5-Aminolevulinic Acid-Mediated Sonodynamic Therapy Alleviates Atherosclerosis via Enhancing Efferocytosis and Facilitating a Shift in the Th1/Th2 Balance Toward Th2 Polarization

Ischemic heart disease (IHD) and stroke due to vulnerable atherosclerotic plaque rupture are the two leading causes of morbidity and mortality worldwide. These conditions are estimated to be responsible for one-quarter of all deaths annually. In addition, the total number of deaths from IHD and stroke are increasing globally [1-3]. Atherosclerosis (AS) is defined as a slowly progressive chronic inflammatory disease occurring in the walls of large and medium-sized arteries that arises from an imbalance in lipid metabolism and a maladaptive inflammatory response [4, 5]. T lymphocytes, a cellular minority, especially CD4+ T helper (Th) cells, are present in human plaques and are known to function decisively in the regulation of inflammation during atherogenesis [5, 6].

According to the signature cytokines produced, as well as their functions, activated Th cells are mainly defined by interferon (IFN)-γ secreted from Th1 cells and interleukin (IL)-4 from Th2 cells [7]. The Th1-cell response mediated by IFN-γ promotes macrophage activation and the production of pro-atherogenic inflammation mediators and counteracts cap formation by enhancing protease and chemokine secretion and collagen degradation. Attenuated atherosclerosis development appears in IFN-γ-deficient atherosclerosis-prone mice and in mice with genetic knockdown of the Th1-cell-associated transcription factor T-box expressed in T cells (T-bet) [8-11]. The Th2-cell-specific transcription factor GATA-binding protein 3 (Gata-3) plays a pivotal role in the differentiation of Th2, upregulates IL-4 and IL-5, and inhibits the production of IFN-γ. Cytokines released from Th2 cells, including IL-10, IL-13 and IL-25, suppress vascular inflammation and atherosclerosis. In addition, promotion of a Th2 cell response in hyperlipidemic mice inhibits atherosclerotic lesion development [12, 13]. Therefore, the equilibrium between these two subsets may influence the development of atherosclerotic plaques, and increasing regulation of Th1/Th2 may provide a novel therapeutic strategy for AS.

Low-intensity ultrasound combined with the chemical compound ‘sonosensitizer,’ termed sonodynamic therapy (SDT), has been widely used in cancer treatments [14-17]. Many studies have demonstrated that SDT has great potential as a non-invasive treatment due to the strong tissue-penetrating and regional focusing properties of ultrasound. Our group previously reported that 5-aminolevulinic acid (ALA)-mediated SDT (ALA-SDT) induces THP-1 macrophage and foam cell apoptosis via mitochondrial pathways [18, 19]. More importantly, we found that ALA-SDT could stabilize atherosclerotic plaques [20] and inhibit the progression of AS in vivo [21]. Although not as abundant as macrophages in human lesions, Th cells have been reported to play key roles in lesion growth and disease aggravation [22]. Information regarding the effects of ALA-SDT on immunologic function, especially on Th cell differentiation, is lacking. Therefore, the aim of this work was to determine whether ALA-SDT induces Th cell apoptosis and differentiation as well as Th cell-associated cytokine secretion in human lymphoblastoid Jurkat T cells and in apoE-/- mice.

