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Centella asiatica modulates cancer cachexia associated inflammatory cytokines and cell death in leukaemic THP-1 cells and peripheral blood mononuclear cells (PBMC’s)



Cancer cachexia is associated with increased pro-inflammatory cytokine levels. Centella asiatica (C. asiatica) possesses antioxidant, anti-inflammatory and anti-tumour potential. We investigated the modulation of antioxidants, cytokines and cell death by C. asiatica ethanolic leaf extract (CLE) in leukaemic THP-1 cells and normal peripheral blood mononuclear cells (PBMC’s).


Cytotoxcity of CLE was determined at 24 and 72 h (h). Oxidant scavenging activity of CLE was evaluated using the 2, 2-diphenyl-1 picrylhydrazyl (DPPH) assay. Glutathione (GSH) levels, caspase (−8, −9, −3/7) activities and adenosine triphosphate (ATP) levels (Luminometry) were then assayed. The levels of tumour necrosis factor-α (TNF-α), interleukin (IL)-6, IL-1β and IL-10 were also assessed using enzyme-linked immunosorbant assay.


CLE decreased PBMC viability between 33.25–74.55% (24 h: 0.2–0.8 mg/ml CLE and 72 h: 0.4–0.8 mg/ml CLE) and THP-1 viability by 28.404% (72 h: 0.8 mg/ml CLE) (p < 0.0001). Oxidant scavenging activity was increased by CLE (0.05–0.8 mg/ml) (p < 0.0001). PBMC TNF-α and IL-10 levels were decreased by CLE (0.05–0.8 mg/ml) (p < 0.0001). However, PBMC IL-6 and IL-1β concentrations were increased at 0.05–0.2 mg/ml CLE but decreased at 0.4 mg/ml CLE (p < 0.0001). In THP-1 cells, CLE (0.2–0.8 mg/ml) decreased IL-1β and IL-6 whereas increased IL-10 levels (p < 0.0001). In both cell lines, CLE (0.05–0.2 mg/ml, 24 and 72 h) increased GSH concentrations (p < 0.0001). At 24 h, caspase (−9, −3/7) activities was increased by CLE (0.05–0.8 mg/ml) in PBMC’s whereas decreased by CLE (0.2–0.4 mg/ml) in THP-1 cells (p < 0.0001). At 72 h, CLE (0.05–0.8 mg/ml) decreased caspase (−9, −3/7) activities and ATP levels in both cell lines (p < 0.0001).


In PBMC’s and THP-1 cells, CLE proved to effectively modulate antioxidant activity, inflammatory cytokines and cell death. In THP-1 cells, CLE decreased pro-inflammatory cytokine levels whereas it increased anti-inflammatory cytokine levels which may alleviate cancer cachexia.

Peer Review reports


The role of inflammation in carcinogenesis has been extensively documented [1]. Although inflammatory responses have shown beneficial effects in tissue repair and pathogen elimination [1, 2], chronic inflammation has been implicated in tumour initiation, promotion and progression [3]. During ideal conditions, the host-mediated anti-tumour activity combats the tumour-mediated immunosuppressive activity and cancerous cells are sentenced to cell death [3]. In the event that the host anti-tumour activity is weakened/inadequate, the persistent and enhanced pro-inflammatory tumour microenvironment will facilitate tumour development, invasion, angiogenesis and metastasis [3].

Many malignancies are associated with the cachectic syndrome [4], a disorder characterised by abnormal weight loss [5] due to adipose tissue (85%) and skeletal muscle (75%) depletion [6]. The enzyme lipoprotein lipase (LPL) hydrolyses fatty acids (FA’s) and transports FA’s into adipose tissue for triacylglycerol (TAG) production, whereas hormone sensitive lipase (HSL) breaks down TAG’s into FA’s and glycerol [6]. Studies have revealed that decreased serum LPL levels/activity [7, 8] and increased HSL levels/activity are associated with cachexia [9]. Additionally, increased proteolysis and decreased proteogenesis have been reported in cachectic patients [10]. The ATP-ubiquitin-dependent proteolytic pathway has been shown to be responsible for the excessive proteolysis seen in cancer cachexia [11].

