Application of Cell Culture in Enhancing Drug Discovery

## **Chapter 6**

## Perspective Chapter: Evaluating New Drugs against K-Ras4B/PDE6δ Using an *In Vitro* Approach

*Dayan A. Carrion-Estrada, Paola Briseño-Diaz, Sandra Delfín-Azuara, Arturo Aguilar-Rojas and Miguel Vargas*

## **Abstract**

Cancer represents the leading cause of global mortality worldwide. Recent estimates have shown that approximately 25% of all cancer types exhibiting *KRAS* mutations, making these mutations one of the most reported so far. Given the important role played by KRas during the progression of different tumors, the search for new therapeutic compounds that can reduce the adverse effects of this oncogene becomes evident. However, discovering effective anticancer compounds is a complex and timeconsuming task. These compounds should ideally exhibit potent anticancer properties at low concentrations, with minimal impact on healthy cells. The validation of potential candidates involves several stages and methods, including *in vitro* techniques such as cell lines or primary cell cultures grown under 2D and 3D conditions. This chapter provides a comprehensive review of methods to support the effectiveness of two compounds, C14 and P8, specifically targeting mutant KRas as potential antitumor agents. Cytotoxicity assays were employed on breast and pancreatic cancer cell lines and primary cell cultures grown in 2D and 3D conditions to evaluate the effectiveness of these compounds. The use of multiple cell culture systems provides more pertinent data, enhancing our understanding and assessment of the potential benefits of new therapeutic molecules.

**Keywords:** human primary cell, 3D cell cultures, K-Ras4B, PDE6δ, therapy

## **1. Introduction**

Cancer is the leading cause of death worldwide. According to GLOBOCAN, every year, over 20 million people are diagnosed with cancer globally, which leads to nearly 10 million deaths reported in 2020 [1]. The increasing incidences of cancer worldwide have created a demand for identifying and developing effective and tolerable therapies for treating different types of cancer [2].

Among the most common causes of cancer death in 2020 were: lung, colorectal, stomach, breast and pancreatic cancer [1]. It is important to highlight that, from these six pathologies, they share in common the presence and dependence of *KRAS* gene mutations [3].

*KRAS* (Kirsten rat sarcoma viral oncogene homolog) was first reported to be a transforming gene related to oncogenesis in humans in the early 1980's [4, 5]. Since then, *KRAS* has emerged as one of the extensively studied proto-oncogenes, owing to its established mutation prevalence of approximately 90% in pancreatic adenocarcinomas, 45% of colorectal adenocarcinomas, 22% of lung adenocarcinomas and 5% of breast cancers [6]. KRAS mutations are found in 30% of all cancers, making it the most frequently mutated oncogene across all cancer types and a promising drug target due to its participation in the progression and maintenance of several types of tumors [7].

*KRAS* belongs to the RAS family of small GTPases and behaves as a molecular switch by cycling between a GTP-bound active state and a GDP-bound inactive state. Wild-type KRAS is a tumor suppressor that is usually activated via specific receptors to regulate different signaling pathways involved in, cell cycle progression, cell growth, proliferation, migration, and apoptosis (**Figure 1**). For K-Ras protein to be active it needs to be transported to the cell membrane, through phosphodiesterase 6 delta protein (PDE6δ) [10, 11]. The interaction between K-Ras and PDE6δ is performed by the recognition of a post-translational modification (farnesyl group) present in the GTPase (**Figure 1**).

#### **Figure 1.**

*RAS effector pathways. Plasma membrane-associated Ras-GTP can directly interact with multiple different effectors to activate different signaling pathways such as cell proliferation, migration, survival/death, differentiation, migration, and adhesion. Previous studies have shown that the administration of C14 and P8 compounds [1] cause a strong binding of the mutant K-Ras4B/PDE6δ complex [2] inhibiting either it dissociation [3], or, and binding to the membrane [4] resulting in a diminution in the activation of its principal effectors Akt and Erk (red arrows). Modified from Refs. [8, 9].*

#### *Perspective Chapter: Evaluating New Drugs against K-Ras4B/PDE6δ Using an* In Vitro *Approach DOI: http://dx.doi.org/10.5772/intechopen.113019*

However, its function has been shown to be frequently lost during tumor progression in many types of cancer. The disruption of KRAS regulation is triggered by specific point mutations occurring at codons 12, 13, 61, and 146 within the *KRAS* gene. These mutations hinder GTP hydrolysis, resulting in continuous activation of the K-Ras protein and abnormal activation of K-Ras signaling pathways. Consequently, this abnormal activation promotes cell proliferation, differentiation, and survival (**Figure 1**) [10].

For decades, several groups made unsuccessful attempts to target K-Ras in cancer treatment due to the protein's unique characteristics, specifically the absence of binding grooves on its surface for small molecules. As a result, K-Ras was considered undruggable [12–14]. However, in 2013, an innovative strategy emerged with the discovery that PDE6δ plays a role in regulating the subcellular location of K-Ras. In this research, inhibiting the interaction between both proteins resulted in K-Ras being retained in the cytoplasm, which impacted its ability to transmit signals. Consequently, the K-Ras-PDE6 complex was identified as a potential pharmacological target [15]. However, these strategies have shown be limited to a single K-Ras mutant, and toxic to non-cancerous cells due to their mechanism affecting both K-Ras WT and other proteins recognized by PDE6 [15, 16].

In 2018, our research group presented a new strategy. We proposed the stabilization of the K-Ras/PDE6 complex by small molecules instead of preventing its interaction (**Figure 1**). Based on this, a series of compounds capable of binding and firmly fixing the interaction between the K-Ras and PDE6 proteins were selected *in silico* (**Figure 2**) [17]. These molecules showed excellent results against *KRAS*-dependent cancers, with elevated affinity towards mutated variants of K-Ras, thereby preserving the signaling activity of K-Ras WT [8, 18–20]. After several assays carried out in a big library of compounds, the molecules denominated as C14 and P8 showed the best ability to specifically bind to and stabilize the mutated K-Ras4B/PDE6δ complex (**Figure 2**) [17].

The antitumoral efficacy of these molecules has been assessed in various *in silico* models, *in vitro* models (cell lines and primary cultures), and *in vivo* models of pancreatic and colon cancer [8, 18]. These have resulted in the decreased activity of the principal K-Ras signaling pathways (e.g., MAPK and PI3K/Akt pathways) (**Figures 1** and **2**), apoptosis-mediated cancer cell death, reduction in clonogenic capacity and decreased tumor volume [8, 17].

Given the high prevalence of cancer, there is a need to find novel drugs to treat this complex disease. These new treatments should have the minimal side effects on the patients, while also being more effective in eliminating malignant cells and showing low rates of resistance to them. For these reasons, it is necessary to evaluate new therapeutic options in many models to achieve their validation.

*In vitro* models play a vital role in drug discovery for cancer, as they afford a high degree of control over numerous variables within a research study, and the experimental design is less complex. Moreover, these models offer a cost-effective approach and serve as alternatives to animal use, thereby addressing ethical considerations [21]. In cancer drug discovery, *in vitro* models include various cell culture techniques, with three primary types demonstrating progressive degrees of complexity: 2D, primary, and 3D cultures. These advancements have significantly contributed to our understanding of cancer cells growth and migration, disease response, and drug interactions within the cell [22]. Currently, drug discovery and testing involve the use of both *in vitro* models and *in vivo*-animal models.

This chapter reviews the most common *in vitro* models used to characterize and validate the efficacy of C14 and P8 as the main drugs against the K-Ras/PDE6δ complex. An overview of cell cultures is provided, including cell lines in 2D conditions, 3D systems, and primary cell cultures.

#### **Figure 2.**

*Anti-cancer compounds identified in-silico search. (A) 2-[(3-chlorophenyl) methyl-methyl-amino]-N-chroman-4-yl-acetamide structure known as C14. (B) 2-[4-(3-chlorophenyl)piperazin-1-yl]-N-[(4R)-chroman-4-yl] acetamide structure known as P8. (C and D) By molecular dynamic assays, both molecules demonstrated strong binding and stabilization of the most common mutant K-Ras4B G12D/PDE6δ complex. The dissociation of this complex is impaired and the GTPase function of K-Ras4B inhibited. Both compounds bind to the complex in different positions, which could have a synergic effect [8].*

## **2.** *In vitro* **models for anti-cancer drug discovery**

The process of drug development and its evaluation can indeed be costly. As a result, *in vitro* cell culture models have been extensively utilized for these purposes due to their numerous benefits, including reduced expenses and time compared to animal studies. These *in vitro* models can be classified into two types: 2D and 3D models, each with its own set of advantages and disadvantages, as described in **Table 1**. Furthermore, the source from which the cells used as models are obtained can be adapted to the study's characteristics. Therefore, the combination of cell lineage or culture conditions provides great versatility to *in vitro* models. For instance, when screening and selecting leads from hundreds of compounds, a 2D assay would be the best option over 3D models, as it allows for the lowest feasible cost due to the reduced requirement of cells and time for these assays. However, for fewer compounds requiring more in-depth characterization, a 3D model would be more suitable [23].

#### **2.1 2D cell culture for anticancer drug testing**

2D cultures are the most employed cell models to test drugs due to two major advantages: the methods to cultivate and preserve the cells under these conditions are uncomplicated and importantly; maintenance in terms of materials that are needed is low-cost and being a simpler model interpretation of results is easier [24].

*Perspective Chapter: Evaluating New Drugs against K-Ras4B/PDE6δ Using an* In Vitro *Approach DOI: http://dx.doi.org/10.5772/intechopen.113019*


#### **Table 1.**

*Advantages and disadvantages of* in vitro *culture methods.*

In this classic model, cells grow as a monolayer in a polystyrene culture flask or petri dish [24]. By definition, a cell line is a collection of cells originated from one cell. These cell lines are *in vitro* systems typically established and propagated in a growth medium in tubes, flasks, or dishes as a monolayer culture. Here, cells can continue to divide indefinitely. Using a cell line allows each cell in an experiment to have a similar response because genetic variation is minimal. This can help researchers to observe differences between test groups more clearly [25].

As previously stated, our research group has evaluated the antineoplastic effects of a group of compounds capable of stabilizing the K-Ras-PDE6 complex. To validate their effects in 2D conditions, we utilized various pancreatic and breast cancer cell lines (**Table 2**). Additionally, antitumoral effects were determinate by viability assays were used.

