Mechanochemical Disruption Suppresses Metastatic Phenotype and Pushes Prostate Cancer Cells toward Apoptosis

The current focus of clinical oncology is on advanced chemical-based medicine that targets specific oncogenes or individual proteins involved in tumor-promoting pathways (1). This approach is often ineffective due to tumor heterogeneity and dormancy (2), and it may result in the unintended development of more aggressive cancer phenotypes with acquired resistance (3). A possible way to overcome this barrier is to combine chemical treatment with therapies that cause the phenotype-independent mechanical disruption of cancer cells. The perturbations of cellular structure and functions induced by such a disruptive therapy may diminish the ability of cancer cells to resist chemical treatment and may potentially lead to a less aggressive cancer phenotype (4).

Mechanical forces are capable of inducing or modulating intracellular signaling associated with gene regulation, protein folding, and the structure and stability of the cell cytoskeleton and membranes (5–8). The mechanotransduction events lead to changes in cell function, phenotype, and viability that strongly depend on the type of mechanical cues the cells are exposed to. For example, mechanical compression of the extracellular matrix during tumor growth makes cancer cells more invasive (9), while tumor exposure to oscillatory forces (vibration) causes cell apoptosis and necrosis in vitro (10, 11).

The focus of this work is on focused ultrasound (FUS), a unique method to noninvasively deliver mechanical (acoustic) energy deep into the tissues in vivo, without exposing the body to harmful ionizing radiation. The acoustic energy deposited locally to a tumor by the FUS beam causes a vibration of molecules inside the tumor, which leads to heating and mechanical disruption of cancer cells and tumor microenvironment (12–14). The ability of standalone FUS to induce tissue hyperthermia is well established, and, at high intensity exceeding 1 kW/cm² (referred to as HIFU), it is used clinically for noninvasive thermal ablation of small and localized tumor masses in the prostate (15, 16). However, the possibility for FUS to induce phenotypic changes in cancer cells through mechanical disruption has not been explored. We hypothesize that the FUS-induced cellular disruption, in presence of chemically induced exogenous stress, prevents uncontrolled proliferation and decreases aggressiveness of cancer cells. Utilizing ethanol, a chemical agent with well understood anticancer effects, we investigate tumor growth and progression in a mouse xenograft model of human prostate cancer and cytotoxic and phenotypic changes in metastatic prostate cancer cells, resulting from this mechanochemical disruption approach.

Primary human umbilical vein endothelial cells (HUVEC) were obtained from ATCC. Cells were cultured in T-75 flasks with Medium 200 (Thermo Fisher Scientific) containing low serum growth supplement (Thermo Fisher Scientific), 10 μg/mL gentamicin, and 0.25 μg/mL amphotericin B (Thermo Fisher Scientific). DU-145, PC-3, and C4-2B prostate cancer cells were obtained from ATCC and grown in high-glucose DMEM (Thermo Fisher Scientific) supplemented with 10% FBS (Thermo Fisher Scientific) and 1% penicillin/streptomycin (Thermo Fisher Scientific). Both DU-145 and PC3 cell lines are androgen receptor (AR) null and represent in vitro models of fully metastatic castration-resistant prostate cancer (CRPC). They were originally isolated from brain and bone metastases, respectively (17). The C4-2B cell line is AR-positive, but it is androgen independent due to castration during cell line development (17). It is known to metastasize to bones in vivo. All cells were cultured in either T-75 or T-175 flasks and maintained in a 37°C, 5% CO₂ incubator.

A 1.1-MHz single-element, concave transducer H102 (Sonic Concepts) with bandwidth from 0.748 to 1.380 MHz, geometric focal length of 63.2 mm, transverse focal width of approximately 1.5 mm, and axial focal length of approximately 8 mm was used in all experiments. The transducer has a stainless steel housing with active diameter of 64 mm. The transducer was coupled to a cone containing degassed water heated to 37°C. In animal experiments, ultrasound gel (Aqua Sonic 100, Parker Labs) was used as a coupling medium between the cone and the skin. A 33220A function generator (Agilent Technology) produced an input sinusoidal signal that passed through a fixed gain (50 dB) ENL 2100L power amplifier (Electronics & Innovation) and then entered the transducer. The FUS signal strength was monitored using a 2 Giga-samples/s InfiniiVision DSO-X-2014A oscilloscope (Agilent Technology). Temperature near a tumor sample was measured during FUS targeting by a mini-hypodermic Copper Constantan type T 200-μm thick bare-wired thermocouple (Omega Engineering) connected to a temperature meter (SDL200, Extech Instruments). FUS was operated in a continuous mode and its exposure time was 30 seconds. Supplementary Figure S16 shows the schematic of the entire FUS setup.

The following treatment groups were considered: (i) untreated cancer cells (control); (ii) 4% ethanol only; (iii) FUS only; and (iv) 4% ethanol + FUS. Prior to experiment, prostate cancer cells were trypsinized and suspended in fresh growth medium. One-hundred microliters of the medium containing 2.7 × 10⁶ cells was resuspended in a thin-wall 0.2 mL centrifuge tube (Bio-Rad) and then centrifuged at 2,000 rpm for 2 minutes to form a dense pellet of cells. In ethanol treatment groups, the growth medium that the cells were centrifuged in contained 4% ethanol. In FUS treatment groups, pellets were positioned within the focus of the ultrasound beam and then exposed to FUS at acoustic output power of 4.1, 8.7, or 12 W (level H3, H4, or H5), which corresponds to the spatial peak temporal average intensity I_SPTA of 0.38, 0.70, or 0.88 kW/cm², respectively. It should be noted that the highest intensity of FUS used in our experiments was nearly twice less than the lower limit of transrectal HIFU intensity (1.3 kW/cm²), required for prostate cancer ablation (18).

