Heterotheca inuloides (Mexican arnica) metabolites protect Caenorhabditis elegans from oxidative damage and inhibit nitric oxide production

Abstract

The accumulation of reactive oxygen and nitrogen species (RONS) leads to oxidative stress, which triggers the onset of various pathological conditions. Appropriate copper chelators and nitric oxide (NO) scavengers represent novel strategies for preventing free radical damage. In this work, we tested a series of compounds derived from Heterotheca inuloides (Mexican arnica), including natural products, semisynthetic derivatives, and an acetone extract of the plant, for their ability to protect Caenorhabditis elegans under stressful conditions. Specifically, we tested their ability to bind copper (Cu²⁺), and inhibit nitric oxide (NO) production in RAW 264.7 macrophages activated with lipopolysaccharides (LPS). The results revealed that several compounds and the acetone extract of H. inuloides exhibited excellent potential for increasing the resistance of C. elegans to stressful conditions induced by copper and juglone. We examined the formation of copper(II) complexes with UV-Vis, ESR, and ESI-MS spectroscopy. We found that quercetin (3,3′,4′,5,7-pentahydroxyflavone) could bind to Cu²⁺. In contrast, natural and semisynthetic cadinene-type compounds showed no ability to bind to copper or protect C. elegans. Additionally, flavonoids isolated from the acetone extract of H. inuloides could inhibit NO production, but cadinane-type compounds showed little or no inhibitory activity.

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1. Introduction

Reactive oxygen and nitrogen species (RONS) such as superoxide anions (O₂˙⁻), hydroxyl radicals (˙OH), singlet oxygen (¹O₂) and nitric oxide (NO) play an important role in many physiological processes as components of intracellular signaling cascades and by regulating several physiological functions thereby affecting cellular homeostasis.¹ However, when the production of highly reactive molecules with unpaired electrons overpowers the scavenging mechanisms, this may also lead to a physiological process known as oxidative and nitrative stress.² This pathological condition has been associated with deleterious effects and oxidative damage to cellular constituents, which in turn is the major determinant of life span and aging as damage accumulates in tissues that govern many senescent functions.^3,4 Primary RONS, such as O₂˙⁻, H₂O₂ or NO˙, are precursors to the formation of highly reactive secondary species; hydroxyl radicals and peroxynitrite (˙OH, ONOO⁻).⁵ Both endogenous and exogenous sources share RONS production catalyzed by transition metals, using a mechanism known as the Fenton-catalysed Haber–Weiss reaction.^6,7 The in vivo Fenton-catalysed Haber–Weiss reaction is initiated by the products of aerobic respiration, such as hydrogen peroxide (H₂O₂) and superoxide (O₂˙⁻).⁸ Transition metal ions such as iron, copper and aluminium have rich coordination and redox chemistry, the ability to vary oxidation states, and their presence at high concentrations plays a significant role in the damage to biomolecules.⁹ Natural products have been investigated as promising options to act as protective agents capable of delaying oxidative damage by scavenging free radicals, inhibiting their formation or interrupting their propagation, and by chelating metal ions; this may lead to a delay in age-related diseases and an extension of lifespan. Heterotheca inuloides is a plant native to Mexico, commonly known as árnica, with a series of additional regional names.^10,11 The infusion of dried flowers has been used to treat contusions, muscular pain, wounds and other painful conditions associated with inflammatory processes.¹² This plant species is characterized by biosynthesizing sesquiterpenes with cadinene skeleton. The major sesquiterpenes isolated from this plant are 7-hydroxy-3,4-dihydrocadalene (1) and 7-hydroxycadalene (2).¹³ The antioxidant and free radical scavenging activity using various in vitro antioxidant assays indicated that cadinenes 1 and 2 protect cells against peroxy radical attack and inhibit the production of lipid peroxides induced by NADPH oxidation. In addition, sesquiterpene 1 protected NADH– and succinate–cytochrome C reductase enzymes against peroxidation. Conversely, cadinenes 1 and 2 showed no inhibition against both enzymatic and non-enzymatic superoxide generation.^14–16 On the other hand, the flavonoids kaempferol and quercetin (5) isolated from H. inuloides inhibited the microsomal lipid peroxidation and showed tyrosinase inhibitory activity.^16,17 Some metabolites isolated from the same source showed an ability to trap radicals such as ABTS˙⁺, DPPH˙, ONOO⁻, O₂˙⁻, ¹O₂, HOCl, H₂O₂ and OH˙.¹⁸ Since RONS play a vital role in the oxidative damage processes, the aim of this study was to investigate whether an acetone extract of H. inuloides and metabolites isolated from this extract could protect Caenorhabditis elegans against stress generated by RONS-inducing substances. In addition, the copper sequestering ability of these metabolites and their ability to inhibit the production of nitric oxide (NO) were explored.

2. Experimental

2.1. Reagent and materials

All chemicals used, including solvents, were analytical grade. Aluminium chloride hexahydrate (AlCl₃·6H₂O), aminoguanidine hydrochloride, catechin, cupric sulphate pentahydrate (CuSO₄·5H₂O), dimethyl sulfoxide (DMSO), E. coli serotype 055:B5 lipopolysaccharide (LPS), ethanol, Folin–Ciocalteu’s reagent, juglone (5-hydroxy-1,4-naphthoquinone), methanol, N-(naphthyl)-ethylene-diamine, orthophosphoric acid (H₃PO₄), phosphoric acid (H₃PO₄), 5-fluoro-2′-deoxyuridine, sodium carbonate (Na₂CO₃), sodium nitrite (NaNO₂), sodium dihydrogen phosphate (NaH₂PO₄), sodium hydrogen phosphate (Na₂HPO₄), sodium hydroxide (NaOH), sulfanilamide, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT), 5-fluoro-2′-deoxyuridine, and trypan blue were obtained from Sigma-Aldrich (St. Louis, MO, USA).

