Combination of Hydroxyl Acetylated Curcumin and Ultrasound Induces Macrophage Autophagy with Anti-Apoptotic and Anti-Lipid Aggregation Effects

While atherosclerotic plaque rupture is a major cause of acute cardiovascular events [1], there is currently a lack of effective treatments for this condition. Macrophages, which accumulate in atherosclerotic plaques, play a crucial role in the occurrence of atherosclerosis [2,3]. Therefore, therapies targeting macrophages have been recognized as beneficial for plaque stability [4]. Accumulating data have indicated that macrophage apoptosis effectively delays the progression of early-stage atherosclerosis [5]. As we have previously confirmed, sonodynamic therapy (SDT) effectively induces the apoptosis of atherogenic macrophage in vitro [6,7,8,9,10,11,12,13]; furthermore, we have previously detected the atherosclerotic plaque stabilization that is associated with increased macrophage apoptosis in vivo after SDT [14]. However, the roles of apoptosis in different stages of atherosclerosis are controversial. While macrophage apoptosis plays a protective role by attenuating plaque formation in early-stage atherosclerotic lesions, some studies have indicated that the occurrence of apoptosis plays a negative role by accelerating plaque inflammation and necrosis in advanced atherosclerotic plaque [5,15]. Additionally, autophagy is gradually becoming the focus of research on atherosclerosis [16].Extensive studies have indicated that the induction of macrophage autophagy plays a protective role in atherosclerosis by preventing cytosolic lipid accumulation and suppressing macrophage foam cell formation [17,18,19]. During the study of the effect of SDT on macrophages, we found that SDT effectively induces macrophage autophagy, suggesting that SDT might play a protective role in atherosclerosis progression by modulating autophagy; as such, autophagy activation provides a promising avenue for treating atherosclerosis.

SDT is a noninvasive therapeutic strategy that is now considered a promising treatment for malignant tumors [20]. Through the synergistic effects of low-intensity ultrasound and a sonosensitizer, SDT effectively produces cytotoxic reactive oxygen species (ROS) [21], which have been reported to play crucial roles in a wide variety of cellular events [22]. In addition, the potential mechanism underlying the cellular responses induced by SDT appears to be highly dependent on this ROS generation [23]. Sonosensitizers, a group of photosensitive materials derived from photodynamic therapy exhibit minimal inherent cytotoxic, are responsible for the sonodynamic damage that occurs after ultrasonic irradiation [12]. Our previous data have showed that hydroxyl acetylated curcumin (HAC) is a relatively safe and effective sonosensitizer at a concentration of 5.0 μg/mL, which clearly generates ROS via ultrasound activation; in addition, we have demonstrated that ROS are indispensable factors involved in the induction of macrophage apoptosis during SDT [13]. Many studies have reported that ROS play a vital role in the activation of autophagy as well as in apoptosis during SDT [24], and some studies have indicated that autophagy induction effectively prevents apoptosis via ROS production [25,26]. However, whether autophagy is involved in HAC-SDT and the corresponding mechanism in macrophages has not yet been thoroughly investigated. Therefore, further research is needed.

In the present study, we aimed to explore whether the autophagy activation was regulated by ROS during HAC-SDT, and we further investigated the mechanism of HAC-SDT action on macrophages, which are considered the most important cell type in all stages of atherosclerosis [27].

Human THP-1 monocytes were purchased from the American Type Culture Collection (Manassas, VA, USA). The cells were cultured in RPMI 1640 medium (HyClone, Logan, UT, USA) containing 10% fetal bovine serum (HyClone, Logan, UT, USA), 20 μg/mL penicillin and 20 μg/mL streptomycin (Sigma-Aldrich Co., St. Louis, MO, USA). They were specifically maintained at 37°C in a humidified incubator containing 5% CO₂. For the experiments, THP-1 monocytes in the log phase were seeded in 96-well plates and 35-mm Petri dishes at a density of 1.0×10⁵ cells/mL with 100 ng/mL phorbol-12-myristate-13-acetate (La Jolla, CA, USA) for 72 h for differentiation into macrophages.

