Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Effects of Tithonia diversifolia (Hemsl.) A. Gray Extract on Adipocyte Differentiation of Human Mesenchymal Stem Cells

Effects of Tithonia diversifolia (Hemsl.) A. Gray Extract on Adipocyte Differentiation of Human Mesenchymal Stem Cells

  • Claudia Di Giacomo, 
  • Luca Vanella, 
  • Valeria Sorrenti, 
  • Rosa Santangelo, 
  • Ignazio Barbagallo, 
  • Giovanna Calabrese, 
  • Carlo Genovese, 
  • Silvana Mastrojeni, 
  • Salvatore Ragusa, 
  • Rosaria Acquaviva


Tithonia diversifolia (Hemsl.) A. Gray (Asteraceae) is widely used in traditional medicine. There is increasing interest on the in vivo protective effects of natural compounds contained in plants against oxidative damage caused from reactive oxygen species. In the present study the total phenolic and flavonoid contents of aqueous, methanol and dichloromethane extracts of leaves of Tithonia diversifolia (Hemsl.) A. Gray were determined; furthermore, free radical scavenging capacity of each extract and the ability of these extracts to inhibit in vitro plasma lipid peroxidation were also evaluated. Since oxidative stress may be involved in trasformation of pre-adipocytes into adipocytes, to test the hypothesis that Tithonia extract may also affect adipocyte differentiation, human mesenchymal stem cell cultures were treated with Tithonia diversifolia aqueous extract and cell viability, free radical levels, Oil-Red O staining and western bolt analysis for heme oxygenase and 5'-adenosine monophoshate-activated protein kinase were carried out. Results obtained in the present study provide evidence that Tithonia diversifolia (Hemsl.) A. Gray exhibits interesting health promoting properties, resulting both from its free radical scavenger capacity and also by induction of protective cellular systems involved in cellular stress defenses and in adipogenesis of mesenchymal cells.


The genus Tithonia comprises 13 taxa, which are distributed in eleven species. Tithonia diversifolia (Hemsl.) A. Gray (Asteraceae) is native to Mexico and also grows in parts of Africa, Australia, Asia, and other countries of North America. Tithonia diversifolia (Hemsl.) A. Gray and its extracts are traditionally used for the treatment of diabetes, diarrhea, menstrual pain, malaria, hematomas, hepatitis, hepatomas, and wound healing [1, 2]. Recently it has been suggested that these effects might be ascribed to terpenoids and flavonoids contained in the aerial parts of Tithonia diversifolia [3, 4]. Several studies investigated anti-inflammatory, analgesic, antimalarial, antimicrobial and antidiabetic activities; although these investigations revealed the potential of this plant and its constituents for different pharmacological/therapeutic activities [4], studies are needed in order to understand the molecular modes of action of Tithonia and its extracts.

Recently, intense interest has focused on the antioxidant properties of natural products. In particular, natural products that may act by preventing the free radical generation, neutralizing free radicals by non-enzymatic mechanisms and/or by enhancing the activity of endogenous antioxidants [5] such as stress-inducible proteins. Heme oxygenase (HO) (EC is a microsomal enzyme that oxidatively cleaves heme and produces biliverdin, carbon monoxide (CO) and iron [6]. To date, two isoforms of HO have been identified: HO-1, or inducible enzyme, and HO-2 or constitutive isoform [610]. A substantial body of evidence demonstrates that HO-1 induction represents an essential step in cellular adaptation to stress subsequent to pathological events [1114]; then HO-1 hyper-expression can be considered both a marker of cellular stress and also regarded as a potential therapeutic target in a variety of oxidant-mediated diseases [15].

Recently it has been reported that polyphenolic natural compounds are able to induce potently HO-1 expression, exercising protective effects. As a consequence, the beneficial actions attributed to several natural substances could be also due to their intrinsic ability to activate the HO-1 pathway [1618].

Adipocytes play a central role in regulating adipose mass and obesity. The increased adipose mass in obesity is caused both by adipose tissue hypertrophy, and also by adipose tissue hyperplasia, which affects the transformation of pre-adipocytes into adipocytes [19, 20]. Thus, adipocyte differentiation and the amount of fat accumulation are associated with the development of obesity.

Obesity was correlated to high levels of lipid peroxidation and/or decreased antioxidant capacities [21, 22]. When free-radical formation exceeds protective antioxidant mechanisms, or the later are compromised, oxidative stress occurs; increasing evidence from research show that oxidative stress is associated with the pathogenesis of obesity and it has been demonstrated that in vitro pre-adipocyte proliferation and differentiation can be controlled by redox metabolism [2326] suggesting that reactive oxygen species (ROS) may be involved in adipocyte differentiation.

The 5'-adenosine monophoshate-activated protein kinase (AMPK) is a heterotrimeric protein consisting of one catalytic subunit (α) and two non-catalytic subunits (β and γ). As AMP binds to the γ-subunit, it undergoes a conformational change which exposes the catalytic domain found in the α subunit; AMPK becomes activated (pAMPK) when phosphorilation occurs at the threonine residue within the kinase domain [27, 28]. AMPK has been proposed to act as a main metabolic switch in response to changes in cellular meabolism [29]. When activated, AMPK leads to an inhibition of energy-consuming biosynthetic pathways and concomitant activation of catabolic ATP-producing pathways [30, 31]. AMPK also acts as a fuel sensor in regulating glucose and lipid homeostasis in adipocytes by many additional effects both on genes and specific enzymes [32, 33].

In the present study the free radical scavenging capacity of different concentrations of aqueous, methanolic and dichloromethane leaf extracts of Tithonia diversifolia (Hemsl.) A. Gray was evaluated by in vitro assays; moreover the ability of Tithonia extract to inhibit plasma lipid peroxidation in a cell-free system was also tested.