2.2. Plant material

Plant material was kindly provided by Laboratorios Mixim (México). Flowers of H. inuloides were collected in 2010 from the locality of San Juan Xoconusco, municipality of Donato Guerra, State of Mexico. The authentication of the plant material was done by M. Sc. Abigail Aguilar-Contreras. A plant specimen of authenticated material was deposited at the Medicinal Plant Herbarium of the Instituto Mexicano del Seguro Social (IMSS, Mexico City) with voucher number IMSSM-16064. The compounds 7-hydroxy-3,4-dihydrocadalene (1), 7-hydroxycadalene (2), 3,7-dihydroxy-3(4H)-isocadalen-4-one (3), (1R,4R)-1-hydroxy-4H-1,2,3,4-tetrahydrocadalen-15-oic acid (4), quercetin (5), quercetin-3,7,3′-trimethyl ether (6), quercetin-3,7,3′,4′-tetramethyl ether (7), and eriodictyol-7,4′-dimethyl ether (8) were obtained from the acetone extract by successive fractionation processes using vacuum liquid chromatography (VLC), while semisynthetic compounds 7-acetoxy-3,4-dihydrocadalene (9), 7-benzoxy-3,4-dihydrocadalene (10), 7-acetoxycadalene (11), 7-benzoxycadalene (12), quercetin penta-acetate (13) and 7-hydroxycalamenane (14) (Fig. 1) were obtained by conventional chemical procedures as were previously described.¹⁹ Due to a shortage, some compounds were not tested in all assays.

Fig. 1 Natural products isolated from H. inuloides flowers and derivatives.

2.3. Caenorhabditis elegans strain

C. elegans wild-type strain Bristol N2 was obtained from the Caenorhabditis Genetics Center (Minneapolis-Saint Paul, MN, USA). The nematodes were maintained at 20 °C on NGM medium containing 3 g NaCl, 17 g agar, 2.5 g peptone, 1 mM CaCl₂, 1 mM MgSO₄ and 25 mM KH₂PO₄, per liter of water, as described previously.²⁰ Age-synchronized worms were generated in all experiments through a sodium hypochlorite method, and were allowed to hatch in Petri dishes with liquid S-medium (a minimal salt solution) seeded with a uracil auxotroph Escherichia coli OP50 (Caenorhabditis Genetics Center) strain as a food resource.²¹

2.4. Cell lines

The murine monocytic macrophage cell line RAW264.7 (ATCC® TIB-71™) was obtained from the American Type Culture Collection (Manassas, VA, USA) and maintained in Dulbecco’s Modified Essential Medium (DMEM), modified to contain 4 mM L-glutamine and 4.5 g L⁻¹ glucose, 1 mM sodium pyruvate and 1.5 g L⁻¹ sodium bicarbonate (ATCC® 30-2002™), and supplemented with 10% heat-inactivated fetal bovine serum (FBS) (ATCC® 30-2020™). Cells were grown at 37 °C, 5% CO₂ in a fully humidified atmosphere and used for experiments between passage 5 and 18. The viability of the cells was determined by Trypan blue exclusion.

2.5. Determination of polyphenolic content

The phenolic content was determined by a reaction with Folin–Ciocalteu (FC) reagent.²² Briefly, 1 mL of H. inuloides acetone extract (0.15 mg mL⁻¹ in DMSO/H₂O 1 : 1) was mixed with 1 mL of FC reagent (diluted 10-fold with water) and incubated at room temperature for 5 min. 1 mL of 75 mg mL⁻¹ Na₂CO₃ solution was added to the reaction mixture. The mixture was vortexed and incubated at room temperature in the dark for 30 min. Aliquots of the final reaction were added into each well of a 96-well plate, and the absorbance was measured at 760 nm using a Synergy-HT multi-detection microplate reader (BioTec Instruments, Inc., VT, USA). Gallic acid was used as a standard, and the results were expressed as μmol of gallic acid equivalents (GAE) per g of the extract.

2.6. UV-Vis analysis of flavonoid content

The flavonoid type compounds present in the H. inuloides acetone extract were distinguished by UV-Vis spectral analysis based on their ability to form complexes with AlCl₃. Due to the low solubility of the extract in aqueous solution, this was dissolved in methanol. Procedure 1: 1 mL of AlCl₃ solution (2%, w/v) was added to 1 mL of the test solution (0.15 mg mL⁻¹ in methanol) and 0.5 mL of water. The mixture was left for 10 min at room temperature and then subjected to spectral analysis in the range of 300–600 nm against the blank, using a UV-Vis Shimadzu U160 spectrophotometer (Shimadzu Co., Kyoto, Japan). Procedure 2: 1 mL of the test solution (0.15 mg mL⁻¹ in methanol) was mixed with 0.3 mL of NaNO₂ (5%, w/v) and after 5 min, 0.5 mL of AlCl₃ (2%, w/v) was added. The sample was mixed and 6 min later was neutralized with 0.5 mL of 1 M NaOH solution. The mixture was left for 10 min at room temperature and then subjected to spectral analysis in the range of 300–600 nm against the blank, wherein the amount of AlCl₃ solution was substituted by the same volume of water. For quantitative analysis, the absorbance of the samples was measured at 425 or 510 nm,²³ using as standards quercetin or catechin, respectively, to build up the calibration curve (concentration range: 30–500 μM). Aliquots of the final reaction were added into each well of a 96-well plate, and the absorbance was measured at 415 and 760 nm using a Synergy-HT multi-detection microplate reader (BioTec Instruments, Inc., VT, USA).

2.7. Stress resistance

Stress resistance assays were assessed in liquid medium at 20 °C in 96-well plates (Corning Life Sciences, NY, USA). Prior to the assays, the nematodes were age-synchronized in accordance with established protocols.²⁴ The nematodes were distributed in wells as L1 larvae (15–20 animals per well) together with Escherichia coli OP50 with a final concentration of 1.2 × 10⁹ bacteria per mL. To prevent self-fertilization, 5-fluoro-2′-deoxyuridine was added 36 h after seeding (0.12 mM final). The compounds whose effect on lifespan was to be tested were dissolved in a DMSO/methanol (1 : 1 v/v) solution (stocks: 100 mM). The final concentration of the DMSO was ≤0.6% in all cases. After the addition of the compounds, the sealed plates were shaken for 2–3 min and returned to the MaxQ 6000 incubator (ThermoFisher Scientific, MA., USA). The observations and counts were performed using an ECLIPSE TS100 inverted microscope (Nikon Instruments Inc., Tokyo, Japan). Worms were scored as dead when they did not respond to tactile or light stimuli. The mean percentage of live worms was calculated from three independent experiments.

