Here, we describe different multicellular spheroid formation methods to perform follow-up multi-parameter live cell microscopy. Using fluorescence lifetime imaging microscopy (FLIM), cellular autofluorescence, staining dyes, and nanoparticles, the approach for analysis of cell metabolism, hypoxia, and cell death in live three-dimensional (3D) cancer and stem cell-derived spheroids is demonstrated.
Multicellular tumor spheroids are a popular 3D tissue microaggregate model for reproducing tumor microenvironment, testing and optimizing drug therapies and using bio- and nanosensors in a 3D context. Their ease of production, predictable size, growth, and observed nutrient and metabolite gradients are important to recapitulate the 3D niche-like cell microenvironment. However, spheroid heterogeneity and variability of their production methods can influence overall cell metabolism, viability, and drug response. This makes it difficult to choose the most appropriate methodology, considering the requirements in size, variability, needs of biofabrication, and use as in vitro 3D tissue models in stem and cancer cell biology. In particular, spheroid production can influence their compatibility with quantitative live microscopies, such as optical metabolic imaging, fluorescence lifetime imaging microscopy (FLIM), monitoring of spheroid hypoxia with nanosensors, or viability. Here, a number of conventional spheroid formation protocols are presented, highlighting their compatibility with the live widefield, confocal, and two-photon microscopies. The follow-up imaging to analysis pipeline with multiplexed autofluorescence FLIM and, using various types of cancer and stem cell spheroids, is also presented.
Multicellular spheroids represent a group of 3D tissue models obtained by the self-aggregation of cells and exhibiting a spherical shape. They are widely used to mimic cell-cell and cell-matrix interaction in vitro and to reproduce a 3D context within a multitude of cancer and stem cell-derived constructs. Several techniques are employed to reduce cell attachment and promote the aggregation. These include the hanging-drop method relying on the surface tension1; cell attachment repelling methods such as ultra-low attachment plates, micro-molds, and microwells2,3; acoustic wave-based approach4; flow-induced aggregation methods (spinner flasks, bioreactor, and microfluidic devices)5; magnetic particles-assisted formation6 and use of the aggregation-promoting synthetic and ECM-based matrices and scaffolds7,8,9.
In cancer research, development, and validation of new drug therapies, spheroids are an attractive model due to their ability to recapitulate the spatial diffusion-limited gradients of nutrients, waste products, and O2, often leading to the formation of a necrotic core, typical to the solid tumors10,11. These more reliable and sophisticated in vitro models challenge the need for extensive use of animal models (Food and Drug Administration [FDA] Modernization Act 2.012), according to the 3Rs principle of animal research (replacement, reduction, and refinement). In addition to cancer, spheroids find their application in stem cell research. For instance, pluripotent stem cells have the capacity to form embryoid bodies (EB), which can be used for the differentiation of induced pluripotent stem cells (iPSCs) towards specialized cell types that are challenging to obtain directly from patients, such as neural precursor cells13 or ovarian granulosa cells13,14. Furthermore, the formation of an EB is often the first step in the development of more complex organoid models, e.g., neural15, retinal16, cardiac17, liver18, stomach19, and intestinal organoids20. Factors including size, reproducibility, throughput, and downstream applications should be considered when choosing an appropriate spheroid formation method for the experiments.
The increased complexity of 3D culture can lead to higher variability compared to 2D culture. Factors such as nutrient composition21, media evaporation22, viscosity23, pH control24, spheroid formation method, and even the time in the culture25,26 can result in obtaining spheroids of varying morphology, sizes, viability, and different chemoresistance27,28. Recent research demonstrated that spheroid oxygen gradients are not always static and are affected by the formation method, spheroid size, and extracellular viscosity, affecting the spheroid heterogeneity29. To improve reproducibility and data accessibility on spheroids, the MISpheroID knowledge base has been developed26, identifying cell line, culture medium, formation method, and spheroids size as the minimal information for a reproducible result. Therefore, a detailed comparison was made of multiple high-throughput (SphericalPlate 5D, lab-made micromolds, and Microtissue molds) and low attachment methods (i.e., Biofloat and Lipidure-coated 96-well plates, both scaffold-free and scaffold-based) (Figure 1 and Table 1), including the well size (given an estimation of the maximum spheroid size), consumables used, preparation time and the possibility of monitoring spheroids without transporting them to microscopy dishes. The latter enables long-term studies, whereas spheroids produced with high-throughput methods often result in endpoint experiments. All methods except for the grids of the 5DspheriPlate do not bring unwanted autofluorescence, hereby enabling their direct use in microscopy.
Figure 1: Spheroid formation methods explained. High-throughput methods such as the SphericalPlate 5D, which has integrated patented microwells in the plate, while the lab-produced micromolds and the MicroTissue molds use stamps to make multiple microwells in agarose (blue). Low-attachment plates such as Lipidure (Amsbio) and Biofloat (Sarstedt) use a non-adherent coating inhibiting cell-surface adhesion and promoting cell self-aggregation. Please click here to view a larger version of this figure.
