# DNA Double-Strand Breaks in Murine Mammary Tumor Cells Induced by Combined Treatment with Doxorubicin and Controlled Stable Cavitation

Cécile Fant, Adeline Granzotto, Jean-Louis Mestas, Jacqueline Ngo, Maxime Lafond, Cyril Lafon, Nicolas Foray, Frédéric Padilla

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## ► To cite this version:

Cécile Fant, Adeline Granzotto, Jean-Louis Mestas, Jacqueline Ngo, Maxime Lafond, et al., DNA Double-Strand Breaks in Murine Mammary Tumor Cells Induced by Combined Treatment with Doxorubicin and Controlled Stable Cavitation. Ultrasound in Medicine & Biology, 2021, 47 (10), pp.2941-2957. 10.1016/j.ultrasmedbio.2021.05.028. hal-05339386

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# DNA DOUBLE-STRAND BREAKS IN MURINE MAMMARY TUMOR CELLS INDUCED BY COMBINED TREATMENT WITH DOXORUBICIN AND CONTROLLED STABLE CAVITATION

CECILE FANT, ADELINE GRANZOTTO, JEAN-LOUIS MESTAS, JACQUELINE NGO, MAXIME LAFOND, CYRIL LAFON, NICOLAS FORAY, and FREDERIC PADILLA

## Abstract

Chemotherapeutic agents such as doxorubicin induce cell cytotoxicity through induction of DNA double-strand breaks. Recent studies have reported the occurrence of DNA double-strand breaks in different cell lines exposed to cavitational ultrasound. As ultrasound stable cavitation can potentiate the therapeutic effects of cytotoxic drugs, we hypothesized that combined treatment with unseeded stable cavitation and doxorubicin would lead to increased DNA damage and would reduce cell viability and proliferation in vitro. In this study, we describe how we determined, using 4T1 murine mammary carcinoma as a model cell line, that unseeded stable cavitation combined with doxorubicin leads to additive DNA double-strand break induction. Combined treatment with doxorubicin and unseeded stable cavitation significantly reduced cell viability and proliferation at 72 h. A mechanistic study of the potential mechanisms of action of the combined treatment identified the presence of cavitation necessary to increase early DNA double-strand break induction, likely mediated by a bystander effect with release of extracellular calcium.

## Introduction

Doxorubicin (DXR), or Adriamycin, has been one of the most common effective chemotherapeutic agents used for the treatment of a wide range of solid as well as hematologi-cal malignancies since 1969 (Bakkali et al. 2003; Wang et al. 2004; Renu et al. 2018). However, the clinical use of DXR is limited by its acute and chronic toxicity in healthy tissues (cumulative cardiotoxicity, hematotoxicity, acute myeloid leukaemia, etc.) (Carvalho et al. 2009). The development of resistance mechanisms in tumor cells, linked to P-glycoprotein and topoisomerase II, can hamper the ther- apeutic efficacy of DXR (Swift et al. 2006). Development of an efficient strategy to potentiate the action of DXR is required to improve the efficacy of chemotherapy while min- imizing the therapeutic dose administered to patients (Dob- son 2006).

DXR is a cytotoxic agent because it induces DNA double-strand breaks (DSBs) through several mechanisms including the intercalation of the planar anthracycline nucleus with the mitochondrial or nuclear DNA double helix (Ashley and Poulton 2009), the creation of a topoisomerase II DXR complex leading to its inhibition (Swift et al. 2006) and release of reactive oxygen species (Doroshow 1983).

Ultrasound (US) can interact with chemotherapeu- tic agents to maximize their biological actions and enhance tumor cell death, while causing minimal dam- age to surrounding healthy tissues (Trendowski 2014). More specifically, the synergetic therapeutic effect of the US DXR combination on tumor cells has been observed in several studies (Saad and Hahn 1989; Loverock et al. 1990; Umemura et al. 1997; Yu et al. 2001; Chettab et al. 2017; Fant et al. 2019).
One of the mechanisms by which ultrasound (US) can potentiate DXR is thought to be cavitation, or the creation of gas bodies in liquid or tissues that will act as oscillating nuclei, being compressed at positive pressures and expanded under negative pressures. Cavitation is a phenomenon that requires an acoustic pressure higher than a certain threshold to be initiated. Two kinds of cavitation can occur: stable cavitation, which is caused by small to moderate growth and oscillations of gas bubbles under sustained acoustic cycles, and inertial cavitation, which refers to the violent growth and immediate collapse of bubbles (Leighton 1994; Wu 2002; Stride and Coussios 2010). For therapeutic applications, cavitation is usually created and maintained by adding synthetic cavitation nuclei such as US contrast agents, gas bubbles that remain in the bloodstream (Ferrara et al. 2007; Sennoga et al. 2017; Kooiman et al. 2020). Alternatives to such agents are being investigated, and we developed a method to create and maintain stable cavitation without the use of these US contrast agents (Yasuda et al. 2016; Fant et al. 2019), a phenomenon referred to as unseeded stable cavitation.

Several studies have indicated that cavitational US may affect DNA structure as well as its function by inducing single-strand breaks and DSBs (Pinamonti et al. 1986; Furusawa et al. 2014). The interaction of US and purified DNA in aqueous solution was documented decades ago, and after sonication, DNA damage has been observed in mouse L cells and Chinese hamster ovary cells (Kondo and Yoshii 1985; Miller et al. 1995). Since then, some studies have reported that cavitational US can induce DSBs both in vitro and in vivo (Ide et al. 2008; Furusawa et al. 2012; Aydin et al. 2019).

Among the three major types of DNA damage that can generally be observed, base damage (BD), single-strand breaks (SSBs) and double-strand breaks (DSBs) (Gastaldo et al., 2008), DSBs are less frequent but the most hazardous for cells if unrepaired (toxicity) or mis-repaired (cancer) (Furusawa et al. 2014; Foray et al. 2016). DSBs are produced within the first microseconds by ionizing radiation and within the first minutes or hours by certain drugs such as bleomycin and doxorubicin (Tounekti et al. 2001; Huelsenbeck et al. 2012). In mammalian cells, the most predominant DSB repair and signaling pathway is non-homologous end-joining (NHEJ). Early after stress, the DSBs managed by NHEJ are recognized by the ATM-dependent phosphorylation of the variant histone H2AX (g-H2AX).

