Summary

Production and Multi-Parameter Live Cell Fluorescence Lifetime Imaging Microscopy (FLIM) of Multicellular Spheroids

Published: August 09, 2024
doi:

Summary

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.

Abstract

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.

Introduction

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
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.

Protocol

1. Generation of multicellular spheroids

  1. Cell culture
    NOTE: Cell cultures can be collected from the American Type Culture Collection (ATCC), Lonza, Sigma-Aldrich, or other vendors. ATCC provides all required handling information, including preferred growth media, subculturing procedures, biosafety level, growth rate, and STR profiles. Here, 500 cells/spheroid of human colon cancer cell line HCT116 is used in McCoy's 5A media (VWR, 392-0420) supplemented with 10% FBS and 1 mM Sodium Pyruvate. For long-term experiments that are monitored daily, 10 mM HEPES, pH 7.2 can be added to the media.
    1. Grow cell culture to reach 70%-90% confluency.
    2. Rinse cells with prewarmed (37 °C) sterile PBS (5 mL for 25 cm2 or 10 mL per 75 cm2 flask).
    3. Add 0.05% trypsin – 1 mM EDTA solution (0.5 mL for 25 cm² or 1 mL per 75 cm2 flask) and incubate for 5-10 min at 37 °C in 5% CO2, 95% humidity to reach cell detachment.
      NOTE: Control cell detachment under the transmission light (brightfield) microscope. The overexposure of cells to the dissociating enzyme solution can affect their viability.
    4. Neutralize trypsin by adding excess cell culture media containing 10% FBS (at least 5 mL of media per 1 mL of dissociation solution).
      NOTE: For cells cultured on low FBS or FBS-free culture media, trypsin neutralization can be done with the addition of 0.5 mL of 100% FBS to the trypsin-treated cell culture, followed by centrifugation to transfer cells to their original culture media.
    5. Dissociate cell aggregates by pipetting to obtain single-cell suspension in media.
      NOTE: Pipetting with a serological pipette with a 1000 µL pipette tip on the top highly improves single-cell suspension generation in a big volume of suspension cell culture.
    6. Use a counting chamber (Neubauer ruled hemocytometer or alternatives) to count cell number per 1 mL of the cell suspension.
    7. Dilute the cell suspension to obtain the desired number of cells per milliliter.
    8. Add concentrated O2 probe (nanoparticles) solution to the cell suspension at a final concentration of 10 µg/mL for ratiometric analysis of O2.
      NOTE: To ensure homogenous cell suspension (with the probe), resuspend multiple times before spheroid formation. If the O2 probe is not required, skip this step and proceed with the formation of the spheroids. A modified protocol should be applied when handling iPSCs. Briefly, iPSCs are grown in colonies on Geltrex-coated plates and passaged using ReLeSR as described in the vendor-provided protocol48. On the day of spheroid formation, colonies should be large, compact, and exhibit multilayered centers with distinct borders. Rinse cells with prewarmed sterile PBS. Add 1 mL of gentle cell dissociation reagent (GCDR) and incubate at 37 °C for 8-10 min. Use a 1000 µL tip to gently detach cells from the well and to obtain a single-cell suspension. Transfer single cell suspension to a sterile 50 mL conical tube and add 4 mL of prewarmed DMEM-F12 medium to neutralize the GCDR. Wash the well with 1 mL of DMEM-F12 and transfer it to the rest of the cell suspension. Centrifuge at 300 x g for 5 min. Resuspend in 1 mL of appropriate media for further experiments. For experiments described in this manuscript, mTeSR + 10 µM Rock Inhibitor was used. Count and dilute the cell suspension to obtain the desired number of cells per milliliter.
  2. Spheroid formation methods
    1. 3D Petri dish micro-molds
      NOTE: This high throughput method is used to simultaneously generate a high number of spheroids (81 spheroids) in a 9 x 9 micromold array with 800 µm diameter and 800 µm depth.
      1. Rinse micro-molds for casting 3D Petri dishes in dH2O and put in autoclavable container.
      2. Measure 2 g of electrophoresis-grade agarose powder and place it into a dry 200 mL autoclave-safe glass bottle.
        NOTE: Be sure the bottle and agarose powder are dry with no liquid or moisture.
      3. Autoclave micro-molds for casting 3D Petri dishes and bottles with agarose powder for 30 min on a dry cycle.
      4. Make a 0.9 w/v% saline solution by addition of 0.9 g of NaCl in 100 mL of ultrapure water and sterilize by autoclaving.
        NOTE: NaCl is recommended by the manufacturer. It increases the agarose stability.
      5. Prepare the agarose solution by adding the sterile saline to the sterilized agarose powder. Screw on the lid loosely fitted to avoid pressure buildup. Swirl the bottle to mix the agarose powder.
      6. Boil and dissolve the agarose powder by using a microwave oven. Stop the microwave frequently (~every 10 s). Swirl the bottle and repeat until agarose is dissolved.
        CAUTION: The agarose solution is hot and requires careful handling. Shaking immediately following the melting procedure may cause the solution to burst out of the vessel. To avoid accidents, use sufficiently large vessels filled to no more than 50% of capacity and use appropriate personal protection (gloves, oven mitt, eye protection, and lab coat).
      7. Let the dissolved agarose solution cool down to 60-70 °C. Using aseptic techniques and conditions, pipette molten agarose into the micro-mold (500 µL for 12-series or 330 µL for 24-series).
        NOTE: Avoid bubble formation while mixing or pipetting agarose. Remove any trapped small bubbles in the small features of the micro-mold by pipetting or gentle scrapping before the agarose solidifies.