5D SpheriPlate | Self-produced micromolds | Microtissue | Low attachment methods | |
Number of spheroids/well | 750 | 1589 | 81 | 1 |
Diameter well | 90 µm | 400 µm | 800 µm | 1 mm |
Culture volume | 1 mL | 5 mL | 1 mL | 200 µL |
Other consumables | / | 7 mL of 3% agarose | 500 µL of 2% agarose | / * |
Preparation time | 10 min | 2 h + 3 days media adaptation | 0.5 h + 15 min media adaptation | 10–30 min + 1 h drying |
Monitoring | Yes | No** | Yes | Yes |
Autofluorescent | Yes | No | No | No |
Reusable | No | Yes | Yes | No** |
Cost | €€ | € | €€€€ | €€€€: Coating and Matrigel |
€€: Commercial 96-well plate | ||||
*Some cell lines need addition of ECM (i.e. 2%–5% Matrigel) to form compact spheroids. | ||||
**The coating is reusable until depleted. However, each plate will consume a small amount of media and dust can accumulate over time. Filter sterilization is regularly needed. |
Table 1: Comparison of multiple spheroid formation methods29. "Monitoring": the ability to monitor spheroid without the need for transfer to a microscopy dish. €: 0-50€, €€: 50-150€ , €€€: 150-500€ , €€€€: >500€
Fluorescence microscopy enables direct monitoring of the key biological aspects within spheroids, including cell death, viability, proliferation, metabolism, viscosity, and even mechanical properties30. Fluorescence lifetime imaging microscopy (FLIM) provides an additional quantitative dimension for studying fluorescent probe interactions within their (micro)environment31,32,33,34, allowing resolving the overlapping emission spectra according to different emission lifetimes35,36 and probing cell metabolism based on intrinsic cellular autofluorescence. Thus, such widespread cellular autofluorescent compounds as nicotinamide adenine dinucleotide phosphate (NAD(P)H), flavin mononucleotide (FMN), flavin adenine dinucleotide (FAD), protoporphyrin IX, and others can be measured with one- and two-photon FLIM and serve as intrinsic 'sensors' of glucose catabolism, oxidative phosphorylation (OxPhos) and provide a general overview of the cell redox state. NAD(P)H exists in free cytoplasmic, or in protein-bound mitochondrial forms37,38. Similarly, the oxidized state of FAD is fluorescent with a longer lifetime of the free form. NAD(P)H and FAD microscopies usually involve two-photon excited FLIM, aiming at preventing sample photodamage39. Frequently, 'optical metabolic imaging' FLIM can be combined with the use of dye-based probes, genetically encoded biosensors, phosphorescence lifetime imaging microscopy (PLIM), and ratiometric intensity-based measurements in order to provide a more complete picture of spheroid or organoid metabolism, oxygenation, proliferation and cell viability29,30,31. In addition, FLIM can also be combined with Förster resonance energy transfer (FRET) method to measure the lifetime variation of the donor fluorophore when in close contact with the acceptor to investigate the binding of a drug with its target domain33,40,41.
The acquired FLIM images are typically analyzed to calculate the lifetime pixel-by-pixel. Currently, there are at least 3 common strategies used to obtain fluorescence lifetime: semi-quantitative 'fast FLIM'42 (sometimes referred to as 'tau sense'43,44), decay curve fitting, using one-, two- or three-exponential fitting, and 'fitting-free' approach with phasor transformation and phasor plot analysis. Depending on the vendor, either provided (LAS X, Symphotime, SPCImage, etc.) or open-source software (e.g., FLIMfit45, FLIMJ46, or others47) can be used to handle measured FLIM data. Typically, vendor-provided software is useful for preliminary data analysis, while open-source solutions can provide for more accurate studies using, e.g., phasor plots and 3D visualization.
Despite the usefulness and attractiveness of FLIM as a method for studying spheroids, very few experimental protocols are available, and there is a general lack of knowledge in choosing the most appropriate formation method for successful live multiparametric microscopy experiments involving FLIM. Here, a detailed comparison of commonly used spheroid formation protocols is presented based on their morphology, viability, and oxygenation with the recently validated and characterized far-red and near-infra-red (NIR) oxygen-sensing nanosensor (MMIR1). The cationic nanoparticle is impregnated with two reporter dyes, the reference O2-insensitive aza-BODIPY (excitation 650 nm, emission 675 nm) and the NIR O2-sensitive metalloporphyrin, PtTPTBPF (excitation 620 nm, emission 760 nm). The MMIR1 enables real-time analysis of oxygen gradients on a conventional fluorescence microscope (using ratiometric analysis) or phosphorescence lifetime microscope (PLIM) without introducing cellular toxicity and allowing for stable signals, long-term monitoring, and multiplexing25,29. Depending on the need to stain with dyes or nanosensors, spheroid throughput, or cell type, the most appropriate formation protocol can be chosen. Since the studies of spheroids viability and oxygenation are relevant for studies of cancer and stem cell-derived spheroids, the presented protocols also include examples and expected typical results of NAD(P)H-FLIM and FAD-FLIM with these models. The presented imaging and analysis pipelines target the most popular time-correlated single photon counting-based FLIM microscopy platforms.
