Amorfrutin C Induces Apoptosis and Inhibits Proliferation in Colon

Article pubs.acs.org/jnp

Amorfrutin C Induces Apoptosis and Inhibits Proliferation in Colon Cancer Cells through Targeting Mitochondria Christopher Weidner,†,∥ Morten Rousseau,†,∥ Robert J. Micikas,‡ Cornelius Fischer,† Annabell Plauth,† Sylvia J. Wowro,† Karsten Siems,§ Gregor Hetterling,§ Magdalena Kliem,† Frank C. Schroeder,‡ and Sascha Sauer*,†,⊥ †

Otto Warburg Laboratory, Max Planck Institute for Molecular Genetics, D-14195 Berlin, Germany Boyce Thompson Institute and Department of Chemistry and Chemical Biology, Cornell University, Ithaca, New York 14853, United States § AnalytiCon Discovery GmbH, D-14473 Potsdam, Germany ⊥ CU Systems Medicine, University of Würzburg, D-97080 Würzburg, Germany ‡

S Supporting Information *

ABSTRACT: A known (1) and a structurally related new natural product (2), both belonging to the amorfrutin benzoic acid class, were isolated from the roots of Glycyrrhiza foetida. Compound 1 (amorfrutin B) is an efficient agonist of the nuclear peroxisome proliferator activated receptor (PPAR) gamma and of other PPAR subtypes. Compound 2 (amorfrutin C) showed comparably lower PPAR activation potential. Amorfrutin C exhibited striking antiproliferative effects for human colorectal cancer cells (HT-29 and T84), prostate cancer (PC-3), and breast cancer (MCF7) cells (IC50 values ranging from 8 to 16 μM in these cancer cell lines). Notably, amorfrutin C (2) showed less potent antiproliferative effects in primary colon cells. For HT-29 cells, compound 2 induced G0/G1 cell cycle arrest and modulated protein expression of key cell cycle modulators. Amorfrutin C further induced apoptotic events in HT-29 cells, including caspase activation, DNA fragmentation, PARP cleavage, phosphatidylserine externalization, and formation of reactive oxygen species. Mechanistic studies revealed that 2 disrupts the mitochondrial integrity by depolarization of the mitochondrial membrane (IC50 0.6 μM) and permanent opening of the mitochondrial permeability transition pore, leading to increased mitochondrial oxygen consumption and extracellular acidification. Structure−activity-relationship experiments revealed the carboxylic acid and the hydroxy group residues of 2 as fundamental structural requirements for inducing these apoptotic effects. Synergy analyses demonstrated stimulation of the death receptor signaling pathway. Taken together, amorfrutin C (2) represents a promising lead for the development of anticancer drugs.

C

found in the roots of Glycyrrhiza foetida Desf. (Leguminosae), a perennial herb endemic to southern Spain (Andalusia) and northwest Africa (Morocco and Algeria), as well as in G. acanthocarpa native to Southern Australia.9,11 It recently was reported that amorfrutins A and B (compound 1) strongly reduce insulin resistance and liver steatosis in metabolic mouse models7,8,12 via selective activation of the peroxisome proliferator-activated receptor (PPAR) nuclear receptor class.13 Amorfrutin B (1) was found to be the most efficient binding molecule of PPARγ among this class of natural products. In addition, amorfrutin A has been reported to inhibit common anti-inflammatory pathways,14,15 in part via modulation of PPARγ.16 In the present study, it was shown that amorfrutin C (2) 2Hydroxy-4-methoxy-3,5-bis(3-methylbut-2-enyl)-6-phenethylbenzoic acid, strongly inhibited proliferation to kill cancer cells. Interestingly, in contrast to the PPAR agonist 1, compound 2

ancer is the third leading cause of death globally, accounting for about 8 million deaths and 13 million new cases per year.1,2 Colorectal cancer (CRC) is the third most common form of cancer, with 1 million new cases and more than 600 000 deaths per year.2 Current chemotherapeutic approaches are hampered by often severe toxic side effects, emergence of drug resistance, and frequent relapse.3 The discovery and development of new promising anticancer agents therefore addresses an urgent need. A large proportion of anticancer agents in current clinical use are based on natural products or their synthetic analogues,4,5 and purified natural products as well as crude extracts have become a recent focus of nutrition research aiming to develop functional food and nutraceuticals with significant health benefits.6 We recently reported the characterization of some members of the amorfrutin family.7,8 These natural products feature a planar 2-hydroxybenzoic acid core with isoprenyl, benzyl, or alkyl residues and a methoxy or hydroxy group at position C-4.9 Amorfrutins were first extracted from the bastard indigo bush Amorpha f ruticosa;10 however, amorfrutins have also been © XXXX American Chemical Society and American Society of Pharmacognosy

Received: January 26, 2015

A

DOI: 10.1021/acs.jnatprod.5b00072 J. Nat. Prod. XXXX, XXX, XXX−XXX

Journal of Natural Products

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did not efficiently activate PPARγ,8 a nuclear receptor that has been proposed as a target for treating both type 2 diabetes and also cancer.9 This study further explored the apoptotic effects of 2 in HT-29 colon carcinoma cells and provided mechanistic insights into potential modes of action of 2, which go beyond interaction with nuclear receptors of the PPAR family.

Table 1. 1H NMR Data (CD3OD, 500 MHz) and 13C NMR Data (CD3OD, Data Deduced from HSQC and HMBC Spectra) proton 1 2 3 4 5 6 7 8 9 10 11 12 13 14 prenyl-1 prenyl-1 prenyl-1 prenyl-1 prenyl-1 prenyl-2 prenyl-2 prenyl-2 prenyl-2 prenyl-2 OMe

Chart 1. Chemical Structures of Compounds 1, 2, 2a, and 2b

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RESULTS AND DISCUSSION Isolation and Structure Elucidation. Amorfrutin C (2) was isolated from the roots of G. foetida as a new derivative of the known amorfrutin B (1) with an additional prenyl residue. Amorfrutin C gave a molecular formula of C26H32O4 as deduced by HRMS. The 1H NMR spectrum (Table 1) showed typical signals of an “AA′BB′C” system of a monosubstituted phenyl ring. From the COSY and HSQC spectra two prenyl groups and an aromatic methoxy group were deduced. Both prenyl moieties were found to be directly attached to a fully substituted aromatic ring with the aromatic rings linked by a CH2−CH2 bridge. The couplings observed in the HMBC spectrum allowed the positioning of the substituents at the fully substituted phenyl ring (Table 1). Particularly, 3JC,H couplings from the methylene groups of the prenyl moieties as well as from the methoxy group to C-4 allowed the placement of the latter between the two prenyl moieties. The other neighbors of the prenyl moieties, a hydroxy group and the ethylene chain, were placed by HMBC correlations to C-6 and to C-2, respectively. The substitution of the phenyl ring was completed by a carboxylic acid unit that showed no correlations in the HMBC spectrum. Antiproliferative Activity. Compounds 1 and 2 were tested for their inhibitory effects on the growth of a small panel of cancer cell lines. HT-29 and T84 colon carcinoma, PC-3 prostate cancer, and MCF7 breast cancer cells were treated with 1 and 2 for 72 and 96 h, respectively. Cell proliferation was determined by measurement of cellular DNA content (Table 2). Both amorfrutin compounds inhibited cancer cell proliferation, with compound 2 slightly more potent than 1, with IC50 values of 8 μM (HT-29), 11 μM (T84), 16 μM (PC3), and 14 μM (MCF7), respectively (Figure 1A), which are lower than the IC50 values observed for cisplatin used as a positive control. Treatment with 1 and 2 above the IC50 concentrations resulted in near-complete death of colon carcinoma cells, whereas the positive controls caused maximally 70% and 90% death (Table 2). Importantly, in primary colon cells, 2 showed clearly weaker antiproliferative effects when

a

2

3.13 m 2.72 m 7.14a 7.19a 7.09a 7.19a 7.14a 3.30 m 4.98 br t (6.5)

