The prune pectin fraction (PWH) was characterized and kindly provided by the Department of Biochemistry at the Federal University of Paraná (Paraná, Brazil). Pitted prunes (dried fruits from P. domestica purchased at a local market in Curitiba, Brazil; LA VIOLETERA^®) were freeze-dried and milled. The extraction of pectic polysaccharides was carried out using hot water in order to obtain the molecules more tightly bound to the cell wall. Moreover, hot-water extraction results in a higher pectin yield compared to cold-water extraction. The water extract was obtained by filtration, and the polysaccharides were recovered by ethanol (3 vol.) precipitation and lyophilization, originating the fractions. A homogeneous fraction was analyzed by sugar composition, high-performance steric exclusion chromatography, methylation, and nuclear magnetic resonance spectroscopy analyses. The PWH comprises rhamnogalacturonans with type I arabinogalactans as side chains, and low-methyl esterified homogalacturonan (9). The freeze-dried pectin fraction was solubilized in the cell growth medium [RPMI-1640 medium (Gibco; Thermo Fisher Scientific, Inc.), supplemented with 10% fetal bovine serum (FBS, Gibco; Thermo Fisher Scientific, Inc.) and 1% antibiotic (penicillin 10,000 U and streptomycin 10 mg/l), MilliporeSigma] and stored at -20˚C until its use.

Cell groups were divided into a control group (CT) and a group treated with prune pectins (PWH). The control group received only the cell growth medium without PWH. For cytotoxicity analysis, doxorubicin (DX, Pfizer^®) at a concentration of 2.5 µg/ml was used as a positive control. In this case, the same control medium was used, supplemented with the chemotherapy drug.

The B16F10 murine melanoma cell line (BCRJ, 0046) and BALB/c 3T3 normal cell line (clone A31, ATCC, CCL-163) were kindly provided by the Laboratory of Inflammatory and Neoplastic Cells, Department of Cell Biology, UFPR, Curitiba, Brazil, and were initiated and maintained according to specific recommendations for each line. The BALB/c 3T3 cell line, derived from fibroblasts of BALB/c mouse embryos, is non-tumorigenic and widely employed as a representative model of normal cells in contrast to tumor cell lines. Given that fibroblasts are distributed throughout the body, this lineage provides an appropriate model for assessing normal cell function in studies of viability and cytotoxicity. Cells were cultured in growth medium and maintained in an incubator (Sanyo Scientific MCO-18AC) at 37˚C, 90% humidity, and 5% CO₂ for 72 h. All experiments were performed in biological triplicates.

To determine the concentrations to be used in the present study, a cell viability and cytotoxicity test was carried out to determine the concentrations that were not cytotoxic to normal cells. For this, the colorimetric tests of reduction of diphenyltetrazolium bromide (MTT) and neutral red (NR) were performed by the protocols proposed by Mosmann (16) and Repetto et al (17), respectively. The B16F10 cells (5x10² cells/well) and BALB/c 3T3 (2x10³ cells/well) were exposed to 0, 2.5, 5, 10, 100 and 800 µg/ml PWH, and 2.5 µg/ml DX for 72 h. Briefly, the MTT cell viability assay was performed by first seeding the cells into 96-well plates and treating them as described in the experimental design. Following treatment, the wells were aspirated to remove the culture medium (RPMI-1640, MilliporeSigma; containing fetal bovine serum 10% Gibco; Thermo Fisher Scientific, Inc.), and 100 µl MTT solution [3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide 5 mg/ml in PBS, MilliporeSigma, pH 7.4] were added to each well. The plates were incubated for 3.5 h at 37˚C in a humidified atmosphere with 5% CO₂ to allow the formation of formazan crystals by metabolically active cells. Following incubation, the MTT solution was discarded, and 100 µl dimethyl sulfoxide (DMSO, MilliporeSigma) were added to each well to solubilize the formazan crystals. The plates were gently shaken to ensure complete solubilization, and the absorbance was measured at 540 nm using a microplate reader (TECAN^® Infinite M200; Tecan Group, Ltd.). The absorbance values obtained were directly proportional to the number of viable cells.

The neutral red cell viability assay was performed by first treating cells cultured in 96-well plates according to the experimental protocol. Following treatment, the medium (RPMI-1640, MilliporeSigma; containing fetal bovine serum 10% Gibco; Thermo Fisher Scientific, Inc.) was removed, and 100 µl Neutral Red solution (MilliporeSigma, 40 µg/ml in culture medium) were added to each well. The cells were then incubated for 3 h at 37˚C in a 5% CO₂ atmosphere to allow dye uptake by viable cells. Following incubation, the dye solution was removed, and the cells were washed gently with 150 µl PBS (MilliporeSigma) to eliminate excess dye. Subsequently, 100 µl destaining solution (50% ethanol, 49% distilled water, and 1% glacial acetic acid, MilliporeSigma) was added to each well to extract the dye from the lysosomes. The plate was shaken until the complete solubilization of the dye, and the absorbance was measured at 540 nm using a microplate reader (TECAN^® Infinite M200). The absorbance values obtained were directly proportional to the number of viable cells. The results obtained from the MTT and neutral red assays (measured by absorbance) were converted into percentages, using the control group (without treatment with the PWH fraction) as having 100% viability.