In order to test the hypothesis that Tithonia leaf extract may also affect adipocyte differentiation, human mesenchymal stem cells (hMSC) were cultured in the presence or absence of Tithonia diversifolia (Hemsl.) A. Gray extracts and adipogenesis was measured by Oil-Red O staining. In the same cell cultures ROS levels were determined, and HO-1 and pAMPK expressions were also evaluated by western blot analysis.

Material and Methods

Ethics Statement

According to Italian law, we have to ask the opinion of the Ethics Committee only in the case of clinical trials; if blood, tissues or cells from donors or patients are used for research purposes, it is necessary that donors/patients give their written consent, not subject to the opinion of the Ethics Committee. The donors/patients are anonymous, because their generality are known only to the doctor who took the sample and that keeps their written consent. However in order to avoid any quandary we obtained the consent by Ethics Committee (N° IRB IOM 07 2012 of 08 February 2012).


Water, methanol and dichloromethane used for the extractions were of analytical grade and were purchased from Merck S.p.A. (Milano, Italy); all the other solvent, chemicals and reference compounds were purchased from Sigma-Aldrich (Milano, Italy).

Plant collection and preparation of extracts

The leaves of Tithonia diversifolia (Hemsl.) A. Gray were collected in August 2010 in the Eastern Democratic Republic of Congo and kindly provided by Sister Kavira Muhyama, Diocese of Butembo, Bunyuka (GPS Coordinates: Lat. 0.127752°; Long. 29.287371. B.P., 179 Butembo, Nord-Kivu). The field studies did not involve endangered or protected species. Sister Kavira Muhyama issued the permit for each location.

A voucher specimen of the plant was deposited in the herbarium of Department of Health Sciences, University “Magna Graecia” of Catanzaro.

Crude methanolic and dichloromethane extracts were obtained by maceration of 5 g of each powdered plant sample three times in 50 mL of solvent, for 45 min under constant shaking at room temperature. For the aqueous extracts, 100 mL of distilled water was used to extract 2 g of powdered plant material and the mixture obtained was boiled for 1 h. The extracts were filtered and evaporated to dryness under reduced pressure with a rotatory evaporator.

Total phenolic and flavonoid content

The concentration of total phenolic compounds was determined spectrophotometrically, using the Folin–Ciocalteu total phenols procedure, described by Ballard et al. [34], with modifications. Known amounts of Gallic acid were used to prepare the standard curve. Appropriately diluted (3.5%, w/v) test extracts (0.1 mL) and the gallic acid standard solutions (0.1 mL) were transferred to 15 mL test tubes. 3.0 mL of 0.2 N Folin–Ciocalteu reagent were added to each test tube and mixed using a vortex mixer. After 1 min, 2.0 mL of 9.0% (w/v) Na2CO3 in water were added and the solution was mixed. Absorbance was determined at λ = 765 nm. The concentration of total phenolic compounds in the extracts was determined comparing the absorbance of the extract samples to that of the gallic acid standard solutions. All samples were determined in triplicate. Total phenolic content was expressed as μMoles gallic acid/L ± S.D. Results represent the mean ± S.D. of 5 determinations.

The flavonoid concentration was measured using a colorimetric assay [35], with modifications. A standard curve of cathechin was used for quantification. Briefly, 25 mL of aqueous, methanolic and dichloromethane extracts and/or cathechin standard solutions were added to 100 mL of H2O. At time zero, 7.5 mL of 5% NaNO2 were added; after 5 min, 7.5 mL of 10% AlCl3 were added and at 6 min, 50 mL of 1 M NaOH were added. Each reaction mixture was then immediately diluted with 60 mL of H2O and mixed. Absorbances of the mixtures upon the development of pink color were determined a λ = 510 nm relative to a prepared blank. The total flavonoid contents of the samples are expressed as μMoles catechin/L. Each result represents the mean ± S.D. of 5 experimental determinations.

Scavenger effect on superoxide anion (SOD-like activity)

Superoxide anion was generated in vitro as described by Acquaviva et al. 2012 [36]. A total volume of 1 mL of the assay mixture contained: 100 mM triethanolamine-diethanolamine buffer, pH 7.4, 3 mM NADH, 25 mM/12.5 mM EDTA/MnCl2, 10 mM β-mercapto-ethanol; samples contained different concentrations of the three (aqueous, methanolic and dichloromethane) extracts of leaves of Tithonia diversifolia (Hemsl.) A. Gray. After 20 min incubation at 25°C, the decrease in absorbance at λ = 340 nm was measured. Results are expressed as percentage of inhibition of NADH oxidation. SOD (80 mU) was used as reference compound. Each result represents the mean ± S.D. of 5 experimental determinations.

Determination of lipid hydroperoxide levels in the plasma of a healthy donor

Heparinized venous blood of a healthy volunteer donor (male, 27 years old), who agreed to take part in the study and gave his written consent. Since this is a non-therapeutic trial it was carried out with the consent of the subjects legally acceptable according our Italian Government (Legge 675/1996 and DL 196/2003, art. 40. Art 32 Codice Italiano di Deontologia Medica).

Heparinized venous blood was collected after overnight fasting. Plasma was separated by centrifugation at 800 g for 20 min. Plasmatic lipid hydroperoxide levels were evaluated by oxidation of Fe2+ to Fe3+ in the presence of xylenol orange at λ = 560 nm [37]. Plasma aliquots (500 μL) were diluted 1:1 with oxygenated PBS and incubated at 37°C for 2 h with or without different concentrations of the aqueous extracts in a total volume of 1 mL. Results are expressed as percentage of inhibition respect to control (plasma incubated in absence of test compounds) and represent the mean ± S.D. of 5 experimental determinations.

Isolation and adipogenic differentiation of human adipose MSCs

Adipose tissue sample was obtained from a patient underwent abdominal plastic surgery (male, 30 years old, 98 kg b.w.); the subject provided his written consent before inclusion in the study. Since this is a non-therapeutic trial, it was carried out with the consent of the subject legally acceptable according our Italian Government (Legge 675/1996 and DL 196/2003, art. 40. Art 32 Codice Italiano di Deontologia Medica).

