SpheroidSync as edge cutting transfer strategy for uniform and robust MCF7 spheroids in 3D culture

Abstract

Three-dimensional spheroid models play a crucial role in cancer stem cell (CSC) research and drug screening. However, traditional methods often struggle with issues such as inconsistent shapes, difficulties in nutrient diffusion, and technical complexities. In this study, we introduce the SpheroidSync (SS) method, an innovative approach that combines conventional techniques to create uniform and size-adjustable MCF7 breast cancer spheroids at a very low cost, without the need for special growth factors or supplements. Our research involved culturing MCF7 cells using both the standard methods and the new SS method. We meticulously assessed various factors, including spheroid shape and cell viability, through fluorescent staining, colony formation assays, and gene expression analysis. The results revealed that the SS method produced spheroids that were not only uniform but also showed better structural integrity and maintained viability over time compared to traditional methods. Fluorescent viability tests indicated that SS spheroids consistently exhibited healthy cell viability as evidenced by sustained intracellular esterase activity throughout more extended culture periods. In contrast, spheroids generated through conventional methods exhibited declining viability, characterized by core deterioration and uneven staining. Furthermore, gene expression analysis showed a significant increase in CSC markers in SS spheroids, with CD44 levels rising over 40-fold, ALDH1 increasing more than threefold, CD24 decreasing, and HIF-1α elevating over 11-fold when compared to two-dimensional cultures. This establishes the typical breast CSC characteristics and confirms that a hypoxic environment was effectively created. Notably, as esterase activity declined, we observed an increase in stem cell populations, indicating a successful shift towards a quiescent, stem-like state. In summary, SpheroidSync represents a significant advancement in three-dimensional cancer modeling. It enables the production of uniform spheroids in a cost-effective manner, while ensuring long-term viability and enriching CSC populations.

Introduction

Cancer stem cells (CSCs) in certain solid tumors, such as breast cancer, are particularly significant for understanding how cancers develop and devising effective therapies^1,2. These rare cells possess unique traits, including the potential for self-renewal, multi-lineage differentiation, and the ability to give rise to malignancies. These features play a significant role in the heterogeneity of breast cancer. Research indicates that CSCs are responsible for recurrence and metastasis to other body parts. They often resist standard treatments like chemotherapy and radiation. The way tumors are organized, with CSCs driving growth and malignancy, underscores their importance as therapeutic targets^3,4.

Spheroids are crucial for studying CSCs, especially in breast cancer, because they can closely mimic tumors’ three-dimensional (3D) architecture. Creating spheroids enables researchers to study how cells interact with each other and the communication pathways that are key for CSC maintenance and tumor progression⁵. The creation of 3D cancer models bridges the gap between 2D and in vivo research, significantly reducing the number of animals sacrificed in preclinical experiments. These models, with their ethical and financial advantages, are powerful tools for predicting tumor responses to treatment. Multicellular tumor spheroids (MCTS), the most popular 3D in vitro model in preclinical cancer research, can replicate key features of tumors, including quiescent cell subpopulations and gradients of nutrients and oxygen (hypoxia)⁶.

MCF7 cells are a widely used breast cancer cell line that was established in 1973 from the pleural effusion of a 69-year-old woman named Frances Mallon, who had metastatic breast cancer. MCF7 stands for Michigan Cancer Foundation-7, reflecting the institution where Dr. Herbert Soule and colleagues developed the cell line. This cell line is significant because it was one of the first to be cultured successfully for extended periods, allowing researchers to study breast cancer biology in a controlled environment⁷.

MCF7 is a hormone-dependent cell with estrogen and progesterone receptors (ER and PR). MCF7 is a hormone-dependent cell with estrogen and progesterone receptors (ER and PR). Regarding metabolism, MCF7 cells are more Pasteur-type, depending on oxidative phosphorylation to produce ATP in normoxic circumstances, but they become more glycolytically active in hypoxic situations. Unlike MDA-MB-231 cells, which are more mesenchymal and have also been shown to be multidrug-resistant, MCF7 cells exhibit the epithelial phenotype⁸. MCF7 spheroids are superior to other cell types and models of breast cancer because they can form well-defined 3D spheroids that closely resemble the in vivo tumor microenvironment, making them a more physiologically relevant model for drug testing and cancer research. In contrast to traditional 2D cultures, these spheroids exhibit a drug resistance profile similar to that of solid tumors, making them particularly useful for assessing therapeutic efficacy and mechanisms of action in a context that reflects the complexities of actual tumor behavior. Furthermore, the portrayal of tumor-stroma interactions is enhanced by co-culturing MCF7 spheroids with fibroblasts, which is crucial for understanding tumor development and therapy response^6,9–11. MCF7 spheroids have several formation issues that affect how well they work as cancer research models. One significant problem is necrosis, which occurs when spheroids exceed a diameter of 700 µm, particularly when cell densities are high⁶. This causes core cells to become oxygen-deprived and nutrient-poor. Furthermore, MCF7 spheroids frequently exhibit non-aggressive behavior and irregular shapes, making it challenging to replicate tumor dynamics¹² precisely. Since changes in cell density and culture conditions can lead to variations in size and shape, achieving repeatability is another significant challenge⁹. The mechanical characteristics of the surrounding matrix are also important; increased rigidity can alter how proteins are expressed and how cells behave within the spheroids¹¹. Ultimately, the aggregation process can lead to an imbalance of reactive oxygen species (ROS), which may compromise cell health and contribute to cell death if not properly regulated¹².