#### *2.1.1 Pancreatic cancer cell lines*

About 90% of pancreatic cancers are histologically classified as pancreatic ductal adenocarcinoma (PDAC) [26], and although it ranks as the 12th most

Pancreatic cancer-derived cell lines

hTERT-HPNE (ATCC, CRL-4023). Cultured in 75% DMEM without glucose (Biowest) with additional 2 mM L-glutamine (Gibco) and 1.5 g/L sodium bicarbonate plus, 25% Medium M3 Base (Gibco). The media is supplemented with fetal bovine serum 5%, 10 ng/ml human recombinant EGF (Sigma-Aldrich), 5.5 mM D-glucose (1 g/L) (Sigma-Aldrich) and 750 ng/ml puromycin (Sigma-Aldrich) at 37°C in an atmosphere with 5% CO2.

BxPC-3 (ATCC, CRL-1687). Cultured in RPMI medium (Biowest, USA) supplemented with 10% fetal bovine serum and 1% antibiotic 100 U/mL (penicillin/streptomycin) and 1.5 g/L sodium bicarbonate at 37°C in an atmosphere with 5% CO2.

Panc-1 (ATCC, CRL-1469). Cultured in DMEM medium (Biowest, USA) supplemented with 10% fetal bovine serum and 1% antibiotic 100 U/mL (penicillin/streptomycin) and 1.5 g/L sodium bicarbonate at 37°C in an atmosphere with 5% CO2.

MIA PaCa-2 (ATCC, CRL-1420). Cultured in DMEM medium (Biowest, USA) supplemented with 10% fetal bovine serum and 1% antibiotic 100 U/mL (penicillin/streptomycin) and 1.5 g/L sodium bicarbonate at 37°C in an atmosphere with 5% CO2.

Capan-1 (ATCC, HTB-79). Cultured in Iscove's Modified Dulbecco's Medium (ATCC) supplemented with 20% fetal bovine at 37°C in an atmosphere with 5% CO2.

Breast cancer-derived cell lines

MCF 10A (ATCC, CRL-10317): Cultured in Dulbecco's modified Eagle's medium F12 (DMEM F12) (Gibco, New York, USA) supplemented with 10% fetal bovine serum (Gibco) and 1% antibiotic 100 U/mL (penicillin/ streptomycin) (Sigma-Aldrich) and 1.5 g/L sodium bicarbonate (Sigma-Aldrich) at 37°C in an atmosphere with 5% CO2.

MDA-MB-231 (ATCC, HTC-26): Cultured in Lebovitz (L-15) medium (Gibco) supplemented with 10% fetal bovine serum and 1% antibiotic 100 U/mL (penicillin/streptomycin) and 1.5 g/L sodium bicarbonate at 37°C in an atmosphere with 5% CO2.

MDA-MB-231 RR (ATCC, HTC-26): Cultured in DMEM F12 medium (Gibco) supplemented with 10% fetal bovine serum and 1% antibiotic 100 U/mL (penicillin/streptomycin) and 1.5 g/L sodium bicarbonate at 37°C in an atmosphere with 5% CO2.

MCF 7 (ATCC, HTB-22): Cultured in DMEM medium (Biowest, USA) supplemented with 10% fetal bovine serum and 1% antibiotic 100 U/mL (penicillin/streptomycin) and 1.5 g/L sodium bicarbonate at 37°C in an atmosphere with 5% CO2.

MCF 7 RR (ATCC, HTB-22): Cultured in DMEM F12 medium (Biowest, USA) supplemented with 10% fetal bovine serum and 1% antibiotic 100 U/mL (penicillin/streptomycin) and 1.5 g/L sodium bicarbonate at 37°C in an atmosphere with 5% CO2.

#### **Table 2.**

*Cell lines employed for drug screening.*

common cancer, for decades it has firmly held the first position as the cancer type with the lowest 5-year survival rate [27]. Although a general mutational profile has been well determined for PDAC, the standardly gene reported as mutated is *KRAS* (>90%) [28]. Among the muted forms that are present in PDAC, the most frequently are G12D (~51%), G12V (~32%) and G12R (~12%) [29]. These mutations have been associated with a poor prognosis due to their ability to induce alterations in cancer cells, enabling them to evade immune response, reprograming their metabolism, and developing resistance to therapy [30]. Given the importance of *KRAS* mutations in the carcinogenesis and tumor progression of PDAC, mutant pancreatic cell lines are an excellent *in vitro* model for anti K-Ras/ PDE6δ drug testing. In our research group, we evaluated the antitumoral effects of the candidates molecules in the pancreatic cell lines (ATCC, USA) as describe in **Figure 3**.

*Perspective Chapter: Evaluating New Drugs against K-Ras4B/PDE6δ Using an* In Vitro *Approach DOI: http://dx.doi.org/10.5772/intechopen.113019*

#### **Figure 3.**

*Representative images of principal pancreatic cancer cell lines that present most of the KRAS mutations. In brief, BxPc-3 is a pancreatic cell line, which does not express the mutant form of K-Ras4B (Figure 1B). Panc-1 expresses the mutation in exon 2 of the KRAS gene, K-Ras4BG12D (Figure 1C). MIA-PaCa expresses the mutation in exon 2 of the KRAS gene, K-Ras4BG12C (Figure 1D) and CAPAN-1 expresses the mutation in exon 2 of the KRAS gene, K-Ras4BG12V. The non-tumoral pancreatic cell line hTERT HPNE, was employed as control (Figure 1A). Scale bar 200 μM.*

#### *2.1.2 Breast cancer cell lines*

According to Globocan, breast cancer (BC) ranks first in incidence and second in mortality among all oncologic malignancies worldwide [1]. This is a heterogeneous group of diseases that can be grouped into four subtypes into four subtypes (luminal A, luminal B, HER2 and Triple negative Breast Cancer (TNBC)) [31, 32]. Regarding the genomic landscape of BC, various alterations have been identified within each subtype. Among them, mutations in *KRAS* play an important role in carcinogenesis or progression of BC [33, 34] and, this gene is mutated in about 7%-12% of the patients (2% luminal A, 20% luminal B, 17% HER2, 0–8% TNBC) [3]. Specifically, the presence of oncogenic K-Ras is associated with the more aggressive luminal B and HER2 subtypes compared to the luminal A type [35]. Significantly, this gene is amplified in approximately 32% of TNBC cases [36] leading to an upregulation of *KRAS* signaling pathways and an enhanced tumor microenvironment that promotes tumor progression [33]. Breast cancer cell lines are described in **Figure 4**.

For initial drug screening it is suggested to determine the proper concentration to use in the desired assays through dose-esponse curve and IC50 determination. This process is often performed in 2D assay of the cell line of interest. The cells are plated and treated with various concentrations of the test compounds, typically in an

#### **Figure 4.**

*Representative images of breast cancer cell lines. Breast cancer cell lines and derived radio-resistant cells growth in monolayer. Scale bar 200 μM. In brief, MCF-7 is an estrogen receptor (ER)-positive cell line, which does not express the mutant form of K-Ras4B (Figure 3B). MDA-MB-231 is a TNBC that expresses the mutation in exon 2 of the KRAS gene, K-Ras4BG13D (Figure 5C) [37]. To explore the effect of C14 and P8 over radioresistant cell lines, MCF 7RR which does not contain the mutation in K-Ras4B (Figure 3D) and MDA-MB-231RR, that possess the mutation K-Ras4BG13D as was observed in its parental cell line were used (Figure 5D) and MDA-MB-231RR, that possess the mutation K-Ras4BG13D as was observed in its parental cell line (Figure 5E) [38]. The non-tumoral breast cell line MCF 10A, was employed as control (Figure 5A).*

eight-point dose range, along with vehicle and media only controls The methods are described in detail in Ref. [39].

#### **2.2 Methods for viability/cytotoxicity methods determination**

Cell viability or cytotoxicity in monolayer cultures can be determined by different methods. Cell viability assays are frequently employed, with a variety of markers as indicators of metabolically active (living) cells, such as measuring ATP levels (e.g., CellTiter-Glo® Luminescent assay), measuring the ability to reduce a substrate (e.g., tetrazolium reduction and resazurin reduction), and detecting enzymatic/protease (e.g., CellTiter-Fluor assay) activities present only in living cells.

The most widely used technique is Tetrazolium Reduction Cell Viability Assays, which measures cellular metabolism as an indicator of cell viability, proliferation, and cytotoxicity. In this method, the positively charged dye, 3-(4,5-dimethylthiazol-2-yl)- 2,5-diphenyltetrazolium bromide (MTT), penetrates viable cells and is metabolized into a purple-colored product known as formazan. As a result, color formation can be a helpful indicator of viable cells. However, the incubation time for this method is long (more than 4 hours). To address this limitation, other negatively charged compounds such as (2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide) (XTT), which cannot penetrate cells, must be combined with intermediate electron coupling reagents that help them enter into the viable cells. Once inside the cells, XTT is reduced to tetrazolium, a soluble formazan product. The incubation time for this method is 1–4 hours, making it more convenient [40]. For our experiments, we used MTT and XTT assays to build a dose-response curves and determine the effect of different concentrations of compounds C14 and P8 in pancreatic and breast cancer cell lines viability.

Anti-cancer drugs C14 and P8 should demonstrate inhibition of cancer cells growth or proliferation, and these is most often determined by IC50. In this work, IC50 is defined as the particular inhibitory drug concentration required to reduce the percentage of viable cells by 50% compared to cells grown without drug exposure [41].

The IC50 value (molar concentration) is important for *in vitro* models because it indicates the amount of a drug is required to inhibit a biological process by half, indicating changes in the population due to increased cell death or decreased cell proliferation. In cancer, using the IC50 concentration of a compounds means killing cancer cells and preventing cancer cell growth while having a less damage on healthy cells in the body or inhibiting tumor growth by half. A low IC50 suggests that the drugs will be effective at lower concentrations and will consequently cause less systemic toxicity when administered to patients for therapy [42].

## **2.3 Evaluation of compound targeting K-Ras4B/PDE6δ in pancreatic cancer cell lines**

A dose-response curve to evaluate the antitumoral effects of compounds of interest was performed using MTT assays. Pancreatic cancer cells were plated in 96-well plate and treated with various concentrations of C14 and P8 compounds, along with vehicle (DMSO) and media-only as controls. The IC50 concentration was determined using Prism 8 software (GraphPad, USA).

