Roswell Park Memorial Institute 1640 (RPMI-1640) medium was obtained from Lonza (Basel, Switzerland). Fetal bovine serum (FBS) was purchased from Gibco via Invitrogen (Lofer, Austria). [³H]-Cholesterol was bought from Perkin Elmer Life Sciences (Vienna, Austria) and pioglitazone was from Molekula (Munich, Germany). Albumin faction V, fatty acid free BSA was obtained from Carl Roth (Graz, Austria). Bisphenol A diglycidyl ether (BADGE), resazurin sodium salt, water soluble unesterified cholesterol, apolipoprotein (apo) A1, phorbol 12-myristate 13-acetate (PMA), lactacystin, calpeptin, chloroquine, and cycloheximide were provided by Sigma-Aldrich (Vienna, Austria). Falcarindiol was isolated from the roots and rhizomes of the traditional Chinese herbal drug Radix Notopterygii, with a purity over 95% as determined by High Performance Liquid Chromatography (HPLC) (Zschocke et al., 1997; Liu et al., 2014). Cathepsin B, L, D, and K Activity Fluorometric kits were purchased from BioVision via THP Medical Products GmbH (Vienna, Austria). Cathepsin S activity assay kit (fluorometric) was bought from Abcam (Cambridge, UK). Antibodies were obtained from the following companies: ABCA1, ABCG1, SR-B1 from Novus Biologicals via THP Medical Products GmbH (Vienna, Austria), anti-actin antibody from MP biologicals (Illkirch, France), horseradish peroxidase-conjugated goat anti-rabbit secondary antibody from New England Biolabs (Frankfurt, Germany), horseradish peroxidase conjugated goat anti-mouse secondary antibody from Upstate (Millipore, Vienna, Austria).

Human monocytic THP-1 cells were acquired from ATCC® and grown in suspension in RPMI-1640 medium supplemented with 2 mM glutamine, 100 U/mL benzylpenicillin, 100 mg/mL streptomycin, and 10% FBS in T175 flasks. For experiments, differentiation was induced by seeding THP-1 monocytes in 96-, 24-, or 6- well plates under treatment of 200 nM PMA for 72 h (seeding cell density of 0.2 × 10⁶ cells per mL). All of the tested compounds were dissolved in dimethyl sulfoxide (DMSO) and stored at −20°C. Control cells were always treated with an equal volume of solvent (DMSO) and its final concentrations did not exceed 0.1%.

The cholesterol efflux assay (Wang et al., 2015) was adapted from previous studies (Chawla et al., 2001; Chinetti et al., 2001). Briefly, THP-1-derived macrophages were incubated for 24 h with the compounds at the indicated concentrations [solvent vehicle (DMSO), falcarindiol (1–20 μM), the PPARγ agonist pioglitazone (10 μM) as positive control] together with [³H]-cholesterol in serum free medium supplemented with 0.1% BSA and 10 μg/mL unesterified cholesterol. On the next day, cells were washed twice with pre-warmed PBS and incubated with the same compounds with or without 10 μg/mL apo A1 or 1% human plasma dissolved in serum-free medium for 6 h. Extracellular as well as intracellular radioactivity were quantified with scintillation counting. The apo A1- as well as human plasma-mediated cholesterol efflux was calculated as follows:

The viability of THP-1-derived macrophages was assessed by resazurin conversion. The assay was performed as previously described (Wang et al., 2015) by quantification of the conversion of the low fluorescent resazurin to high fluorescent resorufin.