      8. Let the agarose solidify for about 2-3 min. Afterwards, carefully flex the micro-mold to remove the 3D Petri dish and transfer to a 12 well tissue culture plate.
        NOTE: Overflexing of the micro-mold might lead to the formation of cracks within the agarose mold.
      9. To equilibrate the 3D Petri dish, add 2.5 mL/well cell culture medium. Incubate for 15 min or longer. Remove the culture medium and replace it with fresh medium. Repeat once more to equilibrate the 3D Petri dish with a culture medium.
        NOTE: The protocol can be interrupted here until cell seeding. For long-term storage (up to 2 weeks at 4 °C), use PBS solution instead of medium.
      10. Remove the 3D Petri dish surrounding the culture medium (or PBS) completely and accurately remove the media inside the 3D Petri dish by tilting the tissue culture plate.
      11. Carefully seed 190 µL of cell suspension containing 40,500 cells dropwise into the cell seeding chamber (see step 1.1).
        NOTE: The number of micro-wells in agarose stamps determines the number of spheroids produced per stamp. In this case, this agarose mold contains 81 microwells (81 x 500 cells/spheroid). Variation of cell concentration in suspension added to a macrowell allows changing the cell number per spheroid, thus controlling the spheroid's size.
      12. Allow ~10 min for cells to settle into the features of the 3D Petri dsh. Then, add 2 mL of medium to the outside of the 3D Petri dish.
      13. Place the tissue culture plate in the cell culture incubator and exchange the medium surrounding the 3D Petri Dish as needed.
    2. Low attachment plate
      NOTE: This method is used to generate a single spheroid per well. Coated plates (Lipidure or Biofloat) are commercially available (skip steps 1.2.2.1-1.2.2.4). Alternatively, the coating can be purchased separately and used to coat untreated multi-well plates. It is recommended to fill wells at the edges of 96-well plates with sterile water or PBS due to faster evaporation in these wells, thereby limiting the number of wells for spheroids to 60. In case fewer spheroids are needed, fill the surrounding empty wells with water or PBS. Use dust-free tips for liquid handling to avoid bringing small particles into the wells, as they interfere with the spheroid formation.
      1. Prepare a 0.5 w/v% coating solution by dissolving 0.25 g of the polymer powder in 50 mL of ethanol in a glass bottle container. Filter-sterilize the coating.
        NOTE: Filter sterilization and all the next steps have to be carried out in sterile conditions under laminar flow.
      2. Add 200 µL of the coating solution to each well of a 96 U-bottom culture plate.
      3. Incubate for 1 min and take off excess coating.
        NOTE: The coating solution can be used multiple times. Store in a glass container at room temperature (RT). Plastic containers are not advised, as the plastic can get partially dissolved and become a part of the solution. If dust is present, filter sterilize with a 0.22 µm polyethersulfone (PES) or nylon syringe filters.
      4. Let the 96-well plate air dry for about 1 h.
        NOTE: If cells need an extracellular matrix, proceed to step 1.2.3. Coated plates can be stored at RT when wrapped in aluminum for up to 1 month. When seeding a lower amount of cells/well, centrifuging the plate for 5 min at 300 g can help pull down the cells.
    3. Extracellular matrix-aided formation protocol
      NOTE: Some cell lines do not produce enough extracellular matrix (ECM) themselves and need the addition of ECM such as Matrigel, Cultrex, or Geltrex in order to form compact spheroids49,50,51. For such cell types as breast cancer MDA-MB-231, human dermal papilla, prostate cancer cells and others, it is possible to use step 1.2.2 with the following modifications52, requiring the addition of ECM. Preferably use dust-free tips for liquid handling to avoid dust interfering with the spheroid formation. Steps 1 and 4-7 must be carried out under a biological safety cabinet (class II).
      1. Proceed with steps 1.2.2.1-1.2.2.4 (low attachment plates) for surface treatment.
      2. Pre-chill the 96-well plate at 4 °C in the refrigerator.
      3. Prepare the centrifuge with the correct adapter for the 96-well plate and prechill it for 4 °C.
      4. Prepare a 5% solution of basement membrane matrix (BMM) in pre-chilled (4 °C) cell culture media.
        NOTE: BMM rapidly crosslinks at RT. When handling, keep stocks and solutions on ice.
      5. Prepare the cell suspension (see step 1.1).
      6. Add 50 µL of the BMM solution into each well.
      7. Gently add 50 µL of cell suspension into each well on top of the BMM solution (25,000 cells/well).
        NOTE: Do not blast this volume into the well, or else the cells will spread up on the sides of the wells and not collect at the bottom. A lower number of cells per well can be obtained by adjusting the cell density accordingly. Not all cells will form spheroids at all seeding densities; the optimization must be performed cell type and desired dimension-wise. With the provided volumes, the final concentration of BMM is 2.5%. If a different concentration is needed, the stock solution has to be prepared at a lower/higher concentration.
      8. Centrifuge the 96-well plate for 5 min at 300 x g and 4 °C.
        NOTE: Without this step, cells do not aggregate properly in the bottom of the well, causing multiple smaller aggregates to form. The centrifuge has to be cooled down to avoid crosslinking at this stage.
      9. Place the plate in the cell culture incubator. Aggregates are considered mature on day 4 after seeding.
        NOTE: For additional protocols on spheroid formation, refer to Supplementary File 1.