As observed below, C14 possess a high cytotoxic property in eliminating cancer cells (**Figure 5B**–**E**, blue line, and **Figure 5F**). On the other hand, P8, an analogue of C14, demonstrates a more potent cytotoxic effect on cancer cells (**Figure 5B**–**E**, red line, and **Figure 5F**) compared to the vehicle (**Figure 5B**–**E** black line).

*Perspective Chapter: Evaluating New Drugs against K-Ras4B/PDE6δ Using an* In Vitro *Approach DOI: http://dx.doi.org/10.5772/intechopen.113019*

**Figure 5.**

*Dose-response curves of pancreatic cell lines treated with compounds C14 and P8. Dose-response curves using 0–200 μM concentrations of compound C14 and P8 tested at 48 hours post incubation measured with MTT. Each curve was done by three independent replicates.*

Another crucial point is the specific effect of C14 and P8 over malignant cells. As the results showed above, *KRAS* mutant cell lines Panc-1-G12D, MIA-PaCa-G12C and Capan-1-G12V exhibit a considerable cytotoxic effect for both compounds compared to the vehicle (DMSO) (**Figure 5C,D** and **E**). This behavior was also observed in BxPC-3WT cell line, although BXPC-3 cells are WT for *KRAS*, the present several other mutations, which contribute to the activation of K-Ras and the dependence to other effectors downstream K-Ras signaling pathway, potentially explaining their susceptibility to the treatment [43].

Remarkably no inhibitory effect was observed in the viability of the control cell line hTERT-HPNE, even at concentrations higher than 200 μM after 48 hours of exposure (**Figure 5A**, red and blue line and **Figure 5F**).

These IC50 values of C14 and P8 in cancer cell lines are lower than 90 μM, which is 2 and 6 times less than the concentrations required to affect the viability of control cells (**Figure 5A** and **F**). This suggests that these compounds could be an excellent treatment alternative for pancreatic cancer, as they exhibit cytotoxicity to cancer cells while sparing normal pancreatic cells from side effects.

As a result, this assay allows to observe the specific cytotoxic effect of compounds on the viability mainly in *KRAS* mutant cancer cells and highly *KRAS*-dependent cells without affecting healthy cells, making them a promising treatment alternative.

## **2.4 Evaluation of compound targeting K-Ras4B/PDE6δ in breast cancer cell lines**

As mentioned earlier, dose-response curves were generated to determine the cytotoxic effect of compounds C14 and P8 on the breast cancer cell lines using XTT assays. The breast cancer cells were plated in 96-well plate and treated with various concentrations of the compounds, with DMSO used as a vehicle and media only as control. The IC50 concentration was determined using Prism 8 software (GraphPad, USA). These data are presents in **Figure 6**.

**Figure 6.**

*Dose-response curves of the breast cell lines treated with compounds C14 and P8. Dose-response curves using 0–200 μM concentrations of compound C14 and P8 tested at 72 hours post incubation measured with XTT each curve was done by three independent replicates.*

Similar to observed in pancreatic cancer cells, no noticeable effect on the viability of the control cell line MCF 10A was observed after its exposure to C14 and P8 (**Figure 6A**, lines red and blue and **Figure 6F**). However, both compounds exhibited a potent cytotoxic effect over the rest of the tested breast cancer cell lines tested (**Figure 6B**–**E**, lines red and blue and **Figure 6F**). According with the IC50 values, even at concentrations higher than 100 μM after 72 hours of exposure, both compounds did not have impact over the viability of non-tumoral cells (**Figure 6A** and **F**). This finding supports the notion that C14 and P8 have a specific effect over *KRAS*mutant cells such as, MDA-MB-231 (**Figure 6C**). This discovery was also observed in the radioresistant cell line MDA-MB-231RR, where the compounds also had a significant inhibitory effect at 72 hours (**Figure 6D**). This finding suggests an alternative course of treatment for patients who have experienced a cancer recurrence after radiation therapy. Finally, in the case of MCF 7 and MCF 7RR, the IC50 values in them were higher than those found in *KRAS* mutant lines (**Figure 6B** and **E**). It is important to note that the compounds have an inhibitory effect on the viability of luminal and radioresistant breast cancer cells, even though that these cell lines do not carry the *KRAS* gene mutation (**Figure 6A, E** and **F**).

Taken together, these findings indicate that the conventional 2D cell-based cancer drug screening on pancreatic and breast cancer cell yields reproducible results of the specific cytotoxic effect over cancer cells without affecting non-tumor control cells. This makes them suitable for further experiment and for evaluating their effects in a more complex cell culture.

Primary cultures generated directly from tumors are a viable alternative to employing cell lines. In comparison to cell lines, in which all cells are genetically identical, primary culture cells gene expression can vary from cell to cell. Primary cultures offer several advantages. Not only are cells isolated directly from the tumor site but

*Perspective Chapter: Evaluating New Drugs against K-Ras4B/PDE6δ Using an* In Vitro *Approach DOI: http://dx.doi.org/10.5772/intechopen.113019*

provide the complete pathology which allows to compare the properties of the culture to those of the original. In general, such cultures can be generated as explants, in which mixed cell populations develop out of small fragments of tissue, or as enriched populations of particular cell types, such as cancer cell or cancer stem cells, being ideal to evaluate the efficacy of a new anticancer drug [44].

## **3. Primary cell culture for anticancer drug testing**

Primary cultures consist of cell cultures derived directly from isolated tissue cells (**Figure 7**). These cultures can proliferate under 2D and 3D conditions without undergoing any genetic modifications. However, the number of passages for their use is highly limited [45]. Primary cultures derived from malignant tumors are particularly of great interest in research. Various protocols are currently available for obtaining primary cell cultures, and the selection of a specific method depends on factors such as tissue type and sample size. In **Table 3**, summarizes the different methods for obtaining primary cell cultures in 2D from pancreatic cancer tumor samples [46–48]. Once the primary cultures are obtained, they must be characterized to determine the presence of the required cell lineage. Currently, various methods facilitate their characterization including, the evaluation of specific markers by immunofluorescence microscopy or Fluorescent Activated Cell Sorter (FACS) (e.g., **Figures 8** and **9**, respectively). The most commonly used antibodies for malignancy markers are: E-Cadherin, Ki-67, Vimentin, B-Catenin, MUC-1, MUC-4, MUC-16, EGFR, CEA, lineage markers: CK19 and CK7, and for cancer stem cells (CSCs) marker: CD24, CD44 and CD133 [8, 49–52].

## **3.1 Isolation and characterization of primary cultures derived from pancreatic cancer**

Primary PDAC cultures were obtain in the laboratory using an enzymatic method involving collagenase and trypsin (1:5 trypsin/EDTA solution, 15 min until the tissue was loose) under sterile conditions (**Table 3** and **Figure 7**). The requirements to maintain the primary culture vary depending on the aggressiveness of the cancer and the stage of the cancer being studied. The percentage of serum and growth factors is gradually decreased until the cells can survive in standard conditions (10% serum and 1% antibiotic) (**Table 4**).

#### **Figure 7.**

*Representative image of the primary cell cultures after three passages denominated as MGKRAS004 (A) and MGKRAS005 (B), obtained from malignant pancreatic tumors. Scale bar (red line) 100 μM.*


**Table 3.**

*Different methods to obtain primary cultures from pancreatic cancer.*

The characterization of those cultures previously was previously performed using immunofluorescence microscopy employing the specific markers, CD44+ (Ambion, USA) (**Figure 8A**), CD24+ (Ambion) (**Figure 8B**) and CD133+ (Ambion)(**Figure 8C**). These assays strongly suggested the presence of cancer stem cells (CSCs), in both cultures. To verify this information, flow cytometry assays were conducted which revealed the presence of CSC in 2.4% of total population of MGKRAS004 cells and in 2.18% of MGKRAS005 population (**Figure 9A**).

Based on these results, there is a high abundance of CSCs in both primary cultures. This finding is significant due CSCs have the ability to contribute to the development of a more aggressive tumor characterized by high rates of proliferation, invasion, and resistance to conventional therapies [53]. Keeping this in mind, C14 and P8 compounds were evaluated on these primary cell cultures.

## **3.2 Evaluation of compounds targeting K-Ras4B/PDE6δ on primary cultures from pancreatic tumors**

After demonstrating the heterogenicity and complexity of pancreatic primary cultures, the efficacy of C14 and P8 compounds was evaluated on 2D conditions using primary cultures obtained from samples of pancreatic malignant tumors, with the same conditions used for cell lines and DMSO as control (**Figure 10A** and **B**). According to our evaluation, both molecules were more effective against the MGKRAS004 cell line. In this case, C14 exhibited an IC50 of 38.72 μM, while P8 showed an IC50 of 80.88 μM with (**Figure 10A** and **C**). These IC50 values are consistent with those obtained for the common pancreatic cell lines (**Figure 5F**). However, in the case of MGKRAS005, the effect of C14 was reduced, and an IC50 of 268.2 μM

*Perspective Chapter: Evaluating New Drugs against K-Ras4B/PDE6δ Using an* In Vitro *Approach DOI: http://dx.doi.org/10.5772/intechopen.113019*

#### **Figure 8.**

*Expression of CSC markers in primary pancreatic cultures. (A) CD44 expression in MGKRAS004 and MGKRAS005 cells. (B) CD24 expression in MGKRAS004 and MGKRAS005 cells. (C) CD133 expression in MGKRAS004 and MGKRAS005 cells.*

#### **Figure 9.**

*Cancer stem cell proportion in pancreatic primary culture. (A) Quantification by FACS of CD44+, CD24+ and CD133+ in 2D conditions of pancreatic cancer primary cultures. (B) Quantification by FACS of CD44+, CD24+ and CD133+ in 3D conditions of pancreatic cancer primary cultures. (C) Comparison of % of CSC in 2D and 3D conditions.*

was observed (**Figure 10B** and **C**). A similar trend was observed with P8, where the IC50 observed was 148.9 μM (**Figure 10B** and **C**). The explanation for this data could be related to the amount of CSC present in each population. As mentioned earlier, both primary cultures contain a similar proportion of CSC. However, in the case of MGKRAS005, the population of cells expressing CD133 is greater (**Figure 8C**). This higher expression of CD133 in MGKRAS005 cells may contribute to their reduced sensitivity to C14 and P8, leading to higher IC50 values observed in the treatment response. These data are in agreement with previously reports that associated this molecule with chemoresistance in various cancer cells [54].