Treated cancer cells were re-cultured in 35 × 10 mm² culture dishes. The number of viable prostate cancer cells were counted at 2, 24, and 72 hours post-treatment using Trypan blue exclusion. Viable, early apoptotic, and late apoptotic/necrotic populations of cancer cells were determined at these time points by flow cytometry employing an Annexin V-FITC Apoptosis Detection Kit (Affymetrix eBioscience). In the latter test, cancer cells in each treated group were first washed by ice-cold PBS and then by × 1 binding buffer. Cell samples were then incubated with 5 μL Annexin V in 195 μL cell suspension at room temperature for 15 minutes and washed with the binding buffer twice. Immediately prior to flow cytometry, 10 μL of propidium iodide (20 μg/mL) was added to the cell suspension. A total of 100,000 events, excluding aggregates and particulates, in the forward- and side-scatter gates were collected using the Attune Acoustic Focusing Cytometer (Applied Biosystems). The Annexin V-FITC Apoptosis Detection Kit stains for apoptotic and necrotic cells through green Annexin V and red propidium iodide, respectively. Early apoptotic cells were identified as propidium iodide–negative and Annexin V–positive, whereas the cells identified as propidium iodide–positive and Annexin V–positive were considered late apoptotic/necrotic.

Cell proliferation was measured using the WST-8 Cell Proliferation Kit (Cayman Chemical). A total of 1.0 × 10⁵ treated cells were plated with 0.1 mL of full growth medium in a 96-well flat-bottom plate (Thermo Fischer Scientific) and incubated for 24 or 72 hours. Then, the cells were exposed to 10 μL of WST-8 reagent reconstituted in Electron Mediator Solution for 2 hours. The plate was then gently mixed for one minute and absorbance of light with a wavelength of 540 nm was measured using a microplate reader (ELx808, BioTek Instruments). Cells were also cultured in 35-mm petri dishes for up to 14 days post-treatment. The growth medium was changed daily and 10 images per sample were taken at 4× magnification to measure the number of adherent cells and assess the cell growth rate and proliferative potential. The average number of cells per image was plotted for different treatment groups (Supplementary Fig. S4).

FITC-conjugated mouse IgG1 and IgG2a and mouse anti-human antibodies to membrane proteins CD29 (MEM-101A), CD44 (MEM-85), CD49d (R1-2), CD95 (DX2), and CD178 (MFL3) were purchased from Thermo Fisher Scientific. The expression level of these proteins was analyzed by flow cytometry. Cancer cells in each group were first washed by ice-cold PBS and washed once by FACS buffer (2% BSA and 0.1% sodium azide in PBS) before adding antibodies. Mouse IgG isotype control and anti-human antibodies were added to cancer cells. After cells were incubated with antibodies on ice for 45 minutes, they were washed by FACS buffer and resuspended in buffer containing 2% formaldehyde. Flow cytometric analysis was conducted at 2, 24, and 72 hours post-treatment using the Attune Acoustic Focusing Cytometer. CD29 is also known as β₁ integrin, CD44 is HCAM, CD49d is α₄ integrin, CD95 is Fas receptor, and CD178 is Fas ligand.

The cell pellet was resuspended immediately after treatment in the culture medium and a drop of the cell suspension was plunged in liquid nitrogen to enable vitrification. The cross-section images of the cells were obtained by fracturing them at −130°C using a flat edge knife. The samples were sublimed at −95°C for 5 minutes to remove surface-vitrified water and expose surface morphology, and then sputtered with a gold−palladium composite at 11 mA for 88 s. Cryo-SEM imaging was done at a voltage of 3 kV and a working distance of approximately 8 mm. ImageJ was utilized to measure diameter of membrane pores after FUS exposure. A schematic of this procedure is shown in Supplementary Fig. S17. A minimum of 100 pores across 50 cells were averaged to determine the pore formation post-treatment.

HUVEC of passage 3 or 4 at a concentration of 0.3 × 10⁶ cells/mL were seeded in each well of a 96-well plate and cultured overnight to ensure confluence. The cells were then activated by 10 ng/mL TNFα (Sigma Aldrich) for 4 hours. Following the HUVEC activation the medium was removed from the wells and replaced with DMEM containing 2.0 × 10⁴ viable DiO-labeled cancer cells per experimental group. After 15-minute incubation, the media in wells were carefully aspirated. The wells were washed three times with PBS to eliminate nonfirmly adherent cancer cells. The stained cancer cells were visualized using an inverted epi-fluorescent microscope (Nikon Eclipse Ti-S) with a 10× objective. Images of adherent cancer cells were captured by a digital CCD camera (Qimaging Retiga EXi) at five different areas of each well. The recorded images were processed by a custom MATLAB (Mathworks) cell counting code to determine the number of firmly adherent cells. The image field size was 904 μm × 675 μm. The static adhesion tests were conducted at 2, 24, and 72 hours post-treatment.

Treated prostate cancer cells were seeded into wells of a 12-well plate at a density of 0.6 × 10⁶ cells/well. They formed a confluent monolayer in 24 hours. At this time point, the cell monolayer was scratched with a 200-μL pipette tip across the center of the well and then washed twice with PBS to remove unadhered cells. Fresh growth medium was added to the cells after the washing procedure, and images of the scratch region (“wound”) were taken daily for up to 5 days. The wound width (distance between edges of the scratched region) was measured by ImageJ.

In this assay, cancer cells were exposed to FUS at power of 1.2 to 2.7 W (level H2) to ensure that FUS has no effect on cell proliferation (Supplementary Fig. S9C). The cell viability assay was conducted prior to each scratch-wound experiment to confirm that none of the treatments led to a greater cell death.

Immunoblotting was performed to detect the intracellular expression of NFκB (total and nuclear) and Hsp70 in treated cancer cells. The total protein was extracted from the cells by using radioimmunoprecipitation (RIPA) lysis buffer (Thermo Fisher Scientific) or Nuclear Extraction Kit ((Cayman Chemical). Equal amounts of protein (50 μg/lane) were resolved by 8%–12% Nu-page gel (Thermo Fisher Scientific) and electrophoretically transferred to Immobilon-P membrane (EMD Milipore). The membrane was incubated with mouse anti-human NFκB (A-12) and Hsp70 (3A3) primary antibodies (Santa Cruz Biotechnology) at 1:5,000 dilution in 5% v/w BSA, 1 × Tris buffered saline (TBS, Thermo Fisher Scientific), and 0.1% Tween 20 (Thermo Fisher Scientific) at 4°C overnight. The solution was then removed and the membrane was washed with TBS and Tween 20, and then incubated with HRP-conjugated goat anti-mouse secondary antibody (LI-COR Biosciences) at 1:10,000 dilution in the BSA/TBS/Tween 20 solution in a dark room at room temperature for 2 hours. Glyceraldehyde 3-phospahate dehydrogenase (GAPDH, Thermo Fisher Scientific) or Histone H3 (1G1, Santa Cruz Biotechnology) was used as a loading control to ensure equal amounts of total added proteins. The molecular weight of proteins was estimated using the PageRuler Prestained Protein ladder (Thermo Fisher Scientific). Protein band images acquired by Odyssey Infrared Imager (LI-COR Biosciences) were then analyzed by Image Studio software (LI-COR Biosciences) to determine the normalized-to-GAPDH or normalized-to-Histone H3 expression levels of proteins.