Adipose tissue sample was removed under sterile conditions, washed in PBS, minced, and digested with 1 mg/mL collagenase type I in 0.1% BSA for 1 h at 37°C in a shaking water bath. The pellet was collected by centrifugation at 650 g for 10 min and then treated with red blood cell lysis buffer (155 mM NH4Cl, 10 mM KHCO3 and 0.1 mM EDTA) for 10 min at room temperature. After centrifugation, the cellular pellet was filtered through a 100-μm mesh filter to remove debris. The filtrate was centrifuged, and the obtained stromal vascular fraction was plated onto 100 mm cell culture dishes in complete culture medium (DMEM containing 20% fetal bovine serum, 100 μg/mL streptomycin, 100 U/mL penicillin, 2 mM L-glutamine, and 1 μg/mL amphotericin-B). Cells were cultured at 37°C in humidified atmosphere with 5% CO2. After 24 h, non-adherent cells were removed, and adherent cells were washed twice with PBS. Confluent cells were trypsinized and expanded in T75 flasks (passage 1). A confluent and homogeneous fibroblast-like cell population was obtained after 2–3 weeks of culture. For all the experiments, only cells at early passages were used. At 50–60% confluence the medium was replaced with adipogenic medium, and the cells were cultured for additional 14 days. The adipogenic media consisted of complete culture medium supplemented with DMEM-high glucose (4.5 g/L), 10% (w/v) fetal bovine serum (FBS), 10 mg/mL insulin, 0.5 mM dexamethasone and 0.1 mM indomethacin. During adipogenic differentiation some flasks were added with different concentrations of the Tithonia extract. Media were changed every 2 days.

Cell viability assay

Cell viability was determined by the MTT assay, measuring the activity of cellular enzymes that reduce the tetrazolium dye, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide, a yellow tetrazole (MTT), to its insoluble formazan, giving a purple color in living cells [38]. The cells were plated at 8x103 cells per well of a 96-multiwell flat-bottomed 200 μl microplate. The optical density of each well sample was measured with a microplate spectrophotometer reader (Synergy HT multi-mode microplate reader, BioTek, Milano, Italy) at λ = 570 nm. Results are expressed as percentage of cell viability; each result represents the mean ± S.D. of 5 experimental determinations.

Oil Red O staining

Oil Red O staining is an assay performed to stain mature adipocytes in adipogenic-induced cultures. The mechanism of the staining of lipids is a function of the physical properties of the dye, which more soluble in the lipid than in the vehicular solvent. For Oil Red O staining, 0.21% Oil Red O in 100% isopropanol was used. Briefly, MSC-derived adipocytes, after 14 days, were fixed in 10% formaldehyde, washed in Oil-red O for 10 min, rinsed with 60% isopropanol, the Oil red O eluted by adding 100% isopropanol for 10 min and Optical Density measured at λ = 490 nm in a microplate reader (Synergy HT multi-mode microplate reader, BioTek, Milano, Italy). Results are expressed as optical density (OD) at λ = 490 nm. Each result represents the mean ± S.D. of 5 experimental determinations.

Determination of ROS

Determination of ROS was performed by using the fluorescent probe 2',7'-dichlorofluorescein diacetate, DCFH-DA, as previously described [39]. Briefly, 100 μL of 100 μM DCFH-DA, dissolved in 100% methanol, was added to the cellular medium, and cells were incubated at 37°C for 30 minutes. After incubation, cells were lysed and centrifuged at 10,000 x g for 10 min. The fluorescence (corresponding to oxidized CDF) was monitored spectrofluorometrically (λex = 488 nm; λem = 525 nm), using an F-2000 spectrofluorimeter (Hitachi) and results were expressed as Fluorescence Intensity (F.I.) x 1 x106/mg protein. Total protein content in each sample was evaluated according to Lowry et al. [40].

Western blot analysis

hMSCs cells were washed with PBS and then trypsinized (0.05% trypsin w/v with 0.02% EDTA). The pellets were lysed in buffer (50 mM Tris-HCl, 10 mM EDTA, 1% v/v Triton X-100, 1% phenylmethylsulfonyl fluoride (PMSF), 0.05 mM pepstatin A and 0.2 mM leupeptin) and, after mixing with sample loading buffer (50 mM Tris-HCl, 10% w/v SDS, 10% v/v glycerol, 10% v/v 2-mercaptoethanol and 0.04% bromophenol blue) at a ratio of 4:1, were boiled for 5 min. Samples (20 μg protein) were loaded into 8 or 12% SDS-polyacrylamide (SDS-PAGE) gels and subjected to electrophoresis (120 V, 90 min). The separated proteins were transferred to nitrocellulose membranes (Bio-Rad, Hercules, CA, USA; 1 h, 200 mA per gel). After transfer, the blots were incubated with Li-Cor Blocking Buffer for 1 h, followed by overnight incubation with 1:1000 dilution of the primary antibody. Primary polyclonal antibodies directed against HO-1, was purchased from Enzo Life Sciences (Farmingdale, NY, USA) while pAMPK was purchased from Cell Signaling Technology (Danvers, MA, USA). After washing with TBS, the blots were incubated for 1 h with secondary antibody (1:1,000). Protein detection was carried out using a secondary infrared fluorescent dye conjugated antibody absorbing at λ = 800 nm or λ = 700 nm. The blots were visualized using an Odyssey Infrared Imaging Scanner (Li-Cor Science Tec) and quantified by densitometric analysis performed after normalization with β-actin (Santa Cruz Biotechnology, Santa Cruz, CA, USA). Results were expressed as arbitrary units (AU).

Statistical Analysis

One-way analysis of variance (ANOVA) followed by Bonferroni’s t test was performed in order to estimate significant differences among samples. Data were reported as mean values ± S.D. and differences between groups were considered to be significant at p<0.005.