Cultivating MCF7 spheroids using agarose and hanging drop methods presents several challenges. In agarose cultures, although the non-adhesive environment promotes cell–cell interactions, it can hinder the formation of perfectly round and repeatable spheroids due to issues with cell accumulation and the relatively non-aggressive nature of these cells^6,13,14. Additionally, the mechanical properties of agarose can restrict the remodeling of the extracellular matrix (ECM), which is essential for accurately mimicking the tumor microenvironment¹³. Conversely, while the hanging drop method facilitates spontaneous spheroid formation, it can be labor-intensive and time-consuming when trying to produce large quantities. This method may also result in inconsistencies in spheroid size and shape due to variations in droplet size and cell culture density. Furthermore, once spheroids are formed, feeding them poses significant challenges; even with additives like methylcellulose to enhance droplet stability, issues can arise with cell interactions and nutrient delivery. Overall, both methods have limitations that can impact the growth and viability of MCF7 spheroids, complicating their use in research^15,16.

On the other hand, the formation of MCF7 spheroids necessitates various essential components, including growth hormones like heat-stable basic fibroblast growth factor (HS bFGF)^17,18, antioxidants such as glutathione (GSH)¹⁹ and, supporting extracellular matrix (ECM)²⁰ constituents. Nevertheless, the integration of these components presents several obstacles. The instability of Specific factors for spheroidogenesis might result in variations in their availability, hence challenging experimental consistency²¹. Moreover, variations in serum-containing medium may lead to contradictory results across trials, complicating the capacity to derive accurate conclusions²². Furthermore, these chemical variables can profoundly modify cellular activity, resulting in responses that diverge from those observed in vivo or in basic two-dimensional cultures, thereby impacting the predictive validity of medication effectiveness studies. The intricacy of cellular interactions within the 3D environment complicates interpretations, as these interactions are only partially duplicated in culture²³. The accompanying expenses and technical problems of sustaining 3D cultures with specific chemical variables may restrict their accessibility for wider research applications, presenting considerable impediments for researchers in this domain²⁴.

By combining several methods, we have developed an effective novel approach that integrates the hanging drop method with a unique transfer mechanism to overcome the limitations inherent in traditional spheroid creation techniques. By cutting the sampler tip to maintain the integrity of the spheroid-like cell sheets, we circumvent the need for viscosity-increasing agents and mitigate the challenges associated with nutrient diffusion and microscopic monitoring. Subsequently, by placing these cell sheets in an agarose gel medium, we generated very homogeneous and spherical spheroids without the need for additional growth agents or supplements. This novel method, SpheroidSync (SS), which we refer to as SS, effectively addresses nutrition and hypoxia issues, thereby enhancing the physiological relevance of MCF7 spheroids for in vitro research. Live/dead fluorescent staining demonstrated sustained shape uniformity and superior viability in SS spheroids, with consistent intracellular esterase function maintained over extended culture periods, while conventional methods showed rapid metabolic decline and compromised cell viability^25,26 (Fig. 1). Additionally, the enrichment of CSCs within the spheroids was confirmed through colony formation assays and quantitative RT-PCR analysis of stem cell markers, revealing a characteristic CD44⁺/CD24⁻/ALDH1⁺ breast CSC phenotype and significant upregulation of HIF-1α, indicative of physiologically relevant hypoxic microenvironments. The progressive decline in esterase activity correlated with enhanced CSC populations, demonstrating successful transition to quiescent, stem-like states that closely mimic in vivo tumor biology. Our methodology offers a comprehensive drug screening and therapeutic research platform, enhancing the efficacy of cancer treatment and CSC therapy options through improved predictive validity and cost-effective implementation.

Fig. 1.

Schematic of the SpheroidSync (SS) workflow. MCF7 cells form compact spheroids in hanging drops and are then transferred to agarose-coated wells for continued culture. Parallel agarose-only and hanging-drop controls run simultaneously. Analyses—including morphology, live/dead Calcein AM/EthD-1 staining, qRT-PCR for CD44/ALDH1/HIF-1α, and extended viability monitoring up to 14 days—validate the SS method’s superior performance.

Materials and methods

Cell culture conditions

The MCF7 human mammary adenocarcinoma cell line, a prevalent model in breast cancer research, was acquired from the Pasteur Institute of Iran (Tehran, Iran). The cells were grown in RPMI 1640 media obtained from Sigma (Germany), augmented with 5% and 10% (vol/vol) heat-inactivated fetal calf serum (FCS, Sigma-Aldrich, Germany). The cultures were sustained under regulated conditions at 37 °C with 5% CO₂ and 90% humidity (Memmert, CO₂ Incubator, Germany). The cells were passaged weekly at 1 × 10⁴ cells/mL to ensure ideal growth conditions.