ROS expression was measured via a chloromethyl (CM) derivative of H2DCFDA (Thermo Fisher Scientific). A total of 100 μmol/L of the CM-H2DCFDA was added and incubated with the cells 2 hours before treatment, and recultured using complete growth medium with 100 μmol/L of CM-H2DCFDA for 24 and 72 hours post-treatment. Cancer cells in each group were first washed by ice-cold PBS and flow cytometric analysis was then conducted using the Attune Acoustic Focusing Cytometer. Samples were excited at 495 nm and emission was observed at 520 nm.

Cancer cells were incubated with 10 μg/mL mouse anti-human TNFR1 mAb (H398, Thermo Fisher Scientific), 10 μg/mL mouse anti-human FasR mAb (ANT-205, Prospec-Tany TechnoGene), or their combination at 37°C for 2 hours prior to treatment to block death receptors TNFR1 and FasR. Blocking experiments were conducted for the following 6 groups: control, 4% EtOH, FUS level H4, 4% EtOH + FUS level H4, FUS level H5, and 4% EtOH + FUS level H5. Treated cancer cells were recultured in 12-well plates with 2 μg/mL TNFR1 antibody, 2 μg/mL FasR antibody, or their combination. Viable, early apoptotic, and late apoptotic/necrotic populations of cancer cells were determined at 24 and 72 hours post-treatment by flow cytometry using an Annexin V-FITC Apoptosis Detection Kit (Thermo Fisher Scientific).

Treated cancer cells were cultured in hanging drops using Perfecta3D Hanging Drop Plates (3DBiomatrix) according to manufacturer's instructions. Cells were seeded into the wells at density of 3.0 × 10⁴ cells/mL and imaged over the course of 3 days. Sample images of multicellular spheroid cultures formed from treated cells were taken at 24 and 48 hours post-seeding. The tumor-forming ability of DU-145 and PC3 cells were blindly scored by five independent scorers as follows: 2 equates to a tightly packed spheroid, 1 is a loosely packed spheroid, and 0 is no spheroid formed.

All procedures using animals were approved by Tulane University's Institutional Animal Care and Use Committee (IACUC).

Four-week-old male Nu/Nu athymic nude mice (J:NU 007850, Jackson Laboratories) were purchased and allowed to acclimate for one week. Tumors were established by subcutaneous injection of 1.0 × 10⁶ DU145 cancer cells using a 28-gauge needle. A 200-μL bolus containing cells in PBS (Thermo Fisher Scientific) mixed in a 1:1 volume ratio with Matrigel Matrix High Concentration (Corning) was injected on left and right flanks of each animal. Tumor dimensions were measured daily using a digital caliper and a diagnostic ultrasound imaging system (180 plus, SonoSite). Tumor volumes were calculated using the formula V = π/6 × 2a × b, where a is the short axis and b the long axis of the tumor. When the volumes reached approximately 200 mm³ (roughly 3 weeks post-injection), the mice were randomly assigned to groups and treatments commenced. A total of 10 tumors per group were tested.

Before treatment, mice were anesthetized using isoflurane gas (Vet One) and constrained in a custom-designed holder that allows for easy access to tumors for treatment and imaging (see Supplementary Fig. S18). Mice in the sham group were injected with 50 μL PBS. Mice in ethanol treatment groups were injected with 50 μL 99% ethanol (25% or less of tumor volume). Using a 3D positioning system (Thorlabs) and diagnostic ultrasound, the focus of the FUS transducer was aligned with the tumor in all FUS treatment groups. The tumors were scanned progressively (point-by-point) by the FUS beam at level H5. Four to five FUS shots at 30-second increments were used to ablate tumors. In the combination treatment group, ethanol was injected immediately prior to FUS exposure. The total procedure time was less than 5 minutes per mouse. The tumor volume was measured with a digital caliper as well as diagnostic ultrasound every day for 14 days post-treatment. No further treatment was given.

Mice were sacrificed at day 5 or day 14 via CO₂ asphyxiation for tumor tissue collection. Collected tissue specimens were fixed in formalin for 24 hours and embedded into paraffin. Paraffin-embedded tissues were sectioned into 4-μm thick slices, placed on glass slides, and subsequently stained with hematoxylin and eosin (H&E). Control and treatment arm slides were randomly mixed and blindly evaluated by the study pathologist (A.B. Sholl). Each slide was evaluated for maximal tumor diameter and for percent necrosis.

The results were evaluated with one- or two-way ANOVA using GraphPad Prism version 5.0.2 (GraphPad Software). Statistically significant differences were set to P < 0.05 between experimental groups. The statistical data were represented as mean ± SEM. The number of independent experiments is listed in each figure.

Figure 1 shows changes in the viability, proliferative potential, and the proapoptotic activity of death receptors in AR-negative (DU-145) and AR-positive (C4-2B) CRPC cell lines exposed to ethanol, FUS, or their combination. Three power levels of FUS (H3: 4.1 W, H4: 8.7 W, and H5: 12.0 W) were selected to ensure that at least 20% of cells remain viable immediately after FUS alone treatment (Supplementary Fig. S1A). For percent (v/v) ethanol at which more than 90% of cells are viable was utilized in all in vitro experiments (Supplementary Fig. S1B). The mean viabilities of DU-145 and C4-2B cells exposed to FUS alone treatment were, respectively, 82% and 72% at H3, 26% and 25% at H4, and 13% and 4% at H5 at 72 hours post-treatment, as measured by a Trypan blue exclusion test (Fig. 1A). When FUS was combined with ethanol, the majority of cancer cells were killed immediately. Specifically, the DU-145/C4-2B cell viabilities in the E+H3, E+H4, and E+H5 treatment groups were 53%/49%, 20%/6%, and 10%/0.3% at 2 hours post treatment and further decreased to 25%/19%, 7%/0.3%, and 0%/0% at 72 hours, respectively (Fig. 1A and B). The change in viability between the individual and combined treatments was statistically significant for both cell lines and all FUS power levels tested (P < 0.001). A similar effect was also observed for PC3 cells (Supplementary Fig. S2A). These data point out that some of the CRPC cells remain viable after exposure to FUS alone, especially if they are AR-negative. However, when FUS is combined with low-concentration ethanol, the percentage of viable cells significantly decreases, reaching zero at level H5 for both AR-positive and AR-negative CRPC.