Oxidative stress, inflammatory cytokines and apoptosis play a pivotal role in the initiation and development of cancer cachexia [12]. Inflammatory cytokine production is increased by lipopolysaccharide (LPS) potently stimulating macrophages [13]. The LPS signal is transduced by LPS binding to LPS binding protein, delivered to CD14 and transferred to Toll like receptor-4 [14]. This subsequently activates nuclear factor kappa B (NF-κB), which regulates the transcription of genes associated with inflammation, proliferation, invasion, angiogenesis and apoptosis [1, 15,16,17]. Previously, IL-1 [18], IL-6 (mice) [19] and TNF-α (rat, mouse and guinea pigs) [20] were shown to decrease LPL activity in adipose tissue. Decreased LPL activity reduces the uptake of exogenous lipids by adipose tissue [20], which decreases lipogenesis. Additionally, previous literature showed that TNF-α increased ubiquitin (concentrations and mRNA), while IL-6 increased the 26S proteasome and cathepsin activities, suggesting the activation of proteolytic pathways [21,22,23,24]. The activation of proteolytic pathways causes extensive muscle wasting through proteolysis. Taken together, an excessive increase in pro-inflammatory cytokine levels may increase tumour immunosuppressive activity [3], as well as tissue wasting [6].

Oxidative stress has been associated with tumour initiation, inflammation [2, 3] and muscle wasting [25]. However, antioxidants have been shown to decrease muscle wasting by neutralizing reactive oxygen species (ROS) [1, 25]. Elevated ROS levels activate apoptotic pathways, ultimately activating caspase-3 [26]. The activation of caspase-3 plays an important role in the execution of apoptosis as well as muscle proteolysis [27]. Additionally, in weight-losing upper gastrointestinal tract cancer patients, deoxyribonucleic acid (DNA) fragmentation and poly (ADP-ribose) polymerase (PARP) cleavage were increased, whereas MyoD protein was decreased [6], suggesting increased apoptosis and decreased muscle replenishment.

There is a constant need for alternative traditional medicines to improve the prognosis of cancer patients and prevent chemotherapy and radiotherapy induced discomfort. The tropical medicinal plant Centella asiatica (Linnaeus) Urban (C. asiatica) is native to India, China, and South Africa [28]. It belongs to the Apiaceae family and is commonly referred to as Gotu kola, Asiatic pennywort and Tiger herb [28]. C. asiatica is widely used in Ayurvedic and Chinese traditional medicines due to its various medicinal properties. These properties include its hepato-protective, cardio-protective, anti-diabetic, antioxidant, anti-inflammatory and anti-tumour potential [28]. The major active compounds in C. asiatica are triterpene saponosides such as asiatic acid, madecassic acid and asiaticoside [28]. C. asiatica also contains flavonoid derivatives, vitamins, minerals, polysaccharides, sterols and phenolic acids [28]. C. asiatica has previously been used in treatment of inflammation due to its promising anti-inflammatory effects [29, 30]. Additionally, C. asiatica extracts have demonstrated high antioxidant [31, 32] and anti-proliferative activity in many cancerous cell lines [33].

There is a need for the discovery of an inexpensive cancer cachectic treatment. The ability of a plant extract to regulate inflammatory cytokines and cell death may elevate cancerous cell death and diminish tissue wasting. We investigated the potential of a C. asiatica ethanolic leaf extract (CLE) to modulate inflammatory cytokines, antioxidants and cell death in leukaemic THP-1 cells and normal peripheral blood mononuclear cells (PBMC’s).