Results and Discussion

Table 1 reports the total phenolic and flavonoid contents in the three different Tithonia extracts. Aqueous extract resulted richer in phenolic and flavonoid compounds compared to methanol and dichlorometane extracts.

Table 1. Total polyphenols and total flavonoids in three different extracts of leaves of Tithonia diversifolia (Hemsl.) A. Gray.

Consistent with their different polyphenol contents, scavenger activities of the three extracts differed depending on the type of extract. The aqueous extract has proven the most effective scavenger, with an effect that, at 0.044 μg/mL, was comparable with 80 mU superoxide dismutase (SOD) (Fig 1). The methanolic and dichloromethane extracts exhibited scavenger activities lower than aqueous; the less active was dichloromethane extract (Fig 1).

Fig 1. Scavenger effect of extracts of Tithonia diversifolia (Hemsl.) A. Gray on superoxide anion; results are expressed as percentage of inhibition of NADH oxidation (rate of superoxide anion production was 4 nmoles/min).

Each value represents the mean ± SD of 5 experimental determinations. * = p<0.001 vs. the same concentration of the aqueous extract.

Based on results regarding phenolic and flavonoid contents, and taking into account findings about scavenger activities, aqueous extract was used for subsequent experiments.

Fig 2 shows results obtained by incubating plasma of a healthy donor in the presence of different concentrations of aqueous extract of Tithonia; as seen, the presence of the extract during incubation caused a significant and dose-dependent inhibition of plasma lipid hydroperoxide formation.

Fig 2. Effect of aqueous extract of Tithonia diversifolia (Hemsl.) A. Gray on LOOH production in plasma; results are expressed as percentage of inhibition of LOOH formation with respect to the same sample incubated in absence of the extract.

Each value represents the mean ± SD of 5 experimental determinations.

These results demonstrated that antioxidants present in aqueous extract of leaves of Tithonia diversifolia (Hemsl.) A. Gray are able to counteract radical chain reactions, preventing peroxidative damage of plasma lipids beyond the action of antioxidants naturally present in plasma.

In the present study we also tested the hypothesis that the aqueous extract of Tithonia diversifolia (Hemsl.) A. Gray might affect adipogenic differentiation of hMSCs. These cells are capable of differentiating into many different phenotypes, including osteoblasts, myocytes, chondrocytes and adipocytes in vivo and in vitro [4143]. The directed differentiation of MSCs can be performed in vitro using the appropriate media and the adipogenic differentiation is confirmed by specific staining.

As seen in Fig 3, none of the concentrations used resulted toxic for hMSC cultures: in fact, no significant difference was observed by MTT test in hMSC cultures exposed to different concentrations of aqueous extract of Tithonia diversifolia (Hemsl.) A. Gray with respect to untreated control (MSCs cultured in the absence of extract).

Fig 3. Cell viability in hMSC cultures exposed to different concentrations of aqueous extract of leaves of Tithonia diversifolia (Hemsl.) A. Gray; values represent the mean ± SD of 5 experiments.

Fig 4 reports data regarding Oil Red O staining; our results demonstrated increased adipogenesis and accumulation of lipid droplets in cultured human adipose tissue-derived mesenchimal cells compared to the same cells treated with the aqueous extract of Tithonia diversifolia (Hemsl.) A. Gray.

Similar results were also demonstrated with other herbal extract such as Momordica foetida Schumach. et Thonn. [44].

Fig 4. Effect of aqueous extract of leaves of Tithonia diversifolia (Hemsl.) A. Gray on adipogenesis of hMSCs as measured by Oil-Red O staining.

Each value represents the mean ± S.D. of 5 experimental determinations. * = p< 0.001 vs control.

It has been suggested that increased levels of ROS, with consequent shifting of the intracellular redox status vs oxidant conditions, promote adipogenesis [24, 26, 45, 46]. Then, the decreased adipogenesis observed in hMSCs cultured in the presence of aqueous extract of Tithonia diversifolia (Hemsl.) A. Gray might be ascribed to its free radical scavenger effects; in order to verify this hypothesis, ROS were determined in hMSC cultures using the fluorescent probe DCFH-DA. After diffusion into the cells, DCFH-DA is deacetylated by cellular esterases to a non-fluorescent compound that can be oxidized by ROS into a highly fluorescent compound, 2',7'-dichlorofluorescein (DCF), whose fluorescence intensity is proportional to the levels of ROS [47]. As reported in Fig 5, the exposure for 72h of hMSCs to 17.5 μg/mL or 175 μg/mL aqueous extract of leaves of Tithonia diversifolia (Hemsl.) A. Gray resulted in a significant decrease in ROS. These data confirmed results obtained using in vitro cell-free systems and further support our hypothesis that antiadipogenic activity of Tithonia diversifolia (Hemsl.) A. Gray might be due to a decrease in intracellular ROS levels.

Fig 5. Determination of ROS by DCFH method in cultured hMSC: effect of aqueous extract of Tithonia diversifolia (Hemsl.) A. Gray.

Each value represents the mean ± S.D. of 5 experimental determinations. * = p< 0.001 vs the same cells cultured in the absence of plant extract (MSC control). Results are expressed as Fluorescence Intensity (F.I.) x 1 x 106/mg prot).

HO-1 (also known as stress protein HSP32) can be over-expressed in many tissues following stressful stimuli and a wide range of conditions characterized by alteration of the cellular redox state [11, 17, 4853]. HO-1 expression might represent an important protective endogenous mechanism; in this regard, there are several reports about the beneficial effects of the induction of this enzyme in several pathological conditions [5457]. In this context, pharmacologic modulation of HO-1 system may represent an effective strategy in several pathologic conditions but it is important to induce HO-1 expression without causing cell damages. Recently the ability of several natural antioxidants to induce HO-1 has been reported [16, 5862].