Such phosphorylation triggers a relocalization of g-H2AX as nuclear foci at the DSB sites.
The number of g-H2AX foci, easily quantifiable by immuno-fluorescence, has been found to be quantitatively correlated with DSBs (Rothkamm and Lobrich 2003).

Here, we hypothesized that combining DXR with unseeded stable cavitation would enhance the therapeutic efficacy of DXR, notably through the induction of additional DNA DSBs. We report the generation of early DSBs, quantified through the g-H2AX immunofluorescence method, by combined treatment with DXR and unseeded stable cavitation of 4T1 murine mammary tumor cells. We report that unseeded stable cavitation synergistically acts with DXR to affect the viability and proliferation of 4T1 cells. We also investigated the underlying mechanisms enhancing DSB induction, notably the roles of a bystander effect and of calcium release.

## Methods

### Cell line and culture conditions

The adherent murine mammary 4T1 tumor cells, with a rapid doubling time of 23 h (Pulaski and Ostrand-Rosenberg 2001), were cultured as a monolayer in T75 culture flasks in supplemented Dulbecco's minimum essential medium (DMEM, 10% fetal calf serum [FCS],
1% L-gluta- mine, 1% penicillin-streptomycin) and incubated at 37°C in an humidified atmosphere (5% CO2, 95% O2). Passages were done every 3 d to prevent cells from exceeding 90% confluence as overgrowth decreases their viability. All reagents were purchased from Invitrogen Life Technolo- gies (Cergy-Pontoise, France). 4T1 cells were tested myco- plasma-free using the Mycoalert kit (Lonza, Cologne, Germany).

## Chemicals

DXR hydrochloride 2 mg/mL (Accord Healthcare, Lille, France) was used at a concentration of 400 ng/mL. This dose was chosen for its ability to induce 20% cell death in the 4T1 cell line at 48 h (Fant et al. 2019).

## Cavitational US exposure

The cavitation device used in this study has been described in detail elsewhere (Fant et al. 2019). Briefly, the device is composed of two confocal transducers operated at 1.1 MHz. This setup generates a small focal spot (2 x 2 x 2 mm) at the intersection of both acousti- cal axes, where the cavitation activity is initiated (Lafond et al. 2017). For treatment, 650 mL of cells was placed in a 2-mL safe-lock micro-test tube (Eppendorf, Hamburg, Germany), positioned at the focal zone (Fig. 1). The emission signal was generated with a National Instruments device (National Instruments, Nan- terre, France) driven by LabVIEW software and powered by a 200-W amplifier (LA-200H EMV, Kalmus, Hong Kong, China). Stable and inertial cavitation was moni- tored by recording the acoustic emission from the bub- bles with an in- house hydrophone made with a polyvinylidene fluoride film (25-mm thickness, 10-mm diameter) in resin (AY103 Araldite + 10% HY956) placed between the two focused transducers, directed at the exposed sample. The signal recorded from cavitating bubbles exhibits specific features in its frequency content that are characteristic of stable or inertial cavitation regimes, and cavitation indexes were defined and calcu- lated based on these features of the acoustic spectrum. The stable cavitation index (sCI) relies on the apparition of subharmonics on the spectrum caused by bubble sta- ble oscillations, and was calculated as the difference between the maximum power level in the sub-harmonic range 540–560 kHz and the mean base level obtained in the inharmonic range 560–600 kHz. During inertial cav- itation events, when bubbles are imploding, a broadband noise is emitted resulting in an increase in the base level of the acoustic spectrum. An inertial cavitation index was calculated as the average of the entire frequency spectrum in the range 0.1–7.1 MHz, first calculated in decibels (relative to 1 mV) and then normalized by the base level, over the same frequency range, and measured when the amplifier is turned on with no input signal. The stable cavitation regulation system has been described previously (Fant et al. 2019) and uses the recorded cavi- tation noise and resulting sCI as input for a feedback loop amplitude control of the emitted signal. Two distinct modes can be emitted: a high-power mode with the generation of higher pressure pulses (Pmax, “bursting pulses”) to initiate cavitation (peak negative pressure [PNP] of 6.7 MPa at the focus) when the sCI is below a threshold of 3 dB— indicating the absence of stable cavi- tation—and a low-power mode (Pmin, “regulation pulses”) to maintain stable cavitation activity (PNP of 2.9 MPa at the focus) when the sCI is >3 dB.

For DNA damage experiments, 6600-cycle pulses were emitted with a pulse repetition frequency of 25 Hz, corresponding to a duty cycle of 15%. Exposures were 30 s long. Those parameters have previously been found to trigger a stable and repro- ducible regimen of cavitation, with limited cell death to <30% (Fant et al. 2019).
To create cell debris for bystander experiments, cells were treated with inertial cavitation. The inertial cavitation regulation system has been previously described, and relies on real-time monitoring of inertial cavitation activity for feedback control of the input electrical power required to sustain inertial cavitation activity during treatment (Sabraoui et al. 2011; Chettab et al. 2015). Cells were exposed to 2750-cycle pulses (pulse repetition frequency = 100 Hz) at an index of inertial cavitation of 12, for 45 s. These parameters have been found to induce approximately 50% immediate cell death.