2.7.1. Copper sulfate assay. Synchronized worms were dispensed into each well of a flat-bottom 96-well plate as L1 larvae (15–20 animals per well) and incubated in liquid S-medium containing an Escherichia coli OP50 strain as a food resource and either metabolites (1 mM) or DMSO/ethanol (1 : 1 v/v) as the solvent control for 1 h. At the tested concentration, no precipitation in the culture medium of the compounds was observed. The pre-treated worms were subjected to oxidative stress by the addition of CuSO₄ (final concentration 1 mM). For the experimental assay, CuSO₄–5H₂O was prepared in sterile distilled water by diluting a stock solution of 20.025 mM. Wells with sterile distilled water and DMSO/ethanol (1 : 1 v/v) were included as a solvent control; a positive control treated with catechin (1 mM) was included. The final volume was 250 μL per well. The plates were incubated for 24 h with shaking at 20 °C in a MaxQ 6000 incubator (ThermoFisher Scientific, MA., USA). After the incubation period with copper, the worms were observed under the microscope and the fraction alive was scored on the basis of body movement. In order to improve counting, the 96 well plates were shaken for 2 min before counting. Comparisons between treatments and controls were performed using the two-tailed, unpaired, Student’s t-test. Approximately 80–120 worms were scored in each experiment. Data were obtained from at least three independent experiments.

2.7.2. Juglone assay. Synchronized L1 larvae worms (15–20 animals per well) were pretreated with 100 μM of H. inuloides metabolites, and after 1 h the worms were treated with 250 μM of the pro-oxidant agent.²⁵ The plates were incubated with shaking at 20 °C in a MaxQ 6000 incubator (ThermoFisher Scientific, MA., USA). The number of dead worms was counted and recorded every hour to determine stress resistance; juglone was applied in 92% ethanol, 8% Tween 80 (v/v). The juglone was prepared by diluting a stock solution of 3.15 mM. The curves obtained were compared for significance using the Mantel–Cox (log-rank) test.

2.8. Spectrophotometric study of the interaction with copper

We evaluated the chelating ability and the interaction with copper of those compounds that showed ability to protect C. elegans using the techniques of UV-Vis spectroscopy, electron spin resonance (ESR) and electrospray ionization-mass spectrometry (ESI-MS).

2.8.1. UV-Vis analysis. 25 μM solutions were prepared in phosphate buffer solution (10 mM, pH 7.5). The UV-Vis-absorbance of the samples was subjected to spectral analysis at a constant temperature in the range of 300 to 600 nm using a UV-2700-CPS-240A spectrophotometer (Shimadzu Co. Kyoto, Japan), equipped with a temperature control unit. UVProbe software (Shimadzu) was used to collect and analyze the data. The same volume of 50 μM of CuSO₄ solution was added to the samples and 10 s later the samples were subjected to spectral analysis and compared with the compounds alone.²⁶

2.8.2. ESR (X-band) spectroscopy. The measurements were carried out in a quartz tube at 77 K in a 1 : 1 MeOH–H₂O (0.5 mM) solution, using a Jeol JES-TE300 spectrometer (JEOL Ltd. Tokyo, Japan) operating in X-Band fashions at 100 kHz modulation frequency and a cylindrical cavity in the mode TE011. The external calibration of the magnetic field was performed using a Jeol ES-FC5 precision gaussmeter (JEOL Ltd. Tokyo, Japan) with a 5350B HP microwave frequency counter (Hewlett Packard, California, USA). Spectral acquisition, manipulations and simulation were performed using ES-IPRITS-TE software (JEOL Ltd.). ESR spectra were recorded as a first derivative, and the main parameters, such as g-factor values, were calculated.

2.8.3. Analysis by electrospray ionization mass spectrometry. The analysis was carried out using an Esquire 6000 mass spectrometer (Bruker Daltonik GmbH, Bremen, Germany) with an electrospray ionization source and ion trap analyzer. Data processing was performed using DataAnalysis™ 3.2 software (Bruker Daltonik GmbH). The samples were diluted before measurement to 0.5 mg mL⁻¹ in methanol/water and were injected into an electrospray chamber by direct infusion, with a constant flow of 240 μL h⁻¹. Mass spectra were obtained in the positive and in the negative modes.

2.9. Nitric oxide production inhibition

In brief, RAW264.7 macrophage cells were seeded in 96 well microtitre plates (Corning Life Sciences, NY, USA) (2 × 10⁵ cells per well) and incubated for 2 h. The adherent cells were incubated with E. coli serotype 055:B5 lipopolysaccharide (1 μg mL⁻¹) in the absence or presence of 50 μL of each test compound dissolved in DMSO and serially diluted at a decreasing concentration of 100–3.1 μM, or aminoguanidine hydrochloride as an inducible NOS (iNOS) inhibitor. The final concentration of DMSO in the cell culture supernatant was ≤0.1% (v/v). Wells without treatment but stimulated with LPS were included as a control and received the same amount of DMSO. Cells were allowed to grow at 37 °C in a 5% CO₂ atmosphere. After 24 h, the inhibitory activity of the test compounds on NO production was determined by measuring the nitrite levels in the supernatants of the cultured RAW 264.7 cells. Briefly, 100 μL of the cell culture supernatant was mixed with the same volume of Griess reagent (1% sulfanilamide in 5% phosphoric acid and 0.1% N-(naphthyl)-ethylene-diamine dihydrochloride in water), and the absorbance of the mixtures was determined at 550 nm,²⁷ using a Synergy-HT multi-detection microplate reader (BioTek Instruments Inc., VT, U.S). The nitrite concentration (μM) was determined by interpolation with the standard curve constructed with the known concentrations of NaNO₂. The percentage of inhibition was calculated against cells that were not treated but were induced with LPS. DMEM alone was used as a blank in all experiments. Experiments were performed 3 times in triplicate.

2.10. MTT assay for cell viability

The viability of cells was determined using the reduction of MTT to formazan in the active mitochondria of viable cells.²⁸ Briefly, after 24 h incubation with the test compounds, 10 μL of MTT was added to the remaining cells in the 96-well plates and incubated for 4 h at 37 °C. The formazan products resulting from the dye reduction were dissolved in DMSO (100 μL). Absorbance was measured on a microplate reader (BioTek Instruments Inc., VT, USA) at 570 nm. The cell viability of the control wells was considered to be 100%. The test compounds were considered to be cytotoxic when the absorbance of the compound-treated wells was less than 80% of that in the control wells.

2.11. Statistical analysis

Data are expressed as mean ± SEM. The data of the stress resistance assays were processed using IBM SPSS Statistics 20, Chicago, IL, USA. The plots were processed using GraphPad Prism 5.0 Software (GraphPad Software, Inc. San Diego, CA, USA). Data were obtained from three independent experiments, and p < 0.05 was considered for significant differences.

