state with a second exposure to UV to provide permanent adhesion. The first and second exposure energy doses (i.e., exposure times if a source of known and constant energy density is used) must be optimized depending on the energy density of the UV source used, the thickness of the layer of glue, the transmission of UV through the optical elements, and the surface texturing (or lack of it) of the surface to be glued.
While the adhesion between the textured film and the adhesive-coated coverslip is required to be strong enough to provide for leak-proof sealing, it also needs to be possible to remove the textured film to retrieve the organoids after imaging, if any further analysis is required, such as genotypic profiling. A correct compromise between leak-proof sealing and the capability to peel off the textured film has been achieved with the protocol described here, but further optimization of the precuring and final curing times for the adhesive thin layer on the coverslip might be required as it depends on the UV-exposure system and film thickness used.
One limitation in the current microwells version is in the seeding procedure; cells must be seeded before the ECM components to allow them to enter the vicinity of the pyramid. Improvements are ongoing to coat the microwells with extracellular matrix components, as a first step. We have not yet performed any electron microscopy (EM) on the fixed organoids within the cavities. Some modification to these fabrication and cell culture protocols will be required before imaging of organoids in the microwells with EM becomes a viable option.
A natural future extension of this method is to provide high-content screening capacities to allow multicondition testing in a single workflow (multiwell plate). This cell culture device provides a unique alternative to existing organoid techniques, offering unsurpassed culturing and imaging throughput and opening new perspectives in the field of organoid research for biomedical application and drug discovery.
The authors have nothing to disclose.
The research is supported by the CALIPSO project supported by the National Research Foundation, Prime Minister's Office, Singapore, under its Campus for Research Excellence and Technological Enterprise (CREATE) programme. V.V. acknowledges the support of NRF investigator NRF-NRFI2018-07, MOE Tier 3 MOE2016-T3-1-005, MBI seed funding, and ANR ADGastrulo. A.B. and G.G. acknowledge the support from MBI core funding. A.B. acknowledges Andor Technologies for the loan of the BC43 microscope.
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 |