5-Aminolevulinic acid (ALA), phytohemagglutinin (PHA), phorbol-12-myristate-13-acetate (PMA) and N-acetyl-L-cysteine (NAC) were obtained from Sigma-Aldrich (St Louis, MO, USA). Qualified fetal bovine serum (FBS) and RPMI 1640 medium were acquired from Gibco (Grand Island, NY, USA). (4R,6S, 8S,10Z,12R,14R,16E,18R,19R,20S,21S)-19, 21-Dihydroxy-22-{(2S,2′R,5S,5′S)-5′-[(1R)-1-hydroxyethyl]-2, 5′-dimethyloctahydro-2, 2′-bifuran-5-yl}-4, 6,8, 12, 14, 18, 20-heptamethyl-11-oxido-9-oxodocosa-10, 16-dienoic acid (Ionomycin) was purchased from Yeasen Biotechnology Co., Ltd. (Shanghai, China). An in situ cell death detection kit was acquired from Roche (Nutley, NJ, USA). We obtained a cell MPTP detection kit from GenMed Scientifics Inc. (Arlington, MA, USA), and a Cell Counting Kit-8 (CCK-8) was acquired from Dojindo Molecular Technologies, Inc. (Kumamoto, Japan). 5, 5',6, 6'-Tetrachloro-1, 1,3, 3'-tetraethylbenzimidazolyl-carbocyanine iodide (JC-1), Mito-Tracker Green, and 4, 6-diamidino-2-phenylindole (DAPI) were obtained from Beyotime Biotechnology (Beijing, China). Mito-Tracker Deep Red FM, Hoechst 33342, CellROX Green reagent and pHrodo^TM Red SE were purchased from Molecular Probes (Carlsbad, CA, USA). A FITC Annexin V Apoptosis Detection Kit II was acquired from BD Biosciences Pharmingen (San Jose, CA, USA). Human IL-4 and IFN-γ ELISA kits were obtained from NeoBioscience Technology Co., Ltd. (Shenzhen, China). Antibodies against Gata-3 (D13C9), T-bet/TBX21 (D6N8B), caspase-3, caspase-9, cleaved caspase-3 and cleaved caspase-9 were acquired from Cell Signaling Technology (Beverly, MA, USA). Antibodies against CD4, IL-4 and IFN-γ were obtained from Abcam (Cambridge, UK). Antibodies against cytochrome c, HSP 60 and actin were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Super ECL reagent was obtained from HaiGene (Harbin, China). All secondary antibodies were acquired from Bioss Inc. (Beijing, China).

Male C57BL/6J apoE^-/- mice (6-8 weeks old) were obtained from Beijing Vital River Laboratory Animal Technology Co., Ltd., (Beijing, China) and housed in a specific pathogen-free environment. The mice were fed a Western-type diet (WD, 0.25% cholesterol, 19.5% casein, 21% fat, 34.16% sugar) for 12 weeks to develop advanced atherosclerotic plaques in the aortic root. The mice were randomly separated into one of two groups: the Control group or the SDT group (n=10 per group). SDT group mice were subjected to ultrasonic radiation for 15 min at 2 h following an injection of ALA (60 mg/kg) into the caudal vein. After SDT treatment, the mice were fed a normal diet. All animal experiments were performed in accordance with the regulations of the Harbin Medical University Human Ethics Committee. The use and care of the animals conformed to the US National Institutes of Health Guide for Care and Use of Laboratory Animals.

The human Jurkat E6.1 T cell line was acquired from American Type Culture Collection (ATCC, Manassas, VA, USA). Jurkat E6.1 cells were cultured at a density of 2×10⁶ cells/mL in RPMI 1640 medium containing 10% FBS, 20 units/mL penicillin and 20 µg/mL streptomycin and maintained at 37°C in a humidified atmosphere containing 5% CO₂. For activation, Jurkat cells were treated with 1 mg/mL PHA for 24 h, washed, and subsequently incubated in the presence of PMA (25 ng/mL) plus ionomycin (1 µg/mL) for another 24 h.

Cells were exposed to ultrasound as described previously. Briefly, the ultrasonic generator, transducer and power amplifier used in this study were designed and assembled by the Harbin Institute of Technology (Harbin, China). The customized ultrasonic transducer (diameter: 3.5 cm; resonance frequency: 1.0 MHz; duty factor: 10%; repetition frequency: 100 Hz) was placed in a degassed water bath 30 cm away from the Petri dishes. The exposure time was 5 min.

Vessel parameters of mice were evaluated through hematoxylin and eosin (H&E) staining. All slices were subjected to H&E staining according to routine histopathological methods.