Generation of spheroids

Hanging drop method

MCF7 cells were cultured under standard conditions until they attained approximately 70–80% confluence. Utilizing sampler tips, 58 μL droplets were deposited on the lids of 10 cm Petri dishes, ensuring each drop contained between 1500 and 15,000 cells. The inverted lid of the suspended drops was placed on a phosphate-buffered saline (PBS)-filled Petri dish to prevent evaporation, allowing the droplets to remain suspended without contacting the bottom. The droplets were categorized into three classes. Some were fed after 24 h. Every 24 h, a limited volume of medium was extracted using a sampler, and fresh medium was introduced. The other group was not fed. The third group was cultured in 0.5% methylcellulose (Sigma-Aldrich) (Fig. 2A). The specimens were incubated at 37 °C in a humid environment with 5% CO₂ for 24–72 h, and spheroid formation was monitored daily by microscopy (Olympus CKX41, Inverted Microscope, product of Olympus, Japan). Methylcellulose was dissolved in sterile water to prepare a 0.5% concentration. Thorough mixing was ensured to prevent clumping.

Fig. 2.

(A) Demonstration of the hanging drop method. Using sampler tips, 58 μL droplets were placed on 10 cm Petri dish lids. The drops were suspended by placing the inverted lid on the PBS-filled Petri dish to prevent evaporation. Plates were incubated for 24–72 h at 37 °C in a humid atmosphere with 5% CO₂. (B) Demonstration of the Agarose method. To form spheroids, cells were resuspended in 150 μL of new culture media with 5% FBS at a density of 1500–10,000 cells per well, depending on the desired size. Plates were centrifuged at 1000 RPM for 5 min after each treatment. We monitored spheroid formation every 24 h on the plate in the incubator.

Agarose method

A 1.5% (w/v) agarose solution was created by dissolving agarose powder (Sigma-Aldrich) in PBS. The solution was heated in a microwave or autoclave until fully dissolved. After cooling and gel binding, the agarose solution was carefully distributed into the wells of a 96-well plate to create a non-stick surface. The gel solidified at room temperature. Before cell cultivation, the plate was sterilized for 60 min using a laminar flow hood and ultraviolet light. Cells were resuspended in 150 μL of fresh culture medium containing 5% FBS at the optimal density for spheroid development, typically between 1500 and 10,000 cells per well, contingent upon the desired spheroid size (Fig. 2B). After every procedure, the plates were centrifuged for 5 min at 1000 RPM. The plate was placed in the incubator, and spheroid development was observed every 24 h. Dimensions and changes were recorded using a microscope and ImageJ software (Version 2.9.0/1.53t; National Institutes of Health, Bethesda, MD, USA; http://imagej.net/Contributors). 50 μL of the used medium was carefully removed from the highest point once every 24 h, and 50 μL of fresh medium was progressively added to the wells from the side.

SpheroidSync (SS) method

Initially, using the Hanging drop Method, the cells were cultivated in quantities ranging from 700 to 12,000 per 58 μL droplets on the petri dish cover (in the absence of methylcellulose) (Fig. 3A). They remained in the incubator for 48 h without any movement. 48 h after cell sheet formation, the sheets were systematically moved to non-adherent wells of 96-well plates containing 1.5% agarose gel. The transfer technique involved edge-cutting the Crystal sampler tips under the laminar hood using a sterile blade, 4 mm from the end of the Crystal tip (Fig. 3B). This was performed because the diameter of the Crystal tips is very small. When the cell sheets are removed, the cells may lose their connection condition, be damaged, or become single-celled, and this must be done gently. Conversely, the limited volume of culture media precluded collecting droplets using the blue and yellow tips. However, with this operation, the diameter of the sampler tip increased, and the cell sheets entered the Crystal tip without getting stuck and messing up the connections. The Petri dish lid was adjusted slightly vertically, and the sampler gradually withdrew the cell sheet (Fig. 3C), transferring it to the wells of the 96-well plate (Fig. 3D). To compensate for minor disruption that may occur during the transfer of cell sheets, the plates were centrifuged for 2 min at 400 RPM. The plate was placed in the incubator, and spheroid formation was monitored every 24 h, with changes in size and alterations documented using a microscope and ImageJ software. Every 24–48 h, depending on the color alteration of the culture media, 50 μL of the media used was carefully extracted from the highest point, and 50 μL of the new medium was gradually introduced into the wells from the side.

Fig. 3.

Demonstration of steps for performing the SS Method. Hanging drop (A). Cutting 4 mL from the end of the Crystal tip (B). Harvesting cell sheets from the drops planted in the lid of the Petri dish (C). Implant the cell sheets in 96-well plates containing agarose gel (D).