To identify the stage of cell damage post-treatment, flow cytometry for Annexin V and PI was conducted on DU-145 cells (Fig. 1C and D) and PC3 and C4-2B cells (Supplementary Fig. S3). With treatment, the cell population shifted from the bottom left corner (viable cells) to the bottom right corner (early apoptotic cells) and to the top right and left corners (late apoptotic/necrotic cells). As seen in the two bottom plots of Fig. 1B, there was an increase in the density of early apoptotic cells and late apoptotic/necrotic cells after treatment with FUS alone (H4) or a combination of ethanol and FUS (E+H4), as compared with control (top left). Combining H4 with ethanol led to a drastic shift of the cell population to the late apoptotic/necrotic region (Fig. 1B, bottom right). The total population percentage of apoptotic/necrotic cells increased as time post-treatment progressed (Fig. 1C). This percentage in E+H groups at 2, 24, and 72 hours statistically increased as compared with control (P < 0.01 or 0.001) and was significantly higher than FUS alone (P < 0.01) at 24 and 72 hours for all FUS powers used. For example, there was a large difference in the population of late apoptotic/necrotic cells between the E+H4 and control groups at 72 hours (66.2% ± 1.9% vs. 11.4% ± 1.7%, P < 0.01). The percent of the apoptotic/necrotic cells in the E+H5 group was 78.3% ± 2.0% at 72 hours, while in the H5 alone group it was decreased to 52.7% ± 0.8% (P < 0.01). Similarly, in the E+H4 group, the percent of apoptotic/necrotic cells at 72 hours was 66.2% ± 1.9%, while only 28.8% ± 9.9% in the H4 alone group (P < 0.01). Overall, the flow cytometric data clearly show that ethanol in combination with FUS achieves a much greater increase in cell death than the simple addition of treatment arms.

To test whether the E+H combination influences cell proliferation, a WST-8 assay was performed (Fig. 1D). As seen in this figure and Supplementary Fig. S2B, the proliferation rate of DU-145, C4-2B, and PC3 cells in FUS alone (H4) and ethanol with FUS (E+H4) groups significantly (P < 0.01 or 0.001) decreased after 24 and 72 hours as compared with the control group. Furthermore, C4-2B and PC3 cells in the E+H4 group had a significantly (P < 0.05) lower proliferative potential at both 24 and 72 hours as compared with the FUS alone group. For DU-145 cells, a significant (P < 0.05) difference between those two groups reached at 72 hours. Thus, ethanol in combination with FUS leads to the greatest reduction in cell proliferation among all treatment arms tested.

In efforts to understand the long-term growth behavior of DU-145 cells post-treatment, surviving cells were cultured with the same seeding density over 14 days (Fig. 1E; Supplementary Fig. S4). Control- and ethanol-treated cells reached confluency in 4 days. Their doubling rates were similar (∼45 hours). H4 alone–treated cells had a drastically reduced proliferation rate until day 7, but they rebounded during the second week of culture and reached confluency at 14 days (doubling rate of 69 hours). On the other hand, E+H4-treated cells did not proliferate for the full 2 weeks. Thus, surviving cells exposed to ethanol + FUS mechanochemical disruption lost their ability to grow in vitro.

Fas and TNFR1 play a key role in programmed death of healthy human prostate cells (19), but the expression of these death receptors is downregulated in prostate cancer cells (20). The upstream TNFR signaling is associated with apoptotic c-Jun N-terminal kinase (JNK) activation as well as the intrinsic (mitochondrial) and extrinsic apoptotic pathways (21). The extrinsic apoptosis pathway involves the TRADD–RIP–FADD complex (TNFR1-associated death domain, Receptor-interacting protein, Fas-associated death domain), which is activated by both TNFR1 and FasR. Prostate cancer cells protect themselves from this pathway by downregulating their expression level of FasR, which is responsible for FADD recruitment. The first two panels in Fig. 1F show flow cytometric data on the expression of FasR and its ligand FasL in DU-145 prostate cancer cells. The ligation of FasR is necessary to initiate apoptosis via this death receptor. A significant increase in FasR expression (with respect to IgG) was only seen in E+H treatment groups beginning at 24 hours post-treatment. For example, FasR had an approximately 3 times higher expression in the E+H5 group than that in the control group at 72 hours (P < 0.001). A small increase in expression, not exceeding 1.5-fold, was observed for the H4 and H5 alone groups at 72 hours post-treatment. The expression of FasL was also dramatically increased post E+H treatment as opposed to control at 2, 24, and 72 hours (P < 0.01 or 0.001). The expression of FasL in E+H5 was approximately 4 times higher than that seen in the control group at 72 hours (P < 0.001). The FasL expression for FUS alone was not significant at 2 hours but then significantly increased to approximately 3-fold at 24 and 72 hours (P < 0.05). Because the expression of both FasR and FasL increases in the E+H treatment groups, it can be concluded that the cells in these groups are more prone to programmed cell death, which was also evident in the data of the cell viability assays (Fig. 1A–C). Although the cells in the FUS alone groups showed increased production of FasL, they were less likely to undergo apoptosis due to their insufficient expression of FasR. The third panel of Fig. 1F displays flow cytometric data of TNFR1 expression in DU-145 prostate cancer cells. The TNFR1 expression increased significantly in cells treated with FUS alone or E+H (P < 0.001), as compared with control and ethanol groups at 24 and 72 hours. At 24 hours, it was approximately 10-fold greater in the E+H5 group than in the control group, and twice the expression in the H5 group. Thus, mechanochemical disruption activates multiple death receptors with different time profiles of expression.

