Magnetic molding of tumor spheroids: emerging model for cancer screening

Iron oxide nanoparticles were produced by PHENIX Laboratory (UMR8234, Paris) using the standard Massart procedure of iron salts co-precipitation [35], with the resulting magnetite (Fe₃O₄) nanoparticles then being oxidized into maghemite (γ-Fe₂O₃). The magnetic nanoparticles were then stabilized in aqueous solution via citrate chelation in order to achieve electrostatic repulsion. The resulting nanoparticles possessed an average diameter of 8 nm.

CT26 murine colon carcinoma and U-87 MG human glioblastoma cells (ATCC^®) were cultured in Dulbecco's Modified Eagle Medium (DMEM, Gibco^®) supplemented with 10% fetal bovine serum (Gibco^®) and 1% penicillin-streptomycin (Gibco^®) and grown at 37 °C in a humidified incubator with 5% CO₂. Upon reaching the desired confluence, the cells were detached with 0.05% trypsin-EDTA (Gibco^®).

The cells were labeled with a solution of magnetic nanoparticles at [Fe] = 1 mM suspended in cell medium and left to incubate overnight before detachment from the culture flasks. The iron uptake per cell was calculated through single cell magnetophoresis, as described previously [36]. Briefly, the measurement is based on the velocity of a single magnetically labeled cell as it is attracted by a known, constant magnetic gradient. The drag force of the cell F_d =3πηDν thus balances the magnetic force F_m = M_cell × gradB, yielding the magnetic moment of a single cell through equation (1):

$$\begin{equation}{M_{{\text{cell}}}} = {{ }}\frac{{3\pi \eta D\upsilon }}{{gradB}}{{ }}\end{equation} \tag{ 1 }$$

where η is the viscosity of the carrier medium, D is the cell diameter, ν is its velocity, M_cell its magnetic moment and gradB the magnetic gradient. The M_cell value is related to the number of loaded particles per cell, from where the total iron mass per cell can be calculated.

The method is based on the magnetic attraction of labeled cells towards a fixed magnet, a force which ultimately drives cell aggregation into a defined shape. Briefly, a 22.1 cm² petri dish (TPP^®) is placed on top of a network of 6 × 2 mm cylindrical neodymium magnets (Supermagnete). Then, stainless steel beads (CIMAP) with a diameter of 0.4, 0.5, 1 or 1.6 mm are placed inside the dish and fixed into position as they are attracted to the magnets. A previously heated and homogenized 2% agarose (A0576, Sigma-Aldrich) solution in phosphate buffered saline is then poured into the dish in a sufficient amount to leave the top part of the steel beads uncovered. After agarose gelation, the beads are carefully removed and a quasi-spherical mold is obtained, which ultimately defines the spheroid shape. The molds are sterilized by UV light exposure for 30 min and filled with cell medium. Then, using the network of cylindrical magnets, previously detached and concentrated labeled cells (100 000 cells µl⁻¹) are seeded into each mold. The attraction of the cells to the magnet drives their aggregation into the spherical mold, forming an easily observable spheroid within minutes. The number of cells attracted into the mold is on average 11 400 ± 1700, 24 000 ± 6000, 106 000 ± 15 000 and 180 000 ± 22 000 for molds of 0.4, 0.5, 1 and 1.6 mm in diameter, respectively, as determined by trypsinization and cell counting of over eight spheroids for each condition. After 3 min of cell seeding, the magnets are removed. The dish is then carefully washed, filled with cell medium and then placed in the incubator to allow spheroid maturation. After overnight incubation, spheroids are removed from the agarose mold with the aid of a micropipette. For all experiments performed, spheroids were placed in plasticware coated with Anti-Adherence Rinsing Solution (07010, STEMCELL™ Technologies) to prevent adhesion, unless otherwise specified.

After cell detachment from the culture flasks, they were counted using the standard trypan blue technique. Single drops of 30 µl of cell medium containing 5000 cells were pipetted onto the lid of a 60 cm² tissue culture dish (TPP^®), evenly separated. The lids were then inverted and placed on the dishes, each one of the latter filled with 10 ml of PBS to prevent evaporation of the drops. The hanging drops were cultured under the previously described conditions, and spheroids were collected after 2 d of incubation. The spheroids were placed in plasticware coated with Anti-Adherence Rinsing Solution (07010, STEMCELL™ Technologies) to prevent adhesion for all experiments, unless otherwise specified.