Apoptotic cells in mouse aortic root sections were detected using an in situ cell death detection kit according to the manufacturer’s instructions. Nuclei were counterstained with DAPI. For in situ efferocytosis assays, cryosections were subjected to immunofluorescent staining of macrophages and TUNEL staining of apoptotic cells simultaneously. All lesions were observed under an Olympus microscope.

The cytotoxicity of ALA was determined by the CCK-8 assay according to the manufacturer's instructions. Concisely, Jurkat cells (2×10⁵ cells/mL) were seeded into 96-well plates, and 0-10 mM ALA was added for 48 h in the dark. Then, 10 µL of CCK-8 solution was added into each well and incubated at 37°C for 1 h. The absorbance at 450 nm was measured using a microplate reader.

To investigate the intracellular metabolic kinetics of ALA-PpIX, Jurkat cells were rinsed three times with phosphate-buffered saline (PBS) and incubated with 1.0 mM ALA in serum-free RPMI 1640 for different time intervals in the dark at 37°C. The cells were then rinsed and fixed with 4% paraformaldehyde for 30 min. The fluorescence of ALA-PpIX in Jurkat cells was observed using an Olympus microscope equipped with a filter with a 405-nm excitation wavelength and a 630-nm emission wavelength. The fluorescence intensity of ALA-PpIX in Jurkat cells was measured with a multimode microplate reader at an excitation wavelength of 405 nm and an emission wavelength of 630 nm. For determination of the subcellular localization of ALA-PpIX, Jurkat cells were co-stained with 200 nM Mito-tracker Green for 30 min and 1 µg/mL Hoechst 33342 for 20 min at 37°C in the dark after incubation with 1.0 mM ALA for 4 and 6 h. Subsequently, the Jurkat cells were visualized by laser scanning confocal microscopy (LSCM).

We stained cells using a FITC Annexin V Apoptosis Detection Kit II for the assessment of cell apoptosis and necrosis by flow cytometry according to the instructions of the supplier.

Seeded cells (6×10⁵ cells/mL) were harvested, gently rinsed with cold PBS, re-suspended in 100 µL 1x binding buffer with 5 µL of Annexin V-FITC and 5 µL of propidium iodide (PI), and then incubated for 15 min in the dark at room temperature. Immediately afterward, the cells were analyzed with a FACSCalibur flow cytometer (BD Biosciences). Data analysis was performed using FlowJo software. For proper statistical analysis, more than 10, 000 cells per group were counted.

The production of intracellular ROS was determined using CellROX Green reagent. Briefly, cells were seeded in six-well plates at 2×10⁵ cells/mL and incubated with CellROX Green reagent (5 µM) for 30 min after sonication. The cells were fixed in 4% formaldehyde solution for 20 min in the dark before detection. The fluorescence intensity of the CellROX Green reagent was measured using a multimode microplate reader with excitation and emission wavelengths of 485 and 520 nm, respectively. Fluorescence images were captured with a fluorescence microscope.

For phagocytosis assays, Jurkat cells from each group were collected and labeled with 100 ng/mL pHrodo^TM Red SE for 30 min with light protection, washed with PBS, resuspended in the supernatant from each group, and then added to bone marrow-derived macrophages (BMDM). After co-incubation with the Jurkat cells for 1 h, phagocytes were detached from Petri dishes, stained with Percp-anti-CD11b for 1 h on ice and then washed thoroughly with PBS. Cells were analyzed with a FACSCalibur flow cytometer immediately and assessed with FlowJo software.

The MMP was determined by staining with the JC-1 fluorescent probe according to the manufacturer's protocol. Jurkat cells were pre-treated with NAC for 1 h after ALA incubation and then exposed to ultrasound. Staining was performed using 10 mg/mL JC-1 for 20 min at 37°C in the dark immediately after ultrasonic treatment. The fluorescence intensity was measured by a multimode microplate reader at an excitation wavelength of 525 nm and an emission wavelength of 590 nm for red fluorescence and an excitation wavelength of 490 nm and an emission wavelength of 530 nm for green fluorescence. Images were then obtained using a fluorescence microscope.

