Potential use of alexidine dihydrochloride as an apoptosis-promoting anticancer agent

Most head and neck cancers begin in the epithelia that line the mucosal surfaces of the head and neck. In the United States alone, it is estimated that >29,370 new cases and 7,320 deaths are attributed to oral cavity and pharyngeal cancers in 2005 (1). At the time of diagnosis, 10% of patients harbor distant metastases, 50% have regional disease, and only 34% have localized tumors (1). Although early detection and diagnosis, along with advances in surgery, chemotherapy, and radiation therapy, have increased survival modestly, the 5-year overall survival rate remains at ∼59% (1), which underscores the need to develop novel therapeutics.

An approach commonly used for antitumor compound lead identification is to conduct a forward chemical biology screen (2). Chemicals are screened for “hits” that induce a particular phenotype before the cellular or protein target is identified. Such approaches often reveal novel targets and pathways. Paclitaxel, for example, was discovered to have antitumor activity in a broad range of rodent tumors several years before it was identified to target microtubules (3, 4). Studies involving rapamycin led to the discovery of the Tor protein nutrient response signaling network (5, 6). Large-scale, phenotype-based assays are currently being done by the National Cancer Institute/National Institutes of Health Developmental Therapeutics Program and the In vitro Cell Line Screening Project for the purpose of discovering novel therapeutic molecules (7).

Our group has extensive experience in identifying new molecular therapies that improve outcome for head and neck cancers, ranging from adenoviral (8–14) to antisense oligonucleotide (15) approaches. To this end, we have recently done a cell-based, phenotype-driven, cell viability–based high-throughput screen for novel head and neck cancer cytotoxics (16). This screen identified several known anticancer compounds from the LOPAC1280 (Sigma-Aldrich Corporation, St. Louis, MO) and Prestwick Chemical Libraries (Prestwick Chemical, Inc., Washington, DC). Significantly, several novel potential anticancer compound leads were also identified.

Alexidine dihydrochloride is perhaps the most interesting of the identified molecules. This chemical was among the most potent cytotoxics screened. In addition, alexidine dihydrochloride is a bisbiguanide compound (17), a class of chemicals never previously evaluated for anticancer properties. The objective of this current study is to evaluate the cancer-specific properties of alexidine dihydrochloride and to assess its mode of action in head and neck cancers.

FaDu (human hypopharyngeal squamous cancer) and NIH/3T3 (untransformed mouse embryonic fibroblast) cells were obtained from American Type Culture Collection (Manassas, VA), whereas GM05757 (primary normal human fibroblast) and HNEpC (primary normal human nasal epithelial line) cells were obtained from the Coriell Institute for Medical Research (Camden, NJ) and PromoCell (Heidelberg, Germany), respectively. All cell lines were cultured according to specifications. C666-1 (human undifferentiated nasopharyngeal cancer) cells were cultured in RPMI 1640 supplemented with 10% fetal bovine serum (Wisent, Inc., St. Bruno, Quebec, Canada) and antibiotics (100 mg/L penicillin and 100 mg/L streptomycin) as previously described (8, 15). All experiments were conducted when cells were in an exponential growth phase.

Alexidine dihydrochloride was obtained from Toronto Research Chemicals, Inc. (Toronto, Ontario, Canada), and dissolved in DMSO at a concentration of 10 mmol/L, with subsequent dilutions done in H₂O. Cisplatin, 5-fluorouracil, and paclitaxel were obtained from the Princess Margaret Hospital Pharmacy Department (Toronto, Ontario, Canada) and diluted in PBS. For a negative control (the vehicle alone), DMSO was diluted in H₂O to a concentration corresponding to the DMSO in the respective drug-treated group.

Cells were seeded in 96-well plates (Corning Life Sciences, Acton, MA) at 5,000 per well in 100 μL of growth medium and allowed to incubate for 24 hours. The chemicals were then added as indicated, in a total volume of 5 μL/chemical. After 48 hours, the CellTiter 96 AQ_ueous One Solution Cell Proliferation Assay (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium, inner salt; MTS; Promega Corporation, Madison, WI) was used to detect cell viability according to the specifications of the manufacturer. A 1-hour MTS incubation time was used and 490 nm absorbance was measured using a SpectraMax Plus³⁸⁴ microplate reader (Molecular Devices Corporation, Sunnyvale, CA). DMSO (0.1%)–treated cells for each respective cell line were used as a negative control, and cisplatin (166.6 μmol/L)–treated cells for each respective cell line were used as a positive control.