Morphometric analysis of tumor spheroids

Based on our previous works^14,27,28, the circularity of spheroids was examined in phase-contrast microscopy images to assess their growth and invasion. During the spheroid formation process, spheroids were imaged using a microscope. Images were taken on days 1, 2, 4, 6, and 8. The diameter of the embedded spheroids in the resulting images was measured using ImageJ software. The outlines of the spheroids were manually extracted, and the diameters of the circles were analyzed using ImageJ software.

Colony formation assay

The spheroid culture was utilized for colony formation following established protocols. The ability to form colonies and self-renew is linked to the stemness capacity of stem cells. Therefore, it is likely that individual colonies capable of forming spheroids (ranging in size from 200 to 400 μm) exhibit characteristics associated with CSCs and their specific markers. Spheroids were passaged (up to three passages) in a serum-free RPMI media, in contrast to Spheroidization, to enrich for CSCs. Research indicates that CSCs can be effectively enriched in a serum-free medium supplemented with essential growth factors, as only CSCs can survive and proliferate under such conditions²⁹. For enrichment, 5000 formed spheroids were collected after 72 h. The cell pellet containing the MCS was gently resuspended in 0.25% trypsin–EDTA and shaken at 37 °C to dissociate the MCS and achieve a single-cell suspension for downstream experiments^30,31. They were then plated in 6-well plates in 0.1% and 0% FBS. As a control, 2D single-cell cultures were also cultured under the same conditions.

LIVE/DEAD viability staining

For live/dead viability staining, the culture medium was carefully drained from the spheroid’s media, and the samples were rinsed three times with sterile PBS (pH 7.4). Hanging-drop spheroids were fixed in 150 µL of 4% paraformaldehyde for 15 min at ambient temperature, followed by washing with PBS; SS-method and agarose spheroids were processed without fixation. The LIVE/DEAD® staining buffer (Calcein AM/Ethidium Homodimer-1 Staining Kit; Molecular Probes, USA) was freshly made by diluting Calcein AM and Ethidium Homodimer-1(EthD-1) stock solutions in PBS to final concentrations of 2 µM and 4 µM, respectively³². For agarose- and SS-method spheroids cultivated in 96-well plates, 150 µL of staining buffer was added per well to submerge the spheroids completely; for hanging-drop spheroids, 50 µL of staining buffer was carefully pipetted onto each drop.

Samples were incubated for 2 h at ambient temperature in darkness, followed by rapid imaging using a Cytation™5 Cell Imaging Multi‐Mode Reader (BioTek Instruments, Inc., Winooski, VT, USA). Images were obtained with GFP filter sets for Calcein AM (Ex/Em 494/517 nm) to identify live cells and Texas Red filter sets for EthD-1 (Ex/Em 528/617 nm) to identify dead cells^32,33. Brightfield, Calcein AM, EthD-1, and merged fluorescence pictures were acquired at 4× magnification for each spheroid. Viability was assessed by quantifying the Calcein AM-positive (green) region as a percentage of the total spheroid cross-sectional area, using ImageJ software by established techniques and the manufacturer’s guidelines³⁴. Spheroids from all three procedures were evaluated at 2 and 7 days; however, SS method spheroids were further studied at 10 and 14 days to determine their long-term viability.

Gene expression analysis via quantitative RT-PCR

Real-time PCR was conducted according to our previous works^{14,27,28,35,36}. Briefly, Trizol (Invitrogen) was used to extract total cellular RNA from monolayer and spheroid cells 7 days after the first culture day with 5000 cells. A first-strand cDNA synthesis kit (Thermo Fisher, Waltham, Massachusetts, USA) was used to reverse transcribe equal amounts of RNA. Then, using SYBR Green RT-PCR Master Mix (RealQ Plus 2× Master Mix Green, Amplicon), qPCR reactions were performed on a 96-well Light Cycler Real-Time PCR system (Roche). The 2^(−ΔΔCT) technique was used to quantify relative gene expression compared to the endogenous control B-actin. The expression of stemness-related genes (CD44, CD24, and ALDH1) and the hypoxia-related gene (HIF-1α) in monolayer and spheroid cultures (2D and 3D cultures) was measured and compared.

Statistical analysis

Statistical analysis was performed using GraphPad Prism software, and all experiments were conducted at least three times to ensure robustness. Unpaired two-tailed Student’s t-tests were employed to compare statistical significance between two groups. A one-way ANOVA with Tukey’s post hoc tests was employed to compare significance among more than two groups. The data are presented as mean ± SD, with statistically significant differences at p < 0.05. This rigorous approach ensured the precise assessment of viability, and CSC cell marker expression in spheroids across both conventional and SS-method cultures.