THP-1 cells were seeded in 6-well plates and differentiated into macrophages with 200 nM PMA for 72 h. The macrophages were treated with the indicated compounds at the corresponding time points, as described in detail in each figure legend. Cellular proteins were extracted using NP40 buffer (150 mM NaCl; 50 mM HEPES (pH 7.4); 1% NP40; 1% protease inhibitor Complete™ (Roche); 1% phenylmethylsulfonyl fluoride (PMSF); 0.5% Na₃VO₄; 0.5% NaF) and quantified with the Bradford method. For western blot analysis, an equal amount of protein (20 μg) was separated via SDS-PAGE and transferred onto a PVDF membrane. The membrane was blocked for 1 h at room temperature with 5% non-fat milk, and then incubated at 4°C overnight with primary polyclonal antibody against ABCA1, ABCG1 or SR-B1 (1:500) or actin (1:10,000). After being washed for three times (15 min each time), the membranes were incubated with a secondary antibody [horseradish peroxidase-conjugated goat anti-rabbit (1:500) or anti-mouse (1:10,000)] for 1 h at room temperature. The signals were detected and visualized using a LAS-3000 luminescent image analyzer (Fujifilm, Düsseldorf, Germany) using AIDA image analyzer 4.06 software (Raytest, Sprockhövel, Germany). The indicated protein band intensity was normalized to that of actin.

THP-1 cells were seeded in 6-well plates and differentiated into macrophages with a 200 nM PMA treatment for 72 h. The macrophages were treated with the indicated compounds at the indicated time points, which are described in detail in the figure legends. Total cellular RNA was extracted from the treated THP-1 macrophages using peqGOLD Total RNA kits (PeqLab, Linz, Austria) according to the manufacturer's instructions. One microgram of total RNA was used for cDNA synthesis according to the protocol of the High Capacity cDNA Reverse Transcription Kit together with RNase Inhibitor (Applied Biosystems). LightCycler® 480 SYBR Green I Master kit (Roche) was used for quantitative real-time PCR (qRT-PCR) with 40 ng cDNA from each sample for triplicate measurements. Amplification cycles were detected using the LightCycler 480 system (Roche). ABCA1 (HS_ABCA1_1_SG QuantiTect primer assay, Cat.no.: #QT00064869, QIAGEN) and 18S (Hs_RRN18S_1_SG QuantiTect Primer assay, Cat.no.: #QT00199367, QIAGEN) primers were used, and the quantification was performed with the ΔC_T method.

The cathepsin activity assays were performed according to the manufacturer's instructions. In general, THP-1-derived macrophages were cultured for 24 h with solvent control (DMSO) or falcarindiol (10 μM). After incubation, cells were washed once with cold PBS and lysed with chilled cell lysis buffer. Protein concentration was determined using the Bradford method. Hundred micrograms of protein per reaction in 50 μL of cell lysis buffer was used for the activity measurement. Fifty microliters of reaction buffer and 2 μL of the respective cathepsin substrate were added to 50 μL cell lysates. The assay plate was incubated at 37°C for 1 h protected from light. The fluorescence was quantified with Tecan Infinite 200® Pro plate reader (Vienna, Austria).

Statistical analysis was performed with data acquired from at least three independent experiments using GraphPad Prism software version 4.03 (GraphPad Software Inc., La Jolla, CA, USA). Figures with bar graphs represent mean ± SD. To determine statistical significance, one- or two- way analysis of variance (ANOVA) were performed and p < 0.05 was considered significant.

In a previous study, we showed that falcarindiol activates PPARγ in HEK-293 cells transfected with a PPARγ-responsive luciferase reporter (Atanasov et al., 2013). Considering that PPARγ is a known regulator of cholesterol efflux (Chawla et al., 2001; Chinetti et al., 2001), we aimed to investigate whether falcarindiol can promote cholesterol efflux in THP-1-derived macrophages. Since apo A1, the nascent and lipid free form of HDL, is the most significant acceptor for cholesterol effluxed from macrophages (Du et al., 2015), we first studied cholesterol efflux mediated by apo A1. Falcarindiol enhanced apo A1-mediated cholesterol efflux in a concentration-dependent manner when applied at 1–20 μM (without effect on cell viability; Figure 1B) with a half maximal effective concentration (EC₅₀) of 5.8 μM (Figure 1C). Moreover, 10 μM falcarindiol also significantly enhanced 1% human plasma-mediated cholesterol efflux (Figure 1D) as plasma containing HDL particles served as cholesterol acceptors in the process of cholesterol efflux. Pioglitazone, a PPARγ agonist known to upregulate cholesterol efflux (Ozasa et al., 2011), was used as positive control both in presence of apo A1 and human plasma (Figures 1C,D).