In order to test the hypothesis that antioxidant acticity of Tithonia diversifolia (Hemsl.) A. Gray may be also mediated by the induction of HO-1, in the present study HO-1 expression was determined by western blot analysis in hMSCs exposed for 72 h to 175 μg/mL Tithonia diversifolia (Hemsl.) A. Gray aqueous extract. Results reported in Fig 6 demonstrated that the presence of the aqueous extract of Tithonia diversifolia (Hemsl.) A. Gray caused a significant increase in HO-1 expression. These results confirmed that antioxidant effect of Tithonia diversifolia (Hemsl.) A. Gray is not merely due to a direct free-radical scavenger activity, but it is also mediated by an induction of protective cellular systems such as HO-1.

We also examined whether AMPK activation might be involved in the inhibition of adipocyte differentiation by aqueous extract of Tithonia diversifolia (Hemsl.) A. Gray; then, hMSCs were exposed to the extract and pAMPK was measured by western blot analysis, using phosphorylated antibody. The results (Fig 6) showed that phosphorylated AMPK levels were significantly increased in hMSC cultures exposed to 175 μg/mL Tithonia diversifolia (Hemsl.) A. Gray aqueous extract compared to untreated hMSCs (control). AMPK is a key enzyme regulating several signals involved in metabolic pathways and appears to be intimately involved in adipocyte differentiation and maturation [63]; our result demonstrated that aqueous extract of leaves of Tithonia diversifolia (Hemsl.) A. Gray is able to inhibit adipocyte differentiation in vitro and its action involves pAMPK.

Fig 6. Representative Western bloting of HO-1 and pAMPK: effect of aqueous extract of Tithonia diversifolia (Hemsl.) A. Gray on HO-1 and pAMPK expression in cultured hMSCs.

Results, expressed as arbitrary units (A.U.), represent the mean ± S.D. of 5 experimental determinations. * = p< 0.005 with respect to tcontrol (C = hMSC control, T = hMSCs treated with 175 μmg/ml of aqueous extract of Tithonia diversifolia (Hemsl.) A. Gray).

Presently intense interest by health care phytomedical reserch has focused on anti-inflammatory [2, 64, 65], antimalarial [66, 67], antidiabetic [6870], and anticancer [71, 72] activities of Tithonia diversifolia (Hemsl.) A. Gray. Results obtained in the present research further support the use of Tithonia diversifolia (Hemsl.) A. Gray and its extracts in the therapy of several diseases and might offer new perspectives for therapeutic strategies in the treatment of obesity and related disorders.


Results obtained in the present study confirmed antiradical capacity of Tithonia diversifolia (Hemsl.) A. Gray suggesting that its antioxidant action may be also due to the ability of affecting the expression of HO-1. In addition, the antiadipogenic activity of Tithonia diversifolia (Hemsl.) A. Gray found in hMSCs offer new stimulating perspective for future therapies, suggesting that its ability to inhibit adipocyte differentiation might be due to the activation of AMPK. Since activated AMPK regulates several metabolic pathways playing a pivotal role in the regulation of carbohydrate and fat metabolism, these findings might be usefull for the treatment of metabolic diseases such as diabetes and obesity. Results obtained in the present study also suggest that the intake of natural preparations of T. diversifolia might lower the risk of human diseases such as atherosclerosis, inflammation, ageing, ischemic reperfusion injury and neurodegenerative diseases.

Supporting Information

S1 Fig. Scavenger effect of extracts of Tithonia diversifolia (Hemsl.) A. Gray on superoxide anion expressed as percentage of inhibition of NADH oxidation (rate of superoxide anion production was 4 nmoles/min).


S2 Fig. Effect of aqueous extract of Tithonia diversifolia (Hemsl.) A. Gray on LOOH production in plasma expressed as percentage of inhibition of LOOH formation with respect to the same sample incubated in absence of the extract.


S3 Fig. Percentage of cell viability in hMSC cultures exposed to different concentrations of aqueous extract of leaves of Tithonia diversifolia (Hemsl.) A. Gray


S4 Fig. Effect of aqueous extract of leaves of Tithonia diversifolia (Hemsl.) A. Gray on adipogenesis of hMSCs as measured by Oil-Red O staining (O.D. at λ = 490 nm).


S5 Fig. Determination of ROS by DCFH method in cultured hMSC, Values represent Fluorescence Intensity (F.I.) x 1 x 106/mg prot.


S6 Fig. Protein expression of HO-1 and pAMPK by western blot analysis in cultured hMSCs; values are expressed as arbitrary units (A.U.).


S1 Table. Total polyphenol content (expressed as μmM Gallic acid) and total flavonoid content (expressed as μmM Catechin) in three different extracts of leaves of Tithonia diversifolia (Hemsl.) A. Gray.



The authors thank Dr. Mike Wilkinson for proofreading the manuscript and Sister Kavira Muhyama for plant collection.

Author Contributions

Conceived and designed the experiments: CDG RA. Performed the experiments: CDG RA LV VS IB. Analyzed the data: RS CG. Contributed reagents/materials/analysis tools: SM SR GC. Wrote the paper: CDG RA.

In Memoriam

This work is dedicated to the memory of Prof. Liliana Iauk.