#### **Table 4.**

*Nutritional requirements for primary pancreatic cultures since its insolation and placed culture passage 0 (P0) until passages 4 to 10 (P4–10).*

*Perspective Chapter: Evaluating New Drugs against K-Ras4B/PDE6δ Using an* In Vitro *Approach DOI: http://dx.doi.org/10.5772/intechopen.113019*

**Figure 10.**

*Effect on the viability of primary cultures in 2D. (A) MGKRAS004 in 2D treated with C14 and P8. (B) MGKRAS005 in 2D treated with C14 and P8.*

It is important to highlight that many drug candidates will fail during clinical trials, leading to the loss of money and time invested in research. *In vitro* models, plays an important role in drug discovery, especially in the early and preclinical stages. Considering this, despite the increase in the IC50 value in pancreatic primary cultures these compounds shown a significant cytotoxic effect in cancer cells, making them candidates for more in-depth characterization.

Although the most used cell culture growth conditions in cancer research are 2D models, they do not accurately reflect the native environment and general physiology of malignant cells and tumors such as, mechanical and biochemical signaling or intercellular communications [55], cell invasion [56] and expression of pathological markers [56]. To address these limitations, three-dimensional (3D) *in vitro* models have been developed. In this models, it has been possible to reproduce a more detailed aspects of tumoral behavior including interactions among tumor cells, the extracellular matrix, and stromal cells [57]. Additionally, cells growing in 3D, conditions have been exhibited a greater resistance to chemotherapeutics agents and mayor invasiveness compared to 2D models [58, 59]. As a result, the use of 3D models has been gaining more popularity every day. Considering this, we present the advances obtained in 3D models to evaluate the effect of C14 and P8 in one more realistic pancreatic cancer cells systems.

## **4. 3D models for anticancer drug testing**

Currently, various types of 3D models are available for cancer research. The most commonly used ones include spheroid models, organ-on-a-chip models, hydrogel models, and bio-printed models [57]. Each of these approaches has its own specific advantages and disadvantages. However, despite the significant time and cost involved in their production, spheroids are the most widely utilized due to their unique characteristics [60].

Spheroids are small-scale 3D models that could self-assemble into spherical cell aggregates (with radii ranging from 100 to 600 μm) [60–62]. These models exhibit various tumoral characteristics, such as, a central necrotic core, surrounded by quiescent cells and an outer layer consisting of actively proliferating cells [60]. Other important features observed are, pH, oxygen, and metabolic compound gradients [63], and in some cases, micrometastases [64].

In this section, the focus is on spheroids, generated from cell lines or cells obtained from primary cultures of tumoral tissues. The following will describe the procedure

for creating spheroids using cell lines (refer to **Table 2**) as well as spheroids from primary cultures, along with the method for characterizing their cellular composition. Subsequently, the cytotoxic effects of compounds C14 and P8 on the 3D models will be discussed.

#### **4.1 Spheroids derived from cell lines and primary cell cultures**

The following method outlines the protocol utilized by our research group to generate spheroids from cell lines or primary cultures (**Figure 11**). Firstly, the process begins with expanding of cultures under 2D conditions. Cell harvesting is conducted using a standard procedure. Briefly, when the cells reach 70-80% confluency in a plate, the medium is removed, and the cells are collected using trypsinization (0.05% trypsin, 0.53 mM EDTA, Gibco, USA). Subsequently, the cells are centrifuged and resuspended in 1–5 ml of spheroid medium, consisting of DMEM/F12 (Gibco), 2 mM L-glutamine, 100 U/ml penicillin, and 100 U/ml streptomycin (Sigma-Aldrich). The spheroid medium is further supplemented with 20 ng/ml recombinant human epidermal growth factor (Sigma-Aldrich), 10 ng/ml recombinant human basic fibroblast growth factor (R&D Systems, USA), and 1X B27 supplement (Sigma-Aldrich).

From this cellular suspension, approximately 3000–5000 cells are seeded per well in 96-well ultra-low adherence plates (Gibco). The plates are then maintained at 37°C and 5% CO2 for a period of 4 to 7 days, depending on the cell line and desired spheroid size. During the initial 5-day growth phase, it is crucial to minimize disturbances to the plates, particularly during the initial 5-day growth phase [65]. At the end of the culture period, spheroids are collected through gentle centrifugation at 800 rpm and utilized in functional assays.

Alternatively, enzymatic dissociation of the 3D models can be achieved by incubating the culture for 10 minutes with 0.05% trypsin (Invitrogen), followed by mechanical pipetting using a flame-polished Pasteur pipette. The dissociated cells are then passed through a 40 μm pore size filter (Corning). Subsequently, the cells can

#### **Figure 11.**

*Representative images of spheroids from cell lines (MDA-MB-231, MDA-MB-231 RR, MCF7 and MCF 7RR) and primary cell cultures (MGKRAS004 and MGKRAS005) after 7 days of culture. Scale bar 200 μM.*

*Perspective Chapter: Evaluating New Drugs against K-Ras4B/PDE6δ Using an* In Vitro *Approach DOI: http://dx.doi.org/10.5772/intechopen.113019*

be analyzed for specific molecule expression using techniques such as FACS [66] or conventional immunofluorescence microscopy.

**Figure 11** shows all the cell lines and cells from primary cell culture, that were able to develop spheroids. In the case of MDA-MB-231, this cell line is able to generate spheroids in ultra-low adherence plates, although they present a less compact appearance and less well-defined edges (**Figure 11**) [61]. Something similar is observed in its derivative radioresistant cell line (**Figure 11**). On the other hand, MCF 7 and MCF 7RR cells, quickly formed created three-dimensional spheroids with a homogenous inner structure when placed on plates with extremely low attachment properties as previously shown [67, 68]. Finally, although the structure of 3D models derived from primary cell cultures MGKRAS004 and MGKRAS005 is looser and more challenging to obtain compared to cell lines, these models provide a closer representation of the *in vivo* microenvironment of the patient's tumor [69].

#### **4.2 Calculation of spheres formation efficiency (%)**

To verify the effectiveness of the aforementioned method, the percentage of spheroids can be determined. This involves counting the number of spheroids (larger than 40 μm) after the culture period using a microscope at 40X magnification. Digital images of 5 random fields are captured using a digital camera connected to an optical microscope, and the size of the spheroids is determined using acquisition software. The spheroid formation efficiency (MFE%) is calculated using Eq. (1).

$$MFE(\%) = \frac{\text{(number of ammonia molecules per well)}}{\text{(number of cells needed per well)}} \times 100\tag{1}$$

#### **4.3 Fluorescent activated cell sorter (FACS)**

As mentioned earlier, the characterization of spheroids involves various methods, including FACS. According to previous reports [70, 71], the generation of spheroids leads to the enrichment of a subpopulation known as Cancer Stem Cell (CSC). These CSCs population, are cell that are distinguished by their high tumorigenic potential, which includes self-renewal, pluripotency, and proliferative abilities. Additionally, they exhibit resistance to conventional treatments such as chemotherapy and radiation [72, 73]. Given these characteristics, CSCs are widely used as therapeutic targets for new drugs.

The following section outlines, the method employed by our research group to evaluate the expression of specific CSC markers using FACS in spheroids. This approach is employed to determine the antitumoral effect of the compounds of our interest. The cells derived from the spheroids are dissociated, as described above, and then stained with the following antibodies: anti-CD24-phycoerythrin (PE) (Abcam, USA), anti-CD44-allophycocyanin (APC) (Abcam), and anti-ALDH1-fluorescein (FITC) (Abcam). The cellular staining is conducted according to a procedure previously reported [66]. In brief, the dissociated cells are passed through a 40 μm cell filter (Corning) and counted to adjust the concentration to 100,000 cells/ml. Each antibody is diluted at (1:100), added and incubated at 37°C for 1 hour. After incubation, the cells are washed three times with PBS. Subsequently, flow cytometry analysis is performed.

### **4.4 Enrichment of CSC in spheroids derived from primary cell cultures**

In the following section, representative data of cells derived from primary cultures of pancreatic tumors are presented. FACS assays demonstrated the presence of CD44+, CD24+, and CD133+ markers in spheroids derived from primary cultures (**Figure 9**) [74]. These findings support the existence of CSCs in spheroids obtained from MGKRAS004 and MGKRAS005 samples [75]. As mentioned earlier, the generation of spheroids selectively enriches the CSC population. The enrichment of CD44+, CD24+, and CD133+ marker expressions in spheroids, compared to 2D cultures, further confirms the selective effect of 3D systems on this cell population (**Figure 9C**). In the case of MGKRAS004, 2.4% of CSCs were found in 2D cultures, whereas this percentage increased to 6.3% in 3D conditions (**Figure 8A** and **B**). Similar results were observed for MGKRAS005 cells, with 2.18% of CSCs detected in 2D cultures and a significant increase to 24% in 3D cultures (**Figure 9A** and **B**). These findings also support the utilization of these 3D models as a more realistic system to evaluate the effects of new therapeutic compounds. It is worth noting that the levels of CD44+ and CD24+ in patients have recently been associated as predictors of mortality [76].

#### **4.5 3D cell viability assay and IC50 determination**

Using the 3D models described above, the cell viability effects of compounds C14 and P8 were determined. For this purpose, the CellTiter-Glo® 3D Cell Viability Assay Reagent from Promega (USA) was employed. This method measures ATP as an indicator of viability and provides a luminescent readout. Briefly, spheroids were generated as mentioned previously, and then exposed to both compounds for 48 hours. Cell lysis and luminescent signal recording were performed according to the manufacturer's instructions, and IC50 values were calculated using Prism 8 software (GraphPad, USA).