1′ 2′ 3′ 4′ 5′ 1″ 2″ 3″ 4″ 5″

1.61 1.73 3.30 5.18

s s m br t (6.5)

1.62 s 1.74 s 3.62 s

carbon 1 2 3 4 5 6 7 8 9 10 11 12 13 14 prenyl-1 prenyl-1 prenyl-1 prenyl-1 prenyl-1 prenyl-2 prenyl-2 prenyl-2 prenyl-2 prenyl-2 OMe COOH

2

1′ 2′ 3′ 4′ 5′ 1″ 2″ 3″ 4″ 5″

109.7 142.2 125.7 161.6 121.1 160.5 32.6 37.0 142.6 128.1 128.1 125.8 128.1 128.1 25.2 125.0 130.9 24.9 19.9 25.2 123.2 130.9 24.9 17.0 60.2 b

b

AA′BB′C system. Not observed in HMBC spectrum.

compared to colon cancer cells (details are shown in Figure 1A). Compound 2 Triggers Cell-Cycle Arrest and Apoptosis. Cellular proliferation is generally linked to cell cycle progression.17 Therefore, the effects of 2 were assessed on key cell cycle modulators by immunoblotting of proteins from HT29 cells that were treated with 10 μM 2 for 48 h. Compound 2 considerably induced the expression of p21/Cip1 and p27/ Kip1, two important cyclin-dependent kinase inhibitors. Compound 2 further reduced the expression of cyclins A2, D3, and E2, as well as the cyclin-dependent kinases (CDK) 2, 4, and 6 (Figure 1B). Additionally, the effects of 2 on the cell cycle were analyzed by flow cytometry. Compound 2-treated cells accumulated in the G0/G1 phase (89% vs 66% for 2 vs control) with concomitant reduction in the S (2% vs 11%) and the G2/M phases (7% vs 21%) (Figure 1C). These results indicated that the growth inhibitory effect of 2 for HT-29 colon carcinoma cells was likely a result of G0/G1 cell cycle arrest. In order to shed more light on the cellular effects of 2, apoptosis was further analyzed in HT-29 cells. In healthy cells, phosphatidylserine (PS) is generally restricted to the inner leaflet of the cell membrane, and the presence of large quantities of PS on the outer leaflet represents a hallmark of apoptosis.18 Treatment with 20 μM 2 for 48 h increased PS externalization from 8% to 44% (Figure 2A). Moreover, early activation of caspases (as markers of the signaling network to connect extrinsic and intrinsic stimuli with downstream apoptotic events) was measured in HT-29 cells.19 Treatment with 20 μM 2 for 2 h significantly induced the enzymatic activity of the effector caspases 3/7 by 2-fold (Figure 2B). B



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Table 2. Antiproliferative Effects of 1 and 2 in HT-29, T84, PC-3, and MCF7 Cancer Cellsa HT-29 (colon cancer) IC50 (μM) 1 2 cisplatin oxaliplatin 5-FU irinotecan etoposide

20 8.1 11 1.6 5.3 1.9 2.2

± ± ± ± ± ± ±

1 0.5 2 0.3 1.2 1.8 0.3

efficacy (%) 98 95 91 70 68 71 97

± ± ± ± ± ± ±

4 3 6 2 5 21 62

T84 (colon cancer) IC50 (μM) 13 11 15 1.1 6.2 1.2 3.0

± ± ± ± ± ± ±

1 1 1 0.3 10.7 0.4 0.3

PC-3 (prostate cancer)

efficacy (%) 99 99 87 87 76 82 90

± ± ± ± ± ± ±

3 3 3 4 18 5 3

IC50 (μM) 32 16 131 0.8 4.9 0.8 1.1

± ± ± ± ± ± ±

3 2 19 0.1 0.4 0.1 0.3

MCF7 (breast cancer)

efficacy (%) 95 95 n.d. 87 80 76 84

±5 ±5 ± ± ± ±

2 2 2 4

IC50 (μM) 33 ± 1 14 ± 1 >100 ± n.d. 0.4 ± 0.0 0.9 ± 0.1 0.9 ± 0.3 0.8 ± 0.2

efficacy (%) 94 92 n.d. 83 80 82 78

±3 ±3 ± ± ± ±

2 2 4 3

a Efficiency is the maximal observed induction of cell death after treatment relative to nontreated cells (set to 0%). HT-29 and PC-3 cells were treated for 72 h; MCF7 and T84 cells for 96 h. Treatment times were adjusted according to cell-specific variation in proliferation. n.d., not determined.

Figure 1. Antiproliferative activity of compound 2 in human cancer cell lines. (A) HT-29 and T84 colon carcinoma, PC-3 prostate cancer, MCF7 breast cancer, and CCD 841 CoN primary colon cells were treated with concentration series of 2 for 72 h (HT-29 and PC-3) and 96 h (T84, MCF7, and CCD 841 CoN), respectively. The relative number of cells was determined. Notably, in CCD 841 CoN primary colon cells, compound 2 (amorfrutin C) showed lower antiproliferative activity than in colon cancer cells. In primary colon cells, an IC50 value of ∼49 μM (and an efficacy of ∼87%) was determined, and in HT-29 colon cancer cells the IC50 value was ∼8 μM (and an efficacy of ∼96%). Data are expressed as means ± SD (n = 4). (B) Whole cell lysates were analyzed for the expression of the cell cycle regulating proteins p21, p27, cyclin A2, cyclin D3, cyclin E2, CDK2, CDK4, CDK6, and GAPDH. (C) Cell cycle analysis of HT-29 cells. Histograms (left) show one representative experiment for each treatment condition. Bar plots (right) show percentages of the cell population in apoptotic SubG1, G0/G1, S, and G2/M phases of the cell cycle and are expressed as means ± SEM (n = 3). *p ≤ 0.05, ***p ≤ 0.001 vs control.