## DXR and stable cavitation combined treatment

Seventy-two hours before the experiment, cells were plated at a density of 2 10⁶ cells per T75 flask to reach 90% confluence on the day of the experiment. Cells were trypsinized with 0.25% trypsin per 1 mM EDTA (Invitrogen Life Technologies), centrifuged at 320g for 3 min, washed once with 5 mL of phosphate-buffered saline (PBS, Invitrogen Life Technologies) to eliminate traces of FCS, to avoid any association between DNA of dead cells and FCS under US exposure, and then resuspended in Opti-MEM Reduced-Serum Medium (Invitrogen Life Technologies), at a concentration of 2 10⁶ cells/mL. The exact volume of Opti-MEM suspension was determined after counting cells using an automated cell counter (Cellometer, Nexcelom Bioscience, Lawrence, MA, USA). The experimental design comprised four experimental conditions: (i) no treatment (NT), (ii) DXR treatment, (iii) US treatment, (iv) combined US and DXR treatment (US + DXR). For treatment, a volume of 650 mL of cell solution was poured into a 2-mL Eppendorf tube. The tubes for the DXR and DXR + US groups were supplemented with doxorubicin at 400 ng/mL. Each tube was placed in the US device, but sonication was applied only to the US and US + DXR groups under the treatment parameters described above.

## In vitro g-H2AX focus immunofluorescence

The g-H2AX biomarker was used to assess DSBs after DXR and/or US treatment. It has been reported that there is a strong correlation between the number of g-H2AX foci per nucleus and the number of DSBs (Sedelnikova et al. 2002; Lammertink et al. 2016). After sonication, cells were transferred and incubated for 10 min, 1 h, 4 h or 24 h (on coverslips in dishes for the 4- and 24-h time points). At 10 min and 1 h, 300 mL of cells was collected, centrifuged at 300g for 5 min, resuspended in 400 mL of PBS and cytospun on coverslips at 650 rpm for 5 min. Cells were then fixed in 100 mL of 4% paraformaldehyde for 10 min at room temperature and permeabilized in permeabilization solution (20 mM Hepes, pH 7.4, 50 mM NaCl, 3 mM MgCl2, 300 mM sucrose and 0.5% Triton X-100; Sigma-Aldrich, Poole, UK) for 3 min at room temperature (Fig. 2). Cells for the 4- and 24-h time points were fixed directly onto the coverslips. After fixation, coverslips were washed in PBS before immunostaining. Anti-phospho-g-H2AXser139 antibody (No. 05636, clone JBW301, Merck Millipore, Darmstadt, Germany) incubation was performed for 40 min at 37°C at 1:800 dilutions (30 mL per coverslip) in PBS supplemented with 4% bovine serum fraction V albumin (BSA, Sigma-Aldrich), followed by washing in PBS (Fig. 2). Incubations with anti-mouse fluorescein isothiocyanate secondary antibodies (No. 12-506, Sigma-Aldrich) were performed at 37°C at 1:100 (30 mL per coverslip) in 2% BSA for 20 min at 37°C (Fig. 2). Nuclei were counterstained with 40,6-diamidino-2-phenylindole (DAPI, Sigma-Aldrich) for 10 min at 4°C. Coverslips were mounted in Vectashield (Vector Laboratories, Burlingame, CA, USA). The number of foci per cell was manually counted using a BX51 Olympus fluorescence microscope with an 100 objective (Olympus, France, Rungis, France), according to a methodology previously
described (Ferlazzo et al. 2020). Three categories of cells with foci were considered: cells
with <10 g-H2AX foci per nucleus, cells with 10 g-H2AX foci per nucleus and highly damaged
cells (HDCs) with
>100 g-H2AX foci. This experiment was done twice.

Cell viability

Cell viability was assessed by flow cytometry using a FACS LSR II (BD Biosciences, Franklin Lakes, NJ, USA) and the FACS DIVA Software for data processing. Cells were stained with the necrosis marker DAPI (dilution 1/500, Sigma-Aldrich, intensity measured with the 405-nm blue laser) and the apoptosis marker Annexin V (dilution 1/100, Annexin V APC, Annexin V Apoptosis detection kit APC, eBioscience, Paris, France, intensity measured with the 633-nm red laser), and assessed viable if negative for both markers. Flow cytometry was performed 1 and 72 h after sonication on 10,000 viable cells according to a protocol already described (Fant et al. 2019). Four experimental conditions were studied (four to six samples per condition): (i) No treatment (NT), (ii) US treatment (US), (iii) DXR treatment (DXR), (iv) combined US and DXR treatment (US + DXR).

Cell proliferation

Immediately after sonication, cell counting was per- formed using a Cellometer (Nexcelom,
Bioscience, Law- rence, MA, USA) to determine for each sample a volume containing 100,000
viable cells. Cells were plated in a well of 24-well plate and supplemented with 1 mL of
culture medium and DXR at 400 ng/mL for the DXR and US + DXR groups. After 72 h of
culture, a cell growth index was calculated as the absolute number of viable cells counted at
72 h, divided by the initial num- ber of cells that were reseeded after treatment (100,000
cells). An index <1 indicates inhibition of cell prolifera- tion and/or cell death, while an index
>1 indicates con- tinuous cell proliferation.

Influence of cavitation

To investigate whether cavitation is required to induce the observed biological response, an experiment was designed to deliver the same acoustic energy in or not in the presence of cavitation. In the first setting (named USCav), US treatment parameters were as

previously described to initiate stable cavitation. In the second setting (USNoCav), the
bursting pulses were omitted to prevent the onset of cavitation, and cells were treated only
with the regulation pulses. Because the bursting pulses were responsible for a very small
fraction of the total acoustic energy delivered to the cells in our settings (Fant et al. 2019),
cells were considered to have been treated with equivalent acoustic energy doses in the two
settings. The presence or absence of cavitation was monitored in both settings via
quantification of the sub- harmonic component in the spectrum of the hydrophone-
recorded signals. The experimental design comprised six conditions: (i) No treatment (NT),
(ii) sonication with cavitation creation (USCav), (iii) sonication without cav- itation creation
(USNoCav), (iv) DXR only (DXR), (v) DXR + US with cavitation (USCav + DXR), (vi) DXR + US
without cavitation (USNoCav + DXR). After sonication, samples were processed and DNA
damage was assessed as described under In Vitro g-H2AX Focus Immunofluorescence. This
experiment was performed only once.