The spheroid morphology and growth was monitored using a Leica DM IL LED microscope (Leica Microsystems) coupled to a Canon EOS 50D digital single-lens reflex camera (Canon^®). Images taken at the desired time points were analyzed using the ImageJ open source software for diameter and sphericity quantification. For the latter, the sphericity index (SI) was defined by equation (2) [15]:

$$\begin{equation}{{\Phi }} = {{ }}\frac{{\pi \sqrt {\frac{{4A}}{\pi }} }}{P}{{ }}\end{equation} \tag{ 2 }$$

where A is the projected area of the spheroid and P its perimeter.

The alamarBlue™ metabolic assay (DAL1100, Invitrogen™) was used to assess the proliferative potential of single spheroids and following the vendor's standard procedure in 96-well plates. A single spheroid was measured for all instances, with an incubation time with the reagent set at 2 h, and the fluorescence was recorded with an EnSpire^® Multimode Plate Reader (PerkinElmer), using a fluorescence excitation and emission wavelength of 570 and 585 nm, respectively.

The LIVE/DEAD™ Cell Imaging Kit (R37601, Invitrogen™) was used to assess cell viability in live spheroids. Following vendor protocols, spheroids at day 3 of maturation were stained with Live Green/Dead Red staining solution in non-adherent dishes for 2 h at 37 °C in a humidified incubator with 5% CO₂. Then, the staining solution was removed and the spheroids were then stained for the cell nuclei with Hoechst 33342 (H3570, Invitrogen™) at a 1:200 dilution in DMEM without phenol red (Gibco^®) for 30 min at room temperature. The spheroids were then imaged using an Olympus IX81F-3 inverted microscope (Olympus^®) coupled with a laser dual spinning disc unit (Yokogawa CSU-X1) and an Andor iXon^EM + CDD camera (Andor™ Technology), with a 10x objective. Images were processed using ImageJ, using the Grid/Collection Stitching plugin for spheroid reconstruction.

Liposomal-encapsulated doxorubicin (300112S-1EA, Avanti^® Polar Lipids) was selected to study drug resistance in magnetically molded spheroids due to its clinical relevancy. After 1 d of maturation, the spheroids were placed in 96-well plates at increasing doxorubicin concentrations in the range of 0.01–100 µg ml⁻¹. Following a 72 h incubation period, the cell death percentage was evaluated using the alamarBlue™ metabolic assay following the previously described protocol used to monitor spheroid growth. 2D cell cultures at a seeding density of 25 000 per well were also prepared in parallel under similar conditions for comparison purposes. The 50% inhibition concentration (IC₅₀) value was determined from the dose-response curves after fitting to a four parameter logistic regression model [37].

For fluorescence imaging, the spheroids were fixed in 4% paraformaldehyde in phosphate buffered saline for 1 h at room temperature after the desired incubation time with doxorubicin at 10 µg ml⁻¹. The spheroids were then washed and embedded in optimal cutting temperature compound (361603E, VWR™) for 1 h at room temperature and shortly after frozen in 2-Methylbutane, GPR Rectapur^® (VWR™) cooled with liquid nitrogen, and then stored at −20 °C. Cryosections of 20 µm were obtained from the center area of each spheroid using a Cryostat (CM3050 S, Leica Microsystems) and then mounted on slides using Fluoromont™ Aqueous Mounting Medium (F4680, Sigma-Aldrich). All samples were stored at 4 °C after gelation of the mounting medium. Imaging of doxorubicin fluorescence was performed using the confocal microscopy setup described previously, with a 60x oil immersion objective. Images were processed using ImageJ, using the Grid/Collection Stitching plugin for spheroid reconstruction.

Matrigel^® Basement Membrane Matrix (356234, Corning^®) at 9.8 mg ml⁻¹ of protein concentration was used to study cell invasion at the periphery of growing spheroids. Pre-chilled 96-well plates were coated with 40 µl of Matrigel following standard vendor procedures. After Matrigel gelation for 30 min at 37 °C, a single spheroid at the desired maturation time was placed in each well and allowed to grow and spread. Images at each stage of invasion were taken using the previously mentioned optical microscope setup, and image analysis to quantify the area of spread was done using ImageJ.