  1. 1. Tona L, Kambu K, Ngimbi N, Mesia K, Penge O, Lusakibanza M, et al. Antiamoebic and spasmolytic activities of extracts from some antidiarrhoeal traditional preparations used in Kinshasa, Congo. Phytomedicine 2000; 7: 31–38. pmid:10782488
  2. 2. Rungeler P, Lyss G, Castro V, Mora G, Pahl HL, Merfort I. Study of three sesquiterpene lactones from Tithonia diversifolia on their anti-inflammatory activity using the transcription factor NF-kappa B and enzymes of the arachidonic acid pathway as targets. Planta Med 1998; 64: 588–593. pmid:9810261
  3. 3. Lee MY, Liao MH, Tsai YN, Chiu KH, Wen HC. Identification and anti-human glioblastoma activity of tagitinin C from Tithonia diversifolia methanolic extract. J Agric Food Chem 2011; 59: 2347–2355. pmid:21332189
  4. 4. Chagas-Paula DA, Oliveira RB, Rocha BA, Da Costa FB. Ethnobotany, chemistry, and biological activities of the genus Tithonia (Asteraceae). Chem Biodivers 2012; 9: 210–235. pmid:22344901
  5. 5. Zhu YZ, Huang SH, Tan BK, Sun J, Whiteman M, Zhu YC. Antioxidants in Chinese herbal medicines: a biochemical perspective. Nat Prod Rep 2004; 21: 478–489. pmid:15282631
  6. 6. Maines MD. Heme oxygenase: function, multiplicity, regulatory mechanisms, and clinical applications. FASEB J 1988; 2: 2557–2568. pmid:3290025
  7. 7. Cruse I, Maines MD. Evidence suggesting that the two forms of heme oxygenase are products of different genes. J Biol Chem 1988; 263: 3348–3353. pmid:3343248
  8. 8. Trakshel GM, Maines MD. Multiplicity of heme oxygenase isozymes. HO-1 and HO-2 are different molecular species in rat and rabbit. J Biol Chem 1989; 264: 1323–1328. pmid:2910857
  9. 9. McCoubrey WK Jr, Maines MD. The structure, organization and differential expression of the gene encoding rat heme oxygenase-2. Gene 1994; 139: 155–161. pmid:8112599
  10. 10. McCoubrey WK Jr, Huang TJ, Maines MD. Isolation and characterization of a cDNA from the rat brain that encodes hemoprotein heme oxygenase-3. Eur J Biochem 1997; 247: 725–732. pmid:9266719
  11. 11. Raju VS, Maines MD. Coordinated expression and mechanism of induction of HSP32 (heme oxygenase-1) mRNA by hyperthermia in rat organs. Biochim Biophys Acta 1994; 1217: 273–280. pmid:8148372
  12. 12. Nath KA, Balla G, Vercellotti GM, Balla J, Jacob HS, Levitt MD, et al. Induction of heme oxygenase is a rapid, protective response in rhabdomyolysis in the rat. J Clin Invest 1992; 90: 267–270. pmid:1634613
  13. 13. Poss KD, Tonegawa S. Reduced stress defense in heme oxygenase 1-deficient cells. Proc Natl Acad Sci U S A 1997; 94: 10925–10930. pmid:9380736
  14. 14. Foresti R, Motterlini R. The heme oxygenase pathway and its interaction with nitric oxide in the control of cellular homeostasis. Free Radic Res 1999; 31: 459–475. pmid:10630670
  15. 15. Li C, Hossieny P, Wu BJ, Qawasmeh A, Beck K, Stocker R. Pharmacologic induction of heme oxygenase-1. Antioxid Redox Signal 2007; 9: 2227–2239. pmid:17822367
  16. 16. Scapagnini G, Foresti R, Calabrese V, Giuffrida Stella AM, Green CJ, Motterlini R. Caffeic acid phenethyl ester and curcumin: a novel class of heme oxygenase-1 inducers. Mol Pharmacol 2002; 61: 554–561. pmid:11854435
  17. 17. Balogun E, Hoque M, Gong P, Killeen E, Green CJ, Foresti R, et al. Curcumin activates the haem oxygenase-1 gene via regulation of Nrf2 and the antioxidant-responsive element. Biochem J 2003; 371: 887–895. pmid:12570874
  18. 18. Motterlini R, Foresti R, Bassi R, Green CJ. Curcumin, an antioxidant and anti-inflammatory agent, induces heme oxygenase-1 and protects endothelial cells against oxidative stress. Free Radic Biol Med 2000; 28: 1303–1312. pmid:10889462
  19. 19. Shepherd PR, Gnudi L, Tozzo E, Yang H, Leach F, Kahn BB. Adipose cell hyperplasia and enhanced glucose disposal in transgenic mice overexpressing GLUT4 selectively in adipose tissue. J Biol Chem 1993; 268: 22243–22246. pmid:8226728
  20. 20. Sorisky A. From preadipocyte to adipocyte: differentiation-directed signals of insulin from the cell surface to the nucleus. Crit Rev Clin Lab Sci 1999; 36: 1–34. pmid:10094092
  21. 21. Bray GA. A concise review on the therapeutics of obesity. Nutrition 2000; 16: 953–960. pmid:11054601
  22. 22. Grattagliano I, Palmieri VO, Portincasa P, Moschetta A, Palasciano G. Oxidative stress-induced risk factors associated with the metabolic syndrome: a unifying hypothesis. J Nutr Biochem 2008;19: 491–504. pmid:17855068
  23. 23. Imhoff BR, Hansen JM. Differential redox potential profiles during adipogenesis and osteogenesis. Cell Mol Biol Lett 2011; 16: 149–161. pmid:21225471
  24. 24. Kanda Y, Hinata T, Kang SW, Watanabe Y. Reactive oxygen species mediate adipocyte differentiation in mesenchymal stem cells. Life Sci 2011; 89: 250–258. pmid:21722651
  25. 25. Vigilanza P, Aquilano K, Baldelli S, Rotilio G, Ciriolo MR. Modulation of intracellular glutathione affects adipogenesis in 3T3-L1 cells. J Cell Physiol 2011; 226: 2016–2024. pmid:21520053