C. asiatica leaves were collected on the 7th of March 2011 (collectors number: Immelman 411) from the Eastern Cape [Langeni forest, roadside (S31°28.135′, E28°32.681′)], South Africa (SA) and identified by Dr. Kathleen Immelman from the Department of Botany at the Walter Sisulu University, SA. Voucher specimens were deposited at the KEI herbarium (13979). The THP-1 cells were obtained from American Type Culture Collection (ATCC, University Boulevard Manassas, Virginia, USA). RPMI-1640 and BD OptEIA enzyme-linked immunosorbant assay (ELISA) cytokine kits were purchased from The Scientific Group (Johannesburg, SA). Foetal calf serum (FCS) and Pen/Strep Amphotericin B (PSF) were acquired from Whitehead Scientific (Cape Town, SA). Dimethyl sulphoxide (DMSO) was purchased from Merck (Johannesburg, SA). Histopaque-1077, LPS and 2, 2-diphenyl-1 picrylhydrazyl (DPPH) were purchased from Sigma (Aston Manor, SA). The 4-[3-(4-iodophenyl)-2-(4-nitrophenyl)-2H-5-tetrazolio]-1,3-benzene disulphonate (WST-1) cell proliferation reagent was purchased from Roche (Johannesburg, SA). Promega (Madison, USA) supplied the caspase (−3/7, −8, −9), adenosine triphosphate (ATP) and glutathione (GSH) kits.

Plant description and extraction

The plants official name is Centella asiatica (L.) Urb and has been confirmed by using the plant list [34]. The English name is Tiger herb. C. asiatica leaves were dried and milled. Ethanol (200–350 ml) was added to milled plant material (10–30 g) and extracted overnight by shaking (4×g, 37 °C). Ethanol extracts were filtered, rotor evaporated, dried (37 °C) and stored (4 °C).

The 2, 2-diphenyl-1 picrylhydrazyl assay

CLE (0.05–0.8 mg/ml) and butylated hydroxytoluene (BHT) (60–300 μM) dilutions were prepared in methanol (99.5% and grade AR). A 50 μM DPPH solution was prepared from a stock solution of 0.135 mM DPPH in methanol. CLE, BHT dilutions and methanol (1 ml, triplicate tubes) were aliquoted into 15 ml polypropylene tubes, followed by the 50 μM DPPH solution (1 ml). Reaction mixtures were vortexed and incubated (room temperature (RT) for 30 min (min)) in the dark. Absorbance of samples was read at 517 nm using a Varine Cary 50 UV-visible spectrophotometer (McKinley Scientific, New Jersey, US).

Isolation of peripheral blood mononuclear cells

Buffy coats containing PBMC’s were obtained from the South African National Blood Service (2011/09). PBMC’s were extracted by differential centrifugation. Buffy coats (5 ml) were layered onto equivolume histopaque-1077 (5 ml) in 15 ml polypropylene tubes and centrifuged (400×g, 21 °C for 30 min). After centrifugation, the PBMC’s were transferred to sterile 15 ml polypropylene tubes, phosphate buffered saline (PBS) was added (0.1 M, 10 ml) and tubes were centrifuged (400×g, 21 °C, 15 min). Cell density of isolated PBMC’s was adjusted (1 × 106 cells/ml) using the trypan blue exclusion test and cryo-preserved (10% FCS, 10% DMSO) using a NELGENE cryo freezing container and stored at −80 °C.

Tissue culture

THP-1 cells were grown in the appropriate tissue culture conditions in a 75 cm3 tissue culture flask (37 °C, 5% CO2). The growth media comprised of RPMI-1640, FCS (10%) and PS (2%). Cells were thawed, seeded into a 75 cm3 tissue culture flask at a concentration of 3 × 105 cells/ml and incubated (37 °C, 5% CO2). THP-1 cells were allowed to grow for 2–3 days before the cells were centrifuged (162×g, 10 min) and re-suspended in fresh growth media. The number of cells should not exceed 8 × 105 cells/ml, therefore the cells/ml was quantified daily by trypan blue staining. Once the cell count reached 8 × 105 cells/ml the THP-1 cells were split/ diluted to 3 × 105 cells/ml with media and incubated. Subsequent experiments were conducted once the cell numbers were sufficient.

Cell viability assay

Cytotoxicity of CLE in PBMC’s and THP-1 cells was measured using the WST-1 assay (Roche, Johannesburg, SA). PBMC and THP-1 cells (10,000 cells/well, 96-well plate, in triplicate wells) were stimulated with LPS (20 μg/ml, 37 °C, 5% CO2, 4 h (h)) before exposure to CLE (0.05–0.8 mg/ml) for 24 and 72 h (37 °C, 5% CO2). Similarly, controls received media containing DMSO (0.2%). Thereafter, plates were centrifuged (162×g, 10 min), supernatant removed, cell pellets re-suspended in growth media (100 μl/well), WST-1 reagent (10 μl/well) added and plates incubated (37 °C, 5%, CO2, 3 h). Optical density was measured at 450 nm (620 nm reference wavelength) with a BIO-TEK μQuant spectrophotometer (Analytical and Diagnostic Products, SA). This experiment was conducted independently on three occasions.