MPTP opening in Jurkat cells was detected using a calcein/Co²⁺ assay with a cell MPTP detection kit according to the manufacturer’s instructions. Briefly, seeded cells (2×10⁵ cells/mL) were rinsed three times with PBS, washed twice with reagent A from the kit, and then incubated with an intermixture of reagent B and C at 37°C for 20 min with light protection. The cells were then stained with DAPI for 20 min in the dark and washed twice with reagent A. Subsequently, the cells were visualized by LSCM.

Whole-cell lysates and mitochondrial and cytosolic fractions were extracted as described previously [23]. The protein samples were mixed with 5x SDS loading buffer, separated via 8%, 10% or 12% SDS-PAGE gels, and electrotransferred to PVDF membranes at 300 mA. After blocking in 6% skim milk for 2 h, the membranes were incubated at 4°C overnight with primary antibodies against caspase-3 (1: 1000), caspase-9 (1: 1000), cleaved caspase-3 (1: 1000), cleaved caspase-9 (1: 1000), cytochrome c (1: 400), T-bet (1: 1000), Gata-3 (1: 1000), actin (1: 2000), and HSP 60 (1: 500). Then, the membranes were rinsed and incubated with HRP-conjugated IgG secondary antibodies (1: 3000) at room temperature for 1 h. The membranes were then rinsed and detection was performed using a BIO-RAD ChemiDoc^TMMP Imaging System (Universal Hood III, Bio-Rad Laboratories, Inc., Hercules, CA, USA) using Super ECL reagent. Actin was used as a loading control for the whole-cell lysates or cytosolic fraction, and HSP 60 was used for the mitochondrial fraction. Densitometric values were quantified using Image Lab software (version 6.0, Bio-Rad).

For cytochrome c translocation analysis, after ALA-SDT treatment, seeded cells were stained with 100 nM Mito-Tracker Deep Red FM for 30 min, fixed with 4% paraformaldehyde for 30 min, and then blocked with 3% bovine serum albumin (BSA) for 30 min at room temperature. Then, the cells were washed and incubated with a rabbit polyclone primary antibody against cytochrome c (1: 500) overnight at 4°C. Subsequently, the cells were washed thoroughly, incubated with a goat anti-rabbit IgG Alexa Fluor 488-conjugated secondary antibody for 1 h and then stained with DAPI for 20 min at 37°C in the dark. Finally, the samples were visualized by LSCM.

The frozen lesion sections of mouse aortic roots were subjected to immunofluorescence staining according to standard procedures. Briefly, cryosections were blocked with 10% goat/rabbit serum for 30 min at 37°C and then incubated overnight at 4°C with primary antibodies against CD4 (1: 500), IFN-γ (1: 500), IL-4 (1: 400), T-bet (1: 500), and Gata-3 (1: 500), followed by co-incubation with FITC-conjugated and TRITC-conjugated secondary antibodies for 1 h and counterstaining with DAPI for 20 min at 37°C in the dark. Finally, the cryosections were visualized using fluorescence microscopy.

For detection of the levels of cytokines, the cell supernatant was collected after treatment. The levels of IFN-γ and IL-4 were measured with ELISA kits according to the manufacturer’s instructions.

All the experiments were repeated at least three times independently. The results are presented as the mean ± standard error of the mean (SEM) values. For the analysis of significant differences between two groups, Student’s two-tailed t-test was applied. The differences among three or more groups were analyzed via one-way analysis of variance (ANOVA) followed by the Student–Newman–Keuls test when appropriate. Statistical evaluation was performed using Statistical Analysis System (SAS) software version 9.2 (SAS institute, Cary, NC). Statistical significance was determined by a P value < 0.05.