Analysis of DXR internalization
DXR is fluorescent (excitation: 470 nm, emission: 590 nm [Laginha 2005]), and its internalization was assessed by measuring the mean fluorescence intensity of cells using flow cytometry, with intensity measured with the 488-nm blue laser. The fluorescence intensity of internalized DXR was assessed 1 and 72 h after sonication.

## Bystander effect

A bystander effect corresponds to indirect induction of DSBs in cells that were not directly affected by a given genotoxic stress. This phenomenon involves various intracellular mechanisms, notably the calcium release in surrounding cells (Prise and O'Sullivan 2009; Mothersill et al. 2018; Lyng 2019). To assess such an effect, cell debris was generated by inertial cavitation, and the supernatant of the corresponding suspension was placed onto non-sonicated cells (Fig. 2). Twenty-four hours before the experiment, 300 mL of non-sonicated 4T1 cells was seeded on coverslips in dishes at a concentration of 2 10⁶ cells/mL and incubated in complete medium at 37°C. On the day of the experiment, 1 mL of 4T1 cells at a concentration of 2 10⁶ cells/mL was treated with inertial cavitation as described under Cavita-tional Ultrasound Exposure and then incubated for 4 h at 37°C. The cells were then centrifuged at 300g for 5 min, the supernatant was collected and non-sonicated cells were exposed to the supernatant of sonicated cells and incubated at 37°C for 10 min, 1 h, 4 h or 24 h. g-H2AX immunofluorescence was then assessed as described above. This experiment was repeated twice.

## Calcium chelation by phosphate buffer saline solution

To further confirm the role of calcium in the bystander effect, the bystander experiment was repeated with the addition of 1 mL of a calcium chelator—PBS— to the supernatant (Fig. 2). This experiment was repeated twice.

## Clonogenic assays

Standard clonogenic assays were used for the assessment of the effect of US treatment on cell prolifer-ation (Franken et al. 2006; Al-Mahrouki et al. 2017). Two experimental conditions were studied: (i) no treat-ment (NT) and (ii) US treatment (US). After unseeded stable cavitational US exposure, 100 cells were re-plated in petri dishes, supplemented with 10 mL of culture medium and incubated for 10 days in an humidified atmosphere (5% CO2, 95% O2). After 10 d, cell medium was removed, and dishes were rinsed with PBS, fixed and stained with 2 mL of a mixture of 75% of 95% etha-nol and 25% of crystal violet (HT90132-1L, Sigma-Aldrich, St Quentin Fallavier, France). The glutaralde-hyde crystal violet mixture was removed and rinsed with tap water. Colonies were then analyzed using a stereomicroscope, and counted if formed of a heap of at least 50 cells. This experiment was repeated twice, with three to five dishes per condition.

## Intracellular calcium chelation with BAPTA-AM

To assess whether intracellular calcium release affected the cell response to the treatment, cells were pre-incubated before treatment with BAPTA-AM (1,2-bis(2-aminophenoxy)ethane-N,N,N0,N0-tetraacetic acid tetrakis(acetoxymethyl ester)), an intracellular calcium chelator (Sigma-Aldrich, St. Quentin Fallavier, France), at a concentration of 10 mM from a 10 mM stock in cell medium (dissolved in dimethyl sulfoxide) for 2 h in a humidified atmosphere (5% CO2, 95% O2) (Ganapathi et al. 1996; Aoyama et al. 1998; Kooiman et al. 2013). After incubation, cell medium was removed, and the cells were
processed and sonicated as previously described. Four experimental conditions were studied (five samples per condition), all pre-incubated with BAPTA-AM: (i) No treatment (NT + BAPTA), (ii) stable cavitation (US + BAPTA), (iii) DXR treatment (DXR + BAPTA), (iv) combined DXR and stable cavitation treatment (US + DXR + BAPTA). Cell viability and proliferation were assessed as described above.

## Statistical analysis

Data are presented as the mean standard deviation. Statistical analyses were performed using t R Software (R Core Team 2015). An analysis of variance with Newman Keuls post hoc test was used. A p value <0.05 was deemed to indicate statistical significance. The asterisks are only intended to flag levels of significance for three of the most commonly used levels: \*p value < 0.05, \*\*p value < 0.01 or \*\*\*p value < 0.001.

## Results

### Combined cavitational US and doxorubicin treatment leads to additive DNA DSBs

As evoked above, the formation of nuclear g-H2AX foci is considered the earliest step of recognition of the DSBs managed by NHEJ, the major DSB repair pathway in mammals. Non-treated cells exhibited a low level of g-H2AX foci (the great majority of cells had <10 foci). However, this number varied with time and reached its maximum at 24 h. These g-H2AX data are expected in our cellular model because they can result from both the cell cycle effect and the genomic instability caused by the immor talization (Fig. 3). With respect to cells treated with DXR alone, the number of g-H2AX foci was found to be drastically higher than that observed in controls at 4 h post-treatment, with 80% of cells exhibiting >10 g-H2AX foci. In addition, under these conditions, another category of cells appeared: cells exhibiting >100 g-H2AX foci, also called HDCs. At 24 h post-treatment, the proportion of HDCs reached 50%, suggesting a massive induction of DSBs caused by DXR. With respect to the cells treated with US alone, 40% exhibited >10 g-H2AX foci at 10 min post-treatment, which was drastically different from the cells treated with DXR that required some times to exhibit DSB. Furthermore, a minority (5%) of HDCs are measurable, suggesting that US can directly induce a massive amount of DSBs in some cells. Similar distributions of foci were observed later after US treatment. At 4 h post-treatment, 35% of US-treated cells had >10 g-H2AX while this proportion reached 90% in DXR-treated cells. At 24 h post-treatment, the distribution of foci observed in US treated cells was found to be similar to that of controls, with a majority of cells having <10 g-H2AX foci. Together, these findings indicate that sonication may result in the production of DSBs, but earlier and to a lower extent than in cells treated with DXR (Fig. 3).