In the 3D models, although C14 and P8 reduced the cell viability of spheroids, their IC50 values were higher compared to the 2D conditions. For MGKRAS004, the IC50 for C14 in 3D was 395.7 μM, whereas it was 38.72 μM in 2D (**Figure 12A**,**C** and **Figure 10A**,**C**). P8 had an IC50 104.4 μM in 3D and 80.88 μM in 2D (**Figure 9A**). In the case of MGKRAS005, C14 exhibited an IC50 of 470.4 μM in 3D and 268.2 μM in 2D (**Figure 12B**,**C** and **Figure 10B**,**C**). On the other hand, P8 showed an IC50 of 96.71 μM in 3D and 148.9 μM in 2D (**Figure 12B**, **C** and **Figure 10B**,**C**). This increase in IC50 values in 3D models is a phenomenon that has been previously demonstrated [59] and is attributed to the greater complexity observed in these systems. In conclusion, despite the increased IC50 values observed 3D culture conditions, both

#### **Figure 12.**

*Effect on the viability of primary cultures in 3D, treated with C14 and P8. (A) MGKRAS004 in 3D treated with C14 and P8. (B) MGKRAS005 in 3D treated with C14 and P8.*

compounds remain active. In this regard, it is evident that compound P8 exhibited better activity, which aligns with its chemical attributes as a compound selected as a C14 analog with enhanced activity. Thus, P8 demonstrates antitumor properties on CSC-enriched populations.

## **5. Drug screening: Challenges and troubleshooting**

The search for new antitumor compounds is a long and challenging process. Demonstrating the effectiveness of candidate compounds must go through different stages. Firstly, they should be highly effective in exerting their effects at low concentrations, while also desirably minimizing their adverse effects on non-tumor targets.

Ensuring the effectiveness of novel antineoplastic candidates requires extensive evaluations in both *in vitro* and *in vivo* models prior to patient administration [24]. This chapter presents the assessment of compounds C14 and P8 in diverse models. The data presented supports the efficacy of both molecules in mitigating the adverse effects induced by various mutant forms of K-Ras in both pancreatic and breast models. However, we encountered several challenges in stablishing the effectiveness of these molecules. One significant challenge was selecting the appropriate method to determine the IC (inhibitory concentration) of each compound. Conventional methods using XTT and/or MTT as cell proliferation quantification techniques have limitations since their response relies on the metabolic rate of the specific cell lines used, which may not accurately reflect the true growth rate of the models being studied. To address this issue, we recognize the importance of implementing models that measure cell growth by quantifying elements independent of cellular metabolism, such as DNA/RNA or ATP [77].

Another crucial aspect that we deem essential during the implementation of the various methods presented in this chapter is their strict standardization to ensure the precision obtained results. This becomes particularly important when working with complex models like 3D cultures. Based on our own experience, incorporating procedures that enhance the homogeneity of mammospheres, such as gently centrifuging cells during seeding, has proven to be highly advantageous.

Finally, we emphasize the significant importance of developing methodologies to assess the physiology of 3D models. Although several effective systems exist for evaluating different aspects in 2D conditions, corresponding tools for 3D cultures are often scarce or nonexistent. Therefore, research groups must take the initiative to develop these methods themselves.

## **6. Conclusion**

*In vitro* models have been of great importance in cancer research and drug screening. The use of these models allows the study of numerous cell process, the effect of drugs on cells, and the processes that trigger apoptosis.

2D models are the standard for drug evaluation keeping the cost lower and requiring fewer cells and less time compared to animal models or human clinical trials. However, this model does not closely mimic the behavior and drug metabolism seen in *in vivo*, resulting in many drug candidates failing during clinical trials.

For more reliable inhibitory effects and better characterization of drugs, primary and 3D models are preferable for analyzing pathologies with the highest mortality,

such as cancer. Primary cells provide broader spectrum of cell types from a greater number of patients to be studied without inducing artificial genetic mutations, and maintaining the same phenotype throughout the culture, while keeping the advantages of a 2D cell line-base assay in terms of time and cost. Nevertheless, management of primary cell culture poses challenges. Obtaining the patient consent, quality of samples, practicing sterility, and maintaining the culture are some of the challenges faced when working with primary cultures. To overcome these difficulties, proper collection with the help of pathologist and selection of appropriate isolation methods and culture media based on tissue type can help to increase the cell viability.

On the other hand, 3D cell culture models offer a number of advantages as they better represent the microenvironment of *in vivo* conditions, with protein and gene expression similar to those found *in vivo.* Due to these advantages, 3D primary cell culture has gained popularity and holds the potential to replace animal *in vivo* models in future, eventually leading to direct human clinical trials.

In conclusion, this chapter reviews several *in vitro* models for assessing of anticancer compounds, demonstrating their selective effectiveness in reducing viability of pancreatic and breast cancer cells in different conditions. These findings are of vital importance, as current chemotherapies often result in severe side effects. The pursuit of alternative therapies with fewer adverse effects could greatly benefit patients with these pathologies. Considering the evidence presented, we propose that compounds C14 and P8 represent novel therapeutic alternatives for the treatment of tumors dependent on oncogenic forms of K-Ras.

## **Acknowledgements**

Dayan Andrea Carrion Estrada and Sandra Delfin Azuara received a predoctoral scholarship from CONACyT (935796 and 1078870). Dr. Paola Briseño Díaz, researcher, thanks to the Faculty of Medicine of UNAM and CONAHCYT for their support in carrying out this work. We thank Dr. Maria del Rocio Thompson Bonilla from the ISSSTE Mexico Hospital 1° de Octubre, since without her support the primary cultures would not have been performed. We thank Dr. Elena Arechega from the UAM Cuajimalpa Mexico, for the donation of radioresistant breast cancer cell lines.

## **Conflict of interest**

The authors declare no conflict of interest.

*Perspective Chapter: Evaluating New Drugs against K-Ras4B/PDE6δ Using an* In Vitro *Approach DOI: http://dx.doi.org/10.5772/intechopen.113019*

## **Author details**

Dayan A. Carrion-Estrada1 , Paola Briseño-Diaz<sup>2</sup> , Sandra Delfín-Azuara1 , Arturo Aguilar-Rojas3 and Miguel Vargas1 \*

1 Department of Molecular Biomedicine, Center for Research and Advanced Studies of the National Polytechnic Institute (CINVESTAV-IPN), Mexico City, Mexico

2 Department of Biochemistry of the Faculty of Medicine of the National Autonomous University of Mexico (UNAM), Mexico City, Mexico

3 Medical Research Unit in Reproductive Medicine, Mexican Institute of Social Security (IMMS), Mexico City, Mexico

\*Address all correspondence to: mavargas@cinvestav.mx

© 2023 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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[67] Chen G, Liu W, Yan B. Breast cancer MCF-7 cell spheroid culture for drug

discovery and development. Journal of Cancer Therapy. 2022;**13**(3):117-130

[68] Pulze L, Congiu T, Brevini TA, Grimaldi A, Tettamanti G, D'Antona P, et al. MCF7 spheroid development: New insight about spatio/temporal arrangements of TNTs, amyloid fibrils, cell connections, and cellular bridges. International Journal of Molecular Sciences. 2020;**21**(15):5400

[69] El Harane S, Zidi B, El Harane N, Krause K-H, Matthes T, Preynat-Seauve O. Cancer spheroids and organoids as novel tools for research and therapy: State of the art and challenges to guide precision medicine. Cell. 2023;**12**(7):1001

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[74] Sumbly V, Landry I. Understanding pancreatic cancer stem cells and their role in carcinogenesis: A narrative review. Stem Cell Investigation. 2022;**9**:1

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## **Chapter 7**

## Using Tumor-Like Spheroids to Study the Effect of Anti-Cancer Drugs *In Vitro*

*Yousuf Alam, Pawel Borowicz, Stefan W. Vetter and Estelle Leclerc*

## **Abstract**

Cell culture techniques have evolved in the last decades and allow now testing anti-cancer drugs using tumor-like spheroids. We describe here issues and troubleshooting solutions when generating spheroids from three human melanoma cell lines (A375, WM115 and WM266). A375 cells generated irregular shape spheroids that were difficult to study due to their fragility. Spheroids generated from all cell lines initially reduced their diameter and increased compacity before increasing in size overtime. Cells present at the periphery of the spheroids showed higher metabolic activity than cells present in the core of the spheroids. When grown as spheroids, a smaller fraction of the A375 and WM115 cells was sensitive to the chemotherapeutic agent temozolomide as compared to cells grown on flat surface. However, this difference was not observed with WM266 cells. Although the presence of spheroids resulted in a smaller fraction of WM155 cells sensitive to the anti-cancer agent vemurafenib, the opposite was observed with A375 cells. Among the cells, WM266 cells were the most resistant to vemurafenib. In conclusion, our study suggests that cell lines behave differently in terms of spheroid formation, and that the effect of the 3D cellular architecture on drug effect is cell type and drug dependent.

**Keywords:** spheroids, cancer, melanoma, vemurafenib, temozolomide

## **1. Introduction**

For many decades, cell culture experiments have been performed with cells attached to the flat surface of plastic wells. These experiments have provided valuable information in many aspects of cell biology, from the identification of new signaling pathways to the screening of new therapeutic anti-cancer agents [1–3]. However, the outcomes of many experiments performed in these *in vitro* conditions could not be reproduced in pre-clinical studies or in clinical trials [4]. One of the reasons for this failure is that two-dimensional (2D) cell culture growth conditions do not mimic the complex three-dimensional (3D) architecture of tumors [5, 6]. Indeed, when cells are grown on flat surfaces, they form only limited contacts with other cells, and the cellular architecture is different from the complex 3D structures observed in tumors

(**Figure 1**). For instance, in 2D growth conditions all cells receive similar levels of oxygen from the environment. However, when grow as spheroids, diffusion of oxygen and nutrients vary according to the position of the cells in the spheroid: cells at the center of the spheroid do not receive the same levels of nutrient and oxygen and are often subject to hypoxia [7, 8].

To overcome the limitations of 2D cell culture conditions, researchers have developed 3D *in vitro* cell culture systems [9–11]. Experimental conditions have been optimized to allow the formation of monoculture or co-culture of 3D assemblies (spheroids) of cells that mimic tumors. Studies have shown that spheroids are valuable 3D cellular models that possess features shared with tumors [12–15]. These features include the presence of gradients of nutrients, waste and gazes (oxygen and CO2) throughout the spheroids, the presence of layers of cells with different metabolic activities (proliferative, quiescent and necrotic cells), as well as stronger deposition of extracellular matrix proteins as compared to 2D cultures of cells [5, 6, 16, 17].