Results

Assessing hanging drop method culture

The spheroid culture was initiated using the hanging drop method (Fig. 4A–C). After 24 h, cell sheets began to form (Fig. 5A–C), and by 48 h, these sheets were visible (Fig. 5Ca). When nutrients were added to the drops, the movement of the solution caused the sheet structures to become messy, resulting in uneven cell adhesion (Fig. 5Cb). Also, if fresh culture medium was not added, these cell sheets began to break apart and died after 72 h (Fig. 5Cc). The increased the viscosity of the droplet media due to the presence of methylcellulose and the non-invasive nature of the cells prevented their movement, aggregation, and the formation of cell sheets (Fig. 5Cd). As a result, this method failed to generate MCF7 spheroids effectively.

Fig. 4.

Culturing of MCF7 spheroids by the hanging drop method in 10 cm Petri dishes.

Fig. 5.

Hanging drop results. Spheroid culture was carried out in three groups (A–C). (a) The first group comprised spheroids that were fed every 24 h. The formation of the spheroids was uneven and incomplete because of the increased flow of the spheroid solution (b). The spheroids in the second group were not fed, which led to cell death after 72 h (c). Because the droplets were more viscous, the third group, whose growth media contained 0.5% methylcellulose, could likewise not aggregate correctly (d) (n = 6).

Assessing agarose method culture

Spheroids were cultivated in agarose gel in 96-well plates. Spheroid development occurred between 24 and 48 h, contingent upon the size and quantity of cells used. Nonetheless, a substantial problem emerged: the spheroids generated by this technique had an incoherent structure and failed to establish the distinctive three-layered architecture commonly linked to properly formed spheroids. Moreover, cell proliferation and aggregation were negatively impacted by slight imperfections on the gel surface, which, despite being considered trivial, damaged the overall morphology of the spheroid mass (Fig. 6). Significantly, these minor abnormalities were reduced throughout the optimization phases of gel formation. The successful production of uniform and consistent spheres in the new method, to be elaborated upon in the next section, indicated acceptable gel formation.

Fig. 6.

Using the agarose gel method to cultivate spheroids with varying cell counts. Because these cells are less invasive and the gel surface is slightly rough, the development of MCF7 spheroids was irregular (n = 9).

Development of the SS method for effective spheroid formation

Photographs of three cultures under identical times and conditions, utilizing distinct techniques, are shown in (Fig. 7A–C). The SS method yields round and uniform spheroids, serving as an effective model for tumor simulation (Fig. 7C).

Fig. 7.

Comparison of three methods for the improvement of MCF7 spheroids. Agarose method (A), hanging drop method (B), SS Method (C).

Representative optical pictures depict spheroids containing varying cell quantities (Fig. 8A). Figure 8B illustrates the variation in average spheroid diameter over time among the five groups. The spheroid diameter of all groups rose progressively after a little initial decline during the first 2 days, attributable to the following factors. During the preliminary phase, the cells autonomously merged to create a spheroid characterized by loose intercellular connections (Fig. 9A). Subsequently, the cells commenced reorganization upon contact, resulting in increased rigidity of the spheroid (Fig. 9A). Furthermore, the spheroids initially exhibited a sluggish development rate, which may have led to their aggregation during the first 2 days (Fig. 8B).

Fig. 8.

Spheroid formation. The process began with cell numbers of 700, 1500, 2500, 5000, and 12,000 per drop for the conventional spheroid generation, which produced perfectly rounded spheroids 24 h after transfer to agarose gel. Five different time points were used to take pictures. Over 8 days, the spheroids’ development was monitored (A). Graph of spheroid size analysis up to day 8 (B) (Mean ± S.D (n = 3)).

Fig. 9.

(A) The spheroid formation can be divided into three stages. Formation of loose cell aggregates through integrin-ECM binding (a), a period for cadherin expression and aggregation (v), formation of compact spheroids through homophilic cadherin-cadherin interactions (c). (B) Different layers of the spheroid. The round spheroids had the predicted three layers: an exterior layer of proliferating cells, a middle layer of difficult-to-identify dormant, non-proliferating cells, and an innermost layer of necrotic cells that covered the entire spheroid. Once moving into the inner layers, CO₂ and waste levels increase, while nutrient, oxygen, and pH levels decrease. (C) Live/dead fluorescence staining further confirms the characteristic three-layer architecture of spheroids: a green-fluorescent outer rim of proliferating, esterase-active cells; an intermediate zone with reduced green intensity and occasional red puncta indicating quiescent cells; and a central core of intense red fluorescence marking necrotic, membrane-compromised cells.

The findings indicated that fully formed spheroids suitable for various applications were produced regardless of the initial cell count used in the procedure (Fig. 9). The standard spheroid production in this procedure started with cell quantities of 700, 1500, 2500, 5000, and 12,000 per drop, resulting in perfectly rounded spheroids after 24 h post-transfer to agarose gel. The spheroids’ development was observed throughout 8 days. The size of spheroids in terms of days and cell number was examined using a graph (Fig. 8B).