The SDT device used in this study was assembled at the Harbin Institute of Technology (Harbin, China), and the parameters of the ultrasound exposure system were as the same as those previously reported [13]. First, THP-1 macrophages were incubated with 5.0 μg/mL HAC for 4 h in complete medium at 37 °C in the dark. For the control and ultrasound groups, an equivalent volume of medium was used to replace the HAC. Next, macrophages in the ultrasound and SDT groups were exposed to ultrasound at an intensity of 0.5 W/cm² for 5, 10 or 15 min. Then, the cells were subjected to different analyses following culture in fresh medium for an additional period. The experiments using emodin (5.0 μg/mL) and curcumin (5.0 μg/mL) were performed under treatment conditions identical to those used for the HAC-SDT experiments.

For the inhibition studies, 1 mM N-acetyl-l-cysteine (NAC; Sigma-Aldrich Co., St Louis, MO, USA), 20 μM z-VAD-FMK (z-VAD; BioVision Inc., USA), or 10 mM 3-methyladenine (3-MA; Sigma-Aldrich Co., St. Louis, MO, USA) were incubated together with HAC.

Macrophage viability was detected using a Cell Counting Kit (CCK)-8 colorimetric assay (Beyotime Biotechnology, Inc., Beijing, China). THP-1 monocytes were seeded in a 96-well plate and incubated for 72 h. At 6 h after the HAC-SDT treatment, the medium in each well was replaced with 100 µL of CCK-8 solution (diluted in RPMI 1640 medium to 10% (v/v)). Following incubation for 2 h at 37 °C in the dark, the absorbance of each well was measured at 450 nm using a Model 680 microplate reader (Varian Australia Pty Ltd., Australia).

Intracellularly generated ROS were detected using the fluorescence intensity of 2', 7'-dichlorofluorescein (DCF) and MitoSOX. For this purpose, THP-1 macrophages were cultured on 35-mm Petri dishes. Six hours after the HAC-SDT treatment, the treated cells were incubated for 20 min with 5.0 μM MitoSOX (Invitrogen, Eugene, OR, USA), a mitochondrial superoxide indicator, and then stained with 20 μM DCFH-DA (Applygen Technologies Co. Ltd., Beijing, China) for another 20 min. After washing, the fluorescence intensity of MitoSOX was measured using a multimode microplate reader at excitation and emission wavelengths of 510 nm and 580 nm, respectively. The fluorescence intensity of DCF was measured at excitation and emission wavelengths of 488 nm and 525 nm, respectively. The fluorescence intensity in the control group was assumed to represent 100% cell viability. The cells were also imaged using laser scanning confocal microscopy (LSCM; LSCM 510 Meta; Zeiss, Gottingen, Germany).

The apoptosis and necrosis of treated macrophages were assessed using an Annexin-V-FITC apoptosis kit (Franklin Lakes, NJ, USA). Six hours after HAC-SDT, the cells were collected by trypsinization and resuspended in 100 μL of 1×binding buffer. Then, the cells were stained with 5 μL of Annexin V and 10 μL of propidium iodide (PI) for 15 min at room temperature in the dark. Then, 400 μL of 1×binding buffer was added, and the samples were analyzed by flow cytometry

To detect autophagy, 1 μg/mL acridine orange (AO; Sigma-Aldrich Co., St. Louis, MO, USA) was added to living cells at the indicated time after different treatments. The cells were then incubated for 30 min in the dark, washed with phosphate-buffered saline (PBS), and harvested by trypsinization. Green (510-530 nm) and red (650 nm) fluorescence emission was recorded from 1×10⁴ cells excited with blue (488 nm) excitation light. The cells and corresponding data were then analyzed using a FACSCalibur flow cytometer (BD Biosciences) and CellQuest software (BD Biosciences), respectively.

