Here, we present a protocol that enables fast, robust, and cheap fabrication of tumor spheroids followed by hydrogel encapsulation. It is widely applicable as it does not require specialized equipment. It would be particularly useful for exploring spheroid-matrix interactions and building in vitro tissue physiology or pathology models.
Three-dimensional (3D) encapsulation of spheroids is crucial to adequately replicate the tumor microenvironment for optimal cell growth. Here, we designed an in vitro 3D glioblastoma model for spheroid encapsulation to mimic the tumor extracellular microenvironment. First, we formed square pyramidal microwell molds using polydimethylsiloxane. These microwell molds were then used to fabricate tumor spheroids with tightly controlled sizes from 50-500 μm. Once spheroids were formed, they were harvested and encapsulated in polyethylene glycol (PEG)-based hydrogels. PEG hydrogels are a versatile platform for spheroid encapsulation, as hydrogel properties such as stiffness, degradability, and cell adhesiveness can be tuned independently. Here, we used a representative soft (~8 kPa) hydrogel to encapsulate glioblastoma spheroids. Finally, a method to stain and image spheroids was developed to obtain high-quality images via confocal microscopy. Due to the dense spheroid core and relatively sparse periphery, imaging can be difficult, but using a clearing solution and confocal optical sectioning helps alleviate these imaging difficulties. In summary, we show a method to fabricate uniform spheroids, encapsulate them in PEG hydrogels and perform confocal microscopy on the encapsulated spheroids to study spheroid growth and various cell-matrix interactions.
Tumor spheroids have emerged as useful in vitro tools in studying cancer etiology, pathology, and drug responsiveness1. Traditionally, spheroids have been cultured in conditions such as low adhesion plates or bioreactors, where cell-cell adhesion is favored over cell-surface adhesion2. However, it is now recognized that to recapitulate the tumor microenvironment more faithfully, in vitro spheroid models should capture both cell-cell and cell-matrix interactions. This has prompted multiple groups to design scaffolds, such as hydrogels, where spheroids can be encapsulated3,4. Such hydrogel-based spheroid models enable the elucidation of cell-cell and cell-matrix interactions on various cell behaviors, such as viability, proliferation, stemness, or therapy responsiveness3.
Here, we describe a protocol for the encapsulation of glioblastoma spheroids in polyethylene glycol (PEG) hydrogels. There are multiple literature reports of glioblastoma cell spheroid encapsulation in hydrogels. For example, spheroids were formed by encapsulating U87 cells in PEG hydrogels decorated with an RGDS adhesive ligand and crosslinked with an enzymatically cleavable peptide to determine the effect of hydrogel stiffness on cell behavior5. U87 cells have also been formed in other PEG-based or hyaluronic acid-based hydrogels to expand the cancer stem cell population6 or to explore matrix-mediated mechanisms of chemotherapy resistance7,8,9. Glioblastoma spheroids have also been encapsulated in gelatin hydrogels to study the crosstalk between microglia and cancer cells and its effect on cell invasion10. Overall, such studies have demonstrated the utility of hydrogel-based in vitro models in understanding glioblastoma pathology and devising treatments.
Further, there are different methods for tumor spheroid fabrication and hydrogel encapsulation11. For example, dispersed cells could be seeded in hydrogels and allowed to form spheroids over time5,12. One drawback of such a method is the polydispersity of the formed spheroids, which could lead to differential cell responses. To produce uniform spheroids, cells could be encapsulated in microgels and cultured for extended periods until they invade and remodel the gel13, or cells could be deposited in templated gels with spherical 'holes' and allowed to aggregate14. The drawback of these methods is their relative complexity, the need for a droplet generator or other means to form microgels or the 'holes' in the gel, and the time it takes for spheroids to grow and mature. Alternatively, spheroids could be pre-formed in microwells9,15,16 or in hanging-drop plates17,18 and then encapsulated in a hydrogel, similar to the technique described here. These methods are simpler and can be done in a higher throughput fashion. Interestingly, it has been shown that the method of spheroid formation can affect spheroid cell behaviors, such as gene expression, cell proliferation, or drug responsiveness19,20.