Where indicated, cells were irradiated immediately before compound treatment. Irradiation was done at room temperature using a ¹³⁷Cs unit (Gammcell 40 Extractor; MDS Nordion, Ottawa, Ontario, Canada) at a dose rate of 1.1 Gy/min.

FaDu cells were seeded (0.3 × 10⁶/T-25), incubated for 1 day, and treated with either alexidine dihydrochloride (4 μmol/L) or vehicle alone for 48 or 72 hours. A 72-hour time point was selected for this assay because DNA degradation occurs late in FaDu cell apoptosis. After treatment, detached and adherent cells were collected (detached cells were decanted, whereas adherent cells were trypsinized and mixed with the respective detached cells), pelleted at 200 × g, resuspended in a 1.5 mL hypotonic fluorochrome solution (50 μg/mL propidium iodide, 0.1% sodium citrate, 0.1% Triton X-100; Sigma-Aldrich), and left in the dark at 4°C overnight. Flow cytometric analysis was then done (FACSCalibur, Becton Dickinson, San Jose, CA) and analyzed using FlowJo (Tree Star, Inc., San Carlos, CA). Sub-G₁ peaks were identified by ModFit LT (Verity Software House, Inc., Topsham, ME).

Cells were seeded (0.3 × 10⁶/T-25), allowed to incubate for 1 day, and treated as described. After 48 hours, detached and adherent cells were collected together, stained with a 10 μmol/L Hoechst 33342 (Invitrogen Corporation, Carlsbad, CA)-4% formalin-PBS solution, and visualized under UV light using a Zeiss Axioskop HBO 40 microscope (Zeiss, Thornwood, NY).

Cells were treated with either alexidine dihydrochloride (4 μmol/L) or vehicle alone as indicated, and then processed by the University of Toronto Faculty of Medicine Microscopy Imaging Laboratory (Toronto, Ontario, Canada). Briefly, harvested cells were fixed with a Karnosky-style fixative [4% paraformaldehyde and 2.5% glutaraldehyde in a 0.1 mol/L Sorensen's phosphate buffer (pH 7.2)]. Cells were postfixed with 1% osmium tetroxide, dehydrated with ethanol, washed with propylene oxide, treated with epoxy resin polymerized at 60°C for 48 hours, sectioned on a Reichert Ultracut E microtome to 80 nm thickness, collected on 300 mesh copper grids, and counterstained with uranyl acetate and lead citrate. Analysis was done with a Hitachi H7000 transmission electron microscope (Hitachi, Tokyo, Japan) at an accelerating voltage of 75 kV.

As previously described, 1,1′,3,3,3′,3′-hexamethylindodicarbocyanine [DiIC₁(5); Invitrogen] was used to determine the mitochondrial membrane potential (ΔΨ_M), cell permeant indo-1 AM (Invitrogen) was used to determine changes in cytosolic calcium, and propidium iodide (Invitrogen) uptake was used to determine cell death (18). Briefly, FaDu cells were seeded (0.3 × 10⁶/T-25), incubated for 1 day, and then treated with alexidine dihydrochloride (4 μmol/L) or vehicle alone. At the indicated time points, all floating and adherent cells (using trypsin) were collected and resuspended in growth medium at a concentration of 10⁶/mL. DiIC₁(5) (40 nmol/L final concentration) and indo-1 AM (2 μmol/L final concentration) were added to the cell suspensions. The cells were incubated at 37°C for 25 minutes, after which propidium iodide (1 μg/mL) was added. The cells were analyzed by flow cytometry using a Coulter Epics Elite [Beckman Coulter; DiIC₁(5) excitation 633 nm, 675 ± 20 nm bandpass; indo-1 AM excitation 360 nm, emission ratio 405 nm/525 nm].