The assay was performed through the crystal violet method, as previously described by Bonnekoh et al (18). The B16F10 cells (2x10³ cells/well) were plated in a new 96-well plate and exposed to 0, 10 and 100 µg/ml PWH. Cell viability was measured in 24, 48 and 72 h. Following the treatment period, the medium (RPMI-1640 MilliporeSigma; containing fetal bovine serum 10% Gibco; Thermo Fisher Scientific, Inc.) was removed, and the cells were gently washed with 150 µl PBS to eliminate non-adherent cells and debris. Subsequently, 100 µl of 4% paraformaldehyde solution (MilliporeSigma) were added to each well to fix the cells for 20 min at room temperature. Following fixation, the wells were washed again with PBS, and 100 µl 0.1% crystal violet solution (prepared in 20% methanol, Merck KGaA) were added to stain the cells at room temperature for 10 min. Excess dye was removed by rinsing the plate under running distilled water until the background was clear. After drying, the bound dye was solubilized by the addition of 100 µl 10% acetic acid (MilliporeSigma) to each well. The plate was gently shaken to ensure complete solubilization. The results were obtained using a TECAN^® Infinite M200 device at 590 nm. The data are presented as a percentage relative to the CT group at 24 h.

Following 72 h of exposure to 0, 10 and 100 µg/ml PWH, the B16F10 surviving cells were plated again (2x10⁴ cells/well) in a new 96-well plate with culture medium (RPMI-1640 MilliporeSigma; containing fetal bovine serum 10% Gibco; Thermo Fisher Scientific, Inc.) to perform the cell migration assay (‘scratch’ method) (19). Briefly, after the cells reached full confluency, cell proliferation was inhibited with the use of RPMI growth medium lacking in FBS (1%), and a straight scratch was made in the monolayer using a sterile 200 µl pipette tip. The wells were then gently washed with PBS to remove detached cells. Images of the scratch area were captured immediately (0 h) and following 24 h of incubation at 37˚C in a 5% CO₂ atmosphere using an inverted microscope (BIOVAL^® XDS-1B). The wound closure area was analyzed using ImageJ^® software 1.52 (National Institutes of Health), and the migration rate was expressed as the percentage of area covered by cells after 24 h compared to time 0(20).

Following 72 h of exposure to 0, 10 and 100 µg/ml PWH, B16F10 surviving cells were plated again at a reduced concentration (1x10² cells/well) in a 24-well plate to perform the colony formation assay analyses (21). This assay is expected to evaluate the ability of a single cell to form new colonies. For this, surviving cells were maintained only in the growth medium (RPMI-1640 MilliporeSigma; containing fetal bovine serum 10% Gibco; Thermo Fisher Scientific, Inc.) and incubated at 37˚C for 96 h. At the end of the incubation period, the colonies were fixed with 4% paraformaldehyde at room temperature for 15 min and then stained with 0.1% crystal violet solution (prepared in 20% methanol) for 10 min at room temperature. Excess dye was washed off with distilled water, and the plates were dried in air. The images were acquired using an inverted microscope (BIOVAL^® XDS-1B), and colonies containing >50 cells were counted using ImageJ^® software 1.52 (National Institutes of Health).

This assay aimed to evaluate morphological changes in B16F10 cells following exposure to the PWH fraction. The cells (5x10³ cells/well) were seeded in 24-well plates and treated with 0, 10, or 100 µg/ml PWH. Cell morphology was assessed at the start of treatment and again after 24 h. At each time point, the cells were fixed with 4% paraformaldehyde at room temperature for 10 min and stained with 0.25% crystal violet (prepared in methanol) at room temperature for 10 min (22). The area occupied by the cells in culture was quantified through digital analysis of images captured using a light microscope (BIOVAL^® XDS-1B) at x400 magnification. Images were processed using ImageJ software 1.52 (National Institutes of Health), applying threshold-based segmentation to differentiate cells from the background. The total cell area was subsequently measured in pixels.