Two main approaches are used for the generation of spheroids: scaffold-based and scaffold-free approaches [12–15, 18]. Each approach has its advantages and disadvantages and the choice of the approach to use is guided by the type of assays the investigator will perform with these spheroids. In the scaffold-based approach, the cells are seeded onto artificial matrices with different porosities, resulting in the growth of cells both around and inside the matrices. This approach is not the most appropriate for drug studies because of the low reproducibility between batches of spheroids, the possible adsorption of the drugs with the matrices, the presence of non-transparent matrices that disturb spectroscopic analysis and the difficulties in detaching and extracting the cells from the scaffold when performing Western blot or flow

#### **Figure 1.**

*Schematic representation of cells grown in standard 2D growth conditions (left) or as spheroids (right). A denser extracellular matrix is present around the cells in 3D growth conditions.*

*Using Tumor-Like Spheroids to Study the Effect of Anti-Cancer Drugs* In Vitro *DOI: http://dx.doi.org/10.5772/intechopen.113857*

cytometry analysis [18]. The second approach and the one that was chosen in this study is the generation of scaffold-free spheroids. Several methods deriving from the scaffold-free approaches have been developed and include the hanging-drop method, the agitation-based approach and the liquid overlay technique [9–11, 18]. This later method is one of the simplest, reproducible and cheapest methods to produce spheroids, and was chosen in the present study for these reasons [19, 20]. In this study, we report the generation of 3D spheroids from three different human melanoma cell-lines (WM115, WM266-4 and A375) using the liquid overlay technique, with the purpose of determining how the growth of cells in 3D conditions affect their sensitivity towards two different types of anti-cancer agents: temozolomide and vemurafenib. To assess the effect of the anti-cancer drugs, we used the Alamar Blue assay that uses changes of oxidative state of resazurin as an indication of changes in metabolic activities. We discuss issues and trouble-shooting solutions throughout our study.

## **2. Materials and methods**

#### **2.1 Cell lines**

The three human melanoma cell-lines, WM115, WM266.4 and A375 were purchased from ATCC (Manas, VA) and were grown in Dulbecco's Modified Eagle's Medium (DMEM, ATCC) supplemented with 10% FBS and penicillin (100 U/ml) and streptomycin (100 μg/ml). The absence of mycoplasma contamination in the cells prior to all experiments was confirmed by PCR. Note that cells in our laboratory are typically tested every 2 months for possible contamination with mycoplasma by PCR using the method described in Tang et al. [21]. For simplicity, WM226.4 cells will be referred to as WM266 throughout the rest of the manuscript.

#### **2.2 Cell culture plates**

For the generation of spheroids using the liquid overlay technique, cells are typically seeded in round-bottom 96-well plates. These plates are made of plastic polymer. Standard tissue culture plates are made from two types of plastic polymer: polystyrene or polypropylene [22]. Because these polymers are hydrophobic in nature, they need to be modified or "treated" by surface functionalization to become more hydrophilic, therefore allowing cells to adhere to the surface of the plates [22]. The generation of spheroids using round-bottom plates requires the use of non-tissue culture treated plastic plates, because cells should not adhere to the plastic but rather form multi-dimensional assemblies between them. However, it is still possible to use tissue-culture treated multi-well plates for the generation of spheroids. In this case, the surface of the wells should be coated with a polymer, such as poly 2-hydroxyethyl methacrylate (poly-HEMA), to prevent attachment of the cells to the plastic [23]. However, multi-well tissue culture plates with low adhesive properties designed for the generation of spheroids are now available from many suppliers.

In the present study, we compared the generation of spheroids from four different types of round-bottom 96-well plates provided by different manufacturers (**Table 1**). One type of plate was made of tissue-culture treated polystyrene (Plate 1). To prevent cell adhesion to this specific plastic surface, this plate was coated with poly-HEMA as described below. The three other plates (Plates 2, 3 and 4) were not tissue-culture treated. Two plates were made of polystyrene (Plates 1 and 2) and one plate of


*\*Nucleon-sphera treated plates have low adhesive properties following a non-disclosed proprietary process.*

#### **Table 1.**

*Characteristics of the round-bottom plates used for the generation of spheroids.*

polypropylene (Plate 3). One type of plate (Plate 4) was specifically marketed for the generation of spheroids.

## **2.3 Poly-HEMA coating**

The wells of the tissue-culture treated 96-well plate 1 (**Table 1**) were coated with poly-HEMA by adding 200 μl of 5 mg/ml poly-HEMA (Sigma-Aldrich, Saint-Louis, MO), prepared in 95% ethanol, to each well of a round bottom 96-well plate [23]. After addition of the poly-HEMA solution to the wells, the plates were left in a Biosafety cabinet for 72 h, allowing ethanol to evaporate completely.

#### **2.4 Methylcellulose preparation**

Methylcellulose was added to the cells to promote spheroid formation by acting as a cell repellent additive [24]. Methylcellulose is an inert polymer that forms a viscous and gel-like solution that promotes cell aggregation [25–27]. The methylcellulose solution was prepared after modification of the procedure described in Longati et al. [26]. Briefly, a 1.2% (w/v) methylcellulose stock solution was prepared by resuspending 6 g autoclaved methylcellulose (M1314, Spectrum) in 50 ml DMEM containing 10% FBS, penicillin (100 U/ml) and streptomycin (100 μg/ml). The solution was stirred overnight at 4° C and centrifuged for 2 h at 7500 RPM. After centrifugation, 20 ml of the clear viscous solution was added to 80 ml of DMEM containing 10% FBS, penicillin (100 U/ml) and streptomycin (100 μg/ml), resulting in a methylcellulose final concentration of 0.24%.

## **2.5 Cell seeding**

Cells grown to 80–90% confluence in T-75 cell culture flasks were detached with 0.05% trypsin, centrifuged for 2 minutes at 500 g and resuspended in DMEM media supplemented with 10% FBS, penicillin (100 U/ml), streptomycin (100 μg /ml), and 0.24% methylcellulose. Cells were then counted using an hematocytometer and added to the wells of the four different 96 well-plates (Plates 1, 2, 3 and 4) at two different seeding densities: 2500 and 5000 cells per well. The wells were observed every day (0, 24, 48, 72,

96, 120, 144 and 168 h) by microscopy, images of spheroids were taken from 12 different wells, and the total number of formed spheroids per plate was counted after 8 days.

#### **2.6 Resazurin assay**

Resazurin or 7-hydroxy-10-oxidophenoxazin-10-ium-3-one is a water soluble and permeable dye [28]. After entering cells, resazurin is reduced by accepting electrons from multiple intracellular electrons donors such as NADPH, FMNH, NADH or cytochromes, without altering the normal transfer of electrons in cells [28]. Non-viable cells will not reduce resazurin. Resazurin is generally non-toxic for many cell-types, but cellular toxicity has been reported [29]. Oxidized resazurin has a dark blue color and has little intrinsic fluorescence, however, reduced resazurin or resofurin, is pink and highly fluorescent with measured excitation and emission fluorescence wavelengths typically of 540 nm and 590 nm, respectively.

Resazurin or Alamar Blue (AB) (Amresco/VWR/Avantor, Radnor, PA.) was prepared at 0.1 mg/ml in 50 mM Phosphate buffer pH 7.4 containing 150 mM NaCl (PBS), sterile filtered using a 0.2 um filter, and stored at 4° C in the absence of light. When added to cells, 10% (vol./vol.) resazurin was added to each well of the plate. Control wells contained resazurin and media only. When measuring changes in metabolic activity of cells grown in flat surface, the AB fluorescence signals were detected after 6 h incubation with the dye. However, after adding AB to the spheroids, we waited a 24 h incubation period before measuring the AB signal.

#### **2.7 Staining of spheroids by immunofluorescence**

Ten-days old spheroids generated from A375, WM115 and WM266 cells were fixed with 4% paraformaldehyde (PFA) for 30 minutes and then stained with 0.1% eosin Y (Sigma-Aldrich, Saint Louis, MO) in 80% ethanol to facilitate the visualization of spheroids in the paraffin block. The spheroids were then paraffin embedded using a Leica ASP300S processor. Briefly, the spheroids were washed with a series of ethanol solutions, starting with 95% ethanol followed by two washes with 100% ethanol. The spheroids were then cleared by two 100% xylene baths and finally embedded in paraffin blocks (Tissue-Tek® Base Mold 4162, 15 x 15 x 5 mm) using a Sakura Tissue-Tek embedding station. The paraffin blocks were sectioned at 5 μm using Thermo Microm 355S microtome. The sections were then floated on a water bath, placed on VWR VistaVision TM Histobond © charged slides and air dried. The spheroid sections were treated with three 100% xylene washes to remove the wax from slides and rehydrated through a series of 100–70% ethanol washed followed by a last wash in deionized (DI) water. An antigen retrieval step was then performed for 20 minutes at 120°C in citrate buffer (at pH 6.0) using a Retriever 2100 (Diagnostic Technology). For the staining procedure, the spheroid sections were first washed with TBS-T for 3 min and blocked for 1 h at room temperature (RT) with 5% NGS in TBS blocking buffer (Vector laboratories, Newark, CA). The sections were then incubated with a Ki67 rabbit polyclonal primary antibody (AbCam # ab15580, Waltham, MA) at a 1:500 dilution in 1% BSA overnight at 4°C. After 3 min washes with TBS-T, the sections were further incubated with CF 633 labeled Goat anti-rabbit secondary antibody (Biotium #20122, Fremon, CA) at a 1:250 dilution in 1% Bovine Serum Albumin (BSA) in TBS for an hour at room temperature. Finally, the sections were washed for 1 min with DI water and stained with 4,6-diamidino-2-phenylindole (DAPI) (Biotium #40043) before being mounted using the EverBrite™ Hardset Mounting Medium

(Vector laboratories #23003). The slides were imaged using a Zeiss Axio Observer Z1 LSM 700 microscope using 20x 0.8NA lens. (Carl Zeiss Microscopy LLC, White Plains, NY).