2. Live microscopy of spheroids

  1. Preparation of spheroids for live imaging analysis
    NOTE: Depending on the experiment design (e.g., long-term monitoring or endpoint analysis, microscope set-up, or spectral properties of the measured fluorescence) or due to incompatibility of the spheroid production method with the microscopy (e.g., sample thickness, autofluorescence of the material, floating of spheroids during imaging) direct monitoring of spheroids in the plate, where they were produced, may not be possible. The protocol explains the preparation of spheroids for imaging, which are suitable for most inverted widefield and confocal microscopes.
    1. Prepare and prewarm (37 °C) imaging media: DMEM supplemented with HEPES-Na, pH 7.2 (10 mM), sodium pyruvate (1 mM), L-glutamine (2 mM), and glucose (5 mM), without phenol red.
      NOTE: Sodium bicarbonate alone or in combination with HEPES-Na can be used, if CO2 control is provided during imaging24. Some cell culture types cannot tolerate the presence of HEPES. Depending on the experimental design, pyruvate, glutamine, and glucose content can be modified.
    2. Prepare sterile microscopy dishes (commercially available or lab-made) with coated (for a strong spheroid adhesion) or non-coated (low spheroid adhesion) cover glass surfaces (thickness #1.5).
      NOTE: The need and the type of coating depend on the cell type, adhesion properties of spheroids, and the rate of their cell migration from 3D to 2D culture interface. This is important to consider, as the coating can facilitate the loss of 3D organization, changing the shape of micro-gradients in spheroids and, as a result, cell behavior. For some experiments (e.g., imaging analyzed the response to drug stimulation), strong spheroid adherence to the surface is required, and a coating with gelatin, BMM, collagen, collagen / poly-D-lysine, or poly-D-lysine is preferred.
    3. Gently wash out the O2 probe-stained spheroids from the microwells of the micropatterned agarose or 96-well plate and transfer still floating spheroids into a 2 mL vial.
      NOTE: To ensure the collection of all spheroids from the high throughput method, rinse the mold 1-3 times with the additional volume of culture media, combining all spheroid suspensions in one vial. For spheroids in a low attachment plate, collect spheroids one by one from individual wells to a vial or directly to a microscopy dish if a small number of spheroids is sufficient for the experiment. When transferring big spheroids, cut the end of the pipet tip to ensure no damage during pipetting.
    4. Leave the vial in a vertical position for up to 5 min to let the spheroids settle down on the bottom of the vial, forming a visible pellet.
    5. Remove the media from the tube, leaving spheroids undisturbed, and gently resuspend them in a sufficient amount of fresh culture.
      NOTE: For the convenience of spheroid handling, transfer of spheroids to the imaging media can be also done at this stage in small batches; see steps 2.1.7 and 2.1.8. Spheroids from the different experimental groups should be treated equally, as the media composition and media preconditioning time can affect their metabolism.
    6. While spheroids are floating, transfer an equal volume of spheroid suspension per well of the microscopy dish.
    7. Incubate spheroids for 1-2 h at 37 °C in a CO2 incubator to ensure their attachment to the surface of the microscopy dish/ well. For imaging, proceed with step 2.1.9. For staining of spheroids with additional probes proceed with step 2.1.8.
      NOTE: The rate of cell migration from the 3D spheroid to the 2D surface interface is a function of time. To avoid a loss of 3D organization, incubation time must be optimized with respect to cell type, surface coating type, and the design of the experiment. For example, HCT116, depending on spheroid size, needs at least 2 h for a proper spheroid attachment to the collagen IV/poly-D-lysine coated surface, while hDPSC attachment and migration to 2D interface is extremely quick, leading to loss of 3D organization in 1-2 h. To avoid imaging '2D spheroids' due to excessive spreading, non-coated glass surfaces are used with the decreased incubation time.
    8. Add fluorescence probe(s) in recommended or empirically optimized concentrations to a known volume of spheroid suspension. Incubate for 1 h at 37 °C CO2 incubator prior to imaging.
      NOTE: For live/dead assay, use propidium iodide and Calcein Green-AM in a standard final concentration 1 µg/mL. To avoid the toxic effect of propidium iodide staining on iPSC embryoid bodies, the final propidium iodide concentration was 0.5 µg/mL. The probe loading time can be prolonged if the diffusion of the probe is not efficient due to the big spheroid size. The loading time should always be considered as a part of the total incubation time needed for spheroid attachment to the surface. If a longer time is needed for spheroid attachment, the staining procedure should be arranged at the end of this period. Be aware that longer incubation time might lead to the loss of 3D organization.