Cells were seeded (0.4 × 10⁶ per well in six-well plates), incubated for 1 day, and treated as indicated. Afterward, cells were stained using the CaspGLOW In situ Caspase Staining kits (BioVision, Mountain View, CA) for caspase-9, caspase-8, caspase-3, and caspase-2 activity according to the specifications of the manufacturer. Analysis was done using flow cytometry (FACSCalibur, CellQuest software, Becton Dickinson).

FaDu cells were seeded in six-well plates (Nalge Nunc International, Rochester, NY) at densities of 10² to 10⁴ per well in 3 mL growth medium and incubated overnight at 37°C to allow for cell attachment. Alexidine dihydrochloride or the vehicle control was added at the specified concentrations in a total volume of 50 μL. After 48 hours, the drug-containing medium was replaced with fresh growth medium and the plates were incubated at 37°C. Eleven days after seeding, colonies were fixed in 70% ethanol, stained with 10% methylene blue, and colonies of ≥50 cells were counted. The surviving fraction was determined as the ratio of the number of colonies in the treated sample to that of the nontreated control sample. Triplicate wells were set up for each condition and the experiment was done twice.

All animal experiments were conducted in accordance with the guidelines of the Animal Care Committee, Ontario Cancer Institute, University Health Network (Toronto, Ontario, Canada). In all cases, 6- to 8-week-old severe combined immunodeficient (SCID) BALB/c female mice were obtained from the Animal Research Colony, Ontario Cancer Institute.

FaDu cells were seeded (2 × 10⁶/T-75 flask), incubated for 1 day, and treated with either alexidine dihydrochloride (4 μmol/L) or vehicle alone. After 48 hours, the cells were harvested and implanted into the left gastrocnemius muscle of SCID mice (2.5 × 10⁵ cells in 100 μL growth medium per mouse). The mice were monitored for tumor formation at least thrice per week for 100 days. Three independent experiments were done, with three mice per group for each experiment.

All experiments were done thrice independently and all data are reported as mean ± SE unless otherwise stated.

We have previously used a cell-based, phenotype-driven, MTS-based, high-throughput screen of the LOPAC1280 (Sigma-Aldrich) and Prestwick Chemical Library to identify novel potential therapeutic agents for head and neck cancer (16). This preliminary screen identified alexidine dihydrochloride as a possible head and neck cancer–specific therapeutic (Fig. 1A). Thus, we did alexidine dihydrochloride dose-response evaluations on two cancer and three untransformed cell lines to confirm the preferential toxicity of the compound in head and neck cancer cells. The dose required to reduce cell viability by 50% (ED₅₀) was ∼1.8 μmol/L in FaDu (∼21 hour doubling time) and ∼2.6 μmol/L in C666-1 (∼48 hours doubling time) cells (Fig. 1B). In contrast, the ED₅₀ values were much higher in untransformed cell lines, specifically at ∼8.8 μmol/L in GM05757 (∼20-hour doubling time), ∼8.9 μmol/L in HNEpC (∼67-hour doubling time), and ∼19.6 μmol/L in NIH/3T3 cells (∼20-hour doubling time). A therapeutic window between malignant cells and nonmalignant cells thus exists with this compound.

To evaluate the effect of combining traditional head and neck cancer therapeutic agents with this novel anticancer compound, FaDu cells were simultaneously treated with a low dose of either cisplatin (1.9 μmol/L), 5-fluorouracil (5 μmol/L), or γ-radiation (10 Gy), along with increasing doses of alexidine dihydrochloride (Fig. 2A). In turn, FaDu cells were simultaneously incubated with a low dose of alexidine dihydrochloride (1.2 μmol/L) and with increasing doses of cisplatin or 5-fluorouracil (Fig. 2B and C). It is apparent from these dose-response curves that alexidine dihydrochloride does not interfere with the actions of cisplatin, 5-fluorouracil, or γ-radiation. The interactions are possibly additive at lower doses of the respective compounds, but seem to be less than additive at most doses.