We thereafter examined the effect of combined US + DXR treatment. At 10 min post-treatment, the percentage of cells with more than 10 g-H2AX foci was found similar to that observed with US treatment alone (40% vs. 45%, p > 0.5) and then reached the same kinetics as DXR treatment alone from 4 h post-treatment (55% of cells with >10 g-H2AX foci and 25% HDCs) to 24 h post-treatment (50% of cells with >10 g-H2AX foci and 50% HDCs), which corresponds to the situation for DXR treatment (Fig. 3). Together, our findings indicated that the DXR + US combination provides an additive effect by combining the early induction of DSBs caused by sonication and the induction of DSBs by DXR at later treatment times. It triggers a faster induction of DSB caused by DXR in cells. Quantitatively, such combined
treatment results in an average of 60% additional cells with >10 g-H2AX foci in the first 4 h of treatment.

### Combined treatment led to synergetic effect on cell lethality

Because unrepaired DSBs are quantitatively correlated to an increase in cell mortality, an increased number of residual DSBs in cells up to 24 h post-treatment is likely to enhance their lethality. To examine this hypothesis, cell viability after US + DXR treatment was assessed in vitro 1 and 72 h after treatment. Sonication, with or without DXR, significantly decreased cell viability at 1 h <10% compared with the non-treated group (NT = 87.33 0.23%, DXR = 86.55 0.41%, US = 79.85 0.96%, US + DXR = 78.98 1.79%, p < 0.0001; Fig. 4). But in the absence of DXR, viability after sonication at 72 h was not statistically different from the viability of the control group.

At 72 h, DXR with or without sonication significantly decreased cell viability compared with controls (NT = 72.3 2.85%, DXR = 33.43 3.8%, US + DXR = 10.15 2.7%, p < 0.001 and p < 0.0001 for DXR and US + DXR groups, respectively; Fig. 4).

Cell viability was significantly lower in the US + DXR group, than in the DXR group at 72 h (p < 0.01; Fig. 4).

In parallel to cell viability, cell proliferation was also quantified by using a cell growth index (CGI) as the ratio of cell number at 72 h to cell number at the time of plating. At 72 h, the CGI did not significantly differ between non-treated and US groups and was close to 6 (NT = 5.54 1.2, US = 6.69 3.73). In the DXR-treated samples, CGI was 0.33 0.17 after 72 h for the DXR group and 0.06 0.004 for the US + DXR group. The CGI at 72 h was significantly lower in the US + DXR group than in the DXR group (p < 0.01; Fig. 4).

### During sonication DSBs may be induced partially because of the presence of a stable cavitation

To investigate the mechanism of formation of DSBs during the sonication, the impact of stable cavitation was examined by sonicating cells with the same acoustic energy, with or without induction of cavitation (by omitting the bursting pulses initiating the cavitation). Similarly to the previous experiments, the g-H2AX foci were used as an endpoint. While between 10% and 20% of non-treated cells had >10 g-H2AX foci, whatever the observation time, stable cavitation treatment led to 40% of cells with >10 g-H2AX foci, with 20% HDCs, and which reached 60% with 10% HDCs at 1 h post-treatment (Fig. 5). When sonication without cavitation was applied, the percentage of cells with >10 g-H2AX foci and the percentage of HDCs appeared to be systematically lower than those obtained with cavitation at all post-treatment times tested (Fig. 5). Together, these findings suggest that cavitation may facilitate the formation of DSBs and the occurrence of HDCs.

When DXR was combined with sonication, the percentage of cells with >10 g-H2AX foci increased, as observed previously. DXR + US without cavitation treatment accelerated the formation of g-H2AX foci and caused HDC cells to appear earlier. In contrast, the addition of stable cavitation led to a stronger impact on the formation of g-H2AX foci until 24 h, with more cells with >10 foci (80%, 100% and 100% of cells with more than 10 foci at 10 min, 1 h and 4 h, respectively, for USCav + DXR treatment compared with 70%, 80% and 90% for USNoCav + DXR (Fig. 5). These findings suggest that sonication with cavitation combined with the DXR treatment leads to the highest yields of DSB and HDC cells.

### Combined treatment did not lead to increased DXR internalization
The fluorescence intensity of DXR was measured to assess whether DSB created by combined US + DXR treatment was owing to an increased internalization of DXR in cells. The fluorescence intensity of DXR was similar for the DXR and US + DXR groups at the two time points (at 1 h: NT = 382.25.74, US = 403.78.47, DXR = 735.543.13, US + DXR = 843.351.78; at 72 h, NT = 561.587.43, US = 796154, DXR = 21850860, US + DXR = 223431024), indicating that drug uptake was similar with and without cavitation (Fig. 6). These findings thus suggested that the increased induction of DSB in the US + DXR group was not owing to an increased internalization of DXR.

### DSBs induced by US treatment can be partially attributed to a bystander effect through calcium release

The mechanism of induction of DSBs by US was investigated further. We hypothesized that it was owing to a bystander effect, via the indirect production of DNA strand breaks by a calcium or cytokine release facilitated by US treatment (Szumiel et al. 1990).

To assess whether such an effect can lead to DSBs, non-treated cells were exposed to the supernatant of sonicated cells for different durations (10 min, 1 h, 4 h and 24 h), and the number of g-H2AX foci was assessed thereafter. Only 10 min after incubation with the supernatant of sonicated cells, 20% of cells had >10 g-H2AX foci (Fig. 7). After 1 h post-incubation, this number increased to 30% and reached its maximum 4 h post-incubation with the occurrence of HDCs (60%) (Fig. 7). At 24 h post-incubation, 40% of cells were still HDCs. Interestingly, when using phosphate-rich medium, such as PBS, to chelate calcium, no cell with >10 g-H2AX foci was observed at any time point (Fig. 7), consistent with our hypothesis that sonication may trigger a calcium release in sonicated cells which, in turn, may induce DSBs.

### Chelation of calcium with BAPTA-AM reverses the synergetic effect of the combined treatment on cell viability and proliferation

To further investigate the role of calcium in DSB induction by combined DXR and US treatment and its impact on cell viability and proliferation, viability assays were performed with the addition of a chelator of intra-cellular calcium, BAPTA-AM, to the cell medium. The cytotoxicity of sonication and combined US + DXR treatment after calcium chelation was then assessed in vitro 1 and 72 h after treatment.