Spheroids cultivated with 5000 cells were chosen because of their appropriate and small hypoxic zone and insufficient size to develop necrotic cores quickly. After about 6 days, these spheroids evolved into structures exhibiting necrotic layers. For the others, the dimensions of the initial cells fluctuated likewise. The necrotic layer in spheroids with 12,000 initial cells formed rapidly. The spheroids included three layers: the outer layer of proliferating cells, the middle layer of dormant and non-proliferating cells that are challenging to identify, and the innermost layer consisting of necrotic cells within the complete spheroid (Fig. 9B). As we progress into the inner layers, there is a reduction in nutrients, oxygen, and pH while levels of CO₂ and waste products rise (Fig. 9B). Live/dead fluorescence staining delineated the canonical three‐layer spheroid architecture: a brightly green‐fluorescent outer layer of proliferating, esterase‐active cells; an intermediate zone of reduced Calcein AM signal with sporadic EthD‐1 staining, representing quiescent cells; and a centrally necrotic core with intense red EthD‐1 fluorescence indicating complete loss of membrane integrity. This stratification was most pronounced in SS‐method spheroids (Fig. 9C).

Viability of spheroids

Live/dead staining revealed significant disparities in spheroid viability across the three culture methods at both early and late time intervals (Fig. 10A,B). After 2 days, SS-method spheroids exhibited high vitality (95 ± 3%), comparable to agarose method spheroids (82 ± 4%) and hanging-drop spheroids (78 ± 5%). On day 7, SS-method spheroids maintained excellent vitality (92 ± 2%). In comparison, agarose-method spheroids had a modest decline to 75 ± 3%. Hanging-drop spheroids displayed essentially no viable cells (0 ± 1%) (Fig. 10B). Statistical study (one-way ANOVA) demonstrated that the survivability of the SS method on day 7 was significantly superior to both the agarose and hanging-drop methods.

Fig. 10.

Functional validation of SS spheroids by live/dead Calcein AM staining. (A) Representative brightfield, Calcein AM (live), EthD-1 (dead), and merged fluorescence images of MCF7 spheroids generated by the SS method, agarose‐coated plates, and hanging‐drop at 2 days and 7 days. (B) Quantification of Calcein AM–positive viability (%) for each culture method at 2 days and 7 days (mean ± SD; n = 5). (C) Time course of SS-method spheroid viability showing brightfield, live, dead, and merged channels at 2, 7, 10, and 14 days. (D) SS-method spheroid viability (%) over 14 days as measured by Calcein AM fluorescence (mean ± SD; n = 5).

The prolonged live/dead assessment of SS-method spheroids over 14 days demonstrated a progressive rise in EthD-1 staining and a decline in Calcein AM signal (Fig. 10C,D). At 10 days, viability decreased to 75 ± 4%, and by 14 days, it further reduced to 50 ± 5%, indicating the progression of central hypoxia and necrosis. The data suggest that the SS-method generates consistently viable spheroids for a duration of up to 7 days, followed by a regulated and foreseeable decrease in viability from days 10–14. The findings reveal that spheroids produced by the SS method exhibit superior survivability over an extended period compared to traditional 3D culture procedures, reflecting an increased functional longevity.

Indirect assessment of metabolic activity via esterase-mediated viability

The fluorescent intensity patterns of Calcein AM provide insights into cell viability and membrane integrity of spheroids across the three culture techniques, as the conversion of Calcein AM to fluorescent calcein is reliant on intracellular esterase activity, which serves as an indicator of cellular viability rather than direct metabolic activity. The qualitative evaluation of green fluorescence intensity revealed varied indirect metabolic activity profiles among the groups³⁷. The SS method spheroids exhibited strong, uniform Calcein AM fluorescence throughout their structure at both 2 and 7 days, indicating significant and consistent esterase activity, which suggests maintained cellular viability and membrane integrity (Fig. 10). Agarose-method spheroids exhibited moderate Calcein AM intensity, with distinct core-periphery gradients, especially apparent on day 7, indicating diminished enzymatic function in the central regions due to nutritional and oxygen constraints. Hanging-drop spheroids exhibited the most impaired cellular viability profile, characterized by a fluctuating and decreasing Calcein AM signal over time, ultimately resulting in a total loss of esterase activity by day 7, which reflects severe loss of membrane integrity and cellular dysfunction²⁶. Indirect patterns of metabolic activity, evidenced by variations in esterase function, corroborate the viability results and demonstrate that the SS method not only preserves cell survival but also maintains crucial cellular metabolic processes necessary for sustaining physiologically relevant three-dimensional culture models^26,38.

Colony formation assay

A colony formation assay was performed to enrich breast CSCs. Enriching CSCs in vitro by exposing them to conditions that enhance their CSC properties is possible. This can be achieved through methods such as prolonged food deprivation, exposure to drugs or radiation, or by increasing their self-renewal capacity through spheroid culture. In the present study, spheroids derived from single cells, as well as multicellular beads, were cultured to enrich for CSCs based on their self-renewal capacity and the structural organization provided by 3D spheroids³⁹. No viable colonies or cells were observed in serum-free or serum-containing media with 0.1% of the 2D culture. In contrast, stem cell colonies were present in both serum-free and serum-containing media, accounting for approximately 0.1% of the 3D culture. Colony formation in serum-free media resulted in a greater number and organization of colonies than serum-containing media (Fig. 11).