Notably, 2 activated caspases 8 and 9 by 3-fold (Figure 2B), indicating activation of the extrinsic (death receptor mediated) as well as the intrinsic (mitochondrial mediated) pathway of apoptosis. Fluorescence microscopy validated cleavage of caspase 3 and further revealed formation of apoptotic bodies in treated HT-29 cells (Figure 2C). As expected, 2 additionally induced significant DNA fragmentation in these cells (Figure 2D). Consequently, treatment with 2 also induced cleavage of the chromatin-associated poly(ADP-ribose) polymerase (PARP) (Figure 2E); thus 2 efficiently activated the apoptotic cascade in human HT-29 colon carcinoma cells. In contrast to amorfurtin B (1), amorfrutin C (2) appeared to be only a low-affinity ligand for the peroxisome proliferator-

activated receptors. Competitive binding studies previously revealed affinity values of 9.1, 5.2, and 0.7 μM for PPARα, β/δ, and γ, respectively.8 Since PPARs are also involved in cell growth and differentiation,20 their potential role was investigated for the observed antiproliferative and apoptotic effects of 2. Transfection of short interfering RNA (siRNA) against all three PPAR subtypes reduced the gene expression of PPARα, β/δ, and γ by 73%, 83%, and 70%, respectively (Figure 3A). However, silencing of the PPARs in HT-29 cells showed no effects on inhibition of proliferation (Figure 3B) and apoptosis (Figure 3C) induced by compound 2, suggesting a PPARindependent mechanism. As knockdown of the PPARs was not complete, contribution to cellular effects owing to low-affinity C



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Figure 2. Activation of apoptosis in HT-29 colon carcinoma cells by compound 2. (A) Phosphatidylserine externalization of HT-29 cells was determined by flow cytometry of annexin V-FLUOS- and propidium iodide (PI)-stained cells. Scatter plots (left) show one representative experiment for each treatment condition. Bar plots (right) show percentage of cell populations in apoptosis, comprising early (annexin positive, PI negative) and late stage (annexin positive, PI positive) apoptotic events. Bars represent means ± SEM (n = 3). (B) HT-29 cells were treated with 30 μM 2 for 2 h. Enzymatic activation of caspases 2, 3/7, 6, 8, and 9 was determined by luminescent assays. Data are normalized to control treatment and are expressed as means ± SEM (n = 4). (C) HT-29 cells were treated with 30 μM 2 for 6 h. Cleavage of caspase 3 was visualized by fluorescence microscopy. Nuclei were stained with DAPI. Apoptotic bodies are marked with white arrows. (D) Effects of 2 on DNA fragmentation. Accumulation of DNA in the cytosol was determined by detection of BrdU-labeled DNA (left) and oligonucleosomes (right) using ELISA. Data are normalized to the control treatment and are expressed as means ± SEM (n = 4). (E) HT-29 cells were treated with 10 μM 2 for 48 h. Whole cell lysates were analyzed for protein expression of total and cleaved PARP. n.s. not significant, *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 vs control.

scriptome of these cells (Figure 4), indicating a mechanistic role of reactive oxygen species (ROS) and intrinsic apoptosis pathways for the effects of compound 2. ROS Formation Is Not Causal for Antiproliferative and Apoptotic Effects of Compound 2. Cancer cells are generally characterized by an imbalance of ROS, and increasing oxidative stress can activate apoptotic pathways.22 It was next asked if compound 2 leads to formation of ROS in HT-29 cells. To measure intracellular ROS formation during treatment with 2, the fluorescence of chloromethyl dichlorofluorescein (CMDCF) was detected in living HT-29 cells. Of note, 30 μM 2 induced significant accumulation of ROS during the first hours of treatment (Figure 5A). Co-treatment with common antioxidants prevented completely the formation of ROS in these cells (Figure 5A). However, the antioxidants did not rescue the cancer cells either from inhibition of proliferation (Figure 5B) or from apoptosis (Figure 5C) induced by 2.

binding of 2 to PPARs remains possible. However, in general activation of PPARs by high-affinity ligands such as amorfrutins A and B usually does not result in strong antiproliferative effects, as was shown recently.7,8 Global Gene Expression Analysis. In order to shed more light on the potential mechanism of action of 2, the transcriptome of HT-29 cells that were treated with 30 μM 2 for 4 h was analyzed using RNA sequencing. Noteworthy, 409 genes were highly regulated after short-term treatment (80 up-, 329 downregulated; Supporting Information Excel file 1). These genes were further subjected to gene ontology (GO) overrepresentation analysis, and an integrative network was built using Cytoscape21 (Figure S1, Supporting Information). As expected, GO terms such as “proliferation”, “cell cycle”, “cell death”, or “apoptosis” were highly enriched in the regulated genes (Figure 4, Supporting Information Excel file 2). Strikingly, “oxidative stress” as well as “mitochondria” related GO terms were further significantly enriched in the tranD



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Figure 3. Role of peroxisome proliferator-activated receptors (PPARs) for antiproliferative and apoptotic effects of compound 2. (A) HT-29 cells were transfected with nonselective siRNA or with an equimolar mixture of PPAR-selective siRNA (against PPAR α, β/δ, and γ) for 48 h. RNAi reduced the gene expression of PPAR α, β/δ, and γ by 74%, 83%, and 70%, respectively. Data are expressed as means ± SEM (n = 4). (B) HT-29 cells were transfected as in part A and treated subsequently with 20 μM 2 for an additional 48 h. The relative number of cells was then determined. Data are expressed as means ± SD (n = 8). (C) HT-29 cells were transfected as in part A and subsequently treated with 30 μM 2 for an additional 24 h. Phosphatidylserine externalization was determined as described in Figure 2. Bar plots (right) show percent of cell population in apoptosis (means ± SEM, n = 6). n.s. not significant, ***p ≤ 0.001 between nonselective and PPAR-selective siRNA.

was partly prevented by preincubation with cyclosporine A (CsA), an inhibitor of cyclophillin D that is a component of the MPTP.25 However, preincubation with CsA was not sufficient for preventing ΔΨm dissipation (Figure 6D) or for preventing the apoptotic effects of 2 (Figure 6E) (actually, CsA potentiates the cytotoxicity of 2). These observations suggest that ΔΨm dissipation and apoptosis are not direct consequences of the MPTP opening induced by 2. Instead, compound 2 may induce mitochondrial membrane depolarization by other mechanisms, e.g., protonophoric uncoupling. In general, perturbed mitochondria should display functional alterations. To investigate mitochondrial oxygen consumption in living HT-29 cells during treatment with compound 2, the fluorescence of an oxygen-sensitive dye was measured. Strikingly, treatment with 2 and 10 μM of compound 2 resulted in elevated mitochondrial oxygen consumption similar to the protonophoric uncoupler carbonyl cyanide 3-chlorophenylhydrazone (CCCP). In contrast, inhibition of the electron transport chain at complex III by antimycin A (AMA) blocked oxygen consumption (Figure 6F,G). Generally, mitochondrial perturbations induce glycolysis to compensate for reduced ATP synthesis. Since the pyruvate-tolactate conversion is the main contributor to extracellular acidification in unsealed cellular systems (in balance with constant levels of atmospheric carbon dioxide), the pH is an indicator for glycolytic flux.26 Therefore, extracellular acidification was measured by fluorescence quenching of a pHsensitive dye. Of note, treatment with 2 and 10 μM 2 resulted in 3- and 6-fold elevated extracellular acidification rates (ECAR) with 1.5 and 2.9 × 10−9 [H+]/min, respectively. Similarly, the mitochondrial perturbators CCCP and AMA also induced extracellular acidification 3- and 7-fold, respectively (Figure 6H,I). Overall, these results demonstrate that compound 2 induced mitochondrial damage in HT-29 colon carcinoma cells, leading to structural and metabolic dysfunction and finally to apoptosis. In order to shed more light on structure−activity relationships, compound 2 was chemically modified by blocking the C1 carboxylic acid and the C-2 hydroxy groups. Both residues can act as hydrogen bond donors, showing acid dissociation constants (pKa) of 3.5 and 11.6, respectively (Figure S2, Supporting Information). However, in nonpolar environments (such as biological membranes) acid dissociation constants of

Figure 4. Global gene expression analysis of HT-29 cells treated with compound 2. Cells were treated for 4 h, and isolated RNA was sequenced subsequently. Expressed RNA transcripts were then globally analyzed for overrepresented gene ontology (GO) terms that were subjected to network analyses. Prominent network clusters were selected and shown with adjusted p values for these terms. Data are expressed as medians ± range.