Here, we focus on glioblastoma since it is a solid tumor whose native environment is the soft, nanoporous brain matrix21, which can be mimicked by a soft, nanoporous hydrogel. Glioblastoma is also the deadliest brain cancer for which there is no available cure22. However, the protocol described here can be used for the encapsulation of spheroids representing any solid tumor. We chose to use PEG hydrogels that are formed through a Michael-type addition reaction23. PEG is a synthetic, non-degradable, and biocompatible hydrogel that is inert and serves as scaffolding and physical cell support but does not support cell attachment23. Cell adhesiveness can be added separately via tethering of whole proteins or adhesive ligands24, and degradability can be added via chemical modifications of the PEG polymer chain or hydrolytically or enzymatically degradable crosslinkers25,26. This allows for biochemical properties to be tuned independently of mechanical or physical hydrogel properties, which could be advantageous in studying cell-matrix interactions. The Michael-type gelation chemistry is selective and happens at physiological conditions; hence, it allows for spheroid encapsulation by simply mixing the spheroids with the hydrogel precursor solution.
Overall, the methodology presented here has several notable characteristics. First, fabricating tumor spheroids in a multiwell assembly is efficient, quick, and the cost of the required materials is low. Second, the spheroids are produced in large batches in a variety of sizes with low polydispersity. Finally, only commercially available materials are required. The utility of the methodology is illustrated by exploring the effect of substrate properties on spheroid cell viability, circularity, and cell stemness.
1. Solutions preparation
2. Fabrication of square pyramidal microwells
3. Multicellular tumor spheroid formation, harvest, and encapsulation in hydrogels
NOTE: The protocol outlined in this section is for U87 human glioblastoma cell line (see Figure 1 and Figure 2), but a similar protocol could be used with other cancer cell types.
4. Fluorescent staining
5. Immunofluorescence fixation, staining, clearing, and imaging of encapsulated spheroids
Spheroid-based drug screening platforms to study chemotherapeutic effects are increasingly sought after due to the emphasis on modulating the tumor microenvironment upon spheroid encapsulation in biomaterials replicating native tissue. Here we developed a method for multicellular tumor spheroid preparation and subsequent encapsulation and imaging in a 3D hydrogel. The spheroids are prepared in microwell molds (Figure 3A,B), which result in spheroids with spherical shapes and tightly controlled polydispersity. For example, for spheroids with 3,300 cells per microwell, the average spheroid size was ~250 µm, and circularity was >0.8, where 1 is a perfect sphere (Figure 3C,D). The percent coefficient of variance (%CV) for spheroid diameter was 19.3%, and %CV for circularity was 4.5%. Spheroid diameters were dependent on the number of cells per microwell, as shown in Table 1. Note that some cells in the microwells might not be incorporated in the spheroid and will be washed off during the spheroid harvesting step.
The spheroid is able to be imaged at varying Z-stack depths, allowing for cell viability for each location within the Z-stack (Figure 4). Note that due to confocal imaging limitations and the high cell density, the spheroid core could not be imaged fully. As discussed later, spheroid clearing or sectioning might be needed for improved imaging throughout the spheroid. Through this method, the spheroid viability was determined to be high (~90%) immediately post-harvest and encapsulation, even though cell viability dipped slightly to 85% in the spheroid core compared to the periphery. To ensure viability throughout the spheroid, spheroids were dissociated using acutase, and the viability of the dissociated cells was calculated using the same method and found to be equally high (>90%). A max projection utilizing the spheroid stack represents the highest point of light intensity within each location of the Z-stack compressed into one image (Figure 4B).