The transmembrane proteins ABCA1, ABCG1, and SR-B1 are transporters mediating cholesterol efflux (Duffy and Rader, 2006). Therefore, we investigated protein expression of these three transporters upon treatment with falcarindiol in THP-1-derived macrophages. As evident in Figure 2A, ABCA1 protein level significantly increased in response to falcarindiol (10 μM). At the same time, there was no significant upregulation of ABCG1 (Figure 2B) or the bi-directional transporter SR-B1 (Figure 2C; Stangl et al., 1999). Next, we investigated whether increased ABCA1 protein level is associated with an increase in ABCA1 mRNA level. ABCA1 mRNA level indeed significantly increased upon falcarindiol (10 μM) treatment (Figure 3B, left).

In macrophages, enhanced ABCA1 mRNA level might result from PPARγ and subsequent LXR activation (Chawla et al., 2001; Chinetti et al., 2001). We, therefore, asked whether the increased ABCA1 mRNA level induced by falcarindiol might be due to PPARγ-activation. To check this possibility, we used the PPARγ antagonist, BADGE (50 μM), to block the activity of PPARγ in THP-1-derived macrophages (Hinz et al., 2003). Indeed, BADGE blocked the falcarindiol-mediated increase of ABCA1 protein as well as mRNA level (Figures 3A,B).

Interestingly, the ABCA1 mRNA up-regulation by falcarindiol was significantly lower than the mRNA induction caused by the PPARγ agonist pioglitazone (Figure 3B, on the left), while the induction of ABCA1 protein level by falcarindiol and pioglitazone was equally potent (Figure 3A, on the left). This inconsistency indicated that an additional effect of falcarindiol might exist, which could enhance ABCA1 protein level besides the increase of ABCA1 mRNA level. To explore this possibility, we measured the rate of ABCA1 protein degradation in the presence and absence of falcarindiol. THP-1-derived macrophages were incubated with falcarindiol or solvent vehicle (DMSO) for 24 h and were then further treated with cycloheximide (140 μM) to block de novo protein synthesis (Arakawa et al., 2009). As evident in Figure 4, ABCA1 protein degradation was significantly inhibited in the presence of falcarindiol showing that the compound interferes with the ABCA1 protein turnover rate.

To further explore how falcarindiol inhibits ABCA1 protein degradation, we first investigated, which protein degradation pathway could be affected by falcarindiol in THP-1-derived macrophages. As presented in Figure 5A, the application of lactacystin, an ubiquitin proteasome inhibitor, failed to affect ABCA1 protein level in our experimental setting. However, the application of calpeptin (a calpain inhibitor) and chloroquine (a lysosome inhibitor) (Figures 5B,C) increased ABCA1 protein level, similarly to falcarindiol. Falcarindiol had an additive effect on ABCA1 protein level upon co-treatment with calpeptin, indicating that its mechanism of action is likely different from the calpain inhibitor calpeptin. However, there was no difference observed with or without chloroquine in the presence of falcarindiol, suggesting that falcarindiol might interfere with protein degradation in the lysosome, mimicking the effect of chloroquine (Figure 5C).

The lysosomal degradation pathway is a well-known regulator of ABCA1 protein turnover (Neufeld et al., 2001; Santamarina-Fojo et al., 2001; Mizuno et al., 2011) and cathepsins are well established proteases mediating the lysosomal proteolysis in general (Turk et al., 2012; Appelqvist et al., 2013). Therefore, we investigated whether the proteolytic activity of lysosomal cathepsins is affected by falcarindiol. Figures 6A–E shows that among the tested cathepsins (B, D, K, L, S) the activity of cathepsin B, S, and K were significantly suppressed by 35, 14, and 8%, respectively. Overall, these data indicate that falcarindiol leads to the suppression of ABCA1 degradation and inhibits lysosomal cathepsins.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.