We first investigated the effects of ALA-SDT on AS in WD-fed apoE^-/- mice. H&E staining of aortic roots revealed significant reduction of the plaque and necrotic core area in the SDT treated mice (P< 0.001 vs. the Control group, Fig. 1A). We used a TUNEL assay to detect apoptotic cells in consecutive sections of the mouse aortic root. As shown in Fig. 1B, ALA-SDT significantly induced apoptosis in atherosclerotic plaques (P< 0.001 vs. the Control group). Furthermore, the percentage of CD4⁺ apoptotic cells among total apoptotic cells was significantly increased in the SDT group (61.3 ± 4.5% vs. 26.6 ± 2.0%, P< 0.001). Efferocytosis of apoptotic cells prevents necrotic core formation and leads to the resolution of inflammation. Indeed, we observed significantly enhanced efferocytosis in SDT treated plaques (P< 0.001 vs. the Control group, Fig. 1C). With further histological analysis, we observed that ALA-SDT reduced the number (per 2000 cells within the atheroma) of IFN-γ-producing CD4⁺ cells (241.8 ± 22.5 vs. 51.3 ± 6.9, P< 0.001, Fig. 1D) and CD4⁺T-bet⁺ cells (292.2 ± 27.1 vs. 100.2 ± 8.0, P< 0.001, Fig. 1E). Furthermore, the numbers of IL-4-producing CD4⁺ cells (199.0 ± 11.2 vs. 366.8 ± 23.3, P< 0.001, Fig. 1F) and CD4⁺Gata-3⁺ cells (65.8 ± 6.3 vs. 227.5 ± 27.7, P< 0.001, Fig. 1G) increased in the plaques of SDT treated mice compared with those of Control mice.

To investigate more cellular specific phenotype, we utilized ALA-SDT on human Jurkat cells in vitro. We first compared the effects of different ultrasonic intensities on cell viability using flow cytometry with Annexin V and PI double staining in vitro. As shown in Fig. 2A and B, there was no significant alteration in apoptotic ratio among different intensity cells. When the intensity of ultrasound was greater than 0.8 W/cm², the necrotic ratio gradually rose as the amount of ultrasonic intensity increased (P< 0.05, the 1.2 W/cm² group vs. the ultrasound untreated group). We subsequently investigated the cytotoxicity of ALA, the biological precursor of protoporphyrin IX (PpIX), on Jurkat cells via a CCK8 assay. As shown in Fig. 2C, the viability was reduced significantly from 88.6 ± 1.6% (2.0 mM, P< 0.001 vs. the ALA untreated group) to 64.0 ± 1.5% (10 mM). Therefore, the application of 1 mM ALA with 0.8 W/cm² ultrasonic intensity represented the optimal treatment parameters.

The fluorescence intensity of 1 mM ALA-PpIX observed under a fluorescence microscope and detected by a multimode microplate reader increased over time and peaked at 6 h in Jurkat cells and then faded over time (Fig. 3A, B). We also observed the subcellular localization of ALA-PpIX in Jurkat cells. As shown in Fig. 3C, the red PpIX fluorescence partially overlapped with the green fluorescence emitted by the mitochondrial probe after incubation with ALA for 4 h and was almost exclusively distributed in the mitochondria after 6 h of incubation when intracellular ALA-PpIX fluorescence intensity reached the maximum.

To investigate whether ALA-SDT elevates the level of ROS in Jurkat cells, the fluorescence intensity of CellROX was measured via fluorescence microscopy and a multimode microplate reader. As shown in Fig. 4B, relative ROS levels did not dramatically increase in either the ALA alone group (118.8 ± 2.6%) or ultrasound alone group (137.3 ± 4.1%) compared with the Control group (100 ± 2.6%), but increased significantly in the SDT group (461.1 ± 17.2%, P< 0.001 vs. the Control). The results of fluorescence microscopy were consistent with those of the multimode microplate reader (Fig. 4A).