  26. 26. Higuchi M, Dusting GJ, Peshavariya H, Jiang F, Hsiao ST, Chan EC, et al. Differentiation of Human Adipose-Derived Stem Cells into Fat Involves Reactive Oxygen Species and Forkhead Box O1 Mediated Upregulation of Antioxidant Enzymes. Stem Cells Dev 2013; 22: 878–88. pmid:23025577
  27. 27. Fisslthaler B, Fleming I. Activation and signaling by the AMP-activated protein kinase in endothelial cells. Circ Res 2009; 105:114–127. pmid:19608989
  28. 28. Hawley SA, Davison M, Woods A, Davies SP, Beri RK, Carling D, et al. Characterization of the AMP-activated protein kinase kinase from rat liver and identification of threonine 172 as the major site at which it phosphorylates AMP-activated protein kinase. J Biol Chem 1996; 271: 27879–27887. pmid:8910387
  29. 29. Yuan HD, Yuan HY, Chung SH, Jin GZ, Piao GC. An active part of Artemisia sacrorum Ledeb. attenuates hepatic lipid accumulation through activating AMP-activated protein kinase in human HepG2 cells Biosci Biotechnol Biochem 2010; 74: 322–328. pmid:20139613
  30. 30. Viollet B, Foretz M, Guigas B, Horman S, Dentin R, Bertrand L, et al. Activation of AMP-activated protein kinase in the liver: a new strategy for the management of metabolic hepatic disorders. J Physiol 2006; 574: 41–53. pmid:16644802
  31. 31. Daval M, Foufelle F, Ferre P. Functions of AMP-activated protein kinase in adipose tissue. J Physiol 2006; 574: 55–62. pmid:16709632
  32. 32. Kim SK, Kong CS. Anti-adipogenic effect of dioxinodehydroeckol via AMPK activation in 3T3-L1 adipocytes. Chem Biol Interact 2010; 186: 24–29. pmid:20385110
  33. 33. Moon HS, Chung CS, Lee HG, Kim TG, Choi YJ, Cho CS. Inhibitory effect of (-)-epigallocatechin-3-gallate on lipid accumulation of 3T3-L1 cells. Obesity (Silver Spring) 2007; 15: 2571–2582. pmid:18070748
  34. 34. Ballard TS, Mallikarjunan P, Zhou K, O'Keefe S. Microwave-assisted extraction of phenolic antioxidant compounds from peanut skins. Food Chem 2010; 120: 1185–1192.
  35. 35. Zhishen J, Mengcheng T, Jianming W. The determination of flavonoid contents in mulberry and their scavenging effects on superoxide radicals. Food Chem 1999; 64: 555–559.
  36. 36. Acquaviva R, Menichini F, Ragusa S, Genovese C, Amodeo A, Tundis R, et al. Antimicrobial and antioxidant properties of Betula aetnensis Rafin. (Betulaceae) leaves extract. Nat Prod Res 2013; 27: 475–9. pmid:22703292
  37. 37. Wolff SP. Ferrous Ion Oxidation in Presence of Ferric Ion Indicator Xylenol Orange for Measurement of Hydroperoxides. Method Enzymol 1994; 233: 182–189.
  38. 38. Mosmann T. Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J Immunol Methods 1983; 65: 55–63. pmid:6606682
  39. 39. Acquaviva R, Di Giacomo C, Sorrenti V, Galvano F, Santangelo R, Cardile V, et al. Antiproliferative effect of oleuropein in prostate cell lines. Int J Oncol 2012; 41: 31–38. pmid:22484302
  40. 40. Lowry OH, Rosebrough NJ, Farr AL, Randall RJ. Protein measurement with the Folin phenol reagent. J Biol Chem 1951; 193: 265–275. pmid:14907713
  41. 41. Da Silva Meirelles L, Caplan AI, Nardi NB. In search of the in vivo identity of mesenchymal stem cells. Stem Cells 2008; 26: 2287–2299. pmid:18566331
  42. 42. Crisan M, Yap S, Casteilla L, Chen CW, Corselli M, Park TS, et al. A perivascular origin for mesenchymal stem cells in multiple human organs. Cell stem cell 2008; 3: 301–313. pmid:18786417
  43. 43. Caplan AI. All MSCs are pericytes? Cell Stem Cell 2008; 3: 229–230. pmid:18786406
  44. 44. Acquaviva R, Di Giacomo C, Vanella L, Santangelo R, Sorrenti V, Barbagallo I, et al. Antioxidant activity of extracts of Momordica foetida Schumach. et Thonn. Molecules 2013; 18: 3241–3249. pmid:23486103
  45. 45. Lee H, Lee YJ, Choi H, Ko EH, Kim JW. Reactive oxygen species facilitate adipocyte differentiation by accelerating mitotic clonal expansion. J Biol Chem 2009; 284: 10601–10609. pmid:19237544
  46. 46. Barbagallo I, Vanella A, Peterson SJ, Kim DH, Tibullo D, Giallongo C, et al. Overexpression of heme oxygenase-1 increases human osteoblast stem cell differentiation. J Bone Miner Metab 2010; 28: 276–288. pmid:19924377
  47. 47. LeBel CP, Ischiropoulos H, Bondy SC. Evaluation of the probe 2',7'-dichlorofluorescin as an indicator of reactive oxygen species formation and oxidative stress. Chem Res Toxicol 1992; 5: 227–231. pmid:1322737
  48. 48. Keyse SM, Tyrrell RM. Heme oxygenase is the major 32-kDa stress protein induced in human skin fibroblasts by UVA radiation, hydrogen peroxide, and sodium arsenite. Proc Natl Acad Sci U S A 1989; 86: 99–103. pmid:2911585
  49. 49. Lautier D, Luscher P, Tyrrell RM. Endogenous glutathione levels modulate both constitutive and UVA radiation/hydrogen peroxide inducible expression of the human heme oxygenase gene. Carcinogenesis 1992; 13: 227–232. pmid:1740012