Fig. 11.

Colony formation assay demonstrating CSC enrichment in 3D SS-derived spheroids. Single-cell suspensions from 2D monolayer and 3D spheroid (SS method) cultures were plated in serum-free and 0.1% FBS-containing media. No colonies formed from 2D MCF7 cells, whereas 3D spheroid–derived cells generated robust, well-organized colonies under both conditions, with greater colony number and size in serum-free media (n = 3).

Analysis of CSC markers in developed spheroids

The expression of genes related to stemness, including ALDH, CD44, CD24, and genes related to the hypoxic induction factor HIF-1α, was examined in MCF7 spheroids and MCF7 2D cells after 48 h (Fig. 12). The results showed that CD44 genes were expressed 43.84 times more in 3D culture than in 2D culture (Fig. 12A). Conversely, CD24 had a decreased expression in 3D culture compared to 2D culture (Fig. 12B). The ALDH gene was also expressed 3.32 times more in 3D culture (Fig. 12C). This level of gene expression was consistent with the stemness phenotype of breast cancer cells (CD44+/CD24−/ALDH1+). HIF-1α also increased in 3D culture by 11.21 times, indicating the creation of a suitable hypoxic environment in the depth of the spheroids (Fig. 12D).

Fig. 12.

Gene expression analysis of CD44 (A), CD24 (B), ALDH (C), and HIF-1α (D) in 3D and 2D MCF7 cells. The (*) symbol corresponds to the significance levels based on the t-test between the 2D and 3D MCF7 cancer cell lines. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001(mean ± SD; n = 3).

Discussion

The SpheroidSync or SS method delivers a transformative improvement in three-dimensional cell culture by integrating the compact self-assembly of hanging drops with the stability of agarose-coated plates, yielding spheroids with exceptional viability, shape uniformity, and enrichment of CSCs. Critically, this approach allows the generation of spheroids of any desired size, from small (less than 150 µm) high-throughput screening models to large (greater than 300 µm) physiologically relevant constructs, simply by adjusting the initial cell number, thereby tailoring the platform to specific experimental goals^6,12,15,40. Our refined and integrated technique yielded balanced, stable spheroids exhibiting improved viability and consistency. Recent research has confirmed the limitations of conventional approaches. Research indicates that MCF7 spheroids generated by agarose gel often display uneven morphologies and non-aggressive characteristics, hindering their efficacy as dependable models for pharmacological evaluation¹³. We have had this problem in previous research. We cultured MCF7 spheroids alongside agarose cultures of MDA-MB-231 spheroids; however, these cells only produced clusters after 72 h, and we did not obtain any round spheroids¹⁴.

Live/dead fluorescence staining demonstrated that SS‐method spheroids maintain high viability (95 ± 3%) at 2 days and 92 ± 2% at 7 days, significantly outperforming agarose‐only (82 ± 4% → 75 ± 3%) and hanging‐drop cultures (78 ± 5% → 0 ± 1%) (Fig. 10A,B). Extended assessment over 14 days revealed a predictable decline in viability, reaching 75 ± 4% at 10 days and 50 ± 5% at 14 days, consistent with the emergence of central hypoxia and necrosis, as characterized in multicellular tumor spheroids^41,42.

The fluorescence of Calcein AM, indicative of intracellular esterase activity and cellular viability, was consistently bright in SS spheroids, signifying strong enzymatic function. Agarose cultures demonstrated peripheral-biased fluorescence with core fading by day 7, indicating the presence of oxygen and nutrient gradients. The hanging-drop spheroids exhibited heterogeneous fluorescence on day 2, with an almost undetectable signal by day 7, indicating esterase activity cessation and the absence of living cells⁴³. SS wells sustained elevated total fluorescence, agarose wells had a reduction of around 10%, and Hanging drops on the petri dish lid nearly completely lost their signal by day 7. The rapid loss of viability in hanging-drop spheroids by day 7 can be attributed to the confined nature of the droplet environment, where nutrient depletion and waste product accumulation occur more readily due to limited medium volume and restricted diffusion. This confined environment creates unfavorable conditions that cannot be easily remediated through medium changes without disrupting the spheroid structure. In contrast, the SS method’s transfer to agarose-coated wells provides a more stable culture environment with improved nutrient diffusion and waste removal, highlighting the mechanistic advantages of our approach over conventional hanging-drop techniques. The progressive decline in esterase activity in SS spheroids after day 7 corresponds with a significant increase in CSC populations, indicating that the surviving cells assume a quiescent, stem-like character in response to the development of hypoxia and metabolic stress. Quantitative RT-PCR analysis confirmed this association: SS spheroids exhibited an 11.21-fold increase in HIF-1α mRNA, indicative of hypoxia signaling, along with 43.84- and 3.32-fold increases in CD44 and ALDH1 expression, respectively. As esterase activity diminished, the relative abundance of CD44⁺/CD24⁻ and ALDH1⁺ cells increased, leading to the formation of tightly clustered colonies of therapy-resistant cells within the spheroid core^44–46.