    9. Remove cell culture media or media containing fluorescent probes and exchange it for a necessary volume of imaging media. To ensure that no media fluorescence background is on the way of spheroid imaging repeat the media exchange (washing) step up to 5 times.
      NOTE: To avoid spheroid removal during media exchange, it is recommended to carefully aspirate media with a 200 µL pipette from the edges of the microscopy dishes and perform the addition of media by the wall or sidewise in the microscopy dish.
    10. Immediately proceed with step 2.2.1 of the imaging protocol.
      NOTE: A too long break between preparation to imaging and actual imaging acquisition can influence cell metabolism (e.g., via changed medium composition), viability (some fluorescent probes used for endpoint analysis have toxic effects, which can stimulate cell death after a long incubation period) as well as lead to a loss of 3D organization. If multiple groups of spheroids or experimental conditions must be compared, the experimental design must be developed accordingly to keep the timing of the treatment, preconditioning, and imaging procedures as equal as possible between the analyzed groups.
  2. Image acquisition
    NOTE: The protocol describes the multiparametric imaging of live spheroids using Stellaris 8 Falcon (Leica) confocal microscope and Leica Application Suite X (LAS X) software version 4.7. However, only minor modifications will be required to perform such an analysis on alternative microscopy platforms.
    1. Turn on the temperature climate control unit 30-60 min prior to imaging. Set the necessary speed of ventilation and temperature (35-37 °C).
      NOTE: If in addition to temperature control, gas concentration (e.g., CO2 or O2) must be controlled during the imaging, the corresponding devices should also be started in advance to reach the necessary conditions prior to imaging.
    2. Turn on the microscope and connected devices (i.e., WLL laser, computer, water pump for the water immersion objective, and other operating electronic blocks). Start the microscope control software (e.g., LAS X Machine Mode or Machine Mode with Environmental Control) provided with the exact microscope set-up and initialize the stage calibration.
    3. Choose the required objective in the software and apply the immersion fluid if required.
      NOTE: For live microscopy, it is recommended to use either water or glycerol immersion objectives having sufficient ('long') working distance, e.g., HC Fluotar L 25x/0.95 W VIS IR (2.4 mm working distance), HC PL Apo 40x/1.25 GLYC (0.35 mm working distance) or at least NA = 0.4 or higher for the air objectives. The choice of magnification and working distance depends on the nature and size of the imaged sample and measured fluorescence signals (brightness, quantum yield, staining efficiency, see e.g., discussion on dyes and nanoparticles53). Big objects (spheroids or organoids, >500 µm size), 'bio-reactor' or microfluidic chips require long working distance objectives and lower magnification, while analysis of individual cells or cell organelles requires high magnification, often achieved via 'mosaic' imaging.
    4. Set up the microscopy dish with spheroids on the stage. Adjust the focus and find an object/region of interest (ROI).
      NOTE: If small, weakly fluorescent, poor contrast, or rare objects must be found and finding the focus is difficult, it is recommended to pre-focus on the walls of the microscopy dish and 'screening' the surface for the object of interest by serpentine, starting from one of the corners of the well.
    5. Choose the Open Project window and click the corresponding icon, Create a New Project. Give a standard name (e.g., starting from "YY-MM-DD+ description") to the research project file. During the imaging, all produced images will be automatically saved into the created project .lif file.
    6. Open the Acquisition window. Set the white light laser (WLL) excitation wavelength and the required range of hybrid or resonance scanning detectors (HyD S, HyD X, or HyD R type) based on the known spectral properties of the measured fluorescence (excitation/absorbance and emission spectra). Choose Line or Frame types of scan.