1. Generation of multicellular spheroids
2. Live microscopy of spheroids
Choosing the appropriate spheroid formation method
The selected spheroid formation method can greatly influence spheroids' size, shape, cell density, viability, and drug sensitivity (Figure 2). Previously, the effects of multiple high-throughput (SphericalPlate 5D, lab-made micromolds, and MicroTissue molds) and the 'medium throughput' low attachment (Biofloat and Lipidure-coated 96-well plates) methods were compared on spheroids viability and oxygenation29.
Here, different formation methods result in spheroids of different sizes, even with the same initial concentration of 500 cells/ spheroid. HCT116 spheroids formed in low attachment 96 well plates were significantly larger compared to high-throughput methods after 5 days of formation (Figure 2 and Table 2). Furthermore, low attachment methods led to the evident development of a necrotic core detected by the propidium iodide staining (red), while spheroids generated with other methods (MicroTissue, lab-made molds, and Sphericalplate 5D) demonstrated the diffused distribution of dead cells across the spheroid body (Figure 2A). This difference in cell viability across the volume of the spheroid can also affect oxygen diffusion through the media, which in turn can be affected by O2 partial pressure, consumption rate, temperature, and O2 solubility. Additionally, in high throughput methods, nutrients get depleted, and waste products accumulate faster as a larger number of spheroids are within the same volume and therefore require more frequent media exchange (Figure 2A).
The overall spheroid oxygenation was measured using the recently characterized and validated MMIR1 probe29, with a detailed protocol previously reported25. Oxygenation could only be compared between the high-throughput methods and low-attachment method separately (Figure 2B), as the signal-to-noise ratio is different in all compared spheroid formation methods. The Spherical plate 5D spheroids showed lower overall oxygenation compared to the MicroTissue (P-value = 0.0697) and the lab-made micromolds (P-value = 0.0005) (Figure 2C). Spheroids produced with the Spherical plate 5D are grown at a bigger distance to the media air interface surface compared to the agarose mold methods, causing a slower O2 delivery in the static culture. This hypothesis is valid as oxygenation values between both low attachment plates and, additionally, between both agarose mold methods are statistically similar (Figure 2D).
Figure 2: Spheroid formation methods affect morphology, size, viability, and oxygenation. (A) HCT116 spheroids (initial seeding density 500 cells) were grown for 5 days prior to 1 h-long staining with Propidium Iodide (cell necrosis, red, 1 µg/mL) and Calcein Green-AM (viable cells, green, 1 µg/mL). Low-attachment formation methods produce bigger spheroids containing a necrotic core (red). Scale bar is 100 µm. (B) Oxygenation in HCT116 spheroids (initial seeding of 500 cells with addition of 20 µg/mL MMIR1 during spheroid formation, 6 days) produced by different methods showed more oxygenated spheroids with the MicroTissue and lab-made micromolds than the SphericalPlate 5D. (C) Overall spheroid oxygenation calculated by the ratio measurements. (D) Spheroids size in area square after 5 days. Results show the average ± SEM of 6-16 spheroids. 5D = SphericalPlate 5D, MM = Micromold method, MT = MicroTissue method. ***P-value < 0.001 and ****P-value <0.0001 Please click here to view a larger version of this figure.
Formation method | Amount of spheroids per well | Total amount of cells seeded per well | Area square (µm²) |
Microtissue | 81 | 40,500 cells/190 µl | 112,558 ± 15,702 |
lab-made microwells | 1859 | 794,500 cells / mL | 53,460 ± 8,332 |
Sphericalplate 5D | 750 | 375,000 cells / mL | 44,048 ± 7,259 |
Lipidure | 1 | 500 / 200 µL | 257,148 ± 28,132 |
Biofloat | 1 | 500 / 200 µL | 254,475 ± 21365 |
Table 2: Different spheroid formation methods lead to differences in spheroid size, measured by area square.
When selecting a specific spheroid formation method for the experiments, it must be kept in mind that the addition of nanoparticles can influence the spheroid's shape. When using the lipid-based coating with round bottom microwells, probe precipitation can occur, leading to the formation of "satellite spheroids" or non-circular spheroids (Figure 3). The addition of the O2 probe to the commercial Biofloat plate results in a non-compact spheroid periphery, suggesting the potential interference of the nanoparticles with the microplate coating. The morphology of the spheroids can also be affected by the dust attached to the pipet tips. Cells in suspension can stick to the dust, hereby affecting the shape. Therefore, it is recommended to use refillable pipette tips already supplied in a rack.