In a different experiment, 72 h old spheroids were stained for the cell viability marker calcein acetoxymethyl (AM) ester (2 μM; Sigma-Aldrich), the dead cell marker Ethidium Homodimer-1 (EthD-1), (3 μM; Invitrogen/Thermo Fisher Scientific, Waltham, MA) and the nuclear marker Hoescht 33,342 (33 μM; Thermo Fisher Scientific) according to a procedure described in [30]. The spheroids were stained simultaneously with the three dyes in the 96 well plate for 30 min at 37°C. After incubation, the spheroids were carefully transferred to a glass bottom dish (Ibidi USA, Fitchburg, WI) for imaging with a Zeiss Axio Observer Z1 epifluorescence microscope equipped with a LSM 700 confocal scanning head, and using a 20×, 0.8 NA objective (Carl Zeiss, Thornwood, NY). Images were captured after excitation/emission of 494/517 nm for the detection of calcein-AM positive cells, 528/617 nm for the detection of EthD-1-positive cells and 360/461 nm for the detection of Hoescht 33,342 positive cells.

## **2.8 Statistical analysis of the data**

Twelve spheroids were imaged for each cell line in each plate. **Figure 2** shows only a representative image from one of these 12 wells. The diameter of 12 independent spheroids were measured each day, for a period of 8 days, using ImageJ software [31]. The mean and the standard deviation were used for the graphs shown in **Figure 3**. A student's t-test was performed to indicate the significance between spheroids'

#### **Figure 2.**

*Images of spheroids formed from A375 cells (A), WM115 cells (B) and WM266 cells (C) at different time points: 0, 24, 48, 72, 96, 120, 144 and 168 h scale bar: 250 μm.*

*Using Tumor-Like Spheroids to Study the Effect of Anti-Cancer Drugs* In Vitro *DOI: http://dx.doi.org/10.5772/intechopen.113857*

#### **Figure 3.**

*Diameters of spheroids formed from 2500 (A) and 5000 (B) A375 cells, from 2500 (C) and 5000 (D) WM115 cells, and from 2500 (E) and 5000 (F) WM266 cells at different time points (0 h, 24 h, 48 h, 72 h, 96 h, 120 h, 144 h and 168 h). \*\* p < 0.001; \*\*\* p < 0.001.*

#### **Figure 4.**

*Staining of spheroids formed from 2500 (A) and 5000 (B) A375 cells, 2500 (C) and 5000 (D) WM115 cells, and 2500 (E) and 5000 (F) WM266 cells with the proliferative marker Ki-67 (pink). The nuclei are stained in blue with Hoescht 33,342. Shown on the figure are the merged images. Scale bar: 50 μm.*

diameters at different time points with \*p < 0.05; \*\*p < 0.001 and \*\*\*p < 0.001. **Figures 4** and **5** show representative images of stained spheroids. The cell-based assays were repeated at least three independent time. The data points from the titration with vemurafenib and temozolomide were fit using Kaleidagraph version 5.0 software (Synergy Software).

#### **Figure 5.**

*Staining of spheroids formed from 2500 (A) and 5000 (B) A375 cells, 2500 (C) and 5000 (D) WM115 cells, and 2500 (E) and 5000 (F) WM266 cells with calcein-AM (green) and EthD-1 (pink). The nuclei are stained in blue with Hoescht 33,342. Shown on the figure are the merged images. Scale bar: 50 μm.*

## **3. Results and discussion**

#### **3.1 The morphology, shape and growth rate vary among cell lines**

Our first goal was to determine if the three different cell lines could form spheroids using the scaffold-free liquid overlay technique. For each cell line, we tested the formation of spheroids in four commercial plates. These plates were made of polystyrene (Plates 1, 2 and 4) or polypropylene (Plate 3) and were either tissue-culture treated (Plate 1) or not (Plates 2, 3 and 4). In case of plate 1, to avoid cell adhesion to the plastic surface, the plate was coated with poly-HEMA prior to seeding the cells. The surface of plate 4 had already been pre-treated by the manufacturer using a non-disclosed proprietary process to enhance spheroid formation. Two different cell densities (2500 and 5000 cells) were seeded for each type of plate. The presence of spheroid was assessed every day for 7 days (0, 24, 48, 72, 96, 120, 144 and 168 h) by microscopy.

We first compared the yield of spheroids produced on the different plates for each cell line. We noticed that in our experimental conditions, a larger number of spheroids was generated using the plates P2, P3 and P4, whereas the poly-HEMA coated plate produced lower yields of spheroid for each of the three cell lines examined (**Table 2**).

Observation of the spheroids by microscopy every day during the time course of 8 days (168 h), revealed that the spheroids generated with the different cell lines had different morphologies and shapes (**Figure 2**). Differences in the ability of cancer cells to form spheroids has been previously reported [32]. A375 cells did not produce spherical spheroids but rather irregular-shape spheroids with small assemblies of cells protruding in different directions. WM155 cells produced *Using Tumor-Like Spheroids to Study the Effect of Anti-Cancer Drugs* In Vitro *DOI: http://dx.doi.org/10.5772/intechopen.113857*


#### **Table 2.**

*Number of spheroids formed per number of examined wells (54 or 60). Poly-HEMA coated plate 1 produced the smallest yield of spheroids. All other plates generated spheroids in every well.*

spheroids with the most regular shape (spheres) whereas WM266 cells generated more oval-shaped spheroids. WM115 cells produced the smallest size spheroids from all three cell lines (**Figure 2**). Note that the WM115 and WM266 cell lines originate from the same patient: whereas WM115 was established from a primary tumor, WM266 was established from a metastatic tumor [33]. Our observation suggests that the choice of the cell line for the generation of spheroid is an important aspect for assuring the success of the study. The size of regular-shape spheroids is more easily measured than that of irregular-shape spheroids. We indeed experienced issues when measuring the diameter of many A375 spheroids as well as some WM266 spheroids whose shapes were not spherical. We also noticed that it was very difficult observing spheroids generated in the P3 polypropylene plate. The plastic presented some opacity that hindered the microscopic observation of the spheroids. If a study aims to follow changes in spheroids' size overtime by brightfield microscopy, these aspects are important to consider, as previously reported by other groups [34].

To assess the changes in spheroids' size overtime, we measured the diameter of 12 spheroids every day for a period of 7 days, in plates 2, 3 and 4. Because the yield of spheroid formation was very variable from plate 1, we chose not to study the change in spheroids' size overtime from this plate. The first measurement was performed 24 h after seeding the cells in the round-bottom wells. The average diameter for these 12 spheroids over time is presented in **Figure 3**. We observed similar growth changes in the size of the spheroids in all three plates. Overall, the size of the spheroids tended to decrease for the first 48 h to 72 h, before slowly increasing over the next 4 days. Observation of the spheroids by microscopy showed that immediately after seeding and centrifuging the cells in the wells, the spheroids initially consisted of an assembly of cells loosely connected. The space between cells seemed then to reduce within the first 3 days, resulting in an apparent shrinkage of the spheroids. The size of the spheroids then increased in the next 4 days, probably because of cell division (**Figures 2** and **3**). This observation was similar for each cell line in each of the three plates. For conciseness, we give below numerical data of the changes in diameter observed for each cell-line in plate 4. As mentioned above, the changes were similar in plates 2 and 3.

From all three cell lines, WM115 cells showed the largest initial size reduction between 24 h and 72 h, with a reduction of 37.5% and 36.1% in the average diameter for spheroids generated in Plate 4 with 2500 and 5000 cells respectively. Between

72 h and 168 h, these spheroids grew and increased their size by 15.5 and 9.5%, respectively. WM115 generated spheroids with the smallest size from the three cell lines examined. After reaching their lowest size at 72 h with a decrease of 5.3 and 13.9% for spheroids generated with 2500 and 5000 WM266 cells respectively, these spheroids later increased their size between 72 h and 168 h by 30.7 and 13.6% respectively (Plate 4). A375 spheroids also saw a decrease in their diameters between 24 h and 72 h 0f 10.9% and 16.3% for spheroids generated from 2500 and 5000 cells respectively, and an increase between 72 h and 168 h by 30.7% and 12.1% respectively. For all three cell lines examined, the growth increase between 72 h and 168 h was larger with 2500 cells than with 5000 cells (**Figure 3**). This suggests that cell seeding density is an important factor to consider if the study aims to use changes of spheroid size when investigating the effect of drugs on spheroid growth. Another important point to consider when performing a drug study is the choice of the age of the spheroids for initiating the drug study. As we just showed, the spheroids generated from the three cell lines all "shrank" in the first 72 h. Starting a drug study earlier than 72 h could hamper the analysis of the data, if one would aim to investigate changes in spheroid size as an effect of the drug. Initial changes of spheroid size have been recently explained by the effect of cell-cell adhesion forces that pull cells towards each other's resulting in more compact structures [35]. In a different study, Thakuri et al. did not observe the initial size reduction in spheroids when dispensing dextran-respuspended HT-29 cells in a poly-ethylene glycol solution [36], suggesting that experimental conditions can greatly modulate the behavior of cells within spheroids.

## **3.2 Spheroids possess proliferative and metabolically active cells at their periphery and edges**

We imaged 8–10 days old spheroids for the presence of proliferative cells using antibodies against the proliferative marker Ki-67. Ki-67 is a nuclear protein that is present in all stages of the cell cycle but is absent in quiescent cells and has been previously used for the staining of spheroids [37]. We first observed differences in abundancy in Ki-67 positive cells in the spheroids generated from the three different cell types. WM266 spheroids showed the highest amount of Ki-67 positive cells (**Figure 4E** and **F**). WM115 showed a lower percentage of Ki-67 positive cells, both with spheroids generated from 2500 and 5000 cells (**Figure 4C** and **D**). The presence of Ki-67 positive cells at the edges of the spheroid was clearly evidenced with the irregular-shape A375 spheroids generated from 5000 cells (**Figure 4A** and **B**).