The macrophages were incubated with 5.0 μg/mL HAC for 4 h, carefully washed twice with PBS, and then stained for 20 min with 0.4 µM Lyso-Tracker Green, 5.0 µM Golgi-Tracker Green, 0.2 µM ER-Tracker Green (lysosome, Golgi and endoplasmic reticulum probes, respectively; Invitrogen, Eugene, OR, USA) and 0.1 µM Mito-Tracker Green (Beyotime Biotechnology, Inc., Beijing, China). After washing, the colocalization of HAC with various fluorescent probes in THP-1 macrophages was analyzed using LSCM. HAC fluorescence was detected using excitation and emission wavelengths of 405 nm and 476 nm, respectively. The Lyso-Tracker Green, Golgi-Tracker Green, ER-Tracker Green, and Mito-Tracker Green fluorescence intensities were measured using excitation and emission wavelengths of 488 nm and 525 nm, respectively.

Cells were seeded in glass-bottom cell culture dishes. To detect the translocation of BAX and cytochrome C and the activation of autophagy, we measured the colocalization of Mito-Tracker Green with BAX and cytochrome C, as well as the fluorescence of LC3-B. The THP-1 macrophages were incubated with 0.1 µM Mito-Tracker Green for 10 min at 37°C at the indicated time after HAC-SDT and were then washed twice with PBS. Next, they were fixed using 4% paraformaldehyde for 10 min and permeabilized with 1% Triton X-100 for 5 min. After being washed, the cells were blocked with 3% bovine serum albumin (BSA) to prevent nonspecific antibody binding and were incubated at 4°C overnight with antibodies against BAX, cytochrome C and LC3-B (diluted at 1:400, Cell Signaling Technology, Inc., USA). The cells were then washed twice with PBS and incubated with diluted (1:200) tetramethyl rhodamine isothiocyanate (TRITC)- or fluorescein isothiocyanate (FITC)-labeled secondary rabbit antibodies in 1% BSA in the dark for 1 h. After washing, the cells were imaged using LSCM. Mito-Tracker Green and FITC fluorescence was measured at excitation and emission wavelengths of 488 nm and 525 nm, respectively. TRITC fluorescence was measured at excitation and emission wavelengths of 555 nm and 570 nm, respectively.

As a marker for lysosomes and autolysosomes, MDC (Sigma-Aldrich Co., St. Louis, MO, USA) was used to observe the development of autophagy in cultured cells. At the indicated incubation time after the different treatments, the cells were incubated with 1 mL of RPMI 1640 medium and 50 μM MDC for 30 min at 37°C in the dark. Subsequently, they were washed twice with PBS and then visualized and imaged using a fluorescence microscope. MDC fluorescence was measured at an excitation wavelength of 380 nm with a 530 nm emission filter.

To measure the expression of BAX, cytochrome C, and cleaved PARP in the mitochondria and the cytosol, mitochondrial and cytosolic proteins were isolated using a mitochondrial/cytosol fractionation kit (Beyotime Biotechnology, Inc., Beijing, China). The cytosolic and nuclear proteins were obtained using a nuclear/cytosolic fractionation kit (Beyotime Biotechnology, Inc., Beijing, China). In brief, cells were collected by trypsinization at the indicated times, washed with PBS, incubated with ice-cold cytosolic separation buffer, agitated and lysed on ice for 15 min. The cytosolic, nuclear and mitochondrial fractions were then harvested by a series of centrifugation steps in accordance with the manufacturer's instructions.

After the designated treatments, the cells were lysed with RIPA buffer, and the protein concentrations in the lysates were measured using a bicinchoninic acid kit (Beyotime Biotechnology, Inc., Beijing, China). The protein samples (50 µg) were then separated by SDS-PAGE and transferred to polyvinylidene fluoride membranes. After being blocked with 5% skim milk powder in Tris-buffered saline-Tween 20 for 1 h, the membranes were labeled with specific primary antibodies at 4°C overnight. After being washed, the membranes were treated with alkaline phosphatase IgG secondary antibody for 2 h. Then, the antibody labeling was detected using enhanced chemiluminescence reagent. The areas of the protein bands were quantified using a Bio-Rad ChemiDoc EQ densitometer and Bio-Rad Quantity One software (Bio-Rad Laboratories, Hercules, CA, USA).