Cell apoptosis and necrosis were determined by flow cytometry with double staining of Annexin V and PI. Cells in the lower-right quadrant (Annexin V⁺/PI^-) represent early apoptotic cells, and those in the upper-right quadrant (Annexin V⁺/PI⁺) represent late apoptotic cells. As shown in Fig. 5A, the apoptotic ratio, particularly the late apoptotic ratio, of the SDT treated cells (45 ± 1.4%) was considerably higher compared to that of Control cells (17.2 ± 0.7%, P< 0.001). Moreover, pretreatment with the ROS scavenger NAC attenuated apoptosis induced by ALA-SDT (21.9 ± 1.1%, P< 0.001 vs. the SDT) effectively. Similarly, enhanced apoptosis was greatly suppressed by pretreatment with the pan-caspase inhibitor Z-VAD-FMK (23.3 ± 1.7%, P< 0.001 vs. the SDT group) as well. The necrotic ratio did not significantly change among the experimental groups. We subsequently examined the phagocytosis of Jurkat T cells in different groups by BMDM and found that apoptotic cell uptakes increased to 63.8 ± 3.2% in the SDT group (P< 0.001 vs. the Control group, Fig. 5B and C). NAC pretreatment (28.5 ± 2.0%, P< 0.001 vs. the SDT group) significantly prevented SDT-induced phagocytosis.

As shown in Fig. 6A, most cells exhibited green fluorescence after SDT treatment, representing dissipation of the ΔΨ_m, whereas cells in the SDT plus NAC group displayed orange-red fluorescence due to a reversion of the ΔΨ_m to normal levels. Consistent with this pattern, the results obtained using the multimode microplate reader showed that when cells were exposed to ALA-SDT, ΔΨm dissipated rapidly, and the ratio of red/green fluorescence intensity of JC-1 was notably increased in the SDT plus NAC group (11.2 ± 1.1) compared to the SDT group (5.5 ± 0.6, P< 0.001, Fig. 6B). We next examined whether SDT could open the MPTP using the calcein/cobalt approach via LSCM. As shown in Fig. 6C, the calcein-dependent green fluorescence was markedly quenched by Co2+ in the SDT group. NAC pretreatment significantly prevented SDT-induced MPTP opening, as indicated by a remarkable reversion of dull calcein fluorescence to a bright signal.

Translocation of pro-apoptotic factor Bax from cytoplasm to mitochondria acts as a trigger of outer mitochondrial membrane permeabilization. Therefore, we analyzed the expression of Bax and cytochrome c in cytoplasm and mitochondria of Jurkat cells by Western blotting. A significant induction in mitochondria-localized Bax and a corresponding reduction in cytosolic Bax were observed in cells upon SDT, whereas the opposite pattern for cytochrome c was observed (Fig. 7A). NAC pretreatment, as expected, inhibited this SDT-induced Bax translocation to mitochondria as well as cytochrome c release into cytosol. Similar anti-apoptotic effects were observed for pretreatment with the singlet oxygen scavenger NaN₃. Cytochrome c release from mitochondria into cytosol was further supported by immunofluorescence staining; cells upon SDT modulation showed diffuse green fluorescence after ALA-SDT treatment (Fig. 7B), and this fluorescence was inhibited by pre-incubation with NAC and NaN₃. Moreover, we assessed the activation of caspase-3 and caspase-9 by Western blotting. ALA-SDT treatment notably induced the activation of both caspases. Pre-treatment with NAC or Z-VAD-FMK inhibited caspase-3 and caspase-9 activation induced by ALA-SDT (Fig. 7C).