In an effort to determine the mode of cell death induced by alexidine dihydrochloride, flow cytometric DNA content analyses was done on FaDu cells treated with 4 μmol/L alexidine dihydrochloride (the dose required to reduce cell viability by 75% after 48 hours, or ED₇₅). There was minimal change in G₁-G₀, S, or G₂-M phase (or aneuploid) populations at 12, 24, 48, or 72 hours (Fig. 3A, data not shown). However, in alexidine dihydrochloride–treated cells, the sub-G₁ population increased, indicative of apoptosis (Fig. 3A). Hoechst 33342 staining of alexidine dihydrochloride–treated (for 48 hours) cells revealed nuclear condensation and blebbing, consistent with apoptotic nuclear morphology (Fig. 3B). Necrosis was not observed. Transmission electron microscopy was used to further visualize the subcellular morphologic characteristics of apoptosis. Chromatin condensation and morphologic abnormalities in the mitochondria were observed (Fig. 3C). The rough endoplasmic reticulum, which can also induce apoptosis (19), did not seem to be affected.

To further investigate the mechanism of apoptosis in alexidine dihydrochloride-mediated cell death, ΔΨ_M depolarization and cytosolic Ca²⁺ increase (which may be due to damage to the endoplasmic reticulum or Ca²⁺ plasma membrane channels) were evaluated. FaDu cells treated (4 μmol/L) for 3, 12, or 24 hours were simultaneously stained with DiIc₁(5) (ΔΨ_M), indo-1 AM (Ca²⁺), and propidium iodide (membrane integrity/cell death; Fig. 3D). The proportion of cells with depolarized mitochondria increased as treatment time increased (Fig. 3D, column A). Increased cytosolic Ca²⁺, indicated by the indo-1 AM 405 nm/525 nm ratio, could be observed in cells with depolarized mitochondria. Propidium iodide uptake, indicating loss of membrane integrity and cell death, also increased with incubation time (Fig. 3D, column C). Notably, in alexidine dihydrochloride-treated FaDu cells, ΔΨ_M hyperpolarization could be observed, a phenomenon commonly observed during apoptosis (20, 21). The relative mean fluorescence readings in polarized cells after 3, 12, and 24 hours of treatment were 44.0, 83.5, and 130.1, respectively (37.8 at 24 hours with vehicle alone; Fig. 3D, columns B and D).

To evaluate the biochemical characteristics of alexidine dihydrochloride–induced apoptosis, caspase activation was evaluated in FaDu cells treated with 4 μmol/L of compound for 12, 24, or 48 hours (Fig. 3E). At 12 hours, caspase-2 activation was observed, along with slight activation of caspase-9. More extensive caspase-9 activation, combined with caspase-8 activation, were observed at 24 hours, with caspase-3 activation following at 48 hours.

To assay the effect of alexidine dihydrochloride on clonogenic survival, FaDu cells (10²–10⁴ per well) were seeded and then incubated with this compound for 48 hours. Nine days later, colonies were counted (Fig. 4A). Colony formation was reduced to <5% with 1 μmol/L alexidine dihydrochloride.

Next, FaDu cells were treated with 4 μmol/L alexidine dihydrochloride for 48 hours (ED₇₅) and then injected into the left gastrocnemius muscle of SCID mice (2.5 × 10⁵ per mouse) to determine whether this compound had the ability to prevent in vivo tumor formation. These mice did not develop tumors even after 100 days, whereas mice injected with 0.1% DMSO–treated FaDu cells started developing tumors after only 10 days (Fig. 4B). Thus, alexidine dihydrochloride effectively eliminates the tumor-forming potential of 2.5 × 10⁵ cells FaDu cells in SCID mice.

This current study, to our knowledge, is the first to evaluate the tumor-specific properties of alexidine dihydrochloride. There has been no known previous report of the anticancer efficacy of such bisbiguanide compounds. Alexidine dihydrochloride possesses anticancer efficacy in human hypopharyngeal squamous (FaDu; head and neck squamous cell carcinoma) and in human nasopharyngeal (C666-1; undifferentiated epithelial) cancers, with minimal toxicity to GM05757, HNEpC, or untransformed mouse embryonic fibroblasts. In vitro, this compound does not interfere with chemotherapy or radiation, and induces apoptosis. In vivo, this compound decreases the tumor-forming potential of FaDu cells.