Antibodies against BAX, cytochrome C, cleaved PARP, HSP60, PCNA, beclin 1, p62, LC3 II, mTOR, p-mTOR, AKT, p-AKT were purchased from Cell Signaling Technology (Cell Signaling Technology, Inc., USA). An antibody against β-actin was purchased from Protein Tech Group (Wuhan, China). The alkaline phosphatase IgG secondary antibody was purchased from ZhongShan Company (Beijing, China). All of antibodies were diluted at a ratio of 1:1000.

Dil-ox-LDL (Yiyuan Biotech., Guangzhou, China) was used to track the cellular uptake of ox-LDL. The macrophages were incubated with Dil-ox-LDL (50 μg/mL) for 24 h at 37°C in the dark. At a predetermined time after the different treatments, the cells were washed twice with PBS and then imaged using a fluorescence microscope. The fluorescence of Dil-ox-LDL was measured at excitation and emission wavelengths of 488 nm and 565 nm, respectively. The fluorescence intensity in the control group was assumed to represent 100% cell viability.

Oil red O (Sigma, St. Louis, MO, USA) staining was used to observe intracellular lipid accumulation. At a predetermined time after the different treatments, the macrophages were incubated for 72 h with RPMI 1640 medium and 50 μg/mL ox-LDL (Yiyuan Biotech., Guangzhou, China). Then, the cells were washed twice with PBS, fixed using 4% paraformaldehyde for 30 min, and dehydrated with 60% isopropanol for 2 min. Next, the cells were stained with filtered 0.3% oil red O for 10 min and then with hematoxylin for 1 min to counterstain the cell nuclei. Subsequently, they were washed twice with 60% isopropanol and ddH₂O and then imaged using a microscope to assess the formation of macrophage foam cells. The obtained images were subsequently processed using Image-Pro Plus software.

All experiments were independently replicated at least 3 times. Differences between groups were determined by ANOVA, and the results are presented as the mean ± standard deviation (SD). P values <0.05 were considered statistically significant.

By measuring cell viability after incubation with various concentrations of HAC, we found that 5.0 μg/mL HAC did not significantly decrease cell viability and may be the optimal concentration for SDT. After 4 h of incubation with 5.0 μg/mL HAC and 15 min of ultrasound irradiation, cell viability significantly decreased (54 ± 8%). In contrast, treatment with 15 min of ultrasound irradiation alone had no influence on cell viability (99 ± 3%) (Fig. 1A). We also found that the sonodynamic effect of 5.0 μg/mL HAC was much greater than that observed using an identical concentration of emodin or curcumin (Fig. 1A). Fig. 1B shows that ROS inhibition by pre-treatment with NAC (85 ± 9%) significantly rescued cell death caused by HAC-SDT. Furthermore, autophagy inhibition via pre-treatment with 3-MA (47 ± 4%) effectively promoted cell death, suggesting that autophagy exerts an anti-apoptotic effect during HAC-SDT.

ROS production was assessed using two fluorescent probes: the intracellular ROS probe DCF and the mitochondrial ROS-specific probe MitoSOX. The fluorescence intensity of both DCF and MitoSOX was significantly increased in the HAC-SDT group (306 ± 16% and 253 ± 21%, respectively) (Fig. 1C). These results indicated that the mitochondria were sites of ROS generation during HAC-SDT. Both of these increases in ROS generation were blocked by pre-treatment with NAC before HAC-SDT with significant reduction of the fluorescence intensity of DCF and MitoSOX to 221 ± 15% and 209 ± 27%, respectively.