3. Results and discussion

The aging process is characterized as a progressive decline in biological functions with time, and a decreased resistance to multiple forms of stress, as well as an increased susceptibility to numerous diseases.²⁹ Several studies have suggested that antioxidants prevent the appearance of age-related diseases and can increase lifespan.³⁰ We evaluate the ability of the metabolites isolated from H. inuloides to protect C. elegans against the damage generated by exogenous stressors, their copper chelating ability and their potential to inhibit NO production by LPS-stimulated macrophages.

3.1. UV-Vis analysis of phenolic and flavonoid type compounds of the H. inuloides extract

Previous studies have reported the phenolic and flavonoid content in acetone and methanol extracts of H. inuloides;¹⁸ however, these studies do not describe the type of flavonoid components. Aluminum chloride produces a bathochromic shift in most flavonoids,³¹ and this property was used to characterize the H. inuloides acetone extract. This extract does not exhibit the characteristic peak around 510 nm associated with the presence of compounds with aromatic rings containing a catechol group, or compounds containing catecholic moieties (Fig. 2). However, both spectra show an absorption maximum around 400 nm, this peak is attributed to flavonol or flavone–aluminum chlorides.²³ This is the first time that the spectral analysis of an extract obtained from H. inuloides is reported regarding the use of an AlCl₃ reaction for determining the type and content of flavonoid compounds. The phenolic content estimated using F–C reagent was 241.61 ± 5.9 μmol gallic acid per g extract. The content of flavonoid expressed as a quercetin equivalent was 137.85 ± 2.25 μmol quercetin per g extract.

Fig. 2 UV-Vis absorption spectra of the H. inuloides acetone extract. The solid line was obtained without NaNO₂, while the dashed line was obtained with NaNO₂, after addition of AlCl₃.

3.2. Copper sulfate assay

Copper is an essential trace element component of metalloenzymes capable of using oxygen or oxygen radicals as substrates by direct interaction with the Cu²⁺ site.³² The deficiency of this metal is associated with disorders such as neutropenia and anemia,³³ bone marrow suppression,³⁴ vascular lesions (Menkes disease),³⁵ hepatolenticular degeneration (Wilson disease)³⁶ and the inability of the organism to perform important biological processes.³⁷ However, exposure to high levels of free copper (non-covalently bound copper) has been associated with damage due its oxygen transferring properties, and its capability to act as a catalyst for oxidative damage by the removal of one electron from O₂ that results in the formation of O₂˙⁻ and the subsequent generation of H₂O₂, ONOO⁻ and ˙OH through the Fenton-catalysed Haber–Weiss reaction.³⁸ Copper also participates in the oxidation of nitroxyl anion radicals to biologically active nitric oxide.³⁹ It has been reported previously that exposure to copper reduces the feeding, and produces locomotion behavioral defects, neurodegeneration and toxic effects in C. elegans.^40,41

In the experiment, Cu²⁺ ions significantly decreased the C. elegans’ survival rate (5.66%), however, pretreatment with the acetone extract and with compounds 1 (7-hydroxy-3,4-dihydrocadalene), 3 (3,7-dihydroxy-3(4H)-isocadalen-4-one) and 5 (quercetin) was able to protect the nematodes from the mortality induced by copper. The survival rate of the worms pre-treated with extract and compounds 1, 3 and 5 was 94.26%, 67.20%, 84.89%, and 93.06%, respectively, compared to untreated ones (p < 0.05). Compounds 4, 10, and 13 showed poor protective activity to the worms. The remainder of the test compounds showed no statistically significant differences on the survival rate compared to the untreated control (Table 1). The protective effects of the H. inuloides acetone extract may be attributed to its ability to trap radicals.¹⁸ In addition, phenolic compounds have been isolated from this plant, some of which have demonstrated the ability to chelate metal ions.⁴²

Table 1 Effect of H. inuloides metabolites on the survival of C. elegans under copper (1 mM CuSO₄) induced stress^a

Compound	No. of			Estimated mean
	Subjects	Dead	Alive	Dead	Alive
a **P < 0.05. Comparisons between treatments and negative control differed significantly using the two-tailed, unpaired Student’s t-test.
1	375	252	123	32.80	67.20**
2	306	0	306	100	0
3	278	236	42	15.11	84.89**
4	333	66	267	80.18	19.82
5	288	268	20	6.94	93.06**
6	376	24	352	93.62	6.38
7	284	0	284	100	0
8	330	25	305	92.42	7.58
9	268	0	268	100	0
10	270	40	230	85.19	14.81
11	320	25	295	92.19	7.81
12	265	0	265	100	0
13	315	39	276	87.62	12.38
14	348	9	339	97.41	2.59
Acetone extract	366	345	21	5.74	94.26**
Catechin	328	212	116	35.37	64.63**
Control (−)	265	15	250	94.34	5.66

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3.3. Spectrophotometric study of the interaction with copper

Previously, the interaction of biologically relevant metal ions with phenolic compounds has been studied using spectroscopic and spectrometric techniques that include photometric titration, IR, Raman, UV-Vis, 2D NMR, mass spectrometry, thermo-gravimetric analysis, and voltammetry among others.^43–45 We used ultraviolet-visible, EPR spectroscopy and electrospray ionization mass spectrometry (ESI-MS) to study the interaction between Cu²⁺ and the acetone extract of H. inuloides, and compounds 1, 3 and 5 which showed a positive effect in the bioassay with C. elegans.

All the ESR spectra in a methanol–water frozen solution of copper–H. inuloides extract complexes (Fig. 3) exhibit an anisotropic ESR spectra characteristic of Cu²⁺ centers with well resolved hyperfine lines in the g_‖, and all signals show the dependence g_‖ > g_⊥ > g_e which is diagnostic of a d_x²−y² ground state for Cu²⁺, the hyperfine structure resulting from the interaction between delocalized π electrons and the nuclear spin I_Cu = 3/2. The values obtained from simulated spectra are shown in Table 2. The ESR spectra of the H. inuloides extract with the free copper ion showed a significant modification of the spectrum (Fig. 3) with spectral values g_‖ = 2.27, and A_‖ = 184 × 10⁻⁴ cm⁻¹ showing a full chelating effect, although the spectrum shows broad signals due to the overlap of signals from the multiple components of the extract including 5, which is the major flavonoid component, and also other compounds including sesquiterpenoids, phenolic compounds, phytosterols, and lipids, among others.^13,46

Fig. 3 Experimentally observed first-derivative ESR spectrum of a frozen solution (methanol/H₂O) of Cu²⁺ (a), Cu²⁺/acetonic extract (b) and Cu²⁺/5 (c).