These results suggest that the formation of ROS might not be (directly) causal for the cellular effects of compound 2. Compound 2 Disrupts Mitochondrial Integrity in HT29 Colon Carcinoma Cells. Mitochondria not only are indispensable for cellular energy production and metabolism but are also regulators of the intrinsic pathway of apoptosis leading to cell death.23 Considering the observed activation of caspase 9 (Figure 2B) and the mitochondria-related gene expression changes (Figure 4), it was assumed that compound 2 could potentially induce mitochondrial dysfunction. In healthy cells, the inner mitochondrial membrane, nearly impermeable to all ions, contributes to the robust formation of an electrochemical gradient leading to the mitochondrial transmembrane potential (ΔΨm) required for ATP synthesis. A long-lasting opening of the mitochondrial permeability transition pore (MPTP) results in permanent ΔΨm dissipation and cell death.23 Cancer cells exhibit increased ΔΨm due to accelerated metabolism, making test compounds that selectively facilitate mitochondrial membrane permeabilization interesting for drug development.24,25 Notably, treatment of compound 2 in HT-29 cells led to potent dissipation of the ΔΨm with an IC50 value of 0.6 μM within a few minutes (Figure 6A). The effects on the MPTP were further tested by flow cytometry. Compound 2 induced a permanent opening of the MPTP in these cells (Figure 6B,C). In contrast, MPTP opening E



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Figure 5. Role of reactive oxygen species (ROS) in antiproliferative and apoptotic effects of compound 2. (A) HT-29 cells were incubated with 30 μM 2 in the absence or presence of the antioxidants N-acetylcysteine (NAC, 1 mM), glutathione (GSH, 2.5 mM), 3H-1,2-dithiole-3-thione (D3T, 25 μM), α-tocopherol (αTOC, 50 μM), and ascorbic acid (AA, 500 μM). Intracellular ROS were kinetically detected (left) by use of CM-H2DCFDA in living cells. Bar plots (right) show the area under the curve (AUC). Data are expressed as means ± SEM (n = 7). (B) HT-29 cells were treated with concentration series of compound 2 in the absence or presence of antioxidants for 48 h. For measurement of antiproliferative effects, the relative number of cells was determined by use of a DNA-binding fluorescent dye. Data are expressed as means ± SD (n = 4). (C) HT-29 cells were treated with 30 μM 2 in the absence or presence of antioxidants for 24 h. Apoptosis was determined as described above. Bars represent means ± SEM (n = 3). ***p ≤ 0.001 vs control, n.s. not significant vs compound 2-only treated cells; ###p ≤ 0.001 vs compound 2-only treated cells.

compounds can shift by up to 7 units.27 Therefore, the 1carboxy and 2-hydroxy methylated synthetic derivatives 2a and 2b were investigated, respectively. Methylation of either of these functional groups was sufficient to completely abolish mitochondrial membrane depolarization (Figure 7A) and apoptosis (Figure 7B) in HT-29 cells. These observations indicate that the 2-hydroxybenzoic acid motif in compound 2 is a relevant feature, potentially by promoting proton transport through the inner mitochondrial membrane, leading to ΔΨm dissipation. Salicylic acid derivatives such as 2 are significantly more acidic than their corresponding benzoic acid analogues (2a). In conjunction with the compound’s high lipophilicity, the acidity of 2 might stimulate protonophoric mitochondrial uncoupling, possibly by transfer of protons over the inner mitochondrial membrane. Additive and Synergistic Effects of Compound 2. Simultaneous treatment with compounds that make use of different molecular mechanisms might produce additive or synergistic effects on cancer cell proliferation and death.28 Compound 2 was tested in combination with various anticancer agents for inhibition of proliferation and apoptosis in HT-29 cells using the Loewe additivity model (see Experimental Section). Interestingly, combinations with the DNA-crosslinking drug cisplatin (Figure 8A) as well as with the topoisomerase I inhibitor irinotecan (Figure 8B) achieved additive effects on proliferation inhibition. These data suggest overlapping molecular mechanisms resulting in antiproliferative effects. However, for all cancer cell lines tested in this study, compound 2 achieved higher maximal cell growth arrest efficiency than cisplatin and irinotecan (Table 2), indicating the

existence of additional molecular events. Potential synergy on apoptotic events was evaluated by analyzing simultaneous activation of the alpha Fas receptor ligand (αFAS) and the TNF-related apoptosis inducing ligand (TRAIL). Synergy was assessed with the Bliss independence model.29 Strikingly, 2 synergistically induced PS externalization with αFAS and TRAIL, suggesting that 2 does not bind to death receptors. However, since compound 2 also activated caspase 8 (Figure 2B), it probably activated the death receptor signaling pathway downstream of the death receptors, as recently described for mitochondrial perturbators.30 Conclusion. In summary, amorfrutin C (2), isolated from the roots of G. foetida, showed potent antiproliferative effects in different cancer cell lines by triggering cell cycle arrest and inducing apoptosis. Mechanistically, the effects of 2 could be correlated with mitochondrial depolarization, although further detailed analysis is needed to identify potentially involved cellular targets. Synergy studies and extrinsic apoptosis activation further indicated stimulation of the death receptor signaling pathway. Since cancer cell mitochondria are structurally and functionally different from their normal counterparts, cancer cells are more susceptible to mitochondrial toxicity than normal cells.31 Thus, targeting mitochondria function appears to be a promising strategy for cancer therapy.25 In line with this conceptual framework, the molecular interference of amorfrutin C (2) with mitochondrial function seems to produce stronger antiproliferative effects in colon cancer cells than in normal colon cells (Figure 1A). On the other hand, mechanistically interesting but lipophilic natural products such as 2 require F