Representative images of free-floating spheroids (no gel) and encapsulated spheroids (PEG gel) stained for stemness markers Nestin and SOX2 are shown in Figure 5. Nestin is an intermediate filament and a stem cell marker present in gliomas. SOX2 is a transcription factor for self-renewal, also present in gliomas. SOX2 was found to be co-localized with DAPI in the nucleus, while Nestin was present throughout the cells. The data shows no difference between the gel and no gel conditions, possibly due to the PEG gel used here being inert and not facilitating cell-matrix interactions through integrin signaling.
A major limitation of imaging is the high spheroid density, making it difficult to image into the spheroid core. Common methods to image the spheroid core include sectioning and clearing the sections. This can work well with free-floating spheroids, but sectioning hydrogels is difficult, and most tissue clearing involves dehydrating samples, which results in deformed samples. Here we adapted a protocol from Kuwajima et al.28 for hydrogel samples to maintain the spheroid structure while still clearing the spheroids. To demonstrate the utility of the clearing method, spheroids were fixed, stained with PI, and cleared (Figure 6). Figure 6A shows representative images of optical sections at ~90 µm into cleared and uncleared spheroids and the total area of the spheroid when the spheroid area is filled. When the clearing was performed, the spheroid core could be imaged ~30 µm deeper, compared to an uncleared spheroid (Figure 6B). Clearing can be very beneficial for immunofluorescence, as the spatial organization of biomarkers can be analyzed into the core of spheroids. This method can only be used for fixed samples as the membrane integrity is disrupted.
Figure 1: PDMS mold fabrication and spheroid formation. PDMS precursor solutions are added to square pyramidal microwell plates to form PDMS molds. The dissociated cell suspension is added to the formed PMDS molds and spun down to allow for spheroid formation after 24 h. Please click here to view a larger version of this figure.
Figure 2: Hydrogel encapsulation, culturing, and imaging. (A) 4-arm PEG-Ac, PEG diSH, and the spheroid suspension is combined in a microcentrifuge tube and (B) mixed well by pipetting up and down to form a gel precursor solution. (C) 20 µL of the gel precursor solution is pipetted onto a glass slide, and a second glass slide is placed on top of the solution and separated by 1 mm spacers. (D) Hydrogel is transferred to 24 well plates with spheroids facing up and is covered with media (500 µL). (E) Hydrogel is transferred to a glass coverslip for microscopy imaging. Please click here to view a larger version of this figure.
Figure 3: Spheroid formation in microwells prior to harvest, 24 h after initial cell seeding in the microwells. (A) Spheroids in microwell stained with DiOC (Green). Scale bar is 200 µm. (B) Brightfield image of spheroids in microwells. (C) Histogram of spheroid diameter in microwells. (D) Histogram of spheroid circularity in microwells. Please click here to view a larger version of this figure.
Figure 4: Representative Z-stack confocal image of live spheroid for live dead analysis. (A) Images from 4 slices of Z-stack with DiOC (green) and PI (red). Scale bar is 200 µm. (B) Max projection of Z-Stack. (C) Cell viability as a function of spheroid depth. Please click here to view a larger version of this figure.
Figure 5: Nestin and SOX2 immunostaining of free-floating (no gel) and hydrogel-encapsulated (PEG gel) U87 cell spheroids at day 5 of culture. Representative confocal images of the spheroids confirm Nestin (red) and SOX2 (green) expression, which were counterstained with DAPI (blue; nucleus). Scale bar is 100 µm. Images cropped and zoomed from the white square show the relationship between the nucleus (blue) and SOX2 (green). Please click here to view a larger version of this figure.
Figure 6: Imaging depth of cleared spheroids. (A) Representative images of spheroids imaged at 91.6 µm into the spheroid and total spheroid area. (B) Percentage of the spheroid area imaged as depth into spheroid increases. Please click here to view a larger version of this figure.