      NOTE: For most commercially available fluorescent dyes, the spectral properties can be found (or added) in the LAS X Dye Assistant package. Choose the detector with the appropriate spectra sensitivity range compatible with the probe spectral properties and, in the case of FLIM, compatible with photon counting (i.e., HyD X or HyD R). For multiparametric imaging set the WLL in multiple excitation positions (e.g., for simultaneous imaging of FAD/Flavins and two fluorescence channels of ratiometric MMIR1 O2 probe – reference and sensitive, the excitation/emission settings might be 460 nm/510-590 nm HyD X1 and 614 nm/631-690 HyD X3 and 724-800 nm HyD R accordingly in one or two sequential scan sequences). It is important to assign the appropriate detector to collect the emission, as detectors can have different spectral sensitivity54.
    7. (Optional for FLIM) In the Acquisition window, choose FLIM mode to perform imaging combined with photon counting (decay collection). Immediately, an additional 'FLIM module in LAS X software' will be opened to navigate and analyze FLIM data.
    8. (Optional for FLIM) Choose the WLL pulse repetition rate based on the expected fluorophore average lifetime.
      NOTE: The frequency of the laser pulse must be adjusted to collect the full fluorescence decay. This can be done using a Pulse Picker feature installed on the microscope. The overlapping of the fluorescence decay with the laser pulse will lead to the shortening of the estimated fluorescence lifetime. It is recommended to have pulse intervals of 4-5 times longer than the expected average fluorescence lifetime (e.g., 25 ns/40 MHz for lifetimes up to 5 ns). Many pulsed lasers have a fixed 80 MHz repetition rate (ideal only for a range of up to 2-3 ns). This is important for choosing the correct fluorophores for the experiment.
    9. Start the preview imaging using FAST LIVE mode and adjust the fine focus of the imaging object on a section of interest.
      CAUTION: Strictly follow the laser safety rules. Always consider laser safety rules and wait until the imaging has stopped prior to turning the transmission light on, looking into the eyepiece or at the sample.
      NOTE: In FAST LIVE mode, a high-speed scan of 600 Hz (corresponds to a maximum frame rate 4.43/s if bidirectional X scanning mode is used), 256 x 256 pixels resolution are applied automatically to the image to keep the fluorescence safe from photobleaching. Open the pinhole (e.g., to 3-4 AU) and/or increase the laser intensity if the fluorescence signal is too weak to focus on the object. Avoid incomplete decay collection.
    10. (Optional for FLIM) By looking at a pixel intensity histogram appearing during the imaging in a FLIM module window (Live mode), adjust the appropriate laser intensity/pinhole size and resolution to achieve the count rate ~1 photon/laser pulse limit (red line). Avoid going significantly higher than 1 to exclude a risk of pile-up effect. If required, adjust the WLL pulse repetition rate to have a full decay collection in a decay window (to avoid incomplete decay collection, see step 2.2.8).
      NOTE: If the number of photons (intensity) is not sufficient to reconstruct a reliable decay for fitting analysis or phasor plot cloud, apply several scan repeats (frames or lines, or set up scanning time), increase laser intensity and/ or sacrifice the resolution (scanned ROI size). Be aware that decreasing the laser repetition rate requires more photons to be collected for a reliable decay reconstruction, and an additional correction of the imaging parameters may be required. Be aware of the potential impact of intense light and long continuous illumination on cell viability and metabolism55. The negative impact on viability and metabolism can be different in each individual case, depending on the intensity, duration, and wavelength of the excitation light, as well as imaging modality (e.g., one-photon confocal vs. multiphoton imaging). Adjust the imaging parameters accordingly and, if necessary, control cell viability/death by Calcein Green-AM or propidium iodide intensity in pilot experiments56. Where possible, further optimizations of the fluorescence probe staining protocol should be considered to reach an adequate fluorescence signal during live microscopy.
    11. (Optional for 3D z-stack) While in Fast Live set the coordinates, scan direction, and attribute them to Begin and End in the Z-stack window (XYZ Scan Mode). Choose Z-step size or number of steps.
      NOTE: While the software automatically calculates the 'optimal' number of steps, based on used resolution and scan parameters, live 3D reconstruction can normally require a smaller number of steps to achieve fast acquisition, e.g., 1-2 µm step size, 50-100 µm stack size, bi-directional scanning, requiring 2-3 min of the total scan time. Be aware that the subcellular organelles, cells and the 3D cell model can also move during measurements. In addition, due to the light penetration depth and scattering limits, typically only 50-100 µm scanning depth on confocal FLIM can be achieved.