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Figure 6. Mitochondrial dysfunction induced by compound 2. (A) Loss of mitochondrial transmembrane potential (ΔΨm) detected by fluorometry. HT-29 cells were incubated with concentration series of 2 for 5 min. Data are expressed as means ± SD (n = 4). (B) Opening of the mitochondrial permeability transition pore (MPTP) determined by calcein/CoCl2 fluorescence. HT-29 cells were treated for 30 min with 2 in the absence or presence of cyclosporin A (CsA, 4 μM). Histograms show one representative experiment for each treatment condition. (C) Mean of calcein/CoCl2 fluorescence intensities. Bars show means ± SEM (n = 6). (D) Effect of MPTP inhibition on compound 2-induced ΔΨm dissipation. Note that blocking the pore with CsA during pretreatment led to hyperpolarization of the mitochondrial membrane. Data are expressed as means ± SD (n = 6). (E) Effect of MPTP inhibition on compound 2-induced cell death. HT-29 cells were treated with 30 μM 2 in the absence or presence of CsA (2 μM) for 24 h. Apoptosis and necrosis was determined by flow cytometry (means ± SEM, n = 3). (F) Mitochondrial oxygen consumption determined by fluorescence quenching. Fluorescence lifetime increased with reduction in extracellular oxygen concentration. Data are expressed as means ± SEM (n = 8). AMA, antimycin A; CCCP, carbonyl cyanide 3-chlorophenylhydrazone. (G) The rate of fluorescence lifetime change was determined between 20 and 60 min of the treatment shown here in F and plotted relative to untreated cells, giving the relative oxygen consumption. Data are expressed as means ± SEM (n = 8). (H) Extracellular acidification determined by fluorescence quenching. HT-29 cells were labeled with a pH-sensitive probe and treated again with the compounds indicated. Fluorescence lifetime was transformed to pH values. (I) Extracellular acidification rate (ECAR) was determined between 20 and 150 min of treatment H and plotted as hydrogen ion concentration change per minute. Data are expressed as means ± SEM (n = 8). n.s. not significant, **p ≤ 0.01, ***p ≤ 0.001 vs untreated cells; #p ≤ 0.05, ##p ≤ 0.01, ###p ≤ 0.001 vs compound 2-only treated cells.

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chemical optimization and/or the use of molecular carriers for efficient application in vivo. Further systematic safety and pharmacokinetic studies of the novel amorfrutin C (2) and analogues will be important to develop potent candidates for preclinical trials. Notably, amorfrutin B (1) has been extensively analyzed in in vivo models, and no toxic side effects were observed.8 Taken together, these data show that amorfrutin C (2) and natural or synthetic analogues thereof may become promising candidates for developing anticancer drugs.

EXPERIMENTAL SECTION

General Experimental Procedures. Chemicals were purchased from the following sources: staurosporine, irinotecan, cisplatin, and antimycin A from LKT Laboratories (Biomol, Hamburg, Germany); oxaliplatin from Cayman Chemical (Biomol). Paclitaxel, 5-fluorouracil, etoposide, doxorubicin, N-acetylcysteine (NAC), glutathione (GSH), 3H-1,2-dithiole-3-thione (D3T), α-tocopherol (αTOC), ascorbic acid (AA), carbonyl cyanide 3-chlorophenylhydrazone (CCCP), and cyclosporine A (CsA) were purchased from Sigma-Aldrich (TaufG



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Figure 7. Role of carboxylic acid and hydroxy group residues of compound 2 on mitochondrial transmembrane potential (ΔΨm) and apoptosis induction. (A) Loss of mitochondrial transmembrane potential (ΔΨm) detected by fluorometry. HT-29 cells were incubated for 5 min with a concentration series of the compounds indicated and labeled with the mitochondria-selective JC-1 dye. Data are expressed as means ± SD (n = 4). (B) HT-29 cells were treated with the compounds indicated for 24 h. Apoptosis was determined by flow cytometry. Bars represent means ± SEM (n = 4). n.s. not significant, ***p ≤ 0.001 vs control; ###p ≤ 0.001 vs compound 2.

Figure 8. Additive and synergistic effects of compound 2. (A) Isobolographic analysis of antiproliferative effects for compound 2 and irinotecan. HT-29 cells were treated with combinations of compound 2 and irinotecan for 72 h. The relative number of cells was determined as described above. IC70 data are expressed as means ± SD (n = 4). CI, combination index. (B) Isobolographic analysis for antiproliferative effects of 2 and cisplatin. (C) Synergistic apoptosis induction of 2 with αFAS and TRAIL. HT-29 cells were treated with 2 (10 μM), αFAS (10 ng/mL), TRAIL (30 ng/mL), or combinations thereof for 48 h. Apoptosis was determined by flow cytometry. Expected combination effects were calculated according to the Bliss independence model and assumed additivity on apoptosis induction. Bars represent means ± SEM (n = 4). **p ≤ 0.01, ***p ≤ 0.001 vs untreated cells; ###p ≤ 0.001 between observed and expected combination effects.

kirchen, Germany). Human αFAS (clone CH11, #05-201) and TRAIL (#GF092) were purchased from Merck (Darmstadt, Germany). Compound 1 was isolated from the fruits of Amorpha f ruticosa and analyzed in detail as described recently.8,12 NMR spectra were acquired on a Bruker 500 MHz and a Varian INOVA 600 MHz spectrometer. ESIMS were recorded on an API 165 LC/MS mass spectrometer coupled with an Agilent 1100 HPLC. Preparative HPLC was performed with a modified Sepbox system (Sepiatec). For further information we refer to Supporting Information - Additional information for the Experimental Section. Plant Material. Roots of Glycyrrhiza foetida were collected in Tiflet, Morocco, in June 2008 by Thomas Friedrich, Friedrich Nature Discovery (FND, http://www.friedrichnaturediscovery.de/en/index. html). AnalytiCon Discovery further processed the roots for compound extraction. AnalytiCon Discovery has neither a direct contract with Morocco nor a permit from the Moroccan authorities but instead purchased the plant material from FND. A voucher specimen with the number ACD-V-20720 is deposited at AnalytiCon Discovery. FND exports several medicinal plants from Morocco to Germany and other countries in the world via the Moroccan-based company Sahara Exporters sarl (http://www.saharaexporters.com/en/ index.html). In Analyticon’s contract with FND, it is stipulated that it is the responsibility of FND to obtain all necessary collection and export permits. FND has an export permit for all plants in the portfolio but no single permit for the collection of G. fetida used in this study. At the time of collection in 2008, before the Nagoya protocol came into force and worldwide practice, this was neither necessary nor applicable owing to a lack of fully established legal and administrative frameworks.

Extraction and Isolation. The procedure for compound 1 was described recently8 whereas compound 2 was mentioned in a previous paper but so far never substantially characterized.8 In this study, to isolate compound 2 (2-Hydroxy-4-methoxy-3,5-bis(3-methylbut-2enyl)-6-phenethylbenzoic acid), 1 kg of air-dried and powdered roots was extracted twice with methyl tert-butyl ether (46.3 g). MPLC fractionation (RP-18, 250 × 50 mm) with a gradient from 50% methanol to 100% methanol yielded five fractions containing resorcinol derivatives. Further separation by preparative HPLC (Select B 10 μm, 250 × 50 mm, flow rate 80 mL/min, gradient with CH3OH/ ammonium formate buffer adjusted with formic acid to pH 4.0) yielded pure compound 2 (22 mg). Amorfrutin C (2): yellow powder; 1H and 13C NMR data, see Table 1; (+)-ESIMS m/z 409 [M + H]+, 391 [M − H2O + H]+, 335; (−)-ESIMS 407 [M − H]−; (−)-HRESIMS m/z 407.2221 [M − H]− (calcd for C26H31O4 407.2221). Compounds 1 and 2 are available as from Analyticon Discovery, product numbers NP-15142 and NP-15934, respectively. A number of H