Herein, we identified that HAC-SDT could significantly induce apoptosis, which was the main cause of cell death. The flow cytometric analysis of Annexin V/PI staining revealed that HAC-SDT induced a significant level of apoptosis (13 ± 2%). Furthermore, the induced apoptosis could be blocked by the ROS scavenger NAC (7 ± 1%) and the broad-spectrum inhibitor z-VAD (8 ± 1%). To further investigate the involvement and role of autophagy in apoptosis during HAC-SDT, we used the autophagy inhibitor 3-MA. The results showed that inhibiting autophagy could further increase the level of apoptosis (20 ± 1%) induced by HAC-SDT (Fig. 1D).

To confirm the sub-cellular localization of HAC in THP-1 macrophages, the cells were loaded with HAC for 4 h and stained with various organelle-specific, fluorescent probes. As shown in Fig. 2, the position of HAC (shown in blue) corresponded well with the positions of Lyso-Tracker Green, Golgi-Tracker Green, ER-Tracker Green, and Mito-Tracker Green (shown in green). The bright blue regions represent an overlap, indicating that HAC was found in the mitochondria and in the ER, lysosomes, and Golgi apparatus.

To further explore the mechanism of HAC-SDT-induced apoptosis, the pro-apoptotic factor BAX and key apoptosis-related proteins were monitored. We measured the intracellular localization of BAX using Western blot, and the results showed that BAX was predominantly localized in the cytosol in the control cells. At 2 h after HAC-SDT, the BAX level was increased in the mitochondria and decreased in the cytosol, and this effect was further enhanced at 4 h and 6 h after HAC-SDT. Meanwhile, cytochrome C levels were significantly increased in the cytosol at 2 h, 4 h, and 6 h after HAC-SDT, and this increase was accompanied by a decrease in the mitochondrial cytochrome C level (Fig. 3A, 3B). Additionally, the activation of PARP was significantly increased at 2 h, 4 h, and 6 h after HAC-SDT (Fig. 3C).

Fluorescence staining was performed to monitor the translocation of BAX and cytochrome C. As shown in Fig. 4A, in the control cells, the Mito-Tracker Green fluorescence exhibited little overlap with the red fluorescence of BAX, indicating that BAX was largely distributed in the cytosol. However, yellow fluorescence resulting from overlapping green and red fluorescence signals was clearly observed after HAC-SDT. This result indicated that BAX was redistributed from the cytosol to the mitochondria. For cytochrome C, untreated control cells displayed yellow fluorescence, signifying that cytochrome C was well distributed in the mitochondria; in contrast, distinctly segregated red and green fluorescence signals were clearly observed after HAC-SDT, suggesting that cytochrome C had moved from the mitochondria to the cytosol after HAC-SDT. The translocation of BAX and cytochrome C and the increased expression of caspase-related proteins during HAC-SDT were both prevented by treatment with NAC (Fig. 4B). These findings suggested that BAX and cytochrome C translocation contributed to the apoptosis induced by HAC-SDT. Furthermore, the translocation of BAX and cytochrome C and the activation of the caspase cascade were regulated by ROS.

To explore whether autophagy was activated during HAC-SDT, we assessed the relative fluorescence level of LC3-B, which is a specific marker of autophagy. Fig. 5A shows that autophagy was activated in a time-dependent manner. The fluorescence intensity of LC3-B increased with time and peaked at 3 h (2.10 ± 0.12) after HAC-SDT. The data suggested that while autophagy could be activated under the same conditions that induced apoptosis, it occurred earlier than apoptosis.

To quantitatively analyze the activation of autophagy during HAC-SDT, we used flow cytometry to measure the fluorescence of cells labeled with AO. We found that autophagy was increased by 20 ± 1% in the HAC-SDT group; this percentage decreased to 12 ± 1% and 14 ± 1% when 3-MA and NAC were added, respectively (Fig. 6B).