Microwell Size (µm) | Number of Cells per Spheroid | Number of Cells per Well | Cell Concentration (cells/mL) | Spheroid Diameter (µm) |
400 | 200 | 120000 | 240000 | 115.4 ± 13.5 |
500 | 300000 | 600000 | 144.6 ± 14.3 | |
1000 | 600000 | 1200000 | 176.5 ± 12.5 | |
800 | 2000 | 300000 | 600000 | 212.4 ± 15.7 |
3000 | 450000 | 900000 | 258.9 ± 16.3 | |
4000 | 600000 | 1200000 | 305.7 ± 21.6 | |
5000 | 750000 | 1500000 | 323.4 ± 29.8 |
Table 1: Speroid diameter and microwell size. The calculated number of cells per spheroid, number of cells per well of a 48-well plate, cell concentration, and resulting spheroid diameter depending on the microwell size of the negative PDMS mold.
Hydrogel-based multicellular tumor spheroid models are increasingly being developed to advance cancer therapeutic discoveries11,13,29. They are beneficial because they emulate key parameters of the tumor microenvironment in a controlled manner and, despite their complexity, are simpler and cheaper to use than in vivo models, and many are compatible with high-throughput screening technologies. The hydrogel biomaterials can be tuned to emulate the tumor extracellular matrix and facilitate cell-matrix interactions, and the spheroid (as opposed to dissociated cells) emulates the cell-cell interactions of the native tumor. Synthetic hydrogels, as demonstrated here, are particularly advantageous because the hydrogel provides the desired structural support and physical and mechanical properties, while adhesive ligands or degradable sequences can be used to independently tune biochemical material properties. Synthetic hydrogels also offer lower batch-to-batch variability and greater ranges of material properties, encompassing most soft tissues in the body.
Here, we describe in detail the use of an inert PEG-based hydrogel that forms via Michael-type addition as described previously25,30. The representative PEG hydrogel used here has Young's modulus of ~8 kPa, similar to glioblastoma tissue stiffness9. The PEG hydrogel is convenient as it has tunable properties and highly specific gelation chemistry and can be formed in the presence of spheroids without compromising cell viability (Figure 4). While the PEG hydrogel is inert and serves as cell scaffolding with defined physical and mechanical properties, cell adhesive ligands can be added to guide cell-matrix interactions, and enzymatically degradable peptide crosslinkers can be added to aid matrix remodeling9. Adhesive ligands and peptide crosslinkers could be selected to emulate the cellular microenvironment to add physiological or pathological relevance to the model. For more details on PEG hydrogel modifications to tune hydrogel degradability, mechanical properties, and adhesiveness, readers are referred to the following published work9,27.
Using the method described here, large quantities of cancer spheroids can be formed quickly and encapsulated in the PEG hydrogel to explore the effect of substrate properties on spheroid cell viability, morphology, or cell stemness (Figure 4 and Figure 5), among other properties. The cells are first allowed to aggregate and form a spheroid and then encapsulated in the hydrogels, as shown in Figure 2 and Figure 3. The aggregation process enables varying the spheroid sizes based on the size of the microwell molds and the concentration of the cell suspension pipetted into the microwell molds (Table 1). The resultant spheroids are relatively monodisperse in size, with an average coefficient of variance of ~10%-20% for all conditions. This is beneficial for a variety of applications, as similar sizes will exhibit similar diffusion limitations, be it of drugs, nutrients, or oxygen, into the spheroid. The spheroids also have a circularity of >0.8, which is comparable to other ultra-low attachment or hanging drop methods31. Note that other methods for spheroid formation can be used first, such as the hanging drop method or rotary cell culture system32, and then encapsulated in the hydrogel. However, the method described here does not require any specialized equipment or expensive consumables, hence, aiding in accessibility for all labs.