A death receptor blocking experiment was conducted to examine specific roles of TNFR1 and FasR in cell apoptosis induced by mechanochemical disruption. DU-145 cells were incubated with anti-human TNFR1 mAb H398, anti-human FasR mAb ANT-205, or their combination at 37°C pre-and posttreatment. Figure 1G shows the effects of single and double receptor blocking on viability of treated and control cells at 24 and 72 hours. Cell viability significantly (P < 0.05) increased with single blocking of FasR or TNFR1 in cells treated with ethanol and FUS. For example, the viability of cells in the E+H5 group at 72 hours post-treatment increased from 11.5% ± 2.8% to 45.2% ± 5.5% with FasR blocking. Double blocking of TNFR1 and FasR led to a consistently higher viability of treated cells at different time points than single blocking did. This confirms that both TNFR1 and FasR are involved in apoptosis of cancer cells exposed to mechanochemical disruption. Interestingly, the contribution of these receptors to cell apoptosis was both time and FUS power dependent. Blocking of TNRF1 at 24 hours, when this receptor reached its peak expression (Fig. 1F, right), significantly (P < 0.05) increased cell viability in the H4 and E+H4 groups, as compared with blocking of FasR (Fig. 1G, left). FasR became as effective as TNFR1 at 72 hours (Fig. 1G, right), when its expression reached the peak value (Fig. 1F, left). At the acoustic power level H4, TNFR1 played a major role in apoptosis of cells treated with FUS or its combination with ethanol, while an increase in acoustic power to level H5 made FasR a dominant death receptor for cells in these groups at 72 hours (Fig. 1G, right). It is important to note that inhibiting death receptor activity cannot fully return cell viability to that of the control group, because a certain percentage of cells exposed to FUS is necrotized at time of treatment instead of undergoing programmed apoptosis. The other interesting fact is the apoptotic activity of TNFR1 in DU-145 cells disrupted by ethanol and FUS does not require TNFR1 ligation by TNFα. As seen in Supplementary Fig. S5, there was no measurable level of TNFα produced by treated cells, yet our blocking study confirms that TNRF1 participates in cell apoptosis. Overall, the activation of TNF- and Fas-mediated apoptotic pathways by mechanochemical disruption cumulatively increases the rate of cancer cell killing and reduces their rate of proliferation.

The images of prostate cancer cells obtained by a cryogenic scanning electron microscope (Cryo-SEM) directly demonstrate the physical destruction of the plasma membrane in the cells exposed to ethanol, FUS, and their combination (Fig. 2A). The membrane of control cells was completely intact. Only small and sparse pores with an average diameter of 50 ± 13 nm were visible in the cells exposed to ethanol alone. FUS treatment (H4) led to larger pores (average diameter of 192 ± 87 nm) that spanned the majority of the cell surface. Substantial destruction to both the plasma membrane and the cytoskeleton, with no clear boundaries between pores, was observed in the cells exposed to the combination treatment. Further representative images as well as evidence for cytoskeletal damage are given in Supplementary Fig. S6. The large cavities are evidence of unrepairable mechanical destruction, which is a cause for the large population of necrotic cells seen in the combination treatment groups (Fig. 1B and C).

Intracellular signaling induced by ligation of transmembrane protein TNFR1, for example, by TNFα, can induce cell proliferation and increase the expression of cell adhesion molecules, but TNFR1 also acts as a death receptor. It is unclear a priori whether the increased expression of TNFR1 after mechanochemical disruption (Fig. 1F) helps cancer cells survive and increase their adhesiveness or it shifts the cells to apoptosis. As seen in Fig. 1, treatment groups in which cells have a high expression of TNFR1 also have an increased population of apoptotic/necrotic cells and a reduced proliferation rate. Figure 2B points out that despite the increased number of TNFR1 molecules, mechanochemical disruption reduces the ability of prostate cancer cells to adhere to vascular endothelium, which is a critical step in cancer metastasis. In this static adhesion assay, DiO-labeled cancer cells at different time points post-treatment were applied to a confluent monolayer of TNFα-activated endothelial cells (HUVEC). Treatment of DU-145 or PC3 cells with ethanol, FUS, or their combination significantly reduced the ability of these cells to attach to vascular endothelium (Fig. 2C; Supplementary Fig. S7). However, this effect was transient for FUS alone, for example, no significant difference in cell adhesiveness was observed between the H4 group and control at 72 hours. The least number of attached cells at 2, 24, or 72 hours was in the combination treatment (E+H4) group (P < 0.01 or 0.001 when compared with control).

Integrin β₁ (CD29) as well as homing cell adhesion molecule (HCAM or CD44) are well known metastatic markers of prostate cancer. These receptors mediate cell adhesion, migration, stem-ness, and overall aggressiveness of prostate cancer (22). In the healthy prostate phenotype, they are expressed at much lower levels. Using flow cytometry we measured the expression of these molecules in treated cancer cells. The mechanochemical disruption (E+H4 and E+H5) dramatically lowered the expression of CD29 and CD44, and the cells exposed to this treatment were able to maintain low levels of those proteins over the course of three days, as opposed to single treatment or control (Fig. 2D; Supplementary Fig. S8). The normalization of prostate cancer cells toward the healthier phenotype after mechanochemical disruption is further demonstrated by an increased expression of α₄ integrin (Fig. 2D), which is known to be downregulated in prostate cancer (23).

To test whether mechanochemical disruption reduces the migratory ability of aggressive cancer cells, we performed a scratch-wound assay with a confluent monolayer of treated or untreated DU-145 cells (Fig. 2E). Treating cells with FUS at nondestructive level H2 led to a significant defect in cell migration; specifically, in the ability of the cells to close the scratch area within 3 days of post-scratch culture (P < 0.05 between H2 and control, P < 0.01 between E+H2 and control). Four days post-scratch, all wounds closed, except for cells treated with a combination of ethanol and FUS (P < 0.001 vs. control, Supplementary Fig. S9B).