We used MDC staining to visualize the development of autophagy, and similar results were also obtained by this method. As shown in Fig. 6C, faint MDC fluorescence was detected in the control group (100 ± 9%), whereas MDC fluorescence was strongly detected in the HAC-SDT group (203 ± 15%). However, the augmented fluorescence intensity was partially reversed by pre-treatment with 3-MA (153 ± 10%) or NAC (145 ± 8%) (Fig. 6C). These results indicated that HAC-SDT effectively induced autophagy in THP-1 macrophages via ROS generation.

To further confirm the activation of autophagy during HAC-SDT, we measured the expression levels of autophagy-specific proteins. Fig. 5B shows that the beclin 1 and LC3 II levels increased significantly along with a decrease in the level of p62 from 30 min to 6 h after HAC-SDT; the maximum effects on these three markers were observed at 3 h. Additionally, the increase in LC3 II induced by HAC-SDT (2.33 ± 0.22) could be blocked by 3-MA (1.71 ± 0.18), an inhibitor of the early phase of autophagy (Fig. 6A). According to the literature, ROS play a crucial role in SDT, and in this study, we found that ROS were an important effector in the induction of apoptosis. Thus, we evaluated whether the activation of autophagy was also related to ROS generation. Fig. 7A shows that the changes in beclin 1, p62 and LC3 II expression induced by HAC-SDT were all inhibited by pre-treatment with NAC. Additionally, the reduced phosphorylation of both AKT and mTOR by SDT was also blocked by pre-treatment with NAC (Fig. 7A). These data demonstrated that autophagy was activated through the PI3K/AKT/mTOR pathway, which was regulated by generation of ROS during HAC-SDT.

To detect the potential relationship between autophagy and the lipid uptake of macrophages during HAC-SDT, we analyzed the Dil-ox-LDL fluorescence intensity. As shown in Fig. 7B, compared with the control group (100 ± 12%), the ultrasound (96 ± 10%) and HAC groups (96 ± 8%) showed no significant differences; the Dil-ox-LDL fluorescence intensity was significantly decreased in the HAC-SDT group (36 ± 8%) but could be recovered by pre-treatment with 3-MA (62 ± 8%) and NAC (54 ± 8%). These results suggested that HAC-SDT effectively weakened the ability of macrophages to uptake lipids and that this trend was inhibited by 3-MA and NAC. Thus, lipid uptake and autophagy were closely related events, and both of them were effectively prevented by ROS reduction.

The macrophage foam cell model was generated to investigate the possible roles of HAC-SDT-induced macrophage autophagy in lipid accumulation. The status of lipid accumulation in the different groups was determined using oil red O staining. As shown in Fig. 7C, there was no obvious difference in oil red O-positive area in the ultrasound (98 ± 9%) and HAC groups (97 ± 8%) compared with the control group (100 ± 7%). Conversely, the oil red O-positive area in the SDT group (44 ± 7%) was significantly decreased, indicating reduced lipid accumulation. Nevertheless, the reduction was partially reversed by pre-treatment with 3-MA (66 ± 9%) and NAC (68 ± 7%), corresponding well with our other results. These findings indicated that HAC-SDT not only effectively impaired the capacity of macrophages to take up lipids but also induced macrophage autophagy, both of which were blocked by 3-MA and NAC in a similar manner.

In this study, the therapeutic effect of HAC-SDT was evaluated in THP-1 macrophages. We furthered our understanding of the mechanism underlying the induced apoptosis by showing that HAC-SDT can activate PI3K/AKT/mTOR pathway dependent macrophage autophagy with an anti-apoptotic and anti-lipid aggregation effects under the same condition of apoptosis induction. The potential mechanisms of SDT-induced autophagy and lipid aggregation attenuation were both evaluated with respect to ROS generation. These findings suggest that HAC-SDT might be an efficient approach for treating atherosclerosis.