While the methods shown here use the U87-MG glioblastoma cell line, the spheroid fabrication and encapsulation method described can be used for various cancer cell types that form solid tumors. If the cells do not readily aggregate to form a spheroid, a mixture of ECM proteins, such as a basement membrane matrix, can be added to the cell suspension to aid the process (as described in the methods). Once spheroids are encapsulated, it is best to be imaged and analyzed directly in the hydrogel instead of being extracted from the hydrogel or the cells being dissociated. This is because spheroids are typically heterogeneous (e.g., a hypoxic core could form due to oxygen and nutrient gradients), and cell responses will differ based on the position within the spheroid. Also, cell extraction might obscure the role of cell-matrix interactions on spheroid fate. For example, we have previously shown that cells in the spheroid periphery versus core have different responsiveness to chemotherapeutics, which is further dependent on the mechanical properties of the substrate27. Hence, we recommend using microscopy techniques for the study of spheroid behaviors, as highlighted in this manuscript.
One issue to consider when imaging dense tissues such as spheroids is their opacity. Here we describe a clearing method to improve imageability through the interior of the spheroid (Figure 6). However, even though spheroid clearing aids in maximizing imaging depth, limitations can be found when placing the encapsulated spheroids in formamide resulting in potential structural damage and affecting spheroid imageability. This process also cannot be performed when observing cell viability through live/dead staining because clearing requires fixation. The clearing process also requires several hours of incubation in formamide following staining, so it could potentially impact the dyes being used. Other techniques, such as cryosectioning and then immunostaining, could also work, provided that the sectioning does not distort or compromise spheroid tissue integrity. In our hands, cryosectioning resulted in "squished" spheroids due to the softness of the hydrogel, which is ~8 kPa in Young's modulus, to emulate glioblastoma tissue stiffness.
Overall, the critical steps in this protocol are successful spheroid fabrication, hydrogel encapsulation and culture, and spheroid imaging and analysis; hence, we have provided notes and troubleshooting strategies for those steps. The hydrogel-encapsulated spheroids described here could be used in a variety of applications, such as drug screening platforms, detailed studies of cell-matrix interactions and their effect on cell behaviors, studies of disease etiology, etc. Such studies can be aided by the tunability of the described system as discussed above and the predictable and controllable properties of the synthetic PEG hydrogel. Some limitations of the described system include a medium throughput, where high throughput is preferable for multiplex or high-volume studies such as drug screening. Another limitation is the need for imaging, such as confocal imaging, for data analysis. While imaging allows for detailed special and temporal analysis, it is also time-consuming and hindered by penetration limitations due to depth and spheroid cell density.
The authors have nothing to disclose.
This work was funded by start-up funds provided to Dr. Silviya P Zustiak by Saint Louis University as well as by a seed grant from the Henry and Amelia Nasrallah Center for Neuroscience at Saint Louis University awarded to Dr. Silviya P Zustiak.
