4-Methylumbelliferone (4-MU) was purchased from Wako Pure Chemicals (Osaka, Japan). An anti-BAX antibody (ABC11) was obtained from Millipore (Bedford, MA, USA). An anti-β-actin antibody (AC-15) was purchased from Sigma (St. Louis, MO, USA). 3-(4,5-Dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide for the MTT assay and hyaluronidase were purchased from Sigma.

Cells from the canine breast cancer cell line CF33 were obtained from the American Type Culture Collection (Manassas, VA, USA). The cells were grown as monolayers, cultured in DMEM (Nissui, Tokyo, Japan) supplemented with 10% heat-inactivated fetal bovine serum (FBS), 4 mM L-glutamine, 10 mg/ml streptomycin and 10,000 =U/ml penicillin G. The cells were maintained at 37°C in a 5% CO₂ incubator. The 4-MU stock solution for experiments was dissolved in DMSO and the final concentration of DMSO in the medium was adjusted to 0.1%.

To determine the effect of 4-MU on CF33 cells, we quantified HA in conditioned medium using a commercially available competitive enzyme-linked immunosorbent assay kit (ELISA) (Echelon Biosciences, Salt Lake City, UT, USA), in which the colorimetric signal was inversely proportional to the amount of HA present in the sample.

To analyze the effect of 4-MU on the areas of the HA-rich pericellular matrices, we used a particle exclusion assay. Fixed sheep erythrocytes (Sigma) were reconstituted in phosphate-buffered saline (PBS) to a density of 5.4×10⁸ cells/ml and used for the particle exclusion assay as previously described (30). HA matrices were visualized by adding 6.0×10⁷ erythrocytes to the growth medium and viewing under an All-in-One Fluorescence Microscope BZ-9000 (Keyence Corp., Osaka, Japan). The HA-rich pericellular matrix-to-cell-area ratios were determined by image analysis using ImageJ. All measurements were made by tracing the digitized images. Non-treated medium and hyaluronidase-containing medium served as negative and positive controls, respectively.

The MTT assay, which is widely used to measure cell proliferation and to screen for anticancer drugs, is based on the reduction of a tetrazolium salt (31–33). We used the MTT assay to evaluate the effects of the 4-MU on cell proliferation. CF33 cells were plated in 96-well plates at 1×10³ cells/well. At each time point (days 0–4), 20 μl of MTT reagent was added and the plates were incubated for 4 h. Subsequently, 100 μl of 10% SDS and 0.01 N HCl were added to each well. Following overnight incubation, the absorbances were measured at 570 nm using a Model 550 microplate reader (Bio-Rad Laboratories, Tokyo, Japan). In these experiments, 5 replicate wells were used for each time point and the results were calculated as the means ± SD.

Cells were harvested and washed with PBS, resuspended in 70% ethanol in distilled water and kept at −30°C overnight. Prior to analysis, cells were mixed and incubated for 30 min in PBS containing 0.05 mg/ml propidium iodide (PI) and 100 U/ml RNase A. The suspension was then passed through a nylon mesh filter and analyzed by FACSCalibur (Becton-Dickinson, Franklin Lakes, NJ, USA). Furthermore, these data were analyzed using FlowJo 7 (Tree Star, Inc., Ashland, OR, USA).

Total-RNA was extracted from cells using the TRIzol reagent (Invitrogen, Carlsbad, CA, USA) and cDNAs were synthesized with a PrimeScript™ RT reagent kit (Takara Bio, Inc., Shiga, Japan) according to the manufacturer’s protocols. Real-time PCR was performed using SYBR Premix Ex Taq™ (Takara Bio) and an ABI Prism 7500 Real-Time PCR system (Applied Biosystems, Foster City, CA, USA) under the following conditions: 30 sec at 95°C; 40 cycles of 5 sec at 95°C and 34 sec at 60°C. Specific primer sets for BAX (forward, 5′-CGCATCGGAGATGAACTGGA-3′; and reverse, 5′-ACCAGTTTGCTGGCAAAGTAGAAG-3′) were purchased from Takara Bio, Inc. HAS1 (forward, 5′-GGACTACG TGCAGGTGTGTG-3′; reverse, 5′-CTCACCTAGGGGAC CACTGA-3′), HAS2 (forward, 5′-CTTAGAGCACTGGGA-3′; reverse, 5′-TCTAAAACTTTCACCA-3′), HAS3 (forward, 5′-AAGTAGGGGGAGTTGG-3′; reverse, 5′-CCCAGAGGC CCACTAA-3′), and GAPDH (forward, 5′-AAGGCTGAGA ACGGGA-3′; reverse, 5′-GGAGGCATTGCTGACA-3′) were obtained from Operon Biotechnologies (Tokyo, Japan). The specificity of each amplification was confirmed by a dissociation curve consisting of a single peak. All samples were amplified in triplicate in each experiment. The values were normalized by the expression of GAPDH.

Whole cell lysates were prepared with ice-cold RIPA buffer (50 mM Tris-HCl pH 7.5, 1% nonidet P-40, 150 mM NaCl, 0.1% SDS, 0.5% deoxycholic acid) containing 1 μg/ml leupeptin, 1 μg/ml pepstatin, 1 μg/ml aprotinin, 1 mM DTT, 1 mM NaVO₄ and 0.5 mM PMSF. The supernatants were saved as total cell lysates following centrifugation. Aliquots of the cell lysates (20 μg of protein) were separated by 12% SDS-PAGE and transferred to PVDF membranes (ATTO, Tokyo, Japan). Primary antibodies that bound to their antigens on the membranes were detected using appropriate HRP-conjugated secondary antibodies (Amersham Biosciences, Piscataway, NJ, USA) and SuperSignal Chemiluminescence or a WesternBright Sirius western blotting kit (Advansta, Menlo Park, CA, USA) according to the manufacturer’s instructions.