    12. When all necessary settings are applied, start imaging.
    13. Give the image an appropriate name.
    14. Search for the next imaging object in transmission light mode and repeat the imaging procedure with previously optimized imaging settings (steps 2.2.8-2.2.12).
      NOTE: For the intensity-based comparison or intensity ratio analysis (e.g., MMIR O2 probe-based oxygenation analysis), always keep the same imaging settings for all analyzed objects (magnification and objective lens type, laser intensity, power and pulse frequency, excitation wavelength, detectors range, pinhole, scan speed, pixel dwell time and resolution). However, as fluorescence lifetime does not depend on fluorescence intensity and requires an appropriate number of photons to be collected for reliable calculation, the FLIM imaging parameters can be readjusted through the course of the experiment to keep the collected photon numbers comparable between different treatments or experimental conditions. Therefore, for multiparametric analysis where intensity-based and fluorescence lifetime-based analysis are both needed, optimized universal imaging settings must be applied for all objects in the compared experimental groups. For "FLIM-only" comparison it is possible to compare images acquired with slightly different imaging settings as LAS X software provides calculation of the IRF for individual image measurement42. However, for FLIM-fitting analysis outside LAS X (e.g., FLIMfit45) the instrument response function (IRF) should be measured for each different imaging condition, as it cannot be exported from the imaging software. Thus, for the simplicity of the experimental design and workload, it is recommended to apply the same imaging settings for all images in the dataset. Then, the corresponding IRF measurements can be done with the use of quenched or fast-fluorescence lifetime fluorophores (within ps range) with the emission properties of the measured spectral channel57,58,59, by gold nanoparticles luminescence60 or by second-harmonic generation signal for multiphoton FLIM61. In LAS X software, the previously optimized imaging parameters can be loaded for a new project by a right click on the file of interest and choosing Apply Image Settings.
    15. When the imaging session is finalized, save the imaging project. To finalize the imaging session, remove the sample from the microscopy stage and clean the objective from the immersion liquid (if used) according to the standard procedure implemented in the imaging facility. Close the project and the software. Switch off the microscope, lasers, and all the connected devices.
    16. Proceed with imaging data analysis (step 2.3).
  3. NAD(P)H/FAD-FLIM phasor image processing with LAS X FLIM module and FIJI
    NOTE: The protocol describes a fluorescence lifetime analysis of imaged spheroids for frequency domain data on examples of NAD(P)H and FAD/Flavins autofluorescence FLIM. NAD(P)H autofluorescence measurement became a gold standard for metabolic analysis, where short and longer NAD(P)H autofluorescence lifetime components are associated with glycolysis or oxidative phosphorylation (OxPhos), respectively. This can be analyzed by the shift on a phasor plot along the metabolic trajectory toward the measured standards of free-NAD(P)H or protein-bound NAD(P)H31,62. To analyze the trajectory of the metabolic shift, as well as to compare the position of phasor clouds (see the NOTE below step 2.3.6) on a plot between experimental groups, a simplified phasor coordinates analysis, based on the calculation of geometrical center (centroid) of the phasor cloud was implemented29. The described protocol demonstrates the calculation of centroid coordinates in FIJI and measuring the distance between two points on a phasor plot using coordinates (e.g., the distance from a centroid of the spheroid NAD(P)H autofluorescence phasor cloud to a "free NAD(P)H" theoretical point). Similarly, FAD and other autofluorescence signals can be analyzed. A dataset 1 with .lif (LAS X software required) or .ptu file formats for learning this procedure is provided (Supplementary File 2, Supplementary File 3, Supplementary File 4, Supplementary File 5, Supplementary File 6, Supplementary File 7, Supplementary File 8, Supplementary File 9, and Supplementary File 10).
    1. Open the FLIM module in LAS X, select Open Project, and load the spheroid image file (.lif) for autofluorescence NAD(P)H/FAD analysis.
      NOTE: Due to potential bugs and intermediate data loss in the FLIM module, use a copy of the original spheroid image file (.lif) for NAD(P)H/FAD analysis, keeping the raw file unchanged.