Table 2 Spectral parameters obtained by simulation at 77 K of ESR spectra of the complexes

Sample	g‖	g⊥	A‖^a	A⊥^a	giso	Aiso^a	f
a Hyperfine coupling constants (A) are expressed in 1 × 10^4 cm^−1.
Cu^2+/ext.	2.272	2.058	184.36	13.48	2.1293	66.95	125.07
Cu^2+/1	2.420	2.076	131.9	9.7	2.1906	50.44	183.38
Cu^2+/3	2.420	2.081	132.8	9.71	2.194	50.53	183.07
Cu^2+/5	2.300	2.062	175.13	9.7	2.1412	64.79	131.32
Cu^2+	2.423	2.079	129.86	9.7	2.1936	49.75	186.58

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Compound 5 showed the greatest protective capacity against Cu²⁺. This flavonoid compound possesses three possible chelating sites: between the 3-hydroxyl group and the 4-oxo group (a), between the 5-hydroxyl group and the 4-oxo group (b), and between the ortho-hydroxyls groups in the B-ring (c). The chelation site involves preferably the positions “a” and/or “b”.⁴⁷ The UV-Vis spectrum for 5 showed two peaks associated with absorption due to the cinnamoyl system (B and C rings), and the benzoyl system (ring A).²⁶ The peak associated with the cinnamoyl system (peak I) undergoes a bathochromic shift of 371 to 434 nm (Δλ = 73 nm) due to the chelation of copper (Fig. 4). This amount of bathochromic shift (Δλ) has been associated with the formation of a 5–copper complex in the position “a”,⁴⁸ implying that deprotonation of the 3-OH takes place to form a 5–Cu²⁺ complex. The interactions determined by ESI-MS indicated that the complex formed between 5 and copper has the stoichiometry of copper–5 1 : 1 and 1 : 2. The fragments corresponding to the complex were observed at m/z = 382 [(M + H) + Cu²⁺ + H₂O]⁺, and m/z = 688 [(M + H) + Cu²⁺ + M + H₂O]⁺. The copper–5 complex with stoichiometry of 2 : 3 [(M + H) + Cu²⁺ + M + Cu²⁺ + M]⁺ was also probably present in solution because the molecular peak at m/z = 1028 can be observed, however, it was observed with very low intensity (Fig. 5). Some studies have been conducted to investigate the products of 5 with copper ions, and complexes with a range of stoichiometries of metal : flavonoid, 1 : 1, 1 : 2, 2 : 2 and 2 : 3 have been observed.^43,47,49,50 This range of results may be associated with the metal ion and the conditions of synthesis,⁴³ however, the stoichiometry 1 : 2 is in general, the preferred one.⁴⁷ For steric reasons the complexes usually include no more than two flavonoid molecules, and methanol solutions favor formation of 1 : 2 metal : flavonoid complexes. The ESR spectra obtained for the pure compound 5 and copper ions showed a complete chelating effect, with g_‖ = 2.30 and A = 175 × 10⁻⁴ cm⁻¹; these spectral changes are consistent with a square pyramid geometry.⁵¹ A complex of two 5 (quercetin) molecules coordinated in position “a” with a single Cu²⁺ ion and one molecule of solvent in the axial position is proposed (Fig. 6). These data were supported by a mass spectrum that showed a peak at m/z = 688. The tetrahedral distortion values (f) calculated for the extract, f = 125, were slightly lower than those of the complex with 5, f = 131. In addition, an increase of g_‖ from 2.270 to 2.30 and decrease of A_‖ from 184.3 to 175.1, respectively, were observed (Table 2); both parameters are indicative of a slight increase in the distortion of the 5–Cu complex, but they are both consistent with a square pyramid geometry.^51,52 The degree of tetrahedral distortion occurring frequently for the square planar copper complexes, is reflected by the ratio of f = g_‖/A_‖ and can be considered as a measure of distortion and the deviation from a perfect geometry depending on the nature of the coordinated ligands.^53,54 An f value in the range 110–120 is characteristic for the signals of a copper complex with planar geometry, whereas its increase to 150 indicates a slight or moderate distortion of the planar symmetry, and a further rise to the values 180–250 suggests a strong deviation from square planar geometry. In general, as a complex becomes more tetrahedral, the g_‖ value and the g_iso value increase, and the A_iso and A_‖ values decrease.⁵⁵ Additionally, a good estimation of the geometry adopted by the compounds can be done by employing the isotropic parameters of the ESR spectra A_iso= (A_‖ + 2A_⊥)/3 and g_iso=(g_‖ + 2g_⊥)/3. Values for planar symmetry are between A_iso = 75–90 cm⁻¹ and g_iso = 2.06–2.11.^56,57 Other compounds that showed a similar activity as protectors against copper were 1 and 3. Compound 3 is an α-hydroxy ketone, but unlike compound 5 (an α,β-unsaturated compound) in which the 3-hydroxyl and 4-oxo groups have been proposed as metal binding sites (site “a”), the hydroxyl group of 3 is attached to an sp³ carbon; furthermore, the presence of an isopropyl group increases steric hindrance and decreases nucleophilicity. The UV-Vis spectral analysis for compound 1 shows three bands attributable to benzene aromaticity with a bathochromic shift due to conjugation with another ring which has double bonds. The first absorption band is found at 217 nm, a secondary band at 265 nm and a third band at 302 nm. The spectrum obtained for compound 1 in the presence of copper showed a hypsochromic shift of the absorption peaks (ESI Fig. 1†). For compound 3, the spectral analysis showed a primary absorption band at 249 nm attributable to benzene aromaticity and a secondary band at 283 nm, associated with η–π*–ketone transitions. (ESI Fig. 2†). The limited capacity of compound 1 and 3 to chelate copper was evidenced by UV-Vis, ESI-MS and ESR spectral analysis, in which 1–Cu and 3–Cu species could not be observed. The UV-Vis spectral analysis does not show a bathochromic shift in the presence of copper. The ESI-MS spectra for 1 and 3 showed the characteristic [M + H]⁺ ions at m/z = 217 and at m/z = 247, respectively, and we also observed cluster molecules charged with Na⁺ and K⁺. No copper-containing species could be observed by ESI-MS. The ESR spectra obtained for 1 and 3 showed the same parameter as the Cu²⁺ aqua free ions g_‖ = 2.423 and A_‖ = 129.86 × 10⁻⁴ cm⁻¹, showing no effect on the ion chelating copper (Table 2). It is possible that these cadinene compounds occurring in H. inuloides may protect C. elegans from oxidative damage through other mechanisms and not by chelation. Previously, it has been reported that compound 1 inhibits lipid peroxidation, and protects against oxidative hemolysis,^14,16 while compound 3 showed moderate ability to trap ROS.⁵⁸ It is important to consider that unbound copper induces lipid peroxidation⁵⁹ and RONS production.³⁸

Fig. 4 UV-Vis absorption spectra of compound 5 in 10 mM buffer phosphate solution at pH 7.5. Solid line represents the compound alone (25 μM), while the dashed line represents the compound (25 μM) plus Cu²⁺ ions (50 μM).