Spheroids were collected after 1 d of maturation in the molds. The measurement of surface tension was either done right away (day 1) or after two more days of maturation (day 3). For the measurement, the spheroids were placed in a 37 °C thermoregulated chamber filled with Dulbecco's Modified Eagle Medium without phenol red (Gibco^®). The chamber was enclosed with glass slides on two of its sides to allow for side view imaging, and at its bottom with a glass slide coated with Anti-Adherence Rinsing Solution (07010, STEMCELL™ Technologies) in order to ensure non-wetting conditions. A cylindrical neodymium permanent magnet (6 × 6 mm), generating a uniform magnetic field gradient (gradB= 170 T m⁻¹) within a cylindric volume of 2 mm in height and 2 mm in diameter, was positioned under the spheroid, in direct contact with the bottom glass slide. Upon magnet application, the spheroids become compressed in the direction of the magnetic field gradient. The spheroid's lateral profile was then monitored with a digital camera (QICAM FAST 1394, QImaging) for 10 min until its equilibrium compressed shape was reached. This equilibrium shape is determined by the balance between magnetic forces per unit of volume, f_v with f_v = M_v × gradB and capillary forces (surface tension). The experimental profile of each compressed spheroid was next computed using ImageJ edge recognition, and fitted to the theoretical profile of a drop submitted to a volume force (such as gravity) derived from the Laplace law of capillarity in non-wetting conditions. The theoretical profile was obtained using Matlab's function ode45 to numerically integrate the Laplace law [38]. The corresponding fitting directly infers the macroscopic tension of the spheroid.

Magnetic cell labeling of CT26 cells translated to an iron loading dose of 8.1 ± 2.6 pg per cell after endocytosis (figure 1(a)), as determined by single-cell magnetophoresis. The magnetic moment of each cell provided by this nanoparticle load corresponds to an average of 5.3 × 10^–13 A m², and it is the driving force for cell aggregation into magnetic spheroids.

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The magnetic molding process is illustrated in figure 1(b). It is based on the preparation of an agarose mold using a steel bead fixed in place by an array of 6 × 2 mm cylindrical magnets placed below the culture dish. The diameter of the selected bead ultimately defines the spheroid diameter, thus allowing the tuning of this parameter from this very early step. Upon removal of the steel beads, and using the same array of magnets, magnetically labeled cells are seeded on top of the molds and, being attracted to the magnets, aggregate in situ and adopt the spherical shape conferred to the mold by the beads. The magnetically-driven cell aggregation takes place instantaneously, and a spheroid can be visibly observed typically within 3 min of cell seeding. This time of aggregation is similar to a centrifugation process: the magnetic field gradient gradB created by each 6 × 2 mm cylindrical magnet below each mold, in the order of 200 T m⁻¹, translates into a volume magnetic force M_v × gradB, M_v being the cell magnetization (magnetic moment per unit of volume, in the order of 500 A m⁻¹). The range of this volume force, around 10⁵ N m⁻³, would be equivalent to 100 g, assuming the cell density close to 1.1 relative to water. Remarkably, it is in the same order of force applied under a routine cell culture centrifugation at 100–300 g, also applied for a few minutes. After cell aggregation in the molds, the magnets are removed. More importantly, mature and cohesive spheroids can be collected under 1 d of incubation within the mold.

We proceeded to show the proof of concept of the methodology by selecting a variety of steel bead sizes, and compared spheroid sphericity, average diameter and cell proliferation with the standard hanging drop technique. Figure 2(a) shows the typical spheroid morphology for an initial bead size of 0.4, 0.5 and 1 mm, as well as for hanging drop spheroids. The magnetic spheroids possess a slightly smaller initial diameter (day 1 of maturation) than that imposed by the mold, at 344 ± 18, 435 ± 15 and 820 ± 25 µm for the initial conditions of 0.4, 0.5 and 1 mm, respectively (figure 2(b)). For all magnetic molding conditions, highly spherical spheroids (SI ≥ 0.9) can be observed from day 1 and up to day 5 of culture, with negligible variability (figure 2(c)). On the other hand, the hanging drop spheroids can only attain the same consistent high level of sphericity towards their 5th day of maturation.