Figure 3: Transmission light microscopy images with examples of non-ideal spheroid formation. Incorrect storage of nanoparticles can cause probe precipitation, interfering with spheroid formation. Examples of interference of probe and dust. Scale bar is 100 µm. Please click here to view a larger version of this figure.
Clearly, spheroid formation methods affect the spheroid's size, morphology, and viability, and lead to differences in their oxygenation. The high throughput method with agarose molds (e.g., commercially available, such as MicroTissue mold) is compatible with most experiments. It results in a more reproducible shape, does not induce nanoparticle precipitation, is reusable, and can be compatible with direct follow-up microscopy. However, for large spheroids (>800 µm) and long-term culturing, the low attachment method is recommended as high throughput methods would require frequent media exchange. In such a case, the nanoparticle precipitation problem and its influence on spheroid formation can be avoided by the overnight pre-staining of the 2D monolayer culture before spheroid formation (Figure 4).
Figure 4: Suggested algorithm for selecting the spheroid formation method for live fluorescence microscopy. Please click here to view a larger version of this figure.
Multi-parameter FLIM of spheroids NAD(P)H imaging via two-photon FLIM microscopy reveals metabolic heterogeneity in HCT116 spheroids
A prominent autofluorescence marker in label-free metabolic imaging, NAD(P)H shows a shorter lifetime for glycolysis and a longer lifetime for OxPhos, enabling deciphering the spatial distribution of metabolic activity in such 3D models as spheroids and organoids31. To conveniently analyze the NAD(P)H distribution in HCT116 spheroids, a phasor-based quantitative method for autofluorescence NAD(P)H analysis was used, revealing glycolysis and oxidative phosphorylation (OxPhos)-linked states. While fluorescence intensity images of HCT116 spheroids did not reveal significant differences between spheroids, fast FLIM imaging of NAD(P)H revealed a clear difference within the spheroid optical sections. In fast FLIM images, spheroids with an internal area displaying a shorter fluorescence lifetime were marked with a white circle, identifying this as a glycolytic core (Figure 5A). Consequently, the spheroids were divided into two groups, the glycolytic core group and the non-glycolytic core group, based on the presence or lack of a glycolytic core. A centroid-based phasor analysis was employed to statistically analyze the differences between two distinct groups and measure the distance from the centroid of the pixel cluster to the free NAD(P)H point. As illustrated in Figure 5B, there was an obvious pixel cluster difference between the glycolytic group and the non-glycolytic group on the phasor plot. To accurately measure the distance, the free NAD(P)H point on the phasor plot was precisely marked using the FLIM module in LAS X software, and then exported into FIJI software for the exact coordinates' determination. Subsequently, following protocol 2.3, centroid coordinates were measured in each group's phasor plot using FIJI software. The distance (D) from the centroid to free NAD(P)H point, using the Pythagorean theorem, was calculated hereby facilitating analysis of NAD(P)H profiles to distinguish OxPhos spheroids and spheroids with a glycolytic core. The results, as displayed in the boxplot, show the significant distance differences between the glycolytic core spheroids and the non-glycolytic core group (***P-value < 0.001) (Figure 5C). This result was consistent with fast FLIM imaging, demonstrating that this phasor analysis method is suitable for quantitative analysis of the NAD(P)H divergence of spheroid populations. Moreover, to validate if this approach was applicable to compare the distance between the two spheroid groups, a linear regression analysis for centroids from both groups and free NAD(P)H point was performed (Figure 5D). The result showed that the centroids are aligned with free NAD(P)H, with a high coefficient of correlation (R2 = 0.997), validating the feasibility of comparing the distance to infer the NAD(P)H difference between spheroid groups.
Figure 5: Phasor analysis of HCT116 spheroids made using low-attachment plates: glycolytic and non-glycolytic cores comparison of the NAD(P)H distribution via two-photon FLIM. Acquisition parameters: laser intensity: 15%, resolution: 512 x 512 pixels, ex. 741 nm/em. 411-491 nm. (A) The fluorescence intensity and fast FLIM images display NAD(P)H distribution in HCT116 spheroids, distinguishing spheroids with glycolytic core (left) from those without (right). White circle in the fast FLIM image displayed a shorter lifetime area, defining a glycolytic core. Initial seeding density: 50 cells (left) and 500 (right), incubation time: 6 days, scale bar: 50 µm. (B) The methodology for measuring the distance from the centroid of HCT116 spheroids to free NAD(P)H point on phasor plots, using LAS X FLIM module and FIJI software across two distinct spheroid groups (D: distance). For HCT116 spheroids, a wavelet filter and threshold value 10 were employed for the phasor plot. (C) Boxplot showing the difference of distance from the centroid to free NAD(P)H point, comparing glycolytic core and non-glycolytic core groups (***P < 0.001). (D) Linear fitting of phasor plot centroids from glycolytic core, non-glycolytic core groups, and free NAD(P)H theoretical coordinates demonstrated accurate linear alignment with R2 = 0.997, allowing for direct comparison of distance difference between glycolytic core and non-glycolytic core spheroids. Please click here to view a larger version of this figure.