70% Ethanol | Fisher Scientific | LC22210-4 | |
15 mL Conicals | FALCON | 352097 | |
24-Well Plate Ultra Low Attachment plates | Fisher Scientific | 07-200-602 | |
35 mm Petri Dish | Amazon | 706011 | |
4-arm poly(ethylene glycol)-acrylate (4-arm PEG-Ac; 10 kDa) | Laysan Bio | ACRL-PEG-ACRL-10K-5g | |
50 mL Conicals | Fisher Scinetific | 3181345107 | |
6-well AggreWell 400 | StemCell Technologies, Vancouver, Canada | 34421 | Square pyramidal microwells |
anti-adherence rinsing solution | StemCell Technologies, Vancouver, Canada | Cat #: 07010 | |
Aspartic Acid-Arginine-Cysteine-Glycine-Valine-Proline-Methionine-Serine-Methionine-Arginine-Glycine-Cysteine-Arginine- Aspartic Acid (DRCG-VPMSMR-GCRD) peptide | Genic Bio, Shanghai, China | n/a | Custom synthesis |
Chemical Fume Hood | KEWAUNEE | 99151 | |
Corning Matrigel Basement Membrane Matrix, LDEV Free | Corning | 356234 | Basement membrane matrix |
DAPI (4',6-diamidino-2-phenylindole, dihydrochloride) | Thermo Scientific | 62247 | |
Detergent – Triton-X | Sigma Aldrich | T8787 | Nonionic surfactant |
Dimethyl sulfoxide (DMSO) | Fisher Scientific | BP231-100 | |
Disposable Pipettes (1 mL, 2 mL, 5 mL, 10 mL, 25 mL, 50 mL) | Fisher Scinetific | 1 mL: 13-678-11B, 2mL: 05214038, 5mL(FALCON): 357529, 10mL: 13-678-11E, 25mL: 13-678-11, 50mL: 13-678-11F | |
Fetal Bovine Serum | HyClone | SH30073-03 | |
Formaldehyde 37% Solution | Sigma Aldrich | F1635 | |
Glass Plates | Slumpys | GBS4100SFSL | |
Glass Transfer Pipettes | Fisher Scinetific | 5 3/4": 1367820A, 9":136786B | |
Glycine-Arginine-Cysteine-Aspartic Acid-Arginine-Glycine-Aspartic Acid-Serine (GRCD-RGDS) peptide | Genic Bio, Shanghai, China | n/a | Custom synthesis |
Hemacytometer | Bright-Line | 383684 | |
Hydrophobic solution – Repel Silane | GE Healthcare Bio-Sciences | 17-1332-01 | |
Incubator | NUAIRE | NU-8500 | |
Inverted Microscope (Axiovert 25) | Zeiss | 663526 | |
Invitrogen DiOC16(3) (3,3'-Dihexadecyloxacarbocyanine Perchlorate) | Fisher Scientific | D1125 | |
Leica Confocal SP8 | Leica Microsystems Inc. | ||
Light and Flourescent Microscope (Axiovert 200M) | Zeiss | 3820005619 | |
Micro centrifuge tubes | Fisher Scientific | 2 mL: 02681258 | |
Microscope Software | Zeiss | AxioVision Rel. 4.8.2 | |
Nestin Alexa Fluor 594 | Santa Cruz Biotechnology | sc-23927 | |
Parafilm | PARAFILM | PM992 | |
PBS (1x), pH 7.4 | HyClone | SH30256.01 | |
Penicillin Streptomycin | MP Biomedicals | 1670046 | |
Pipette Aid | Drummond Scientific Co. | P-76864 | |
Pipette Tips (1–200 µL, 101–1000 µL) | Fisher Scinetific | 2707509 | |
Plastic Standard Disposable Transfer Pipettes | Fisher Scientific | 13-711-9D | |
Plastic Weigh Boats (100 mL) | Amazon | mdo-azoc-1030 | |
poly(ethylene glycol)-dithiol (PEG-diSH; 3.4 kDa) | Laysan Bio | SH-PEG-SH-3400-5g | |
Polydimehylsiloxane (PDMS) [Slygard 182 Elastomer Kit] | Elsworth Adhesives | 3097358-1004 | Polydimethylsiloxane |
Powder Free Examination Gloves | Quest | 92897 | |
Propidium iodide, 1 mg/mL aqueous soln. | Fisher Scientific | AAJ66584AB | |
RPMI-1640 Medium (1x) | HyClone | SH30027-02 | |
Silicone spacers – Silicone sheet, 0.5 mm thick/13 cm x 18 cm | Grace Bio-Labs | JTR-S-0.5 | |
SOX2 Alexa Fluor 488 | Santa Cruz Biotechnology | sc-365823 | |
Tissue Culture Hood | NUAIRE | NU-425-600 | |
Triethanolamine, ≥99.0% (GC) | Sigma Aldrich | 90279 | |
Trypsin 0.25% (1x) | Sigma Aldrich | SH30042.01 | |
U-87 MG human glioblastoma cells | American Type Culture Collection | HTB-14 |