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Quantitative cell proliferation data was obtained using the alamarBlue™ metabolic assay. Magnetic spheroids of sizes of 0.4, 0.5, 1 and 1.6 mm, as well as hanging drop ones, were prepared in order to assess the correlation between the alamarBlue™ fluorescent signal of a single spheroid and the total number of cells within it, after its dissociation with trypsin. The correlation curve is shown in figure 3(a). Remarkably, the proliferation data are perfectly comparative for each spheroid size and/or condition, demonstrating the robustness of the assay in providing a reliable estimation of the number of cells within a spheroid. Figure 3(b) shows the cell proliferation in 0.4, 0.5 and 1 mm magnetic spheroids, as well as hanging drop ones, in relation to their maturation in days. For all conditions, the spheroids were still in a proliferative state up to day 5 of maturation, with proliferative rates correlating to spheroid size. Interestingly, the proliferation rate of CT26 cells with the hanging drop method can be matched by that of the magnetically molded spheroids with a diameter of 0.4–0.5 mm of initial mold size.

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The viability of the magnetic molding spheroids was additionally characterized using the LIVE/DEAD™ Cell Imaging Kit. Spheroids at day 3 of maturation were stained with the Live Green/Dead Red staining solution, as well as with Hoechst 33 342 dye to observe the cell nuclei, and then imaged by confocal microscopy. Characteristic images of the fluorescent distribution for each spheroid condition is shown in figure 3(c). Only the Live Green (calcein) signal was observed, homogeneously throughout the spheroid for all conditions. However, it should be noted that the imaging corresponds to the outer and first layers of cells within the spheroid, as it remains a difficult prospect to image a spheroid's core in live settings, especially ones with the diameters shown here.

Finally, we demonstrate that the magnetic molding method can accommodate other cell lines by using the U-87 MG glioblastoma cell line. Figure 4 shows the resulting spheroids obtained from 1 mm molds, featuring a very similar diameter and cell proliferation profiles to those of CT26, highlighting the robustness of the method.

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We investigated the cell penetration and distribution of liposomal-encapsulated doxorubicin (DOX) in magnetically molded spheroids of 1 mm initial size. Figure 5(a) shows the fluorescent intensity of DOX of a typical cross-section at the center of the spheroid, as determined by confocal microscopy imaging. Spheroid maturation times of 4 h, 1 d and 3 d were analyzed at different DOX incubation times. The maturation time point of 4 h was chosen for its relevancy as a tumoral model of early maturation and thus of less cohesiveness. For this early maturation condition, DOX penetration can be observed after 2 h of incubation throughout the area of the spheroid in a discrete manner, with full cellular penetration taking place. The drug penetration increases considerably after 1 d of incubation, and reaching higher levels after 3 d. For spheroids at day 1 of maturation, there is little to no drug penetration after 2 h incubation with DOX. However, at 1 d of incubation drug penetration can be observed at the periphery of the spheroid, at approximately 50 µm from its border periphery, with a significant increase after 3 d of incubation. Lastly, spheroids at day 3 of maturation saw no DOX penetration after 2 h of incubation, but some drug fluorescent signal was observed after 1 d of incubation all around the periphery of the spheroid, at 50 µm in depth from its border. We then observed the penetration of DOX in hanging drop spheroids at two days of maturation and incubated with the drug for 1 d (figure 5(b)). Similarly to the longer DOX incubation times in the 1 mm magnetic spheroids, a high DOX fluorescent signal was detected throughout the entire hanging drop spheroid. This can be explained by the spheroid cohesiveness: whereas the magnetic one is well into its maturation state, the hanging drop spheroid has barely achieved a cohesive state by this maturation time, which translates to a more uniform drug penetration profile.