At 1 h, the impact of sonication appeared significant, as previously, with a decrease in cell viability of approximately 10% after treatment (NT = 85.49, 0.78%, US = 73.32, 2.56%, US + DXR = 79.24, 0.9%, p < 0.001; Fig. 8).

At 72 h, DXR significantly decreased cell viability compared with non-treated groups, as expected (NT = 72.92.85%, DXR = 33.24.51%, p < 0.001;

Fig. 7) but, after calcium chelation, there was no longer a significant difference between the DXR and US + DXR group (US + DXR = 31.72.13%, p = 0.84; Fig. 8).

Cell proliferation had a CGI of approximately 5 for controls and US groups (CGI = 4.74 § 1.2 and 5.17 § 1.94, respectively), and no significant difference was found between the DXR and US + DXR groups (CGI = 0.34, 0.27 and 0.54, 0.32, respectively, p = 0.22; Fig. 8).

### Ultrasound without DXR has a feeder impact on cell proliferation

As sonication does not have an impact on cell viability and proliferation at 72 h despite the induction of DSBs until 24 h after treatment, we examined whether sonication may produce
a feeder effect on the surviving cells. In other terms, we sought to verify if the cells stressed by US and entering senescence would serve as feeders to healthy cells to proliferate to a physiological level at 72 h (Llames et al. 2015). Because this hypothesis involved cellular clonogenic capacity rather than cell viability, clonogenicity assays were performed by plating untreated and treated cells at similar initial number. The assays revealed that a significantly larger number of colonies were formed after sonication, as compared with the non-treated group (NT = 54 3.57, US = 120.6 5.97, p < 0.0001; Fig. 9), validating the assumption of a feeder effect.

Discussion

The aim of this study was to evaluate the impact of combined DXR and cavitational US treatment on DNA damage in murine 4T1 mammary tumor cells in vitro and to investigate the underlying mechanisms. Using g-H2AX immunofluorescence, we revealed that US can elicit DNA DSBs as early as 10 min after sonication, which, when combined with DXR, results in enhanced cytotoxicity.

Combined US + DXR treatment led to early DSBs that increased until 4 h to reach the same kinetics as after a treatment with DXR alone. Moreover, the combined DXR and US treatment led to a synergetic effect on cell viability and proliferation, with a decrease observed at 72 h compared with DXR alone, while the treatment with cavitation alone did not affect cell viability or proliferation. In a previous study, we had reported that stable cavitation can increase the cytotoxic effect of DXR, resulting in decreased cell viability and proliferation. However, none of the potential underlying mechanisms investigated independently, which included increase in intracellular DXR, temperature elevation, production of reactive oxygen species and changes in cell morphology, could explain this enhanced lethality (Fant et al. 2019). The early DSBs created by the combination could be one possible mechanism explaining this synergy.

To investigate the underlying mechanisms by which US, in combination or not with DXR, can induce DSBs, we chose the quantification of g-H2AX foci visible by immunofluorescence routinely used in radiobiology (Foray et al. 2016). The mechanism by which DSBs are induced and repaired under sonication is not fully understood. Indeed, one of the earliest responses to DNA damage is phosphorylation by kinases belonging to the PIKK family of histone H2AX. The first step of DNA repair therefore consists of the phosphorylation of histone H2AX to form the protein g-H2AX, which will then recruit other proteins involved in the DSB repair and signaling pathways. The major proteins involved in the phosphorylation of H2AX are ATM and DNA-PKATR. We performed ATM immunofluorescence experiments (data not shown) but the results were not conclusive. However, it is noteworthy that Furusawa et al. (2012, 2014) suggested that DNA-PK rather than ATM may be involved in the formation of DSBs by US exposure. Further investigation is therefore needed to distinguish the specific roles of DNA-PK and ATM kinases in the formation of DSB by US.

The impact of US in combination with chemotherapeutic agents has already revealed increased DNA damage compared with cells treated with chemotherapeutic agents alone when microbubbles are used to induce stable cavitation and to increase the uptake of the DNA-toxic chemotherapeutic agent cisplatin by cells in vitro (Lammertink et al. 2016). In our setting, however, no increase in DXR uptake was revealed, but we previously reported that stable cavitation was necessary to enhance DXR efficacy (Fant et al. 2019). To evaluate the impact of stable cavitation in the induction of DSBs, cells were sonicated with or without
induction of cavitation but with similar acoustic energy delivered. Without cavitation (USNoCav), sonication also induced some DSBs compared with non-treated cells, but the number of these DSBs decreased to the same level as in non-treated cells 24 h post-sonication, indicating that sonication without stable cavitation led to fewer DSBs compared with sonication with stable cavitation. We determined that sonication with cavitation (USCav) resulted in the appearance of DSBs at 10 min that were still present at 24 h after sonication, indicating that stable cavitation was partially responsible for the induction of DSBs in sonicated cells.

In the cells treated with US alone, despite the early induction of DSBs revealed at 10 min post-sonication, the DSBs tended to decrease over time. Furthermore, despite an early impact on cell viability 1 h after exposure for sonicated cells, the US treatment did not result in a significant difference in viability at 72 h when compared with non-treated cells. This lack of long-term impact of the US treatment on viability, despite the early appearance of DSBs and an immediate impact on cell viability, could be explained by a feeder effect as revealed by our clonogenic assays.