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amorfrutin-related natural products, which were tested in pilot experiments and which displayed similar properties to amorfrutin C, were obtained from Analyticon Discovery, namely, product numbers 2-Hydroxy-4-methoxy-3,5-bis(3-methylbut-2-enyl)-6-pentylbenzoic acid (3), 2,4-Dihydroxy-3,5-bis(3-methylbut-2-enyl)-6-pentylbenzoic acid (4), 2,4-Bis(3-methylbut-2-enyl)-5-pentylbenzene-1,3-diol (5), 2(2-Hydroxy-3-methylbut-3-enyl)-4-(3-methylbut-2-enyl)-5-pentylbenzene-1,3-diol (6), and 2-(2-Hydroxy-3-methylbut-3-enyl)-4-(3-methylbut-2-enyl)-5-phenethylbenzene-1,3-diol (7) (see Supporting Information, 1H NMR spectra for 2−7 and LC-MS spectra for 2−7). Preparation of Derivatives 2a and 2b. Compound 2 (100 mg) was dissolved in anhydrous acetone (5 mL) under argon. Potassium carbonate (339 mg, 10 equiv) was added to the stirring solution, and the resulting suspension was then cooled to 0 °C. To the chilled suspension was added methyl iodide (35 mg, 2 equiv), and the reaction was immediately allowed to warm to room temperature. The reaction was stirred at room temperature for 4 h, followed by neutralization at 0 °C using 1 M HCl. The acidic solution was extracted using diethyl ether and hexanes (1:1, 3 × 1 mL), and the combined organic collection was washed with H2O (5 mL). The organic phase was dried over Na2SO4 and concentrated under reduced pressure, yielding intermediate 2c as a yellow oil (as indicated in Supporting Information, Experimental Section). Without further purification, intermediate 2c was transferred using DMSO (0.3 mL) to a solution of potassium hydroxide (249 mg, 20 equiv) in 5% aqueous DMSO (5 mL). The basic solution was heated to 120 °C for 8 h under argon in a sealed container. Subsequently, the reaction was cooled to 0 °C and acidified with 1 M HCl. The acidic solution was extracted using diethyl ether (3 × 20 mL), and the combined organic collection was dried over Na2SO4 and concentrated under vacuum. Column chromatography (silica gel; hexanes/EtOAc = 10:1 progressing to 2:3) afforded acid 2a (44 mg, 47% yield) as a colorless oil: 1H NMR (600 MHz, CDCl3) δ 7.29−7.15 (5H, m,), 5.24 (1H, m), 5.08 (1H, m), 3.84 (3H, s), 3.73 (3H, s), 3.40 (4H, d), 2.99−2.81 (4H, m), 1.79 (3H, br s), 1.76 (3H, br s), 1.71 (3H, br s), 1.69 (3H, br s); 13 C NMR (150 MHz, CDCl3) δ 172.4, 159.4, 155.0, 141.9, 138.1, 131.82, 131.76, 130.6, 128.4, 128.3, 127.1, 126.0, 124.0, 123.6, 123.0 62.8, 61.5, 37.4, 33.0, 25.7, 25.6, 25.5, 23.7, 18.2, 18.0; (+)-HRESIMS m/z 445.23395 [M + Na]+ (calcd for C27H34NaO4+, 445.23493). A scheme of the synthesis strategy is outlined in Supporting Information (Experimental Section). Compound 2b was obtained as a byproduct of methylation of compound 2 and isolated by column chromatography (silica gel; hexanes/ethyl acetate = 10:1 progressing to 2:3). 1H NMR (600 MHz, CDCl3) δ 7.33−7.27 (2H, m), 7.24−7.18 (3H, m), 5.26 (1H, m), 5.04 (1H, m), 3.98 (3H, s), 3.73 (3H, s), 3.42−3.37 (4H, m), 3.20−3.16 (2H, m), 2.82−2.77 (2H, m), 1.80 (3H, br s), 1.74 (3H, br s), 1.70 (3H, br s), 1.68 (3H, br s); 13C NMR (125 MHz, CDCl3) 172.1, 161.5, 160.2, 142.3, 141.4, 131.8, 131.5, 128.4, 128.1, 126.2, 126.0, 124.2, 122.8, 121.2, 109.5, 61.5, 52.3, 37.6, 33.2, 25.7, 25.6, 25.3, 23.5, 18.1, 18.0; (+)-HRESIMS m/z 423.25220 [M + H]+ (calcd for C27H35O4+, 423.25299). Standard Biological Procedures. Standard procedures such as cell culturing, cell cycle analysis, immuno-(Western) blotting, and caspase activation assay are summarized in the Supporting Information (for the Experimental Section). pKa Determination. Acid dissociation constants were measured at Sirius Analytical (Forrest Row, UK). In brief, compound 2 was titrated from pH 12.0−2.0 at concentrations of 32−26 μM under methanol/ water cosolvent conditions and from pH 12.5 to 8.9 at concentrations of 30−21 μM under aqueous conditions, respectively. Titration was performed in the presence of 0.15 M KCl at 25 °C. Analysis was done using the UV metric method. Proliferation Assay. To investigate the growth arrest of HT-29, T84, PC-3, and MCF7 cancer cells and normal CCD 841 CoN cells, the cells were seeded in black 384-well plates (#3712, Corning, Fisher Scientific, Schwerte, Germany) with a density of 750 cells/well (HT29), 250 cells/well (PC-3), and 900 cells/well (T84, MCF7, and CCD 841 CoN), respectively. One day later, the cells were treated with concentration series of the tested compound, as indicated for 72 h