Multiplexed analysis of FAD autofluorescence and hypoxia in human dental pulp stem cell (DPSC) spheroids
Dental pulp stem cell spheroids are an attractive experimental tool for the biofabrication of different tissues, including osteoblasts25,66,67. However, their viability and metabolism are rarely studied. These stem cell-derived spheroids display rather small sizes (<200 µm), bright green autofluorescence from FAD/Flavins (later referred to as FAD) (ex. 460 nm, em. 550 nm) and the presence of 'direct' and 'inverted' hypoxic gradients29. Figure 6 demonstrates the results of combined confocal ratiometric imaging of spheroids oxygenation (with the help of O2-sensing nanoparticles MMIR) and autofluorescence FLIM of FAD. The 'classical' direct oxygenation gradient was observed in spheroids of 69 mm diameter, while larger (141 µm) spheroid showed an 'inverted' gradient. Adding FAD-FLIM to these measurements helps validate differences in oxidative metabolism, as the fast FLIM and phasor plots demonstrate: small spheroids displayed a more prominent fraction of longer FAD lifetimes, potentially indicating higher glycolytic activity in the smaller-size spheroids.
Figure 6: Example of multiparametric imaging of flavin autofluorescence (ex. 460 nm/em. 510-590 nm) and oxygenation (intensity ratio analysis with MMIR O2 sensitive nanoparticles: ex. 614 nm/reference em. 631-690 nm / sensitive em. 724-800 nm) in hDPSC spheroids of different size (small: 188 cells/spheroid, big: 820 cells/spheroid), produced by high-throughput self-produced agarose micromolds method (confocal FLIM). (A,B) Representative example of oxygenation intensity ratio and flavin-autofluorescence lifetime imaging in big (A, "B Sphs") and small (B, "S Sphs") hDPSC spheroids. (C,D) Comparative analysis of flavin-autofluorescence lifetime using phasor plot cloud analysis, based on the comparison of phasor cloud geometrical centers. (E) The pixel centroids (geometrical centers) coordinates of flavin autofluorescence from big hDPSC spheroid (B Sphs) and small hDPSC spheroid (S Sphs) on phasor plot. Two different color codes applied for different FLIM analysis approaches, where color coding with τ shows average photon arrival time (FAST-FLIM images) distribution, while color coding with τ φ – tau phase corresponds to phasor-FLIM analysis of fluorescence lifetime in the same spheroids. Please click here to view a larger version of this figure.
Multiplexed analysis of FAD autofluorescence and cell death in human iPSC spheroids
Human induced pluripotent stem cell (iPSC) spheroids are often used as a first step during the induction of tissue-specific differentiation and the production of organoids, e.g., in the case of neural organoid culture. Depending on their handling and the specific generation protocol, viability and subsequent reproducibility of organoids can be severely affected. Non-destructive investigation of organoids and neural progenitor cell-containing spheroids is crucial for the structural assessment, monitoring, and predicting the quality of growing and assembled neural tissues during their development68,69. Figure 7A shows that the non-destructive FLIM of autofluorescent Flavin/FAD molecules can give us information about the viability of iPSC spheroids. Imaging was performed 4 days after seeding 9,000 iPSCs in commercial ultra-low attachment plate wells (Corning). Before imaging, spheroids were stained for 1 h with 0.5 µg/mL Propidium Iodide (PI) to visualize dead cells. No necrotic cores were observed in these iPSC spheroids with a seeding density of 9,000 cells after 4 days. Three main patterns can be distinguished on the corresponding phasor plot of flavin/FAD autofluorescence (Figure 7B left panel): pattern of media autofluorescence (magenta color), pattern (with average τ φ – tau phase, ~2.6 ns) corresponding to no PI regions (ROI1) and pattern (with average τ φ ~3.1 ns) of regions correlating with the cell death marker – staining (ROI2). An additional comparison of PI-treated and non-treated 2D cultures of iPSCs demonstrated no impact of PI on the appearance of long-lifetime flavin / FAD phasor patterns (Figure 7C), indicating that this increase of flavins / FAD lifetime can be an independent marker of compromised viability of iPSCs and their derivative cells. While this observation is in line with the recent report on intensity-based imaging70, we cannot completely rule out the presence of propidium iodide intensity with FAD emission channel (506-582 nm). A further investigation is needed to prove the link between flavin / FAD lifetime changes and potential events of cell death. The simple multiparametric phasor-based analysis of ROI regions demonstrates an elegant way for quick screening and data analysis.