    2. Select a single image and navigate to the FLIM analysis interface. Click Phasor to access the phasor plot and activate the phasor analysis mode. Apply filter (Median or Wavelet) and set Threshold to minimize the noise and improve the data visibility for all phasor analyses. Choose the harmonics. For analysis of the metabolic shift based on NAD(P)H data, proceed with steps 2.3.3-2.3.5. For the general comparison of phasor plots, proceed from step 2.3.6.
      NOTE: Apply analysis settings (filter type, harmonics, threshold, binning, and phasor ROIs) equally to all images in a compared data set.
    3. (Optional for NAD(P)H analysis) Choose any dataset-related image and use the Draw Ratio Cursor for Two Components option to accurately locate the position of 0.45 ns on a universal circle of a standard phasor plot space. This position will be assigned to an average fluorescence lifetime of a pure homogeneous solution of free-NAD(P)H, which normally is close to the mono-exponential decay62.
      NOTE: Free-NADH and free-NAD(P)H have similar spectral properties and similar values of fluorescence lifetime in water solution, with two short lifetime components, 0.3 ns and 0.7 ns63. Thus, for the simplicity of the phasor-based analysis and due to a small difference between lifetime components their fluorescence decay is accepted to be mono-exponential, which allows allocation of the phasor cloud on a universal circle. The reference average lifetime of a free-NAD(P)H form can also be measured and plotted in a phasor space for a similar analysis. The reference lifetime was chosen based on the literature62; note that in other sources, slightly different value of a free-NAD(P)H in solution can be found (0.4 ns64).
    4. (Optional for NAD(P)H analysis) Export the phasor plot with allocated free-NAD(P)H lifetime (see step 2.3.3) by right-clicking on the plot and selecting Export Data. Export the phasor plot as a .tiff format file to a designated storage folder.
      NOTE: The original pixel size of the exported phasor plot image from the LAS X FLIM module is always 1024 x 600 pixels. If another software is used for data export and pre-analysis, be sure that all phasor plot images are exported with the same size and resolution.
    5. To export spheroid-related phasor cloud, utilize the Draw Cursor tool in the LAS X FLIM module to choose the spheroid ROI on the image. Export generated phasor plot as outlined in step 2.3.4.
      NOTE: The corresponding g and s (similar to x and y) coordinates of the phasor space will be assigned to every pixel of the chosen ROI, according to their lifetimes, transformed into a frequency domain data set64,65. The cluster of pixels with similar values of tf (phase lifetime) and tm (modulation lifetime) will form a cloud pattern (phasor cloud) on a plot, where color coding (with a range from deep blue to red) will reflect the abundance of lifetime values. By the position of the cloud on a universal circle or inside, the mono- or multi-exponential decays can be distinguished. Some measurement media exhibit strong (auto)fluorescence, leading to the appearance of a corresponding cloud on a phasor plot, which cannot be simply removed with the means of intensity threshold. This pattern will influence centroid coordinates calculation and must be excluded from the exported phasor plot. Working with the spheroid ROI allows the exclusion of the non-related pixels from further phasor analysis.
    6. Repeat the phasor plot export procedure for all spheroid ROIs in a data set (see steps 2.3.4 and 2.3.6). Additionally, check the exported set of .tiff images to guarantee the full set of data for further comparative analysis and ensure that all exported images have the same pixel size (see NOTE in step 2.3.4).
      NOTE: At this stage of the protocol, the image set must include phasor with the free-NAD(P)H location (based on literature or empirically obtained data) and all spheroid ROI (or alternative ROI patterns if needed for the specific analysis) phasor plots. From this step, further analysis will be done in FIJI and afterward in a spreadsheet. Using the Analyze tool window, Set Scale option in FIJI, and ensure all phasor plot images are calibrated with the same unit type, e.g., only in pixels. If not, set the unit length in the Set Scale window for the chosen one (e.g., for pixel-based scale, put 1 in the Distance in Pixels field and set the unit of length to Pixel). For further comparison, measure all exported data using the same unit type.
    7. (Optional for NAD(P)H analysis) Determine the pixel point position of a free-NAD(P)H average fluorescence lifetime on the corresponding exported phasor plot image (see step 2.3.3): Open the phasor image with FIJI, magnify the image to clearly visualize with a pixel resolution the intersection between universal semi-circle and the Ratio Cursor for Two Components line; use Rectangle ROI tool to select the intersection.