As shown in Fig. 8A and B, the level of IFN-γ in supernatants of PMA plus ionomycin-activated Jurkat cells exposed to ALA-SDT (461.2 ± 55.9 pg/mL) was lower than that of control cells (917 ± 42.7 pg/mL, P< 0.001 vs. the SDT group), but the cells treated with ALA-SDT (12.9 ± 1.3 pg/mL) secreted a higher level of IL-4 than the Control group (3.3 ± 0.5 pg/ mL, P< 0.001 vs. the SDT group). To further explore the mechanism of ALA-SDT-regulated Th cell polarization, changes in the expression of T-bet as well as Gata-3 are shown in Fig. 8C. ALA-SDT downregulated T-bet expression but upregulated Gata-3 expression in vitro. All of these effects were blocked by NAC or NaN₃ (Fig. 8A-C).

Intriguing results from the Canakinumab Anti-Inflammatory Thrombosis Outcomes Study (CANTOS) recently showed that the IL-1 beta monoclonal antibody Canakinumab significantly reduced the incidence of adverse cardiovascular events in patients with myocardial infarction compared with the incidence in those treated with placebo, providing clinical trial evidence and indicating the significance of inflammation in AS pathology [24, 25]. Further investigation to identify novel efficient regimens that lessen inflammation in AS, therefore, is of great importance. We demonstrated in the present study that ALA-SDT efficiently regresses atherosclerotic plaque and improves the inflammatory environment within the atheroma by enhancing cytotoxic effects on Th cells, subsequently stimulating efferocytosis and facilitating a shift in the Th1/Th2 balance toward Th2 cells.

Our previous studies demonstrated that ALA-SDT induces apoptosis of macrophages and foam cells and stabilizes AS plaques in animal models [18-20]. In addition to macrophages, lymphocytes , primarily CD4-positive T cells are also vital participants throughout the course of atherosclerosis development [4]. In this study, we found that ALA could be metabolized into PpIX and accumulated in the mitochondria of Jurkat cells by 6 h after administration of 1 mM ALA (Fig. 3). The pattern of subcellular localization and the total level and peak time of PpIX accumulation in these cells was the same as that in the THP-1-derived foam cells [19], suggesting that ALA-SDT may simultaneously modulate both major types of cell function in atherosclerotic plaques and providing further evidence to utilize ALA-SDT in the lesion area on multi-dimensional targets.

In addition to synthesis rate, the specific subcellular localization patterns and tissue characteristics of the photosensitizer are important determinants of the efficacy of sonodynamic therapy [26-28]. We previously reported that ALA-derived PpIX mainly accumulates in the mitochondria of THP-1 macrophages and foam cells to induce mitochondrial damage [18, 19]. In the present study, we found that PpIX derived from the exogenous administration of ALA consistently aggregated in the mitochondria and activated the mitochondrial pathway of apoptosis in Jurkat cells. SDT-induced cellular damage seems to mainly depend on the specific subcellular localization patterns of the sonosensitizers. Activation of the sonosensitizer enhanced ROS formation in the binding site, determining the initially damaged subcellular organelles.

Expansion of the necrotic core contributes to the formation of vulnerable plaques and plaque disruption. Adequate phagocytosis of apoptotic cells by macrophages prevents necrotic core expansion and curbs atherosclerosis progression. However, if apoptotic cells are not efficiently removed by efferocytosis, these cells may undergo secondary necrosis, leading to larger necrotic cores and inflammation [29, 30]. In this study, ALA-SDT not only induced CD4+ cell apoptosis but also stimulated macrophage-mediated phagocytosis of these apoptotic cells concurrent with the regression of lesion size and necrotic core size (Fig. 1A, B and C and Fig. 5). ALA-SDT directly or indirectly resolves inflammation via multiple mechanisms. First, ALA-SDT induces Th cell elimination, resulting in a decrease in inflammatory mediators, including IFN-g. Second, enhanced efferocytosis inhibits post-apoptotic necrosis and prevents dead cells from releasing inflammatory contents. Third, the recognition of apoptotic cells activates anti-inflammatory mediators and anti-inflammatory eicosanoids. Finally, ALA-SDT enhances the Th2 response, thereby suppressing vascular inflammation.