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The alamarBlue™ metabolic assay was used to determine DOX cytotoxicity in CT26 cells cultured in standard 2D conditions, in magnetic spheroids of 0.5 mm and 1 mm of initial size at day 1 of maturation, and in hanging drop spheroids at day 2 of maturation (figure 5(c)). The IC₅₀ value was calculated to be at 7.4 µg ml⁻¹ (13.6 µM) for CT26 cells cultured in 2D, in the same range as reported elsewhere [39]. After 3 d of DOX incubation, and for the three spheroid types, the cells respond better to the treatment at low doses than cells cultured in 2D, but the effect is not amplified at higher doses, with the spheroids' metabolic activity saturating at 40% and 60% for the 0.5 mm and 1 mm spheroids, respectively. The hanging drop spheroids' drug response is very similar, and almost parallel, to that of the 0.5 mm magnetic ones. It should be remarked that the magnetic spheroids were placed in the DOX drug at day 1 of maturation, whereas the hanging drop ones could only be collected and placed in the drug after 2 d of maturation. The cells in the 3D spheroid configuration thus show a higher resistance to DOX compared to those in 2D culture, with the 1 mm spheroids showing the highest one. This indicates a limited penetration of the drug towards the spheroid core, with an observable increase in 0.5 mm spheroids.

We next evaluated the cell invasion potential of magnetically molded spheroids in order to further characterize the suitability of the method for assessing cell invasiveness potential as an experimental output. We selected Matrigel as a culture basement membrane matrix for this purpose.

Magnetic spheroids of 0.4, 0.5 and 1 mm initial size were placed on Matrigel after 4 h, 1 d and 2 d of maturation, and their area of growth and cell invasion into the layer of Matrigel matrix was quantified. Spheroids formed through the hanging drop method are also shown along with the 2 d of maturation condition for comparison purposes. The 4 h maturation time was chosen to show the advantage of the magnetic attraction ability of the magnetic molding method to produce cohesive spheroids at early incubation time points. This particular experimental setup shows how the cell proliferation under this early maturation condition compares to that of cell invasion studies where longer maturation times are used, a limitation which is imposed by the typical spheroid formation methods.

Spheroids formed through the magnetic molding method and cultured on Matrigel at 4 h of maturation (figure 6(a)) show a similar invasion area of spread compared to that at day 1 (figure 6(b)) and day 2 (figure 6(c)). Magnetic spheroids of the two smaller sizes appear to reach an overall invasion area as high as that of 1 mm ones under all three maturation conditions. Akin to the parameters of spheroid diameter and proliferation, the cell invasion potential of CT26 hanging drop spheroids (day 2 of maturation) shows a similar pattern to that of the magnetically formed spheroids in the lower range of sizes. Remarkably, all spheroids present similar invasion areas over a long term, one week of culture.

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The spheroid's magnetic moment, created due to the magnetic nanoparticle loading, provides it with magnetic responsiveness and allows for its remote stimulation by external magnets. One direct application is the measurement of the surface tension, mediated by magnetic flattening. Indeed, by placing a spheroid on top of a 6 × 6 mm cylindrical magnet developing a magnetic field gradient of gradB= 170 T m⁻¹, the spheroid experiences a volume force created by its magnetization (magnetic moment per unit of volume M_v), which was found to be in the same range for CT26 and U-87 MG spheroids, at 362 ± 48 A m⁻¹ and 414 ± 45 A m⁻¹, respectively. This magnetic volume force can be seen as super-gravity, in the range of 100 g, which compresses the magnetic spheroid in the z-direction and ultimately reaching an equilibrium shape that can be fitted to capillary Laplace equations. From this equilibrium profile of a spheroid in super-gravity, the macroscopic surface tension can be calculated. This parameter, signature of tumor cohesiveness [40], is generally unreachable for typical non-magnetic spheroids without the use of more complicated setups such as micropipettes [41] or microplate devices [42].

Figure 7 shows the surface tension measurement for spheroids made from the two tested cancer cell lines: CT26 colon carcinoma and U-87 MG glioblastoma. Figure 5(a) shows the spheroid lateral profile for CT26 under control conditions and when placed on top of the magnet, at day 1 and 3 of maturation. The surface tension value increased almost two-fold from day 1 to day 3, at 8.8 ± 1.6 and 16 ± 1.8 mN m⁻¹, respectively, indicating a tightening of cell interactions and thus an increase in spheroid cohesiveness within this timeframe (figure 7(b)). U87-MG spheroids show a similar flattened profile when placed on top of the magnet (figure 7(c)). The surface tension at day 1 was about half the one of CT26, with a value at 4.9 ± 1 mN m⁻¹, and its increase towards day 3 was more than 2-fold, to 12.4 ± 1.8 mN m⁻¹ (figure 7(d)).

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