Fig. 5 Electrospray ionization mass spectra in the positive mode of the 5–Cu²⁺ complex. Sections showing the distribution isotope patterns for positive ion clusters of (a) [(M + H) + Cu + H₂O]⁺, (b) [(M + H) + Cu + M + H₂O]⁺ and (c) [(M + H) + Cu + M + Cu + M]⁺.

Fig. 6 Complex forming sites and proposed structure for [CuL₂] (L = 5). S = solvent.

3.4. Juglone assay

On the other hand, some metabolites of H. inuloides showed protective effects to C. elegans against stress generated by juglone. Agents such as juglone are cytotoxic due to their capacity to produce ¹O₂; and H₂O₂ through O₂ reduction by semiquinone intermediates, which enables them to form adducts to cellular constituents, and may thereby cause oxidative stress.^60,61 Pretreatment of the C. elegans N2 wild-type with 100 μM of certain metabolites is indeed able to protect the organism against lethal oxidative stress generated by juglone and increase survival rates (Table 3). Compounds 5, 3 and 6 caused an increased survival during juglone-induced oxidative stress compared to the untreated controls. Compounds 4, 7, 10, 12 and 13 showed no statistically significant differences compared to the control, whereas compounds 9, 11 and 14 had a negative effect on the survival of the worms.

Table 3 Protective effects of H. inuloides metabolites on C. elegans under juglone-induced oxidative stress^a

Compound	Juglone 250 mM
	Mean ± SE (h)	N treated	Log-rank χ^2	P-value^b
a **P < 0.01, *P < 0.05.b Determined by log-rank test.
3	2.058 ± 0.053	242	4.004	0.0454*
4	1.898 ± 0.051	310	0.0496	0.8239
5	2.074 ± 0.041	335	51.67	<0.0001**
6	2.066 ± 0.047	280	3.858	0.0495*
7	1.905 ± 0.043	322	0.045	0.8312
8	2.045 ± 0.047	301	4.373	0.0365*
9	1.248 ± .037	214	110.6	<0.0001**
10	1.859 ± 0.046	311	0.001	0.9745
11	1.481 ± 0.034	329	95.75	<0.0001**
12	1.791 ± 0.051	289	0.904	0.3416
13	1.951 ± 0.048	307	0.843	0.3586
14	0.688 ± 0.018	255	415.6	<0.0001**
Control	1.915 ± 0.042	248	—	—

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Compound 5 showed the greatest protective effects. In this compound an ortho-dihydroxy group and a 2,3-double bond in conjugation with a 4-oxo function confer the structural characteristics necessary to be considered good for radical trapping.⁶² The mean life-span of the worms after treatment with 250 μM juglone was 1.91 ± 0.042 h, and for the group pretreated with 5 for 1 h was 2.074 ± 0.041 h (p < 0.001). The O-methylation and O-acetylation of 5 resulted in minor activity in 6, 7 and 13. These results are consistent with reports that indicate that the RONS scavenging activity of flavonoids is highly dependent on the presence or modification of hydroxyl substituents.⁶³ We found no significant effect of cadinene type sesquiterpenes against the stress caused by juglone, and this can be explained by the fact that the H. inuloides cadinenes are not good scavengers of O₂˙⁻.¹⁶

3.5. Nitric oxide production inhibition

NO participates in the control of pathogens and in the regulation of blood pressure and cardiovascular health, and it acts as a neurotransmitter and modulates many functions.⁶⁴ However, this lipid- and water-soluble radical gas can react with oxygen and generate NO₂, NO₂⁻, NO₃⁻, N₂O₃, and the strong oxidant peroxynitrite ONOO⁻, which in turn induce inflammatory cytokines, which lead to induced cell death by apoptosis and necrosis.^65,66 Natural products can act as alternative inhibitors of nitric oxide synthase. H. inuloides is a plant widely used in Mexican traditional medicine for the treatment of bruises and injuries associated with inflammatory processes.¹² From this plant various metabolites with anti-inflammatory activity were isolated,^13,58 however, the ability of these metabolites to inhibit nitric oxide synthesis has not been reported. We examined the inhibitory activity of the main components from an acetone extract and some semisynthetic derivatives. Table 4 shows the inhibitory activity of H. inuloides compounds towards NO production by LPS-activated macrophages. Compounds 3, 5, 6, 8, 10 and 12 showed greater than 25% inhibition of NO production at a concentration of 25 μg mL⁻¹. Among these six compounds, 5 (quercetin) and 8 (eriodictyol-7,4′-dimethyl ether) significantly decreased the levels of NO production from LPS-stimulated RAW264.7 cells, by 100%, and 87.3%, respectively. This result is congruent with the previous report on the iNOS inhibitory effects of flavonoids.⁶⁷ The numbers of viable activated macrophages were not significantly altered, thereby indicating that the inhibition of NO synthesis by compounds 5 and 8 was not due to cytotoxic effects. The substitution of the hydroxyl groups in 5 produces a decrease in the ability to inhibit NO production in O-methylated derivatives 6 and 7. Compounds 1 and 2 were found to be toxic to RAW264.7 cells as determined by an MTT assay. On the other hand, the benzoylated derivatives 10 and 12 showed moderate effects without affecting the viability of the activated RAW264.7 macrophages. Significant differences in NO production between 10 and 12 were not observed. The cadinene 4 showed the lowest inhibitory activity without affecting the viability of the macrophages. However, acetylated derivatives 9 and 11 showed similar toxic effects to 1 on the viability of the macrophages.