The SS method employs a two-phase approach, with initial gravity-driven compaction followed by transfer to agarose, which enhances cell–cell interactions and nutrient diffusion. This alleviates the diffusion barriers and mechanical strains associated with single-method cultures. By maintaining esterase-mediated cell viability pathways and enhancing hypoxia-induced CSC enrichment, SS spheroids more accurately replicate in vivo tumor microenvironments, making them particularly advantageous for mechanistic investigations and pharmacological screening⁴⁷.

In comparison to contemporary 3D platforms such as, microfluidic spheroid arrays⁴⁸, magnetic bioprinting technologies⁴⁹, and polysaccharide hydrogels⁵⁰, the SS method demonstrates enhanced vitality, uniform cell viability, and CSC enrichment without the need for specialized equipment. The capacity to generate spheroids of various quantities and dimensions, preserve significant esterase activity, and maintain physiologically pertinent gene expressions across prolonged cultures sets a new standard for three-dimensional tumor modeling. The simplicity, reproducibility, and functional accuracy of the SS method render it an indispensable platform for translational cancer research, personalized medicine, and pharmaceutical development.

Future direction

The SS method’s versatility extends beyond MCF7 cultures, offering a tunable platform for generating spheroids of any size, from high-throughput small-diameter models to large, physiologically relevant constructs across diverse tumor types and co-culture systems. Future studies should integrate SS spheroids with high-throughput drug and nanoparticle screening, leveraging real-time fluorescence assays to capture dynamic changes in viability, metabolism, and microenvironmental signaling. This multiplexed imaging approach, combined with adjustable mechanical and biochemical culture parameters, will enable precise dissection of biophysical and biochemical influences on spheroid behavior, CSC enrichment, and therapeutic responses. Embedding SS spheroids within microfluidic and perfusion systems will further refine nutrient delivery and waste removal, prolonging functional lifespan and replicating intratumoral fluid dynamics. The addition of fluorescent metabolic probes (e.g., 2-NBDG, TMRM) and immunofluorescent markers (e.g., Ki-67) will enrich biological readouts, creating a comprehensive in vitro model that closely mimics the complexity of in vivo tumors. Collectively, these advancements will elevate the SS platform into a powerful tool for mechanistic tumor biology studies, personalized drug development, and predictive oncology applications.

Study limitations

Our evaluation relied on live/dead and Calcein AM/ EthD-1staining, providing only an indirect assessment of metabolic activity, and did not include proliferation markers (e.g., Ki-67) or quantitative metabolic assays such as AlamarBlue or CellTiter-Glo, limiting insights into cell cycle dynamics and metabolism. Assessments were also restricted to discrete epifluorescence images rather than continuous high-resolution monitoring with confocal plate readers. Finally, the SS method requires significant technical skill for precise edge-cut transfer and spheroid handling.

Conclusion

This paper presents the SS method as an effective and reproducible technique for producing MCF7 breast cancer spheroids characterized by homogeneous shape, adjustable size, and prolonged functional lifespan. Through the integration of hanging-drop self-assembly and careful transfer to agarose-coated plates, SS spheroids demonstrate enhanced circularity, homogeneity in size, and structural integrity relative to traditional hanging-drop or agarose techniques. The SS approach facilitates the creation of spheroids of any specified diameter, ranging from < 150 µm for high-throughput models to > 300 µm for medically relevant constructions, simply by modifying the initial cell quantity.

In addition to morphological integrity, SS spheroids exhibited strong functional performance. Live/dead Calcein AM/ EthD-1 staining demonstrated maintained viability (95 ± 3% at day 2; 92 ± 2% at day 7), considerably surpassing agarose (82 ± 4% → 75 ± 3%) and hanging-drop (78 ± 5% → 0 ± 1%) controls. Calcein AM fluorescence demonstrated sustained intracellular esterase activity and bioenergetic integrity for a minimum of 7 days, unlike the swift cessation of activity observed with conventional methods. Quantitative RT-PCR revealed an 11.21-fold elevation of HIF-1α and increases of 43.84-fold and 3.32-fold in CD44 and ALDH1 expression, respectively, linking Esterase activity adaptation to the enrichment of CSCs in hypoxic environments. These functional validations confirm SS spheroids as physiologically pertinent in vitro tumor models. The SS method provides exceptional control over spheroid structure and viability, establishing a robust basis for advanced cancer biology research, high-throughput drug and nanoparticle screening, and personalized medicine applications.

Author contributions

MAM: conceptualization, investigation, visualization, paper drafting, HH and RJE: supervision, funding acquisition, reviewing, editing.

Funding

RJE is supported by Tabriz University of Medical Sciences, Tabriz, Iran, Grant Number 71052.

Data availability

The data analyzed during the current study is not publicly available because the data is mostly qualitative and data can be available from the corresponding author on reasonable request.

Declarations

Competing interests

The authors declare no competing interests.