Cancer cells are known for their ability to adhere to each other and form uncontrollable tumor colonies. The inhibition of this tumorigenic potential is critical for effective treatment of cancer. A three-dimensional tumor spheroid model was utilized to investigate the tumorigenic potential of DU-145 and PC3 cells exposed to various treatment arms. One day post-treatment, 2.5 × 10⁵ viable cells were placed within each well of a perfect 3D spheroid culture plate. They were cultured for two days, while being suspended in growth medium droplets to allow for multicellular tumor spheroid formation (DU-145, Fig. 3A; PC3, Supplementary Fig. S10A; C4-2B, Supplementary Fig. S11). From this experiment three possible outcomes were identified: (i) tight spheroid (compact and dense cluster of cells), (ii) loose spheroid (less compact and more dispersed aggregate), and (iii) no spheroid. The cells in the control and ethanol only groups were more likely to form tight spheroids, roughly 500–600 μm in diameter (Fig. 3A). The cells in the FUS only groups (H4 and H5) preferentially formed loose spheroids. The cells in the combination treatment groups (E+H4 and E+H5) were characterized either by loose spheroid formation or did not from spheroids at all. These observations were quantified by assigning a score of 0 for no spheroid, a score of 1 for a loose spheroid, or a score of 2 for a tight spheroid. The ethanol + FUS combination is the only treatment arm for which on average no spheroid formation can be observed after 2 days of culture for both DU-145 and PC3 cancer cells (P < 0.01 or 0.001; Fig. 3B; Supplementary Fig. S10B).

The in vitro data we reported so far indicate that prostate cancer cells experience greater death and reduced tumorigenic potential upon exposure to a combination of ethanol and FUS. We anticipate that this combination treatment would have a similar destructive effect on tumors in vivo. To test this hypothesis, athymic nude mice bearing established DU-145 subcutaneous tumor xenografts were treated with sham, one time injection of 50 μL of 70% ethanol, FUS (level H5), or ethanol injection immediately followed by FUS. Procedures involved in treatment of tumor xenografts are illustrated in Supplementary Fig. S12. Animals were observed for 14 days post-treatment and representative images for each treatment arm at 1, 5, and 14 days can be seen in the top panel of Fig. 4A. In the sham and ethanol alone–treated xenografts, rapid tumor growth can be observed as they near 14 days post-treatment (first and second columns in top panel of Fig. 4A). Deep discoloration as well as hematomas can be observed in the tumors exposed to ethanol alone (second column). The tumors exposed to FUS alone showed obvious scarring from the initial ablation, and they continued to enlarge in the areas surrounding the ablated region (third column, see also Supplementary Fig. S13). The representative images of tumors in the combination treatment group show tumor regression from a volume of approximately 260 mm³ one day post-treatment to a non-caliper–measurable tumor at 14 days (fourth column). The sham, ethanol alone, and FUS alone treatments were incapable of inhibiting tumor growth over a two-week period based on diagnostic ultrasound and caliper measurements (Fig. 4B). Only the ethanol + FUS combination (E+H5) showed significant and consistent reduction in relative tumor size starting at 5 days post-treatment, reaching, on average, one-third of the original pretreated size at 14 days (Fig. 4B).

Mice were euthanized at 5 and 14 days post-treatment, and tissues were stained with H&E for histologic examination. Standard sections of H&E-stained tumors were blindly analyzed and scored for % viability and % necrosis by the study pathologist. H&E stains for the representative tumors at 14 days post-treatment are shown for the different treatment arms in the middle and bottom panels of Fig. 4A at 1× and 20× magnification, respectively. These stains illustrate that nearly 99% of the tumor tissue was viable in the sham and ethanol alone groups. Tumors exposed to FUS alone were also mostly viable but had small regions of necrosis highlighted in the 20× H&E stain (third column in bottom panel of Fig. 4A). The ethanol + FUS combination led to necrotic tumors (fourth column) containing clearly visible ablation regions void of cells at 1× magnification (middle). The 20× magnification image on the bottom panel highlights that the tissue was necrotic even in the regions where the cancer cells were not fully lysed. Supplementary Figure S14 provides further images of stained tumor tissues for each of the treatment arms. Statistical data plotted in Fig. 4C and D indicate that the combination treatment group had a significantly higher percent necrosis (54% ± 17% at 5 days and 42% ± 19% at 14 days, P < 0.05) than any other treatment. Moreover, the tissues treated with sham, ethanol alone, or FUS alone were nearly 99% viable at 14 days, while the combination treatment group showed marked tumor necrosis (Fig. 4D). These data clearly demonstrate that mechanochemical disruption causes significant tumor regression in vivo.

In addition to having a direct mechanical effect on gene expression (24), FUS causes rapid heating of tumor tissue, which initiates a heat shock response in cancer cells (25). Somehow, the molecular changes induced by the mechanical and thermal effects are amplified in the cells by chemical effects due to ethanol treatment, resulting in diminished tumorigenic and metastatic potentials. To elucidate the molecular mechanisms behind the synergy between FUS and ethanol, we measured the expression level and activity of intracellular proteins potentially responsible for observed changes in cell adhesiveness, viability, proliferation, and tumorigenicity.

One of the molecules we studied was NFκB, a primary transcription factor that plays a central role in cell proliferation, proinflammatory signaling, and production of cytokines, chemokines, and adhesion molecules including CD29 and CD44 (26). The total and nuclear expression levels of NFκB were measured in DU-145 cells at 2, 24, and 72 hours post-treatment using Western blot analysis (Fig. 5B). The total NFκB expression at 2 hours significantly and monotonically reduced between control, ethanol alone, FUS alone, and ethanol + FUS treatments, respectively. It returned to its control level at 24 hours in all treated cells excluding those exposed to FUS at a sufficiently high acoustic power (H5). Notably, the cells in the E+H5 group had no measurable NFκB expression at 2 hours (Fig. 5B, top left) and maintained a significantly low NFκB expression even at 72 hours as compared with control (0.16 ± 0.1 vs. 1.0 ± 0.1, P < 0.01), while the cells in the H5 group incompletely rebounded to the control level of expression (0.5 ± 0.2 vs. 1.0 ± 0.1, P < 0.05). The nuclear NFκB expression data (Fig. 5B, top right and bottom) inform us about the NFκB transcriptional activity, which requires translocation of this protein to the nucleus. The cells exposed to combination treatment (E+H4, E+H5) had very low nuclear expression of NFκB at 24 hours (P < 0.05), which did not rebound at 72 hours (P < 0.01). The cells in the ethanol alone or H4 alone group had a reduced NFκB transcriptional activity than control cells at 72 hours (P < 0.05), but their NFκB activity was consistently higher than that of the cells in the combined treatment groups. NFκB signaling is stimulated by TNFR1 activity (27). However, the ethanol + FUS treatment decreases NFκB expression and inhibits its activity (Fig. 5B), thus drastically diminishing the contribution of TNFR1 to cell proliferation and metastasis. Thus, mechanochemical disruption causes TNFR1 to act solely as a death receptor.