(HT-29 and PC-3) or 96 h (T84, MCF7, and CCD 841 CoN), respectively. For cotreatment of HT-29 cells with antioxidants, the treatment time was reduced to 48 h. The relative number of cells was determined by measurement of cellular DNA content applying the fluorescent CyQUANT NF cell proliferation assay kit (Life Technologies), according to the manufacturer’s instructions. Fluorescence intensity was measured with the POLARstar Omega (BMG Labtech, Ortenberg, Germany) and was transformed to relative cell number. For concentration series, data were fitted using GraphPad Prism 5.0 according to the equation Y = Top + (Bottom − Top)/(1 + 10(LogIC50−X) × Hill slope) with variable Hill slope. Efficiency is the maximal observed induction of inhibition (100% − Bottom) relative to nontreated cells (set to 0%). IC50 is the concentration that is required for 50% inhibition related to the compound’s specific (maximal) efficiency, which often is not sufficient to inhibit 50% of all present cells. For a better comparison of proliferation inhibition to determine additive effects, the IC70 value was calculated as the concentration required for 70% inhibition related to the total number of cells. Additive effects were determined by treatment with compound mixtures with the following ratios: 7:0, 6:1, 5:2, 4:3, 3:4, 2:5, 1:6, and 0:7. HT-29 cells were treated with different concentration series of these compound mixtures, and IC70 data were calculated and plotted as isobolograms according to the Loewe additivity model.32,33 The combination index (CI) for compounds x and y in each mixture (M) was calculated as follows: CI(M) = IC70(Mx)/IC70(x) + IC70(My)/ IC70(y). Annexin V Assay. Phosphatidylserine externalization was analyzed using the annexin-V-FLUOS staining kit (Roche Diagnostics, Mannheim, Germany), according to the manufacturer’s instructions. Flow cytometry was performed using an Accuri C6 apparatus (BD Biosciences, Heidelberg, Germany), and data were analyzed with FlowJo 7.6 (Tree Star). Apoptosis was defined as annexin positive/PI negative and annexin positive/PI positive, whereas necrosis was defined as annexin negative/PI positive. Synergistic effects were investigated by use of the Bliss independence model,29 which is defined by the equation Exy = Ex + Ey − ExEy, where E is the fraction of cells in apoptosis and Exy is the (expected) additive effect of drugs x and y as predicted by their individual effects Ex and Ey. DNA Fragmentation Assays. Accumulation of DNA in the cytoplasm of treated HT-29 cells was determined by use of the Cellular DNA Fragmentation ELISA and the Cell Death Detection ELISA kits (both Roche Diagnostics). For the Cellular DNA Fragmentation ELISA, HT-29 cells were seeded in 96-well plates (TPP, Biochrom, Berlin, Germany) with a density of 13 000 cells/well in the presence of 10 μM 5-bromo-2′-deoxyuridine (BrdU) and incubated for 2 days at 37 °C. After removal of unincorporated BrdU, cells were treated for 4 h. The cytosolic fractions were harvested and analyzed on a 96-well half-area clear high-binding microplate (#3690, Corning), according to the manufacturer’s instructions. For the Cell Death Detection ELISA, HT-29 cells were seeded in 96-well plates (TPP) with a density of 20 000 cells/well and incubated for 1 day at 37 °C. Cells were then treated for 4 h. The cytosolic fractions were harvested and analyzed according to the manufacturer’s instructions. Absorbance was measured using the POLARstar Omega (BMG Labtech). Cell-free samples were used as background control for subtraction, and data were normalized to vehicle-treated cells. Fluorescence Microscopy. For visualization of cleaved caspase 3, HT-29 cells were seeded on 13 mm coverslips (Sarstedt, Nürnbrecht, Germany) and placed in 24-well plates (Nunc, Wiesbaden, Germany) with a density of 125 000 cells/well, 1 day before treatment. Treated cells were washed with PBS, fixed with 4% formaldehyde/PBS for 15 min, permeabilized with 0.3% Triton X-100/PBS (PBS-T) for 10 min, and subsequently blocked with 5% goat serum (Sigma-Aldrich) in PBS-T (0.3%) for 60 min (at room temperature each). Cells were then incubated with primary antibody against cleaved caspase 3 (#9664, Cell Signaling Technology, Merck, Darmstadt, Germany) diluted (1:200) in 1% BSA/PBST (0.3%) at 4 °C overnight. Cells were washed in PBS-T (0.3%) labeled with anti-rabbit IgG (H+L) and F(ab′)2 fragment Alexa Fluor 488 conjugate (#4409, Cell Signaling Technology) diluted (1:1000) in 1% BSA/PBS-T (0.3%) at room I



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temperature for 1 h. Finally, coverslips were counterstained with ProLong Gold Antifade Mountant solution (containing DAPI) (Life Technologies) and incubated at room temperature for 24 h. Fluorescence microscopy was performed on the LSM700 (Zeiss, Jena, Germany). RNA Interference (RNAi). HT-29 cells were transfected with an equimolar mixture of Silencer Select Validated siRNA against PPAR α, β/δ, and γ (10 nM each) or 30 nM nonspecific Negative Control siRNA (all Life Technologies) using the Trans-IT TKO transfection reagent (Mirus Bio, VWR, Darmstadt, Germany), according to the manufacturer’s instructions. Transfection reagent/siRNA complexes were prepared in opti-MEM (Life Technologies), incubated at room temperature for 30 min, and added to the HT-29 cell suspension in antibiotic-free DMEM/F12/FBS directly before seeding. Cells were seeded with a density of 850 cells/well (384-well plate) and 150 000 cells/well (12-well plate) for proliferation and annexin V assays, respectively. After 48 h, the cells were harvested for gene expression analysis or additionally treated by adding 3-fold concentrated compound or vehicle control. Since the cells that were treated with DMSO as vehicle control did not show any growth-inhibiting effects, the total incubation time for subsequent experiments with these cells had to be reduced to prevent confluence. A pool of siRNAs against all PPAR subtypes was used to prevent any potential compensatory effects. Cells were analyzed for proliferation and annexin V after 48 and 24 h of compound treatment, respectively. Gene Expression Analysis. Expression of PPARs was analyzed by quantitative real-time PCR, as described recently.34 Global RNA expression was analyzed by RNA sequencing using four biological replicates for each sample. A 1 μg amount of total RNA was used to generate the cDNA library by use of the TruSeq RNA Sample Prep kit v2 (Illumina, San Diego, CA, USA) according to the manufacturer’s instructions. The libraries were sequenced using HiSeq 2500 (Illumina) in paired-end-50nt mode. Sequencing quality was assessed using FastQC.35 Sequence reads data were deposited at the European Nucleotide Archive (accession number: PRJEB7551), Web-link http://www.ebi.ac.uk/ena/data/view/PRJEB7551. Reads were mapped using STAR,36 and quantification of reads mapping to gene features (Gencode 16) was done using HTSeq.37 Differentially expressed genes were defined as those that differed by more than 4fold with corrected p-value ≤ 0.0001 and were identified using edgeR.38 Differentially expressed genes were analyzed using Cytoscape21 and BiNGO.39 Detection of Reactive Oxygen Species. Formation of intracellular ROS was detected with the ROS-sensitive dye 5-(and-6)chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate (CMH2DCFDA, Life Technologies) according to the manufacturer’s instructions. One day before treatment HT-29 cells were seeded in 96-well plates (TPP) with a density of 15 000 cells/well. Before treatment, adherent cells were washed with prewarmed PBS and loaded with 50 μM dye diluted in PBS. Cells were then incubated at 37 °C for 30 min to allow incorporation and activation of CMH2DCFDA, followed by removing free dye and washing with prewarmed PBS. Phenol red-free DMEM/F-12 medium was added, and cells were again incubated at 37 °C for 60 min. Compounds were added as indicated, and fluorescence intensity (485/530 nm) was measured with the POLARstar Omega (BMG Labtech) at 37 °C for 22 h. Data were analyzed using GraphPad Prism 5.0. Mitochondrial Transmembrane Potential Assay. The mitochondrial transmembrane potential (ΔΨm) was investigated with the JC-1 assay (Cayman Chemicals, Biomol, Hamburg, Germany), according to the manual. This assay makes use of a lipophilic cationic dye (5,6, 5′,6′-tetrachloro-1,3,1′,3′-tetraethylbenzimidazolylcarbocyanine iodide), which selectively enters into mitochondria and changes reversibly its color from red (JC-1 aggregates) to green (JC-1 monomers) as the membrane potential decreases. Thus, HT-29 cells were seeded in 96-well plates (TPP) at a density of 40 000 cells/well. One day later, cells were treated with the test compounds for 5 min at 37 °C, followed by addition of the JC-1 dye for an additional 10 min. Cells were then washed twice with prewarmed JC-1 assay buffer to remove free JC-1 dye. Red (excitation 560/10 nm, emission 590/30