To further evaluate the viability of cells within the spheroids, we stained 72 h old spheroids with calcein-AM, ethidium homodimers and Hoescht 33324 as described in [30, 38]. In these conditions, nuclei from both viable and dead cells are labeled with Hoescht 33342, the viable cells are labeled with calcein-AM (green) and the dead cells with ethidium homodimer-1 (pink). Imaging of the stained spheroids revealed that the calcein-AM positive cells were at the periphery of the spheroids (**Figure 5**). After the staining procedure and during the transfer of the spheroids from the roundbottom well of the 96 well plate to the imaging dish, many spheroids from A375 and WM115 cells broke, and only incomplete fragments could be imaged (**Figure 5**). However, these fragments do show that the metabolically active and calcein-AM positive cells are located at the edge of the spheroids in agreement with the observation of Ki-67 positive proliferative cells present at the edge of the spheroids as well (**Figure 5**). Furthermore, the non-fragmented WM266 spheroids also showed the

*Using Tumor-Like Spheroids to Study the Effect of Anti-Cancer Drugs* In Vitro *DOI: http://dx.doi.org/10.5772/intechopen.113857*

presence of metabolically more active cells at the periphery of the spheroids. Staining of the spheroids with EthD-1 revealed only few dead cells (pink) in the 72 h old spheroids (**Figure 5**). The presence of the external layer of proliferative and metabolically active cells in the spheroids generated from all melanoma three cell lines (**Figures 4** and **5**) is in agreement with previous studies (reviewed in [39]).

## **3.3 Sensitivity of melanoma cells towards temozolomide and vemurafenib in 2D and spheroid 3D growth conditions**

After identifying the optimal properties for the generation of the spheroids from the three different melanoma cell lines, our goal was to compare the sensitivity of the three melanoma cell types, grown in standard 2D growth conditions and as spheroids, towards two cancer drugs, temozolomide (TMZ) and vemurafenib. TMZ is an alkylating agent that alkylate guanine bases in DNA leading to DNA cross-linkages and impaired cell division, thus reducing melanoma cell growth [40].

When comparing the effect of TMZ on cells either grown in 2D conditions in cell culture flasks and as spheroids, we observed similar patterns of sensitivity. In general, A375, WM115 and WM266 all showed some resistance to TMZ as the highest concentration tested (1000 μM) only reduced the signal by 50% with WM115 cells (**Figure 6D**). For A375 and WM266, the signal observed at 1000 μM was only 37.5% and 30% lower than that in the absence of TMZ (**Figure 6A** and **G**). In our experimental conditions, we were not able to increase the TMZ concentration at higher levels due to the toxic effect of DMSO. For A375 and WM115, when comparing the effect of TMZ on the spheroids to that on cells, we observed that TMZ effected a smaller proportion of cells grown as spheroids than on flat surfaces (**Figure 6B** and **C** for A375 spheroids and E and F for WM115 spheroids). Only WM266 spheroids appeared to be slightly more sensitive towards TMZ with only 30% signal reduction by 1000 μM TMZ in cells and about 50% and 45% reduction by the same concentration of TMZ in the spheroids generated from 2500 and 5000 cells, respectively (**Figure 6G, H** and **I**). These results reflect the pattern and distribution of the proliferative marker Ki-67 in the A375, WM115 and WM266 spheroids. Indeed, because TMZ affects cell division, it is only effective in cells that divide, thus being most effective in proliferative cells, such as those staining positive for Ki-67. As observed in **Figure 5**, the spheroids generated from WM266 are those that possess the largest percentage of proliferative cells, from the three cell lines examined, and therefore, should be the spheroids that respond the most to TMZ. However, we also observed that a larger fraction of the WM266 cells responded to TMZ when assembled as spheroids than as monolayers (**Figure 6G**, **H** and **I**).

The second drug that we tested in our study was vemurafenib. Vemurafenib is a BRAF kinase mutant inhibitor [41]. This small molecule drug only inhibits the mutant form of the kinase and is therefore only effective when the melanoma cells carry the appropriate BRAF V600E mutation. By blocking the BRAF kinase, vemurafenib inhibits the growth of the melanoma cells by inhibiting the BRAF/Map kinase pathway. All three cell lines used in our study carry the BRAF V600E mutation and were thus suitable for the study [33, 42].

We first observed that when testing vemurafenib on cells grown on 2D flat surfaces, WM115 showed the strongest IC50 for vemurafenib with a calculated IC50 = 0.057 ± 0.021. Based on the shape of the curve, it appeared that most cells in the dish were affected by vemurafenib (**Figure 7D**). This was not the case for A375 (**Figure 7A**) and WM266 (**Figure 7G**) where only a portion (less than 50%) of the

**Figure 6.**

*Effect of temozolomide on: 2D grown A375 cells (A) and 3D A375 spheroids formed from 2500 (B) and 5000 (C) cells; 2D grown WM115 cells (D) and 3D WM115 spheroids formed from 2500 (E) and 5000 (F) cells; and 2D grown WM266 cells (G) and 3D WM266 spheroids formed from 2500 (H) and 5000 (I) cells.*

cells seemed to be affected by the BRAF inhibitor. A similar effect of vemurafenib on only a portion of the cells was also observed in spheroids generated from WM115 and WM266 cells (**Figure 7E**, **F**, **H** and **I**). Interestingly, vemurafenib appeared to affect most of the cells within the A375 spheroids, with the fluorescence signal decreasing by up to 80% at the highest vemurafenib concentration used (**Figure 7B** and **C**).

Vemurafenib inhibits the BRAF/Map kinase pathway, and by doing so, affects cell proliferation [41]. Based on the larger population of proliferative cells in the WM266 spheroids (**Figure 5E** and **F**) when compared to the A375 and WM115 spheroids, it was expected that these spheroids would respond the best towards vemurafenib. However, we observed that for these spheroids, the fluorescence signal decreased by about 37 and 25% in the presence of the highest concentration of vemurafenib, for spheroids generated from 2500 and 5000 cells respectively. We are currently investigating the reasons of this discrepancy. As future studies, it would be particularly interesting to stain the vemurafenib treated spheroids with Ki-67, as well as with calcein-AM and ETHD-1 to identify the regions of the spheroids affected by the drug. *Using Tumor-Like Spheroids to Study the Effect of Anti-Cancer Drugs* In Vitro *DOI: http://dx.doi.org/10.5772/intechopen.113857*

**Figure 7.**

*Effect of vemurafenib on: 2D grown A375 cells (A) and 3D A375 spheroids formed from 2500 (B) and 5000 (C) cells; 2D grown WM115 cells (D) and 3D WM115 spheroids formed from 2500 (E) and 5000 (F) cells; and 2D grown WM266 cells (G) and 3D WM266 spheroids formed from 2500 (H) and 5000 (I) cells.*

Our study showed that the response of the cancer cells towards the two cancer agents TMZ and vemurafenib was affected by the architecture of the cells (3D spheroids versus 2D monolayers), but that the response to the drugs depended on both the cell type and the drug itself. Indeed, although the A375 cells appeared to be more resistant towards TMZ when present as 3D spheroids than as 2D monolayers (**Figure 6A–C**), they were more sensitive to vemurafenib when present as 3D spheroids than as monolayers (**Figure 7A–C**). In an earlier study, Filipiak-Duliban et al. had also shown that the drug sensitivity of cancer cells was affected by the mode of culture of the cells [43]. In their study, the authors showed that the B16F10 murine melanoma cells were more resistant to the three anti-cancer drugs everolimus, doxorubicin and cisplatin when grown as spheroids than as 2D monolayers [43]. However, the authors also showed that the effect of the 3D structure on drug sensitivity was not observed when studying a renal cancer cell line [43]. These data, in agreement with our own observations, strongly suggest that the effect of the 3D structure of the cell assemblies depends on both the cell type and the drug that is studied. In addition, the

size of the spheroids has also been shown to affect drug resistance, with spheroids of larger sizes being in general more resistant than spheroids of smaller sizes [44]. In our study, we also observed that spheroids generated from 5000 WM115 cells were more resistant towards TMZ than spheroids generated from 2500 cells. Similarly, Spheroids generated from 5000 WM266 cells were more resistant to vemurafenib than spheroids generated from 2500 cells. However, the size of the spheroids did not seem to affect the sensitivity of WM115 cells towards vemurafenib, or the sensitivity of WM266 cells towards TMZ, making the relationship between each cell line and each drug unique.

## **4. Conclusions and perspectives**

Research performed in the last decade has clearly demonstrated that cells grown as 3D spheroids represent a more accurate model of tumors than cancer cells grown as monolayers in cell culture flasks [11, 39]. As shown in our study, the spheroids generated from the three human melanoma cells all showed features of tumors. For instance, all three types of spheroids exhibited a layer of proliferative and metabolically active cells at the edge of the spheroids, as shown by their positive staining with Ki67 and calcein-AM. However, we are also aware that our spheroid-based model presents weaknesses and needs to be improved to better mimic tumors. One of the weaknesses is that our spheroids were generated from cancer cells only. Tumors are formed not only from cancer cells, but also from other cell types such as fibroblasts, macrophages, pericytes and endothelial cells. All these cells communicate with each other's in tumors [45]. Therefore, in tumors, responses to anti-cancer drugs can be very different than the response to the same drugs in spheroids generated from a single type of cells. To solve this issue, new generation of multicellular spheroids are being developed and studied [10, 46–49]. For example, Klicks et al. recently generated spheroids composed of a mixture of fibroblasts, keratinocytes and melanoma cells [48]. However, challenges remain when using multicellular spheroids for the study of anticancer drugs. One of those challenges reside in the generation of consistent spheroids of homogeneous size and shape [50–52]. As we have shown in our study, certain cell lines such as A375, generate irregular shape spheroids that are difficult to study. In addition, in an effort to better mimic the complexity of tumors' architecture and cellular heterogeneity, spheroids are now combined with bioprinting technologies, resulting in additional challenges due to, among other factors, the mechanical instability of spheroids [53, 54]. However, with the fast advancement of new technologies, these challenges will be surely overcome in a near future [55–59].

## **Acknowledgments**

We would like to thank Virginia Montgomery from the Advanced Imaging Microscopy (AIM) Core Facility at NDSU for her assistance in staining the spheroids. This study was supported by NIH-NIGMS Award number U54GM128729 (DaCCoTA Scholar Award to E.L.), the National Science Foundation under NSF EPSCoR Track-1 Cooperative Agreement OIA#1946202 and the College of Health Professions at NDSU.

## **Conflict of interest**

"The authors declare no conflict of interest."

*Using Tumor-Like Spheroids to Study the Effect of Anti-Cancer Drugs* In Vitro *DOI: http://dx.doi.org/10.5772/intechopen.113857*

## **Author details**

Yousuf Alam, Pawel Borowicz, Stefan W. Vetter and Estelle Leclerc\* North Dakota State University, Fargo, NC, United States

\*Address all correspondence to: estelle.leclerc@ndsu.edu

© 2023 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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Section 4