Alexidine dihydrochloride has been previously used in mouthrinses (17). This compound is positively charged, and such cations are attracted to the negatively charged bacterial cell walls (17). In fact, gram-positive bacteria are more sensitive to cations because they are more negatively charged (17). Alexidine dihydrochloride induces lipid-phase separation and domain formation at bacterial membranes (17). Interestingly, mitochondria are thought to be evolutionarily derived from bacteria, and are one of the most negatively charged biological membranes (22, 23). Many cationic lipophilic toxins preferentially accumulate in the mitochondrial matrix of cancer cells, which generally have higher plasma and mitochondrial membrane potentials, as well as more mitochondria compared with normal epithelial cells (24, 25). With alexidine dihydrochloride treatment, we observed loss of ΔΨ_M after only 3 hours, caspase-9 activation after 12 hours, and transmission electron microscopy–visible mitochondrial alterations at 24 hours. Thus, damage to the mitochondria occurs early in alexidine dihydrochloride–induced death. Alexidine dihydrochloride–induced lipid-phase separation and domain formation potentially represents a novel anticancer therapeutic mechanism of action.

Caspase-2 was more active after 12 hours of treatment than the mitochondria-activated caspase-9. This may reflect either a differential sensitivity in the caspase activation assay, differing amounts of active caspase types necessary for cell death, or an alternate alexidine dihydrochloride target. Caspase-2 has been controversially implicated in genotoxic stress, endoplasmic reticulum stress, death receptor ligation, and in heat shock–induced apoptosis (26). Endoplasmic reticulum damage was not visible in our transmission electron microscopy studies, although further studies will be required to identify other potential targets of alexidine dihydrochloride.

Head and neck cancers are treated with traditional therapeutic modalities, including surgery, radiation, cisplatin, and/or 5-fluorouracil (27). Alexidine dihydrochloride did not interfere with the action of cisplatin, 5-fluorouracil, or radiation, but the interaction seemed to be less than additive. Clonogenic-based assays, a more complete range of dose-response curves, and further analyses will be required to fully elucidate the efficacy of the combinations in vitro (28). Further studies will also be required to evaluate the efficacy of combining alexidine dihydrochloride with other therapeutics. The observation that alexidine dihydrochloride led to caspase-8 activation (after 24-hour treatment) and apoptosis in FaDu cells, a cell line known to be resistant to tumor necrosis factor–related apoptosis-inducing ligand therapy due to a homozygous deletion in the death receptor DR4 gene (preventing caspase-8 activation; ref. 29), suggests that an alexidine dihydrochloride–tumor necrosis factor–related apoptosis-inducing ligand combination might have significant therapeutic potential.

FaDu cells represent a highly aggressive cell line in vivo. These cell lines form tumors after only 10 days with the injection of 2.5 × 10⁵ cells, and increase tumor-plus-leg diameter by ∼0.45 mm/d. Merely 4 μmol/L alexidine dihydrochloride was able to eliminate the tumor-forming ability of 2.5 × 10⁵ cells in SCID mice. Using this compound as a drug lead, future studies should focus on enhancing its solubility and bioavailability for further in vivo treatment and safety studies.

Alexidine dihydrochloride is chemically similar to chlorhexidine, another bisbiguanide compound. Chlorhexidine possesses chlorophenol end groups, as opposed to the ethylhexyl end groups in alexidine (17). Chlorhexidine affects membrane integrity at low concentrations, and causes the cytoplasm to congeal at higher concentrations (17). Cellular leakage induced by this compound is not responsible for cellular inactivation, but is a consequence of cell death (17). The difference in end-groups between these two compounds is thought to be responsible for the more rapid bactericidal action of alexidine dihydrochloride, as well as the ability of this biguanide to produce lipid domains (30, 31). Evaluating the anticancer potential of chlorhexidine, other bisbiguanide compounds and alexidine analogues will provide for a more complete structure-activity relation profile.

In conclusion, our study has identified alexidine dihydrochloride as a novel anticancer drug lead. This compound, as well as other compounds from the bisbiguanide family and alexidine analogues, warrant further study.

We thank Melania Pintilie for her statistical advice.