To our knowledge, our study is the first to report induction of DSBs by unseeded stable cavitation. Evidence of induction of DSBs in DNA after therapeutic US has already been observed in vitro in different cell lines (Kondo and Yoshii 1985; Miller et al. 1995; Furusawa et al. 2012), in vivo in thermally ablated lesions of human prostate cancer (Ide et al. 2008) and in melanoma and mammary murine tumor models (Aydin et al. 2019). However, US-induced DSBs have not been investigated in relation to changes in immediate or delayed cell viability. It appears that DSBs induced by mechanical US would rather be caused by US-mediated mechanical stress, such as oscillating nuclei, rather than other US-mediated mechanisms such as sonochemical effect (Furusawa et al. 2012). After radiotherapy treatment, the induction of DSBs is dependent on the dose (Borrego-Soto et al. 2015). Our data suggest that induction of DSBs by US may also be dose dependent, with the presence of a threshold mechanism, requiring the presence of cavitation. However, this threshold might be highly dependent on the parameters and cells or tumor models used, with DSBs observed, for example, at intensities >0.1–0.2 W/cm² (Furusawa et al. 2012), pulse repetition below the order of seconds (Kubota et al. 2017) or PNP >6 MPa (Aydin et al. 2019).

We also hypothesized that DSBs created by unseeded stable cavitation were partially owing to a bystander effect. The bystander effect was first described after radiotherapy treatments, where DNA damage, chromosomal aberrations, changes in gene expression and cell death observed in a non-targeted cell population were induced by exposure of targeted cells to radiation (Prise and O'Sullivan 2009; Lyng 2019). The signaling pathways of such a bystander effect can be activated through gap junctions or by the release in the extracellular medium of exosomes, calcium, reactive oxygen species or even NO (Mothersill et al. 2018; Lyng 2019). The induction of DSBs in non-treated cells after exposure to the supernatant of sonicated cells and the neutralization of this effect by PBS medium rich in calcium chelator phosphates confirmed the existence of a bystander effect and provided a rationale for a possible role of calcium in the mechanism of DSB induction. To further investigate the role of calcium in DSB induction and its impact on cell viability and proliferation, viability assays were performed with the addition of a chelator of intracellular calcium, BAPTA-AM, to the cell culture medium. The addition of BAPTA-AM counteracted the synergetic impact of combined DXR + US treatment on cell viability and proliferation. Calcium is known to be a key regulator of cell proliferation and cell apoptosis (Li et al. 2000; Munaron et al. 2004; Hassan et al. 2010). Consequently, the inhibition of the synergy of the combined DXR + US
treatment on cell viability and proliferation by addition of the calcium chelator BAPTA-AM reinforces the hypothesis of a role for calcium in our results.

Increased intracellular calcium levels ([Ca2+]i) after sonication constitute a US-mediated previously reported mechanism (Mortimer and Dyson 1988; Honda et al. 2004; Zhang et al. 2012; Suarez Castellanos et al. 2016; Beekers et al. 2020). Most of the studies have investigated the role of microbubble oscillations in driving this increase (Pan et al. 2005; Schlicher et al. 2006; Juffermans et al. 2009). As unseeded stable cavitation also creates oscillating gas nuclei, it may be possible that unseeded stable cavitation leads to increased intracellular calcium levels, and a direct visualization or quantification of calcium intracellular levels after unseeded stable cavitation treatment would be of interest.

Variations in calcium levels in the extracellular medium after US exposure also remain to be investigated. Cells can pump calcium out of the cell membrane, and it has been reported that intracellular calcium concentrations return rapidly to physiologic levels post-sonication.

Moreover, some calcium flux has been seen in cells distant from the sonication focal zone (Schlicher et al. 2006; Kumon et al. 2009; Fan et al. 2010). It has been reported that, during sonication, the changes in [Ca2+]i are owing more to outflux and influx through the membrane than to the release from internal stores or buffers (e.g., endoplasmic reticulum, mitochondria, buffering proteins) or calcium channels (Kumon et al. 2007). Calcium pumped into the extracellular medium after sonication could be a mechanism of intercellular communication and an initiator of a bystander effect.

DXR can also induce an increase in [Ca2+]i (Kalivendi et al. 2001; Asensio-Lopez et al. 2017; Renu et al. 2018; Bong et al. 2020). The main source of [Ca2+]i after DXR treatment has not been clearly identified, and several mechanisms have been proposed, such as oxidative stress (Asensio-Lopez et al. 2017), mitochondrial Ca2+ overload (Abdoul-Azize et al. 2018), endoplasmic reticulum calcium (Aziz et al. 2019) and efflux pumps (Peters et al. 2016).

Moreover, it is not clear if the addition of extracellular calcium with DXR leads to a synergetic or antagonist effect on DXR cyto-toxicity and, thus, on cell viability, as different studies have produced contradictory results (Huang et al. 2015; Nguyen et al. 2016).

Both unseeded stable cavitation and DXR can potentially lead to increased intracellular and/or extracellular calcium concentrations and, when the treatments are combined, could have an impact on DNA damage formation through a bystander effect, resulting in decreased cell viability.

## Conclusions

Using g-H2AX immunofluorescence, we revealed that US, in combination or not with DXR, elicits DNA double strand breaks in 4T1 murine mammary tumor cells in vitro. Without DXR, the early formation of DSBs does not have an impact on cell viability and proliferation 72 h post-sonication, probably because of a feeder effect by senescent cells. The sonication-induced early DSB induction combines synergistically with DXR and results in an enhanced therapeutic effect, quantified by decreased cell viability and proliferation. The presence of stable cavitation is required for the potentiation of DXR by US treatment and is partly mediated by a bystander effect through calcium release.

Acknowledgments—This work was supported by the LabEx DEVwe- CAN (ANR-10-LABX-0061) of the University of Lyon, within the program "Investissements d'Avenir" (ANR-11-
IDEX-0007) operated by the French National Research Agency (ANR). Frederic Padilla was supported by the Merkin Fellowship from the Focused Ultrasound Foundation.

Conflict of interest disclosure—The authors declare no competing interests.