RNA isolation from the B16F10 cell samples following 72 h of PWH treatment was performed using TRIzol^® reagent (Bio-Rad Laboratories, Inc.) according to the manufacturer's recommendations. The extracted RNA samples were then converted to cDNA using the iScript Reverse Transcription Supermix for RT-qPCR kit (cat. no. 1708841, Bio-Rad Laboratories, Inc.). The reactions were taken to the thermocycler (Applied Biosystems; Thermo Fisher Scientific, Inc.), and the samples were incubated in three cycles: 1st cycle of 5 min at 25˚C, 2nd cycle of 20 min at 46˚C, and 3rd cycle of 1 min at 95˚C. The RT-qPCR reaction was performed using the one-step RT-PCR SYBR^®-Green kit (cat. no. 1725270, Bio-Rad Laboratories, Inc.) and the appropriate primers for each reaction, as follows: Focal adhesion kinase (FAK) forward, 5'-TCTGTGGAATTGGCAATCGG-3' and reverse, 5'-TGGATGGTCTGCACTTGGTT-3'; beta actin (ACTB) forward, 5'-CTGTATTCCCCTCCATCGTG-3' and reverse, 5'-GGGTCAGGATACCTCTCTTGC-3'; hypoxanthine-guanine phosphoribosyltransferase (HPRT1) forward, 5'-GTTGGGCTTACCTCACTGCT-3' and reverse, 5'-TAATCACGACGCTGGGACTG-3'. The samples were analyzed in a QuantStudioTM 5 device (Applied Biosystems; Thermo Fisher Scientific, Inc.). For each reaction, a control RT (without conversion of RNA to cDNA) was performed. The relative quantification was measured according to the pre-set threshold fluorescence level of the target gene (FAK) compared to the endogenous controls used (ACTB and HPRT1). ACTB and HPRT1 were utilized for normalization. The quality and purity of total RNA were evaluated using a NanoDrop spectrophotometer (Thermo Fisher Scientific, Inc.). Samples exhibited A260/A280 ratios between 1.8 and 2.0 and A260/A230 ratios >1.8, indicating minimal contamination by proteins or phenolic compounds and falling within commonly accepted parameters for gene expression analysis. For each RT-qPCR reaction, 4.5 µl cDNA, synthesized from a standardized RNA concentration of 10 ng/µl, was used, corresponding to 45 ng total RNA per reaction. The assay performance was monitored using two endogenous reference genes (ACTB and HPRT1), selected for their stability across the analyzed Mus musculus samples. No-reverse transcriptase (RT-) controls were included to confirm the absence of genomic DNA contamination. Each sample was tested in triplicate, and the relative gene levels were normalized using the 2^-ΔΔCq method (23).

The normal distribution of values was verified using the Shapiro-Wilk test. One-way ANOVA followed by Tukey's test was performed in all analyses, apart from cell proliferation, which was analyzed using two-way ANOVA (mixed model) followed by the Bonferroni post hoc test, both through GraphPad Prism8 Software^® (Dotmatics). P<0.05 was considered to indicate a statistically significant difference. Tukey's test was used as a post hoc test. Data are presented as the mean ± SD of at least three independent experiments.

In the NR assay, all PWH concentrations significantly reduced the viability of the B16F10 cells compared with the CT group (Fig. 1A). In the BALB/c 3T3 cells, a significant reduction was observed only following treatment with 2.5 and 800 µg/ml PWH in relation to the CT group (Fig. 1B). The MTT assay revealed a similar viability profile for the B16F10 cells as in the NR assay (Fig. 1C). In the BALB/c 3T3 cells, cytotoxicity was detected only following treatment with 800 µg/ml PWH, although it remained lower than that induced by DX (Fig. 1D). Overall, PWH was less cytotoxic than DX across all concentrations and assays (Fig. 1A-C), apart from the concentration of 800 µg/ml in the NR assay (Fig. 1A). These assays suggest that the PWH fraction exhibits greater cytotoxicity toward tumor cells than toward normal cells.

Based on the cell viability data, concentrations were selected to avoid cytotoxic effects on normal cells while effectively reducing tumor cell viability. Subsequent experiments were performed using 10 and 100 µg/ml PWH, with the control (CT) group remaining polysaccharide-free (0 µg/ml). The results demonstrated the cytotoxicity of the PWH fraction on tumor cells over time. PWH did not significantly affect cell viability at 24 or 48 h compared to the CT group. However, after 72 h, viability was markedly reduced at both concentrations, with decreases of 23.6 and 40.3%, respectively (Fig. 2).

In the migration assay, tumor cells treated with the PWH fraction closed the scratch to a similar extent as untreated cells, indicating that PWH did not significantly impair cell migration at either concentration within 24 h (Fig. 3A).

By contrast, PWH effectively reduced colony formation at both concentrations. Compared to the CT group, the number of new colonies decreased by 19.6% in the cells treated with 10 µg/ml PWH and by 25.9% in those exposed to 100 µg/ml PWH (Fig. 3B). These data indicated that tumor cells exhibited a reduced proliferation and diminished colony formation capacity following treatment with PWH.

Morphological changes were observed in the cells exposed to PWH under an inverted microscope. To confirm and further investigate these changes, a detailed morphological analysis was performed. Immediately after treatment (0 h), no noticeable differences were detected between the treated and control cells (Fig. 4A). However, following 24 h of exposure, the cells treated with 100 µg/ml PWH exhibited a 30% reduction in the area occupied by cultured cells compared to the control group (Fig. 4B), suggesting that contact with the polysaccharides in the PWH fraction interfered with the cytoskeletal organization of tumor cells.

Based on the observed effects on cell proliferation, colony formation and morphology, it was hypothesized that PWH may influence the focal adhesion points of cells. To determine this, FAK gene expression was quantified. FAK is a central regulator that promotes focal adhesion to the extracellular matrix and cytoskeletal remodeling. The results revealed the altered expression of the FAK gene in the B16F10 cells treated with PWH, with a 57% increase at the concentration of 10 µg/ml and a 3-fold increase at the concentration of 100 µg/ml (Fig. 5).