The constitutive expression of HSP 70 kDa (Hsp70) inhibits apoptosis (28). However, Hsp70 overexpression suppresses NFκB signaling (29) and sensitizes the cells to Fas-induced apoptosis (30). This suggests that the overproduction of Hsp70 may be responsible for reduced expression of NFκB and increased apoptosis in prostate cancer cells treated with ethanol and FUS. Figure 5C shows the effect of treatment conditions on Hsp70 expression in DU145 cells, measured using Western blot at 2, 24, and 72 hours post-treatment. There was an immediate increase in Hsp70 expression in the E+H4 group (1.5 ± 0.3 vs. 0.9 ± 0.0 in control group, P < 0.05), followed by increased Hsp70 expression in all cells exposed to FUS at 24 hours post-treatment (1.79 ± 0.1 in E+H4 vs. 1.12 ± 0.0 in control group, P < 0.05).

Chemical drugs that induce significant production of reactive oxygen species (ROS) cause TNF- and Fas-mediated cell death (31, 32). We investigated the effect of various treatments on ROS expression in DU-145 and C4-2B cells. As seen in Fig. 5D, the expression of ROS in the cells of the E+H5 group was significantly higher than the positive control (100 μmol/L of H₂O₂) at 24 hours (8.4 ± 0.06 vs. 5.2 ± 0.14, P < 0.001). The ROS expression was greater or within range of the positive control in all FUS-treated groups at 72 hours, with significant differences observed in groups treated with or without ethanol [4.7 ± 0.8 (H4) vs. 8.2 ± 0.9 (E+H4), P < 0.01 and 7.5 ± 0.8 (H5) vs. 10.7 ± 0.2 (E+H5), P < 0.01]. Similarly, the combination treatment groups led to significant (P < 0.05 or less) overproduction of ROS in C4-2B cells at both 24 and 72 hours post-treatment, as compared with FUS alone, ethanol alone, or control (Supplementary Fig. S15). Overall, the data presented in Figs. 1F and G and 5 suggest that mechanochemical disruption activates proapoptotic pathways and inhibits NFκB-dependent survival in prostate cancer cells by overproducing Hsp70 and ROS.

The median age of a cancer diagnosis, including prostate cancer, is 66 years (33). Because of old age, invasive therapies are not recommended for the majority of patients with prostate cancer even with indications that they may decrease disease progression and increase life expectancy (34). Most of the elderly patients undergo radiation treatment, sometimes together with androgen deprivation therapy, if their disease reaches a high-risk localized state. It has been shown that high-risk localized prostate cancer is often resistant to this curative therapy, leading to the death of approximately 50% of patients within 10 years after treatment (35). There is a clear need for alternative, minimally invasive treatment options for locally advanced cancer, with a reduced risk of complications and high effectiveness. In this regard, the use of HIFU for cancer treatment is spreading in recent years but standalone HIFU has its own challenges such as small treatment area, long procedure time, and incomplete tumor ablation (36, 37) that prevent its use for advanced cancer treatment. The clinical study conducted in Germany (38) revealed that only 52% of patients with intermediate- or high-risk localized prostate cancer (clinical stages T2b–T4) were disease free at 5 years after standalone HIFU treatment, while disease-free survival was observed in 82% of patients with low-risk cancer. A drop from 85% to 47% in disease-free survival between low- and high-risk prostate cancer patients was also detected within 21-month follow-up of standalone HIFU treatment (39).

As demonstrated in our study, when FUS at intensities much less than required for clinical HIFU was used in combination with ethanol injection, it dramatically reduced and even eliminated tumors in the xenograft mouse model of human prostate cancer (Fig. 4). It is important to mention that this strong sensitizing effect of ethanol on FUS cannot be fully recapitulated by targeted chemotherapy. Not only ethanol synergizes with FUS in cancer cell apoptosis and reprograming, but ethanol also modifies biophysical properties of the tumor that enhances FUS ablation. According to our earlier studies (12, 13, 40), a combination of ethanol and FUS synergistically increases the tumor ablation volume due to biophysical effects such as FUS-induced acoustic streaming and membrane erosion as well as ethanol-induced reduction of the inertial cavitation threshold. Such biophysical effects are not expected from chemotherapeutic drugs, which target a specific part of the cancer cell's machinery. Ethanol is also a chemical agent that directly affects multiple structures inside the cell, including the cell membranes, endoplasmic reticulum, mitochondria, and nucleus. This increases the likelihood of overlapping and complimentary effects of ethanol and FUS on multiple signaling pathways, leading to enhanced cell death and cell reprograming. These effects include mitochondrial production of reactive oxygen species (ROS) that activate mitochondrion-based cell apoptosis through cytochrome C and caspases (41), translocation of FasR and FasL to the plasma membrane (42), and suppression of NFκB activation by IKK complex inactivation (43), to name a few.

Our molecular analysis reveals that the mechanical perturbations induced by FUS also trigger molecular events within the cell, leading to a less aggressive cell phenotype (Figs. 1F, 2B, and 5B and E). Specifically, our data indicate that the decline in cell viability (Fig. 1A–E) coincides with increased expression of Fas and TNFR1 (Fig. 1F), and the viability is recovered via blocking of these death receptors (Fig. 1G). Thus, Fas and TNFR1 play a vital role in the observed synergistic elimination of cancer cells by ethanol and FUS. ROS is likely a major player in the increased apoptotic activity via Fas and TNFR1. Its drastic increase in the combination treatment group (Fig. 5D) coincides with increased expression of death receptors (Fig. 1F) and a large population of early and late apoptotic cells (Fig. 1B and C). Death receptor activity alone, however, cannot explain the observed decrease in proliferation rate (Fig. 1E) and adhesion to endothelium (Fig. 2B and C), reduction in cell surface expression of markers for adhesion and migration (Fig. 2D), and the inability to form tumor spheroids (Fig. 3) in the combined treatment group. These effects are most likely caused by impairment of NFκB activity (Fig. 5B) due to overproduction of both ROS and Hsp70 (Fig. 5C and D). Cellular uptake of ethanol directly produces ROS (41) and is known to induce Hsp70 production via Heat Shock Factor 1 (HSF1; refs. 44, 45). Similarly, tissue vibration and heating induced by FUS are shown to increase ROS production (46) and stimulate Hsp70 production either directly or also through HSF1 (24, 47). The overproduction of each of these molecules has been implicated in inhibiting key metabolites in the NFκB signaling pathway (29, 48, 49) and thus switching the cells from a surviving/proliferating/adhesive state to an apoptotic/nonadhesive state. Our proposed schematic of the molecular pathways induced in cancer cells after exposure to ethanol and FUS is shown in Fig. 6. Nontoxic chemical agents that target the pathways identified in this figure (NFκB, ROS, Hsp70, etc.) can be used as safer and more accessible alternatives to ethanol for FUS-mediated treatment of cancer.