      NOTE: Ensure that the rectangular selection is a small area around the point of intersection for accurate determination of its coordinates in the next step (step 2.3.9).
    8. (Optional for NAD(P)H analysis) Open the Analyze tool, choose Set Measurements window, and select Centroid as a measurement parameter. Click on Measure in the Analyze tool window to determine the centroid coordinates of the free-NAD(P)H reference point. Export these coordinates to a spreadsheet.
      NOTE: Free-NAD(P)H coordinates will be used as the reference point to compare the distances from this point to the spheroid phasor cloud position in a data set (the way to characterize the metabolic shift between glycolysis and OxPhos in NAD(P)H FLIM autofluorescence analysis)
    9. Using FIJI, open the image of the spheroid phasor cloud. Open the Image window tool, choose Adjust, and select Color Threshold in the toolbar. Select the Thresholding Method of choice (e.g., Otsu), and set the Hue Value and Brightness Value to narrow the parameters for the selection of a phasor cloud part with the most abundant pixel coordinates. Click Select to define the cluster area.
      NOTE: Keep the same threshold parameters for all phasor plot images, which must be analyzed. For the presented NAD(P)H-FLIM data, the thresholding method Otsu with set Hue Value 9 and Brightness Value 160 was chosen and applied to all phasor images. Alternatively, the selected area can be copied to ROI Manager (follow the path Edit > Selection > Add to Manager) to create a library of phasor ROIs for further analysis.
    10. While keeping the selection, calculate the centroid coordinates of the selected area following the procedure described in step 2.3.9. Export these coordinates to the spreadsheet file.
    11. Repeat steps 2.3.9 and 2.3.10 to determine centroid coordinates for all ROI phasor images to create a data set in the spreadsheet.
      NOTE: Using ROI Manager ROI library helps to simplify and organize the ROI analysis (see step 2.3.10)
    12. (Optional for NAD(P)H analysis) Open the spreadsheet with exported reference free-NAD(P)H and spheroid ROIs coordinates from different comparison groups. Calculate the distance between each individual Spheroid phasor centroid to reference free-NAD(P)H position using determined coordinates and the following equation:
      Equation 1
      Where, Xc and Yc are centroid coordinates, Xf and Yf are the reference coordinates.
      NOTE: Application of centroid parameter for determination of the shift toward the reference lifetime is appropriate only in case all centroids from a data set are lying on the same linear trajectory toward the reference point. To check this, all centroid points from the data set have to be plotted together with the reference point in the same coordinate space, and the linear trend alignment should be performed. If the R2 coefficient of the linear trend line drawn through all points is close to 1 (e.g., R2 is 0.8-0.99), the distance analysis is assumed to be appropriate.
    13. Organize all data accordingly for comparison and perform statistical analysis with the use of any corresponding software (e.g., Origin, MatLab). Choose the appropriate statistical test according to the data set characteristics (distribution normality, statistical units' number, etc.).
      NOTE: For NAD(P)H analysis, compare distance values to characterize the metabolic shift depending on experimental conditions. For comparison of any phasor plots between experimental groups, perform the comparison of ROI phasor cloud centroids coordinates.

Representative Results

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
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
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
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
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
Figure 6Example 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
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.

Discussion

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.

Disclosures

The authors have nothing to disclose.

Acknowledgements

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.

Materials

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

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Cite This Article
Debruyne, A. C., Ferrari, G., Zhou, H., Van Loon, N., Heymans, N., Okkelman, I. A., Dmitriev, R. I. Production and Multi-Parameter Live Cell Fluorescence Lifetime Imaging Microscopy (FLIM) of Multicellular Spheroids. J. Vis. Exp. (210), e66845, doi:10.3791/66845 (2024).

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