Figure 7: Flavin autofluorescence confocal FLIM and propidium iodide staining in iPSC spheroids to assess cell viability. (A) Flavin autofluorescence (left) and Propidium Iodide – PI (right) FLIM of an iPSC spheroid 4 days after seeding 9,000 cells in an Ultra-low attachment plate (Corning) and PI staining (0.5 µg/mL, 1 h) prior to imaging. (B) Phasor-plot analysis of flavin autofluorescence (left) and propidium iodide (right) overall images and corresponding ROIs (ROI1 – green and ROI2 – red, media autofluorescence – magenta). ROIs were selected based on the presence of propidium iodide labeling. (C) fast-FLIM and phasor plot analysis of flavins autofluorescence in 2D culture of iPSC treated and non-treated with PI (0.5 µg/mL, 1 h). No impact of PI staining on the appearance of the long fluorescence lifetime of flavins (corresponds to a phasor pattern within a pink circle) was detected when collecting the Flavin excitation within a range from 469-542 nm. This detector range does not overlap with the excitation spectrum of PI (550-720 nm). When a broader excitation range for FAD is collected (469-590 nm) an impact of PI can be noticed. Image color-coding corresponds to the average photon arrival time values (fast-FLIM). Imaging parameters were T = 35 °C, 25X/0.95 water-immersion objective (A-B), 40X/1.25 Glycerol objective (C). Flavins: ex. 460 nm, em. 506-582nm (A-B), pinhole 4 AU. PI: ex. 535 nm, em. 584-667 nm, pinhole 1 AU. Channels: Intensity and Tau. The repetition rate of 40 MHz to collect full decay of Flavin autofluorescence. Phasor analysis settings; Harmonic 1, Threshold 1 (A-B), 47 (C), Median Filter 17 (A-B), 11 (C). Please click here to view a larger version of this figure.
Supplementary File 1: Additional protocols on spheroid formation. Please click here to download this File.
Supplementary File 2: FLIM data of glycolytic core and non-glycolytic core.lif. Please click here to download this File.
Supplementary File 3: FLIM data of glycolytic core.ptu. Please click here to download this File.
Supplementary File 4: FLIM data of non-glycolytic core.ptu. Please click here to download this File.
Supplementary File 5: Glycolytic core fast FLIM.tif Please click here to download this File.
Supplementary File 6: Non-glycolytic core fast FLIM.tif Please click here to download this File.
Supplementary File 7: Glycolytic core free NAD(P)H point.tif Please click here to download this File.
Supplementary File 8: Glycolytic core phasor plot.tif Please click here to download this File.
Supplementary File 9: Non-glycolytic core free NAD(P)H point.tif Please click here to download this File.
Supplementary File 10: Non.glycolytic core phasor plot.tif Please click here to download this File.
Multicellular spheroids are becoming a method of choice in the studies of tumor and stem cell niche microenvironments, drug discovery, and development of the 'tissue building blocks' for biofabrication. Spheroids' heterogeneous internal architecture, gradients of nutrients and oxygenation can mimic those of in vivo tissues and tumors in a relatively simplified and accessible setting. With the need for more methodological transparency26,28 and standardization71, these protocols are expected to help researchers choose the best spheroid formation method for the live quantitative fluorescence lifetime imaging microscopy experiments. The methods presented take advantage of cell staining nanoparticle sensors and intrinsic cellular autofluorescence, which can be easily combined with staining cell death.
The most critical steps are in spheroid formation protocol; thus, for the high throughput agarose-based method, it is essential to sufficiently equilibrate the agarose molds for the culturing media. Insufficient equilibration time will lead to the dilution of nutrients in the remaining PBS and bring additional factors for spheroid heterogeneity21,26,72,73. The freshly made agarose mold has to be also visually checked for the presence of air bubbles, micro-cracks and other defects.
Although efficient for imaging, nanoparticle suspension can display self-aggregation and cause the formation of irregularly shaped spheroids in low-attachment formation methods. This can often be mitigated by the addition of serum to the culturing media or by pre-staining the monolayer cell culture before spheroid formation. Frequently, during spheroid compactization, an addition of a small amount of nanoparticles (e.g., 10% of the initial staining concentration) can also help staining the proliferating zone of the spheroids. Lastly, it is important to use "dust-free" sterile pipette tips with all formation methods.