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A. US device

<figure><img src="image_3.png" /></figure>

B. Real-time control of stable
cavitation activity

<figure><img src="image_4.png" /><figcaption>Fig. 1. Cavitation setup. (A) US device: (a) The tank was filled with de-gassed water. A hydrophone (c) was placed in the water between the two confocal transducers (b) and directed toward the exposed sample in a 2-mL Eppendorf tube (d). (B) Real-time control of stable cavitation activity. The control system was based on the alternation of high-power pulses (Pmax) and low-power pulses (Pmin). The level and type of cavitation were characterized by several indicators on the spectrum of the signal recorded by the hydrophone: The occurrence of subharmonics (half the emission frequency) indicated stable cavitation, and a high level of broadband noise indicated inertial cavitation. If the sCI (index of stable cavitation) fell below a user-defined threshold (empiri-cally defined as 3 in our case), a high power pulse was triggered to revive the cavitation cloud. The number of bursting pulses was recorded. DOX = doxorubicin; PVDF = polyvinyl difluoride.</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>Fig. 2. g-H2AX focus immunofluorescence and bystander effect. (a) Immunofluorescence protocol. Briefly, after incubation, cells were fixed in paraformaldehyde, permeabilized, fixed with primary phosphor-H2AX and secondary antibodies and finally mounted on slides for scoring of g-H2AX foci. (b) Bystander effect. H2AX phosphorylation is the first step in DNA repair. H2AX foci in the nucleus are correlated with the number of double-strand breaks. A bystander effect is an indirect induction of DNA double-strand breaks through various intracellular mechanisms such as calcium or cyto-kin release. (c) Bystander protocol. Cell debris was generated by inertial cavitation, and the supernatant of the corresponding suspension was collected and placed onto non-sonicated cells. g-H2AX immunofluorescence was performed as described in the method section. DSBs = double-strand breaks; PBS = phosphate-buffered saline; PFA = paraformaldehyde.</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>Fig. 3. g-H2AX foci in cells at 10 min, 1 h, 4 h and 24 h after no treatment, DXR alone, unseeded stable cavitation alone or the combination of both. (a) Kinetics of the appearance of g-H2AX foci in 4T1 cells over time. Data are representative of two independent experiments. (b) Representative examples of cells stained by g-H2AX antibodies for the control and combined groups (10). (c) Representative stacked graphs of the distribution of percentage of cells with less than 10 g-H2AX foci, more than 10 g-H2AX foci and HDCs for the different conditions and the different time points of analysis. This experiment shows the additive effect of unseeded stable cavitation and doxorubicin treatment on DSB induction. DSB = double-strand break; DXR = doxorubicin; HDC = highly damaged cells; US = ultrasound.</figcaption></figure>
<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /><figcaption>Fig. 4. Cell viability and proliferation at 1 and 72 h. (a) Cell viability at 1 and 72 h of cells exposed to DXR alone, controlled stable cavitation alone or the combination of the two. (b) Effect of US, DXR and combined treatment on 4T1 cell proliferation (log scale on boxplot chart). Cells were counted at t = 0 and 72 h. This experiment illustrates the synergy of the combined treatment on cell viability and proliferation. CGI = cell growth index; DXR = doxorubicin; US = ultrasound.</figcaption></figure>
<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /><figcaption>Fig. 5. Influence of cavitation. (a) Kinetics of the appearance of g-H2AX foci in 4T1 cells over time after no treatment (Controls), doxorubicin alone (DXR), unseeded stable cavitation (USCav), US treatment without cavitation (USNoCav) or combined ultrasound and doxorubicin treatment (USNoCav + DXR and USCav + DXR) (c) Representative stacked graphs of the distribution of percentage of cells with <10 g-H2AX foci, >10 g-H2AX foci and highly damaged cells (for the different conditions and the different time points of analysis. HDC = highly damaged cells.</figcaption></figure>
<figure><img src="image_13.png" /><figcaption>Fig. 6. Mean fluorescence intensity of DXR in DXR and US + DXR groups at 1 and 72 h. This experiment indicates that combined treatment did not lead to increased DXR internalization. DXR = doxorubicin; US = ultrasound.</figcaption></figure>

<figure><img src="image_14.png" /><figcaption>Fig. 7. Bystander effect. (a) Kinetics of the appearance of g-H2AX foci in 4T1 cells after exposure of the supernatant of sonicated cells onto non-sonicated cells with (PBS) or without (Bystander) the addition of the calcium chelator PBS. Data are representative of two independent experiments. (b) Representative examples of nuclei stained by g-H2AX anti-bodies for Bystander and PBS groups (40). (c) Representative stacked graphs of the distribution of percentage of cells with <10 g-H2AX foci, >10 g-H2AX foci and highly damaged cells for Bystander group and different time points of analysis. This experiment indicates that induction of DNA double-strand breaks by cavitational ultrasound could be partially explained by a bystander effect through calcium release. DXR = doxorubicin; HDC = highly damaged cells; PBS = phosphate-buffered saline; US = ultrasound.</figcaption></figure>
<figure><img src="image_15.png" /><figcaption>Fig. 8. Cell viability and proliferation at 1 and 72 h in the presence of BAPTA-AM. (a). Cell viability at 1 and 72 h of cells pre-incubated with BAPTA-AM at a concentration of 10 mM and exposed to DXR alone, controlled stable cavitation alone or the combination of the two. Data for controls, US, DXR and US + DXR groups are the same that those in Figure 4. (b). Effect of US, DXR and combined treatment on 4T1 cell proliferation (log scale on boxplot chart). Cells were counted at t = 0 and 72 h. This experiment illustrates the impact of doxorubicin treatment on cell viability and proliferation. However, the addition of BAPTA-AM counteracted the synergetic effect of combined treatment compared with doxorubicin treatment alone, suggesting a role for calcium in the synergy. BAPTA-AM = 1,2-bis(2-aminophenoxy) ethane-N,N,N0,N0-tetraacetic acid tetrakis(acetoxymethyl ester)); CGI = cell growth index; DXR = doxorubicin; PBS = phosphate-buffered saline; US = ultrasound.</figcaption></figure>
<figure><img src="image_16.png" /><figcaption>Fig. 9. Clonogenicity assays. (a) Number of colonies formed after 10 d of incubation for non-treated and unseeded stable cavitation groups. Data are representative of two independent experiments. (b) Representative examples of formed colonies for both conditions. US = ultrasound.</figcaption></figure>