Based on our previous data, either 5.0 μg/mL HAC alone or 15 min of 0.5 W/cm² ultrasound alone had little effect on cell viability, while their combination significantly decreased cell viability, which was the optimal effect [13]. In addition, we observed that 5.0 μg/mL HAC showed a more satisfactory sonodynamic effect under low-intensity ultrasound than did the same concentrations of emodin and curcumin, indicating that HAC was the preferred sonosensitizer. Meanwhile, the apoptosis rate increased significantly in the HAC-SDT group, and the expression levels of caspase-related proteins increased in a time-dependent manner, peaking at 6 h after HAC-SDT. In addition, the apoptosis induced by HAC-SDT was prevented by a broad-spectrum inhibitor, z-VAD, indicating that the caspase cascade played a pivotal role in this process.

Studies have shown that the site of sonosensitizer localization is critical for SDT due to the very short half-life and diffusion distance of certain radical products derived from the ultrasound-activated sonosensitizer [28]. Therefore, the sub-cellular localization of HAC in THP-1 macrophages was evaluated in the current study. Our results showed that HAC localized in the ER, lysosomes, Golgi apparatus and mitochondria. These findings clearly indicated that the mitochondria incurred apoptotic damage and corresponded well with the findings of related apoptosis studies [6,10]. While the present study demonstrated that HAC-SDT is capable of causing direct damage to mitochondria, the sonodynamic effect of HAC on the ER, lysosomes, and Golgi apparatus needs further investigation.

Our previous findings indicated that HAC-SDT induced THP-1 macrophage apoptosis accompanied by increased ROS generation [13]. Consistent with the mitochondria being the main site for intracellular ROS generation, we found that the ROS generated by HAC-SDT diffused throughout the cells, including in the mitochondria. Moreover, the induced apoptosis could be rescued by pre-treatment with the ROS scavenger NAC; this result supported the predominant role of ROS in HAC-SDT-induced apoptosis. Due to the previously reported preferential accumulation of HAC in the mitochondria of THP-1 macrophages and decrease in mitochondrial membrane potential, we evaluated what types of changes take place in the mitochondria during HAC-SDT-induced apoptosis.

Mitochondria play a key role in apoptosis by regulating the permeabilization of the outer mitochondrial membrane and releasing cytochrome C; numerous studies have suggested that this process is regulated by BAX translocation [29]. Several studies have reported that BAX is a crucial pro-apoptotic factor that can undergo a conformational shift and become integrated into the mitochondrial membrane, leading to cytochrome C release from the intermembrane space of the mitochondria into the cytosol [30]. This event is known to activate the caspase cascade, causing irreversible apoptosis [31]. Here, we observed that the translocation of BAX and cytochrome C and the subsequent activation of the caspase cascade were induced after HAC-SDT. However, all of these events were blocked by NAC, suggesting that ROS are upstream factors in the mitochondrial-caspase pathway, which is consistent with previous findings [32,33].

With increasing knowledge about therapies targeting atherosclerosis, many studies suggest that the induction of macrophage autophagy plays a protective role in plaque stabilization [34,35,36]. In addition, a growing body of data suggests that autophagy and apoptosis are activated under the same conditions, yet the crosstalk and interplay between them appear to be complex and controversial [37,38]. In this study, it was intriguing to find that the apoptosis rate in the HAC-SDT group was much higher with the 3-MA pre-treatment than without, which demonstrated that autophagy inhibition aggravated HAC-SDT-induced cell apoptosis. Thus, we speculated that HAC-SDT was able to induce autophagy as a cytoprotective mechanism against apoptosis. Given the controversial role of apoptosis in atherosclerosis progression that apoptosis is benefit for early stage plaque while bad for advanced plaque, HAC-SDT-induced apoptosis maybe benefit for early stage plaque. Moreover, HAC-SDT-induced autophagy with anti-apoptotic effect can overcome disadvantages of apoptosis for advanced plaque, and is benefit for the whole progress of atherosclerosis corresponding well with the findings of previous studies [25,26]. However, little is currently known about the specific mechanism of autophagy and its effect on macrophages during HAC-SDT.