Cancer cells are under endogenous chromosomal and oxidative stresses due to their high proliferation rate and oxygen- and nutrient-deprived environment (50). In response to these stresses, they constitutively produce chaperones, including HSPs, which refold damaged proteins or mediate disassembly of irreparable proteins (51). Through protein repair and interference with apoptotic pathways, chaperones allow for cancer cells to survive and provide an opportunity to develop resistance to therapy (52). However, exogenous stresses due to physical force fields, such as electromagnetic radiation and FUS, may overload even the most aggressive cells by inducing extensive disruption of cellular structures. These cause the cells to be senescent (53) to either repair cell-wide damage or apoptose. During senescence, the cells halt the transcription and translation of general proteins but produce a large amount of chaperones (54). When overproduced, some of these chaperones, like Hsp70, begin to interfere with NFκB signaling, thus reducing the ability of the cells to proliferate, adhere, and migrate. It is important to note that FUS-induced mechanical vibrations not only broadly and directly disrupt cellular structure, but they also induce conformational changes in macromolecules (55), which may limit the binding ability and efficacy of repair proteins. The FUS-induced broad disruption of cellular activity is equally experienced by diverse populations of cancer cells, and the mechanisms of response to such stresses are conserved even among lower organisms (54). Redirecting the cancer cells to cell-wide repair diminishes their ability to develop resistance to a targeted chemical treatment. This interplay of exogenous stresses, via a combination of physical force fields and chemical exposure, could explain a dramatic regression of tumor size in vitro (Fig. 3) and in vivo (Fig. 4) after treatment with FUS and ethanol.

Several studies have shown the possibility to reverse cancer via mechanisms such as unpacking of chromatin (56), inhibition of ROCK (57), and regulation of PLEKHA7 (58) and Myc (59). These mechanisms can be activated by mechanochemical disruption via FUS and ethanol. For example, strong vibrations of molecules in the nucleus due to FUS exposure can unpack DNA structures (60). The collapse of actin cytoskeleton, as evident in Fig. 2, may impair the functions of ROCK and PLEKHA7 in regulating cell contractility and homotypic cell-cell adhesion. Finally, heat shock induced by mechanochemical disruption may downregulate the expression of transcriptional factor Myc (61). By integrating multiple mechanisms, FUS-induced mechanical perturbations coupled with chemical agents, which amplify and guide remedial cellular responses, are capable of transforming heterogeneous and drug-resistant populations of cancer cells to a healthier, native phenotype. This unique, in vivo–accessible mechanochemical treatment platform could be the holy grail in the fight against cancer.

Chemical ablation via percutaneous ethanol injection (62) is one of few curative treatments available for patients with early-stage liver cancer. The transurethral application of ethanol has been tested successfully for treatment of benign prostatic hyperplasia (63). Because both ethanol and FUS are already used clinically for treatment of prostatic diseases and their combination requires subclinical doses to induce therapeutic effects on aggressive prostate cancer cells, we anticipate that mechanochemical disruption (MCD) via ethanol and FUS can be quickly translated into a clinical therapy. Ethanol is more beneficial to use with FUS in treatment of locally aggressive cancer because it cannot be delivered systemically at doses required to induce cytotoxicity. To expand the MCD therapy for curative treatment of metastatic CRPC, systemic drugs that increase ROS production and inhibit NFκB signaling can be used instead of ethanol. It should be noted that CRPC preferentially metastasizes to the bone. FUS has shown great promise and approved by FDA for palliative treatment of bone metastases (64, 65). Using advanced PET/CT technology (66), metastatic CRPC foci can be detected in the bone and treated by a drug–FUS combination. There is still a long road to this exciting opportunity, which requires establishing feasibility of the MCD therapy in patient-derived xenografts, immunocompetent mouse models (to particularly test whether the intact immune system can interfere with MCD-induced ROS production and NFκB signaling), and clinical trials. The MCD therapy development will be the major focus of our future studies.

D.B. Khismatullin is a president and chief scientific officer and has ownership interest (including stock, patents, etc.) in Levisonics, Inc. No potential conflicts of interest were disclosed by the other authors.

Conception and design: H.Y. Murad, H. Yu, D. Luo, G.M. Halliburton, D.B. Khismatullin

Development of methodology: H.Y. Murad, H. Yu, D. Luo, G.M. Halliburton, D.B. Khismatullin

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): H.Y. Murad, H. Yu, D. Luo, E.P. Bortz, G.M. Halliburton

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): H.Y. Murad, H. Yu, D. Luo, E.P. Bortz, G.M. Halliburton

Writing, review, and/or revision of the manuscript: H.Y. Murad, H. Yu, A.B. Sholl, D.B. Khismatullin

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): H.Y. Murad, H. Yu

Study supervision: D.B. Khismatullin

This study was supported by grants from the U.S. NIH (R01HL127092), U.S. National Science Foundation (1438537), American Heart Association Grant-in-Aid (13GRNT17200013), Louisiana Board of Regents (LEQSF-EPS(2012)), Tulane University Office of the Provost, Tulane Center for Engaged Teaching and Learning, and Newcomb-Tulane College. H.Y. Murad was supported by Louisiana Board of Regents Graduate Fellowship (LEQSF(2014-2018)-GF-14). We thank W.T. Godbey for assistance with diagnostic ultrasound imaging, Z. You for help with Western blot experiments, P. Chandra for help with the scratch-wound assay, and D. Mondal, C. Abshire, and N. Ramanujam for fruitful discussions. We also thank J. Arora and J. He for Figs. 2A and 5A and Supplementary Fig. S6.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.