Table 4 Nitric oxide inhibition by H. inuloides metabolites

Compound (25 μg mL^−1)	Nitrites^a (μM)	NO inhibition (%)
a Value represents mean ± S.E.M.b Cytotoxic effect in murine RAW 264.7 macrophages was observed with compounds 1, 2, 9 and 11. With the remaining compounds, the observed viability was greater than 80%. (—) = not determined.
LPS 1 μg mL	14.99 ± 0.86	0
1	—	^b
2	—	^b
3	10.51 ± 0.52	29.9
4	13.25 ± 0.66	11.6
5	0	100
6	6.55 ± 0.32	56.30
7	12.67 ± 0.60	15.47
8	1.9 ± 0.90	87.3
9	—	^b
10	7.04 ± 0.35	53.03
11	—	^b
12	7.91 ± 0.39	47.23
Oleic acid	12.0 ± 0.60	19.94

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4. Conclusions

In this work C. elegans was exposed to the oxidative stressors copper and juglone. We find that some compounds of H. inuloides cause a significant increase in the C. elegans life span. Particularly, quercetin (5) showed the highest activity. This activity is important because metal chelation is considered as another mechanism for the antioxidant effect. The spectrophotometric analyses, using the techniques of UV-Vis, ESR and ESI-MS provide evidence for complex formation between 5 and Cu²⁺, with a metal : ligand stoichiometry of 1 : 2. Cu²⁺ ions are coordinated with 5 in position “a”. Cadinane type compounds have a minimal protective effect against oxidative stress inductors in the cultured C. elegans. The flavonoids of H. inuloides showed a greater ability to inhibit nitric oxide synthesis than the compounds with a cadinene skeleton. Generally the modification of the hydroxyl groups of the flavonoids and cadinene compounds modifies their ability to inhibit the synthesis of nitric oxide.

Conflict of interest

The authors declare that there are no actual or potential conflicts of interest in relation to this article. The authors alone are responsible for the content and writing of the paper.

Funding

This work was partially supported by UNAM-DGAPA-PAPIIT (Project IG200514) and by the Consejo Nacional de Ciencia y Tecnología (CONACyT, México) through a scholarship granted to J. L. Rodríguez-Chávez to pursue his Doctor Sc. Degree. The funders had no role in study design, data collection and analysis, decision to publish or preparation of the manuscript.

Acknowledgements

We thank Chem. J. Ebrard (Laboratorios Mixim) for providing the plant material and M. Sc. Abigail Aguilar-Contreras (Instituto Mexicano del Seguro Social) for the identification of the species. We also thank Antonio Nieto Camacho, Teresa Ramírez Apan, Rocío Patiño Maya, Lucía del Carmen Márquez, Eréndira García and Lucero Ríos (Instituto de Química, UNAM) for technical assistance.