LC3 II is a well-known and reliable biomarker of autophagy, and its level directly varies with autophagy because LC3 II anchors the membranes of autophagosomes [39]. Recent studies have indicated that SDT leads to increased autophagy activity by causing more LC3 II/I conversion [40]. Similarly, we observed that autophagy activity increased in a time-dependent manner via the accumulation of LC3 II in THP-1 macrophages during HAC-SDT. In addition, the increased conversion of LC3 II/I was accompanied by a decrease in p62 and an increase in beclin 1 expression, and the effects on these classic autophagy markers peaked at 3 h after HAC-SDT. Furthermore, the MDC and AO staining for acidic vesicular organelles at 3 h after HAC-SDT corresponded well with the visible autophagy phenomenon.

Our study contributed toward elucidating the mechanism of autophagy activation via HAC-SDT. Various molecular mechanisms are involved in the regulation of autophagy, including those of AMPK, PI3K and p53 [41]. In this study, we focused on the PI3K/AKT/mTOR pathway. We discovered that the phosphorylation of mTOR and AKT decreased after HAC-SDT but increased with a 3-MA pre-treatment, which suggested that HAC-SDT activated autophagy through the PI3K/AKT/mTOR pathway. Whether other autophagy-related pathways also participate in this process will be studied in the future.

Recently, numerous studies have indicated that ROS are also involved in autophagy activation, aside from starvation, oxidized lipids and other factors [42,43]. ROS are known to play a crucial role in the mechanisms underlying the biological effects induced by SDT [10,33]. In the present study, we found that autophagy was inhibited by the ROS scavenger NAC, yielding almost the same results as 3-MA. Simultaneously, the changes in autophagy-related proteins mentioned above were also blocked by NAC, coinciding well with the results of a previous study [25]. These findings suggested that the initiation of autophagy through the PI3K/AKT/mTOR pathway was most likely regulated by ROS generation, which is believed to be a key factor of HAC-SDT.

As a conserved catabolic process that involves the transport of cellular organelles and proteins, autophagy has been shown to play a variety of important roles related to cell survival, differentiation, and development [44]. Accumulating evidence indicates that macrophage autophagy exerts a protective effect in atherosclerosis by attenuating lipid accumulation, suggesting that the induction of macrophage autophagy might be a potential target for plaque stabilization [16]. In this study, the Dil-ox-LDL binding assay results revealed an autophagy-dependent lipid uptake capacity in THP-1 macrophages, which implied that HAC-SDT-induced autophagy significantly reduced the lipid uptake capacity of THP-1 macrophages. Based on this finding, we further examined the lipid aggregation in macrophages using oil red O staining, and observed that lipid aggregation and uptake were similarly prevented by autophagy induction. Correspondingly, these events were also regulated by NAC. All these findings highlighted the fact that the autophagy induced by HAC-SDT could effectively suppress macrophage foam cell formation in the development of atherosclerosis by regulating lipid aggregation, and as expected, ROS were the key to the entire process.

Overall, we demonstrated that as a preferential sonosensitizer, HAC could not only induce mitochondrial-caspase pathway dependent apoptosis, but also activate PI3K/AKT/mTOR pathway dependent autophagy under ultrasonic irradiation. Furthermore, HAC-SDT-induced autophagy exerted an anti-apoptotic effect and relieved lipid accumulation in macrophages. Therefore, this study revealed a possible relationship between apoptosis and autophagy induced by HAC-SDT in macrophages and provided a potential therapeutic strategy for atherosclerosis.

This study was supported by the National Natural Science Foundation of China (81000688, 81271734, 81571833), the Heilongjiang Provincial Science Foundation (H2015006), the Foundation of Science and Technology Innovation Talent of the Harbin Science and Technology Bureau (2015RAQXJ100), the Wu Liande Youth Science Foundation of Harbin Medical University (WLD-QN1104), the Postdoctoral Science-Research Development Foundation of Heilongjiang Province (LBHQ12049), and the China Postdoctoral Science Foundation (20090460911).

None.

L. Zheng and Y. Li contributed equally to this work.