  50. 50. Balla J, Jacob HS, Balla G, Nath K, Eaton JW, Vercellotti GM. Endothelial-cell heme uptake from heme proteins: induction of sensitization and desensitization to oxidant damage. Proc Natl Acad Sci U S A 1993; 90: 9285–9289. pmid:8415693
  51. 51. Choi AM, Alam J. Heme oxygenase-1: function, regulation, and implication of a novel stress-inducible protein in oxidant-induced lung injury. Am J Respir Cell Mol Biol 1996; 15: 9–19. pmid:8679227
  52. 52. Tyrrell R. Redox regulation and oxidant activation of heme oxygenase-1. Free Radic Res 1999; 31: 335–340. pmid:10517538
  53. 53. Sacerdoti D, Colombrita C, Ghattas MH, Ismaeil EF, Scapagnini G, Bolognesi M, et al. Heme oxygenase-1 transduction in endothelial cells causes downregulation of monocyte chemoattractant protein-1 and of genes involved in inflammation and growth. Cell Mol Biol (Noisy-le-Grand, France) 2005; 51: 363–370. pmid:16309586
  54. 54. Ryter SW, Alam J, Choi AM. Heme oxygenase-1/carbon monoxide: from basic science to therapeutic applications. Physiol Rev 2006; 86: 583–650. pmid:16601269
  55. 55. Cuadrado A, Rojo AI. Heme oxygenase-1 as a therapeutic target in neurodegenerative diseases and brain infections. Curr Pharm Des 2008; 14: 429–442. pmid:18289070
  56. 56. Novo G, Cappello F, Rizzo M, Fazio G, Zambuto S, Tortorici E, et al. Hsp60 and heme oxygenase-1 (Hsp32) in acute myocardial infarction. Transl Res 2011; 157: 285–292. pmid:21497776
  57. 57. Acquaviva R, Campisi A, Raciti G, Avola R, Barcellona ML, Vanella L, et al. Propofol inhibits caspase-3 in astroglial cells: role of heme oxygenase-1. Curr Neurovasc Res 2005; 2: 141–148. pmid:16181106
  58. 58. Abraham NG, Kappas A. Pharmacological and clinical aspects of heme oxygenase. Pharmacol Rev 2008; 60: 79–127. pmid:18323402
  59. 59. Scapagnini G, Butterfield DA, Colombrita C, Sultana R, Pascale A, Calabrese V. Ethyl ferulate, a lipophilic polyphenol, induces HO-1 and protects rat neurons against oxidative stress. Antioxid Redox Signal 2004; 6: 811–818. pmid:15345140
  60. 60. Li Volti G, Sacerdoti D, Di Giacomo C, Barcellona ML, Scacco A, Murabito P, et al. Natural heme oxygenase-1 inducers in hepatobiliary function. World J Gastroenterol 2008; 14: 6122–6132. pmid:18985801
  61. 61. Ferrandiz ML, Devesa I. Inducers of heme oxygenase-1. Curr Pharm Des 2008;14: 473–486. pmid:18289074
  62. 62. Cho HS, Kim S, Lee SY, Park JA, Kim SJ, Chun HS. Protective effect of the green tea component, L-theanine on environmental toxins-induced neuronal cell death. Neurotoxicology 2008; 29: 656–662. pmid:18452993
  63. 63. Zhou Y, Wang D, Zhu Q, Gao X, Yang S, Xu A, et al. Inhibitory effects of A-769662, a novel activator of AMP-activated protein kinase, on 3T3-L1 adipogenesis. Biol Pharm Bull 2009; 32: 993–998. pmid:19483304
  64. 64. Owoyele VB, Wuraola CO, Soladoye AO, Olaleye SB. Studies on the anti-inflammatory and analgesic properties of Tithonia diversifolia leaf extract. J Ethnopharmacol 2004; 90: 317–321. pmid:15013196
  65. 65. Siedle B, Garcia-Pineres AJ, Murillo R, Schulte-Monting J, Castro V, Rüngeler P, et al. Quantitative structure-activity relationship of sesquiterpene lactones as inhibitors of the transcription factor NF-kappaB. J Med Chem 2004; 47: 6042–6054. pmid:15537359
  66. 66. Do Ceu de Madureira M, Paula Martins A, Gomes M, Paiva J, Proenca da Cunha A, Do Rosario V. Antimalarial activity of medicinal plants used in traditional medicine in S. Tome and Principe islands. J Ethnopharmacol 2002; 81: 23–29. pmid:12020924
  67. 67. Goffin E, Ziemons E, De Mol P, De Madureira Mdo C, Martins AP, da Cunha AP, et al. In vitro antiplasmodial activity of Tithonia diversifolia and identification of its main active constituent: tagitinin C. Planta Med 2002; 68: 543–545. pmid:12094301
  68. 68. Miura T, Nosaka K, Ishii H, Ishida T. Antidiabetic effect of Nitobegiku, the herb Tithonia diversifolia, in KK-Ay diabetic mice. Biol Pharm Bull 2005; 28: 2152–2154. pmid:16272709
  69. 69. Miura T, Furuta K, Yasuda A, Iwamoto N, Kato M, Shihara E, et al. Antidiabetic effect of nitobegiku in KK-Ay diabetic mice. Am J Chin Med 2002; 30: 81–86. pmid:12067100
  70. 70. Zhao G, Li X, Chen W, Xi Z, Sun L. Three new sesquiterpenes from Tithonia diversifolia and their anti-hyperglycemic activity. Fitoterapia 2012; 83: 1590–1597. pmid:22986291
  71. 71. Gu JQ, Gills JJ, Park EJ, Mata-Greenwood E, Hawthorne ME, Axelrod F, et al. Sesquiterpenoids from Tithonia diversifolia with potential cancer chemopreventive activity. J Nat Prod 2002; 65: 532–536. pmid:11975495
  72. 72. Kuroda M, Yokosuka A, Kobayashi R, Jitsuno M, Kando H, Nosaka K, et al. Sesquiterpenoids and flavonoids from the aerial parts of Tithonia diversifolia and their cytotoxic activity. Chem Pharm Bull (Tokyo) 2007; 55: 1240–1244. pmid:17666852