Statistical comparisons between two groups were made using an unpaired Student’s t-test. Values of P<0.05 were considered to indicate a statistically significant difference.

4-MU was originally found to inhibit HA synthesis in cultured human skin fibroblasts (25). To determine the effect of 4-MU on HA synthesis in the CF33 cells, we quantified the levels of HA in conditioned medium using the ELISA assay. As shown in Fig. 1, 4-MU was associated with decreased levels of HA production in a dose-dependent manner at all time points, whereas there was no change in the HA levels between parent cells and vehicle (DMSO)-treated controls. It was previously reported that the cell-associated HA matrix plays a critical role in the progression of several malignancies (13,34). We confirmed the influence of 4-MU on HA-matrix formation by particle exclusion assay. Abundant accumulation of the cell-associated HA matrix was shown in CF33 cells following incubation with control medium with or without DMSO (Fig. 2A and C). The HA matrix disappeared after treatment with Streptomyces hyaluronidase, indicating that these matrices were comprised of retained HA (Fig. 2B). Cells treated with 4-MU showed a significant dose-dependent decrease in the area of the cell-associated HA matrix and ratio of matrix area to cell area at each time point (Fig. 2D–G). Furthermore, 4-MU treatment caused a change in cell morphology. As seen in Fig. 2C–F, within 72 h of 4-MU treatment, CF33 cells tended to become flat in form and there was an expansion of the cell area compared with control. These findings suggested that 4-MU inhibited HA synthesis and weakened the cell-associated HA matrix. The observations that 4-MU effectively reduced HA production in canine mammary tumor cells were comparable to those previously reported in human and mouse cells.

Three HAS isoforms (HAS1, HAS2, and HAS3) have been identified in mammalian cells. It has been reported that inhibition of HA synthesis by treatment with 4-MU reduced mRNA expression by HAS2, HAS3, or both, in various cell lines, including melanoma, breast and ovarian cancers (27). To assess the effect of 4-MU on the expression of the HAS genes in CF33 cells, we investigated the expression of HAS using real-time RT-PCR. We found that the expression of HAS1 mRNA was undetectable in the CF33 cells (data not shown), which mainly expressed HAS2 and HAS3 mRNA. CF33 cells treated with 4-MU showed a tendency to decrease levels of HAS2 mRNA, while the expression of HAS3 was increased (Fig. 3A and B). These data indicated that 4-MU-related inhibition of HA synthesis was mainly associated with downregulation of HAS2. The increase in the levels of HAS3 mRNA may signify a back-up system for HA production and these events are likely to occur in HA-related genes (17,35).

The rate of cell proliferation is often associated with that of HA synthesis (35). Furthermore, it was previously reported that cell proliferation is inhibited by 4-MU in various cancer cells (27,28). Cell proliferation was assayed in the CF33 cells by MTT assay upon addition of 4-MU and at 0, 1, 2, 3 and 4 days later (Fig. 4). The number of cells in control cultures increased steadily during the days following plating, while the proliferation rate was dose-dependently suppressed by 0.2, 0.6 and 1.0 mM concentrations of 4-MU (Fig. 4). Notably, by day 4 of the 1.0 mM 4-MU treatment, proliferation was completely blocked (Fig. 4). This result demonstrated that 4-MU was able to inhibit the growth of canine mammary tumor cells as it has in the several other cancer cell types mentioned.

4-MU caused marked inhibition of the proliferation of CF33 cells in our experiments, and to determine whether 4-MU is associated with alteration of cell cycle progression in the CF33 cells, cells were grown to 70% confluence and the cell cycle distribution was analyzed by flow cytometry after 24-, 48- and 72-h exposure to 4-MU (0.2, 0.6 and 1.0 mM). Among the cells treated with 1.0 mM 4-MU at each time point, the percentage of cells in the S phase was lower than the corresponding value for the control cells, and the percentage of cells in the G2/M phases in cultures treated with 1.0 mM 4-MU was significantly higher than in the corresponding control cells (Fig. 5A–C). Following G2/M arrest, several cancer cell lines, notably certain breast cancer cell lines, exhibit morphologic changes consistent with apoptosis, i.e., plasma membrane blebbing, the appearance of a rounded morphology and eventual detachment from the surface of the tissue culture dish where the epithelial cancer cell lines normally grow as an adherent monolayer (36). The percentage of apoptotic cells in our specimens was quantified with PI staining and flow cytometric analysis, with the sub-G0/G1 peak representing apoptotic cells. As shown in Fig. 5D, apoptosis was induced more frequently in cells treated with 4-MU compared to control cells at all time points examined. The percentages of cells treated with 1 mM 4-MU in the sub-G0/G1 phase were 4.04, 9.44, and 12.27% after 24, 48 and 72 h of incubation, respectively (Fig. 5D). In particular, samples with cells treated with 1 mM 4MU showed percentages of apoptotic cells that were approximately 24 times higher than control samples (Fig. 5D). Next, to clarify the effect of 4-MU on apoptosis-related proteins, we measured BAX mRNA and protein expression levels using real-time RT-PCR and western blotting. The 4-MU-treated cells demonstrated higher levels of expression of BAX mRNA than control cells at all-time points examined (Fig. 6A). In addition, we observed that 4-MU (0.6 mM and 1.0 mM) caused dose-dependent increases in BAX protein levels at 48 and 72 h (Fig. 6C and D). These results indicated that 4-MU decreased cell survival mainly through an extrinsic apoptosis pathway. It is conceivable that the 4-MU-treated cells showed no change in the levels of BAX protein at 24 h (Fig. 6B) as a reflection of a lapse in time from mRNA expression to protein synthesis.