Accumulating evidence indicates that oxidative stress acts as an obligate second messenger, and the oxidative status of Th cells controls their effector functions, including cytokine production and subset differentiation [31-34]. Investigations both in vivo and in vitro have shown that ROS can preferentially promote Th2 skewing [35-37]. Moreover, natural antioxidant vitamin E suppresses Th2 responses upon T cell activation [38]. These studies highlight that ROS is an essential determinant of T helper differentiation and can dictate T helper bias. In the current study, ALA-SDT was shown to facilitate a shift in the Th1/ Th2 balance toward Th2 in vivo and in vitro (Fig. 1D-G and Fig. 8). Here, we report for the first time the direct effects of ALA-SDT on naïve T cell differentiation. Moreover, our results demonstrated that ALA-SDT stimulated Th2-predominant responses, including modulation of the associated transcription factors that may be dependent on the production of ROS, particularly singlet oxygen (Fig. 8). Notably, recent studies have shown that ROS also function as intercellular messengers, and the extracellular oxidative microenvironment can influence cytokine generation in T cells. Activated macrophages can act as antigen-presenting cells for T cell priming, and the diffusion of ROS, such as H₂O₂, from macrophages to the T cells is necessary for this process during their close contact phase [31]. Thus, ROS derived from ALA-SDT in macrophages may be involved in the activation and polarization of Th cells. Further investigation regarding the intercellular signaling is undergoing in our laboratory.

Excessive levels of ROS lead to detrimental reactions with proteins, nucleic acids and lipids, playing an indispensable role in various cellular processes, including cell life and death decisions [39]. It is now clear that when exposed to ultrasound, the porphyrin-based sensitizers generate ROS, causing cytotoxicity [40]. Consistent with this finding, we found that ALA-SDT increased the intracellular ROS levels in Jurkat cells (Fig. 4). In addition, pre-incubation with NAC, an ROS scavenger, reversed ALA-SDT-mediated mitochondrial apoptotic signaling and T cell differentiation (Fig. 5-5-8). The ultrasound field is heterogeneous and ultrasonic waves can be reflected, and attenuated through a medium [41] and human tissue towards an atherosclerotic plaque. Cells at different distances or depths are exposed to different ultrasound fields. The different energies of SDT produce different ROS levels, thereby causing two distinct effects simultaneously, namely, pro-apoptotic effects and stimulation of Th1/Th2 shifting. We found that ALA-SDT induced Th cell apoptosis and enhanced macrophage-mediated phagocytosis of those apoptotic cells (Fig. 1A and B). Because Th1 predominates in advanced atherosclerotic plaques [42], ALA-SDT induced mostly Th1 cells apoptosis, but stimulated Th2-predominant responses, which could subsequently inhibit the Th1 response. Taken together, the results show that ALA-SDT plays a dual and synergistic role in modulating the Th1/Th2 equilibrium. In future, we plan to conduct more in vivo and in vitro experiments to distinguish the thresholds and spectra associated with distinct ROS level as well as their equivalent biological effects.

In summary, our results confirm that the optimization of ALA-SDT exerts cytotoxic effects on Th cells by selectively activating the mitochondrial apoptotic signaling pathway and enhancing macrophage-mediated phagocytosis. By inhibiting a Th1 response and favoring a Th2 response, ALA-SDT activated anti-inflammatory reactions, mainly through the generation of ROS. Our studies provide new insights into the mechanisms of SDT treatment as a novel therapeutic strategy for combating atherosclerosis.

This study was supported by the Major Research Equipment Development Project of National Natural Science of China (81727809), the State Key Project of National Natural Science of China (81530052), the National Natural Science Foundation of China (81371709) and the Major State Basic Research Development Program of China (2017YFB0403800). This study was also supported by the Graduate Student Innovative Research Project of Harbin Medical University (YJSCX2015-68HYD).

The authors declare that they have no conflicts of interest to disclose.

Y. Yang, Y. Liu and X. Chen contributed equally to this work.