The potential impact of illumination on cell viability must always be considered during imaging experiments. Thus, it is important to know the risk factors negatively affecting each live imaging experiment. In general, photodamage involves several mechanisms: (1) high local rise of temperature, (2) mechanical stress, and (3) photochemical stress (oxidative and free-radicals reaction)74. Depending on the imaging modality (e.g., one-photon confocal vs. multiphoton imaging, continuous vs. pulse illumination modes, the sensitivity of detectors and power of the light source at different wavelengths if the excitation tuning is possible), cell and tissue type (e.g., tissue type-dependent accumulation of efficiently light absorbing pigments or other autofluorescence molecules, 3D vs. 2D organization, growth and imaging media composition), the origin of the fluorescent probe (e.g., photosensitizing properties) different combination of risk factors can provoke phototoxicity during imaging. In addition, even if no increase in cell death is observed after illumination, light power-dependent earlier pre-death effects on mitochondria and metabolism can potentially interfere with experiment analysis and interpretation55.
To exclude the phototoxic impact on experimental results the researcher should perform general monitoring of cell viability and changes in cell morphology in comparison to intact control or using cell death probes to estimate the increase of cell death rate after illumination. If the metabolic experiments are performed, the optimization of imaging parameters should be done accordingly in the pilot experiments to estimate and minimize the imaging impact on measured metabolic parameters. In the case of oxygen-sensitive probes, the light overdose can be easily tracked by a sudden increase of O2-sensitive dye phosphorescence lifetime and intensity (corresponding changes in the intensity ratio of reference to O2-sensitive dye) due to so-called 'photoinduced O2 consumption' effect53,75.This can be easily done by kinetic curve analysis of time-lapse experiments with different light power parameters using, for example, the following published protocol25. In 'defense' of the pulse-illumination (e.g., utilized in TCSPC-F(P)LIM microscopy), the recent studies demonstrated its minimal impact on ROS production and probe photobleaching in comparison to steady-state illumination measurements76,77, pointing to additional advantage of TCSPC-FLIM vs. conventional intensity-based imaging approach.
The main limitation of the presented spheroid formation methods lies in direct and long-term monitoring applications. The agarose molds require a long working distance objective (or additional upright microscope or fluorescence stereo microscope) due to the thick layer of agarose and the material of the 5D Spheriplate may interfere with measurements in red fluorescent channels (O2 probe). The spheroid size produced by high throughput methods is limited by microwell diameter and requires frequent media exchange to avoid media depletion and acidification. With all these limitations, we recommend the 3D Petri Dishes (MicroTissue) for experiments requiring high-throughput (e.g., for 3D bioprinting applications) and low-attachment methods for "medium-throughput" basic studies, also requiring lower cell numbers.
Ratiometric oxygenation monitoring and conventional live cell fluorescence dye-based microscopy experiments are affordable on widely available inverted microscopes if equipped with a near-infrared (NIR)-sensitive camera, light-emitting diode (LED)-based excitation, and appropriate objective25. For more appropriate optical sectioning and FLIM, off-the-shelf available commercial confocal microscope systems (such as described here) are recommended. While we also describe a method for detecting NAD(P)H autofluorescence, requiring a more costly two-photon FLIM microscope, there are many different approaches, compatible and useful for confocal FLIM systems, equipped ideally with white light laser and hybrid or SPAD detectors. These include assessment of cell metabolism, FRET-FLIM biosensor proteins, imaging cell and tissue mechanics, autofluorescence of specific cell and tissue types, and receptor-drug interactions visualized via FLIM-FRET31.
The vendor-provided FLIM acquisition and analysis software can affect the success of the experiment, as it is not always possible to export the IRF or the images in a format compatible with the downstream open-source analysis. Post-processing is an important part of the FLIM analysis workflow, and it is still not universally standardized or accepted. Curve fitting requires a higher number of photons and more computational power compared to the phasor approach. FLIMfit45 (global and pixel-by-pixel fitting) and Flimview78 are among the open software used for curve fitting analysis, while PAM79, FLUTE64, and AlliGator80 are used for phasor analysis. FLIMJ46 is a useful ImageJ plugin that works with both approaches. Altogether, these options help improve the post-processing routine and make it faster, more versatile, and user-friendly.
The authors have nothing to disclose.
This work was supported by the Special Research Fund (BOF) grants of Ghent University (BOF/STA/202009/003; BOF/IOP/2022/058), Research Foundation Flanders (FWO, I001922N) and the European Union, fliMAGIN3D-DN Horizon Europe-MSCA-DN No. 101073507.