nm) and green (excitation 485/30 nm, emission 520/10 nm) fluorescence was measured in the POLARstar Omega (BMG Labtech). The ratio of red to green fluorescence was used as an indicator of mitochondrial transmembrane potential (ΔΨm). Data were fitted using GraphPad Prism 5.0 according to the equation Y = Bottom + (Top − Bottom)/(1 + 10(LogIC50−X) × Hill slope), with a variable Hill slope. Mitochondrial Permeability Transition Pore Assay. Effects on the MPTP were analyzed by use of the MitoProbe transition pore assay kit (Life Technologies), according to the manufacturer’s instructions. Cells were loaded with a fluorescent calcein dye that accumulates in the mitochondria. The fluorescence of cytosolic calcein was quenched by addition of CoCl2, while mitochondrial fluorescence was maintained. Opening of the MPTP leads to loss of mitochondrial calcein fluorescence. For this purpose, trypsinized HT-29 cells were suspended in HBSS/Ca buffer containing 10 nM calcein and 400 μM CoCl2 with a density of 106 cells/mL. Cells were pretreated with 2 μM cyclosporin A or vehicle control for 60 min at 37 °C and subsequently treated with the tested compounds for a further 30 min at 37 °C. Cells were finally washed with HBSS/Ca buffer and analyzed by flow cytometry (Accuri C6, BD Biosciences), according to manufacturer’s instructions. Analysis was performed using FlowJo 7.6 (Tree Star) and Prism 5.0 (GraphPad). Oxygen Consumption Measurements. Oxygen consumption was determined by time-resolved fluorescence of an oxygen-sensitive probe (MitoXpress-Xtra HS, Luxcel Biosciences, Cork, Ireland). The probe fluorescence is quenched by molecular oxygen, so that fluorescence lifetime increases with reduction in extracellular oxygen concentration. One day before treatment, HT-29 cells were seeded in 96-well plates (TPP) at a density of 80 000 cells/well. Medium was removed, and cells were incubated with 140 μL of prewarmed probe diluted in phenol red-free DMEM/F-12 medium. Cells were incubated at 37 °C for 10 min, and the test compounds were added. Finally, cells were sealed with 100 μL of prewarmed HS mineral oil (Luxcel Biosciences) to prevent back diffusion of ambient oxygen. Timeresolved fluorescence was measured in the POLARstar Omega (BMG Labtech) apparatus with the following settings: temperature = 37 °C; TRF optic Z height = 6 mm; excitation = 380/20 nm; emission = 655/ 50 nm; window 1 (w1): 30 μs delay and 30 μs integration time; window 2 (w2): 70 μs delay and 30 μs integration time; interval time = 90 s; measurement time = 150 min. Background fluorescence was measured in wells with medium and oil, but without cells and a probe. For data analysis, background fluorescence was subtracted for each well. Fluorescence lifetime (τ) was calculated by τ = 40/(ln(w1/w2)) and plotted over treatment time. For comparing oxygen consumption between treatments, the rate of probe fluorescence lifetime was determined between 20 and 60 min and expressed relative to untreated cells. Data were analyzed using Prism 5.0 (GraphPad). Extracellular Acidification Measurements. Extracellular acidification, which is mainly due to lactate production reflecting glycolytic activity,26 was determined by time-resolved fluorescence of a pHsensitive probe (pH Xtra, Luxcel Biosciences). Fluorescence lifetime of this probe increases with decrease in pH, so that it allows measurement of extracellular acidification. One day before treatment, HT-29 cells were seeded in 96-well plates (TPP) with a density of 80 000 cells/well and incubated at 37 °C overnight in a CO2-free incubator. For subsequent measurements, the following low-buffering aspiration medium was used according to the manufacturer’s instructions: 1 mM PBS (pH 7.4), 20 mM glucose, 75 mM NaCl, 54 mM KCl, 2.4 mM CaCl2, and 0.8 mM MgSO4. Before treatment, cells were washed twice with 200 μL of aspiration buffer and incubated with 140 μL of prewarmed probe diluted in aspiration medium. Cells were incubated at 37 °C for 10 min, and the test compounds were added. Wells were not sealed with oil to avoid trapping of CO2 reflecting Krebs cycle activity. Time-resolved fluorescence was measured in the POLARstar Omega (BMG Labtech) apparatus with the following settings: temperature = 37 °C; TRF optic Z height = 6 mm; excitation = 380/20 nm; emission = 615/50 nm; window 1 (w1): 100 μs delay and 30 μs integration time; window 2 (w2): 300 μs delay and 30 μs integration time; interval time = 100 s; measurement time = J



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200 min. Background fluorescence was measured in wells with medium without cells. For data analysis, background fluorescence was subtracted for each well. Fluorescence lifetime (τ) for each sample was calculated by τ = 200/[ln(w1/w2)] and transformed to absolute pH values according to the equation pH = (1687.2 − τ)/199.12, as reported previously.26 The pH values were plotted over treatment time. For comparing different treatments, the extracellular acidification rate was determined between 20 and 150 min. Data were analyzed using Prism 5.0 (GraphPad). Statistical Analysis. Data are expressed as means ± standard error of mean (SEM), if not otherwise noted. Statistical tests were performed using GraphPad Prism 5.0. For comparison of two groups, statistical significance was examined by unpaired two-tailed Student’s t test. For multiple comparisons, data were analyzed by one-way ANOVA with subsequent Dunnett’s and Bonferroni post-test, respectively. A p value ≤ 0.05 was defined as statistically significant as given in the figure legends.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jnatprod.5b00072. Additional information for the Experimental Section (PDF) GO network analysis for 2 (Figure S1) and the pKa determination for 2 (Figure S2) (PDF) Global gene expression data (XLS) Global gene expression data (XLSX) 1 H NMR spectra for 2−7 (PDF) LC-MS spectra for 2−7 (PDF)

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AUTHOR INFORMATION

Corresponding Author

*Tel: +49 30 8413 1661. Fax: +49 30 8413 1960. E-mail: [email protected]. Author Contributions ∥

C. Weidner and M. Rousseau contributed equally to this study. Notes

The authors declare the following competing financial interest(s): K. Siems and G. Hetterling are employees of Analytion Discovery, a company that sells natural products.

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ACKNOWLEDGMENTS The present work was supported by the German Ministry for Education and Research (BMBF, grant numbers 0315082, 01EA1303) and the European Union [FP7/2007-2011], under grant agreement no. 262055 (ESGI).

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REFERENCES

(1) Anonymous. The Global Burden of Disease: 2004 Update; World Health Organization: Geneva: 2008. (2) Jemal, A.; Bray, F.; Center, M. M.; Ferlay, J.; Ward, E.; Forman, D. Ca-Cancer J. Clin. 2011, 61, 69−90. (3) Shekhar, M. P. Curr. Cancer Drug Targets 2011, 11, 613−623. (4) Li, J. W.; Vederas, J. C. Science 2009, 325, 161−165. (5) Newman, D. J.; Cragg, G. M. J. Nat. Prod. 2012, 75, 311−335. (6) Muller, M.; Kersten, S. Nat. Rev. Genet. 2003, 4, 315−322. (7) Weidner, C.; de Groot, J. C.; Prasad, A.; Freiwald, A.; Quedenau, C.; Kliem, M.; Witzke, A.; Kodelja, V.; Han, C. T.; Giegold, S.; Baumann, M.; Klebl, B.; Siems, K.; Muller-Kuhrt, L.; Schurmann, A.; Schuler, R.; Pfeiffer, A. F.; Schroeder, F. C.; Bussow, K.; Sauer, S. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 7257−7262. K


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