References

1. N. Bashan, J. Kovsan, I. Kachko, H. Ovadia and A. Rudich, Physiol. Rev., 2009, 89, 27–71.
2. S. Zhao, Z. Xiong, X. Mao, D. Meng, Q. Lei, Y. Li, D. Meng, Q. Lei, P. Deng, M. Chen, M. Tu, X. Lu, G. Yang and G. He, PLoS One, 2013, 8, e73665.
3. I. Martin and M. S. Grotewiel, Mech. Ageing Dev., 2006, 127, 411–423.
4. R. A. Roberts, R. A. Smith, S. Safe, C. Szabo, R. B. Tjalkens and F. M. Robertson, Toxicology, 2010, 276, 85–94.
5. A. Weidinger and A. V. Kozlov, Biomolecules, 2015, 5, 472–484.
6. H. J. H. Fenton, J. Chem. Soc. Proc., 1894, 10, 157–158.
7. F. Haber and J. Weiss, Proc. R. Soc. London, 1934, 147, 332–351.
8. A. Lemire, J. J. Harrison and R. J. Turner, Nat. Rev. Microbiol., 2013, 11, 371–384.
9. J. Prousek, Pure Appl. Chem., 2007, 79, 2325–2338.
10. A. Argueta, L. Cano and M. E. Rodarte, Atlas de las plantas medicinales de la medicina tradicional mexicana, Instituto Nacional Indigenista, Mexico City, 1994, vol. 1–3.
11. M. Martínez, Catálogo alfabético de nombres vulgares y científicos de plantas que existen en México, ed. Botas, Mexico City, 1937.
12. X. Lozoya, A. Aguilar and J. R. Camacho, Rev. Méd. IMSS., 1987, 25, 283–286.
13. G. Delgado, M. S. Olivares, M. I. Chávez, T. Ramírez-Apan, E. Linares, R. Bye and F. J. Espinosa-García, J. Nat. Prod., 2001, 64, 861–864.
14. H. Haraguchi, T. Saito, H. Ishikawa, Y. Sánchez, T. Ogura and I. Kubo, J. Pharm. Pharmacol., 1996, 48, 441–443.
15. I. Kubo, S. K. Chaudhuri, Y. Kubo, Y. Sánchez, T. Ogura, T. Saito, H. Ishikawa and H. Haraguchi, Planta Med., 1996, 62, 427–430.
16. H. Haraguchi, H. Ishikawa, Y. Sánchez, T. Ogura, Y. Kubo and I. Kubo, Bioorg. Med. Chem., 1997, 5, 865–871.
17. I. Kubo, I. Kinst-Hori, S. K. Chaudhuri, Y. Kubo, Y. Sánchez and T. Ogura, Bioorg. Med. Chem., 2000, 8, 1749–1755.
18. E. Coballase-Urrutia, J. Pedraza-Chaverri, R. Camacho-Carranza, N. Cárdenas-Rodríguez, B. Huerta-Gertrudis, O. N. Medina-Campos, M. Mendoza-Cruz, G. Delgado-Lamas and J. J. Espinosa-Aguirre, Toxicology, 2010, 276, 41–48.
19. J. L. Rodríguez-Chávez, Y. Rufino-González, M. Ponce-Macotela and G. Delgado, Parasitology, 2015, 142, 576–584.
20. S. Brenner, Genetics, 1974, 77, 71–94.
21. J. A. Lewis and J. T. Fleming, in Methods in cell biology, ed. H. F. Epstein and D. C. Shakes, Academic Press, San Diego, 1995, vol. 48, p. 3.
22. J. C. Sánchez-Rangel, J. Benavides, J. B. Heredia, L. Cisneros-Zevallos and D. A. Jacobo-Velázquez, Anal. Methods, 2013, 5, 5990–5999.
23. A. Pekal and K. Pyrzynska, Food Anal. Method., 2014, 7, 1776–1782.
24. G. M. Solis and M. Petrascheck, J. Visualized Exp., 2011, 49, 1–6.
25. E. de Castro, S. H. de Castro and T. E. Johnson, Free Radical Biol. Med., 2004, 37, 139–145.
26. B. Bukhari, S. Memon, M. Mahroof-Tahir and M. I. Bhanger, Spectrochim. Acta, Part A, 2009, 71, 1901–1906.
27. L. C. Green, D. A. Wagner, J. Glogowski, P. L. Skipper, J. S. Wishnok and S. R. Tannenbaum, Anal. Biochem., 1982, 126, 131–138.
28. T. Mosmann, J. Immunol. Methods, 1983, 16, 55–63.
29. K. C. Kregel and H. J. Zhang, Am. J. Physiol.: Regul., Integr. Comp. Physiol., 2006, 292, 18–36.
30. S. Casani, R. Gómez-Pastor, E. Matallana and N. Paricio, Free Radical Biol. Med., 2013, 61, 151–160.
31. T. B. Gage and S. H. Wender, Proc. Okla. Acad. Sci., 1949, 30, 145–148.
32. R. A. Festa and D. J. Thiele, Curr. Biol., 2011, 8, R877–R883.
33. W. Harless, E. Crowell and J. Abraham, Am. J. Hematol., 2006, 81, 546–549.
34. G. J. Bbrewer, J. Cell. Mol. Med., 2003, 7, 11–20.
35. Z. Tümer and B. Møller, Eur. J. Hum. Genet., 2010, 18, 511–518.
36. B. E. Reis, M. A. A. Costa, C. E. L. Rachid, D. M. Mitiko and M. Scaff, Arq. Neuro-Psiquiatr., 2009, 67, 539–543.
37. J. Y. Uriu-Adams and C. L. Keen, Mol. Aspects Med., 2005, 6, 268–298.
38. A. N. Pham, G. Xing, C. J. Miller and T. D. Waite, J. Catal., 2013, 301, 54–64.
39. S. Nelli, M. Hillen, K. Buyukafsar and W. Martin, Br. J. Pharmacol., 2000, 131, 356–362.
40. G. L. Anderson, W. A. Boyd and P. L. Williams, Environ. Toxicol. Chem., 2001, 20, 833–838.
41. P. J. Chen, E. Martinez-Finley, J. Bornhorst, S. Chakraborty and M. Aschner, Front. Aging Neurosci., 2013, 5, 1–11.
42. P. Mladenka, K. Macáková, T. Filipský, L. Zatloukalová, L. Jahodár, P. Bovicelli, I. P. Silvestri, R. Hrdina and L. Saso, J. Inorg. Biochem., 2011, 105, 693–701.
43. S. Domínguez, J. Torres, P. Morales, H. Heinzen, A. Bertucci and C. Kremer, in Coordination Chemistry Research Progress, ed. T. W. Cartere and K. S. Verley, Nova Science Publishers Inc., NY, USA, 2008, ch. 10, pp. 305–328.
44. K. Brodowska, Biotechnol. Food Sci., 2013, 77, 45–53.
45. M. M. Kasprzak, A. Erxleben and J. Ochocki, RSC Adv., 2015, 5, 45853–45877.
46. R. O. F. Mijangos, J. Ruiz-Jiménez, L. Lagunez-Rivera and M. D. de Castro, Phytochem. Anal., 2011, 22, 484–941.
47. M. T. Fernandez, M. L. Mira, M. H. Florêncio and K. R. Jennings, J. Inorg. Biochem., 2002, 92, 105–111.
48. Y. Takamura and M. Sakamoto, Chem. Pharm. Bull., 1987, 26, 2291–2297.
49. J. E. Brown, H. Khodr, R. C. Hider and C. A. Rice-Evans, Biochem. J., 1998, 330, 1173–1178.
50. A. Pękal, M. Biesaga and K. Pyrzynska, BioMetals, 2011, 24, 41–49.
51. M. A. Maldonado-Rogado, E. Viñuelas-Zahínos, F. Luna-Giles and A. Bernalte-García, Polyhedron, 2007, 26, 1173–1181.
52. C. Karunakaran, K. R. J. Thomas, A. Shunmugasundaram and R. Murugesan, J. Chem. Crystallogr., 1999, 29, 413–420.
53. M. Łabanowska, E. Bidzińska, A. Para and M. Kurdziela, Polymers, 2012, 87, 2605–2613.
54. R. Pogni, M. C. Baratto, A. Diaz and R. Basosi, J. Inorg. Biochem., 2000, 79, 333–337.
55. S. J. Cline, J. R. Wasson, W. E. Hatfield and D. J. Hodgson, J. Chem. Soc., Dalton Trans., 1978, 1051–1057.
56. M. Joseph, M. Kuriakose, K. M. R. Prathapachandra, E. Suresh, A. Kishore and S. G. Bhat, Polyhedron, 2006, 25, 61–70.
57. E. Suresh, M. M. Bhadbhade and D. Srinivas, Polyhedron, 1996, 15, 4117–4310.
58. J. L. Rodríguez-Chávez, E. Coballase-Urrutia, A. Nieto-Camacho and G. Delgado-Lamas, Oxid. Med. Cell. Longevity, 2015, 2015(843237), 1–11.
59. W. Y. Boadi, P. A. Iyere and E. Adunyah, J. Appl. Toxicol., 2003, 23, 363–369.
60. K. Ollinger, J. Llopis and E. Cadenas, Arch. Biochem. Biophys., 1989, 275, 514–530.
61. J. M. Van Raamsdonk and S. Hekimi, Proc. Natl. Acad. Sci. U. S. A., 2012, 109, 5785–5790.
62. D. Procházková, I. Boušová and N. Wilhelmová, Fitoterapia, 2011, 82, 513–523.
63. M. A. Soobrattee, V. S. Neergheen, A. Luximon-Ramma, O. I. Aruoma and T. Bahorun, Mutat. Res., 2005, 579, 200–213.
64. M. Rosselli, P. J. Keller and R. K. Dubey, Hum. Reprod. Update, 1998, 4, 3–24.
65. D. Burgner, K. Rockett and D. Kwiatkowski, Arch. Dis. Child., 1999, 81, 185–188.
66. M. N. Hughes, Biochim. Biophys. Acta, 1999, 1411(2–3), 263–272.
67. S. K. Kim, H. J. Kim, S. E. Choi, K. H. Park, H. K. Choi and M. W. Lee, Arch. Pharmacal Res., 2008, 3, 424–428.

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Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c5ra21646j

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