0.05% Trypsin-EDTA | Gibco | 25300-054 | Also available from Sigma |
10 mL serological pipets | VWR | 612-3700 | Similar products are also available from Sarstedt, Corning, VWR and other companies |
12 well cell-culture plates, sterile | Greiner bio-one | 665-180 | Similar products are also available from Sarstedt, Corning and other companies. |
12 Well Chamber slide, removable | Ibidi | 81201 | Also available from Grace Bio-Labs, ThermoFisher Scientific and others |
15 mL centrifuge tubes | Nerbe plus | 02-502-3001 | Similar products are also available from Sarstedt, Corning, VWR and other companies |
3D Petri Dish micromolds | Microtissue | Z764000-6EA | |
6 well cell-culture plates, sterile | Greiner bio-one | 657160 | Similar products are also available from Sarstedt, Corning, VWR and other companies |
70% ethanol | ChemLab | CL02.0537.5000 | |
Biofloat | Sarstedt | 83.3925.400 | Commercial available coated 96-well plate for spheroid formation |
Calcein Green-AM | Tebubio | AS-89201 | Apply in dilution 1:1000 |
CellSens Dimension software | Olympus | version 3 | |
Collagen from human placenta, type IV | Sigma | C5533 | For the preparation of 0.07 mg/mL Collagen and 0.03 mg/mL Poly-D-lysine coated microscopy dishes |
Confocal FLIM Microscope | Leica Microsystems | N/A | Stellaris 8 Falcon inverted microscope with white-light laser, HyD X detectors, climate / T control chamber (OkoLab), 25x/0.95 W objective |
D(+)-Glucose | Merck | 8342 | Prepare 1 M stock solution, 1:100 for preparation of imaging medium (final concentration 10 mM) |
Dulbecco's modified Eagle's medium (DMEM), phenol red-, glucose-, pyruvate- and glutamine-free | Sigma-Aldrich | D5030-10X1L | For preparation of imaging medium |
Fetal Bovine Serum (FBS) | Gibco | 10270-098 | Also available from Sigma. Needs to be heat-inactivated before use. |
HEPES (1M) | Gibco | 15630-080 | Dilution 1/100 for preparation of imaging medium (final concentration 10 mM) |
Human colon cancer cells HCT116 | ATCC | ||
ImageJ | NIH | version 1.54f | |
Leica Application Suite X (LAS X) | Leica Microsystems | version 4.6.1.27508 | |
L-glutamine | Gibco | 25030 | Also available from Sigma. Apply in dilution 1:100. |
Lipidure-CM5206 | Amsbio | AMS.52000034GB1G | |
McCoy's 5A, need addition of 1 mM Sodium Pyruvate and 10 mM HEPES | VWR | 392-0420 | Standard growth medium for HCT116 cells |
micro-patterned 3D-printed PDMS stamps | N/A | N/A | Provided by the Centre for Microsystems Technology, Professor Dr. Jan Vanfleteren, Ghent University |
NaCl | Chemlab | CL00.1429.100 | |
Neubauer couting chamber | Fisher Scientific | 15980396 | |
O2 probes: MMIR1 | N/A | N/A | Full characterization, validation and some applications can be found at: https://www.biorxiv.org/content/10.1101/2023.12.11.571110 v1 |
PBS | Fisher scientific | Gibco18912014 | Dissolve PBS tablet in 500 mL of distilled water. |
Pen Strep :Penicillin (10,000 U/mL) / streptomycin (10,000 μg/mL) 100x solution | Gibco | 15140-122 | Also available from Sigma. Apply in dilution 1:100. |
Poly-D-lysine | Sigma | P6407-5mg | For the preparation of 0.07 mg/mL Collagen and 0.03 mg/mL Poly-D-lysine coated microscopy dishes |
Propidium Iodide | Sigma-Aldrich | 25535-16-4 | Cell death staining, use 1 µg/mL at 1h incubation |
PVDF syringe filter 0.22 µm | Novolab | A35149 | Similar products are also available from Sarstedt, Corning, VWR and other companies |
Sodium pyruvate (100 mM) | Gibco | 11360-070 | Dilution 1/100 for preparation of imaging medium (final concentration 1mM) |
SphericalPlate 5D 24-well | Kugelmeiers | SP5D-24W | |
sterile petridish | Greiner bio-one | 633181 | Similar products are also available from Sarstedt, Corning, VWR and other companies |
Tissue culture flask (25 cm² ) | VWR | 734-2311 | Similar products are also available from Sarstedt, Corning, VWR and other companies |
Tissue culture flask (75 cm²) | VWR | 734-2313 | Similar products are also available from Sarstedt, Corning, VWR and other companies |
U-bottom 96-well plate | VWR | 10062-900 | Similar products are also available from Sarstedt, Corning, Greiner Bio-one and other companies |
Ultrapure Agarose | Invitrogen (Life Technologies) | 16500-500 | Other types of Agarose such as Agarose low melting point (A-9414, Sigma), Agarose for routine use (A-9539, Sigma) |
Widefield fluorescence inverted microscope | Olympus | N/A | Inverted fluorescence microscope IX81, with motorised Z-axis control, CoolLED pE4000 (16 channels, 365-770 nm), ORCA-Flash4.0LT (Hamamatsu) cMOS camera, glass warming plate Okolab, CellSens Dimension v.3 software and air objectives 4x/0.13 UPlanFLN and 40x/0.6 LUCPlanFLN. (Optional, for high-resolution imaging) 60x/1.0 LUMPLFLN water |