Stimulation and treatment of cells

PBMC’s and THP-1 cells (1 × 105 cells/ml) were transferred into 24-well plates, stimulated with LPS (20 μg/ml, 37 °C, 5% CO2, 4 h) before exposure to CLE (0.05–0.8 mg/ml) for 24 h (TNF-α) and 72 h (IL-1β, IL-6, IL-10) (37 °C, 5% CO2). After incubation, plates were centrifuged (162×g, 10 min) and supernatant was collected and stored (−80 °C) for cytokine analysis. Cell pellets were used to conduct the caspase (−8, −9, −3/7) activity, ATP and GSH assays. The experiments were conducted independently (twice for all subsequent assays).

Quantification of cytokines

Cytokine levels were estimated using the BD OptEIA ELISA kits (The Scientific Group, SA) and the procedure was followed as per the instruction manual. ELISA plates were coated with capture antibody overnight (100 μl/well, 4 °C). Thereafter, plates were washed (3×) with wash buffer and blocked with assay diluent (200 μl/well, 1 h, RT). Standard solutions were prepared by diluting a stock solution [TNF-α, IL-10 (500 pg/ml), IL-6 (300 pg/ml), IL-1β (250 pg/ml)] serially [TNF-α, IL-10 (500–7.8 pg/ml), IL-6 (300–4.7 pg/ml), IL-1β (250–3.9 pg/ml)]. Plates were washed (3×), standards and samples (100 μl/well, triplicate wells) were aliquoted into appropriate wells and plates were incubated (2 h, RT). Plates were washed (5×), working detector (100 μl/well) added and plates incubated (1 h, RT). The plates were washed (7×), substrate solution (100 μl/well) added and plates were incubated (30 min, RT) in the dark. Finally, stop solution (50 μl/well) was added and the absorbance was read at 450 nm (570 nm reference wavelength) with a Multiskan FC micro-plate reader (Thermo Scientific). Cytokine concentrations were calculated by extrapolation from a standard curve.

Glutathione assay

The GSH-Glo™ assay (Promega, Madison, WI, USA) was used to measure GSH levels. Standard GSH solutions were prepared by diluting a 5 mM stock solution serially (1.56–50 μM) and PBS (0.1 M) was the standard blank. Cells (50 μl/well, 2 × 105 cells/ml) and standards were added into an opaque 96-well plate (duplicate wells), followed by GSH-Glo™ reagent (25 μl/well) and allowed to incubate (30 min, RT) in the dark. Subsequently, luciferin detection reagent (50 μl/well) was added and plates incubated (15 min, RT) in the dark. The absorbance was read on a Modulus™ microplate luminometer (Turner Biosystems, Sunnyvale, USA) and GSH concentrations were calculated by extrapolation from a standard curve.

Caspase and ATP assays

Caspase activity and ATP levels were determined using the Caspase-Glo®-3/7, −8, −9 and ATP assay kits (Promega, Madison, WI, USA). Caspase-Glo®-3/7, −8, −9 and ATP reagents were reconstituted according to the manufacturer’s instructions. Cells (100 μl, 2 × 105 cells/ml) were added into duplicate wells of a microtitre plate for each assay, thereafter caspase −3/7, −8, −9 and ATP reagents (100 μl/well) were added into appropriate wells. The plate was incubated (30 min, RT) in the dark. Luminescence was measured on a Modulus™ microplate luminometer (Turner BioSystems) and expressed as relative light units (RLU).

Statistical analysis

Statistical analysis was performed using the STATA and GraphPad Prism (v5) statistical analysis software. The one-way analysis of variance (ANOVA) was used to make comparisons between groups, followed by the Tukey multiple comparisons test, with p < 0.05 indicating significant results.


The oxidant scavenging potential of CLE

The oxidant scavenging activity of CLE using the DPPH assay is shown in Fig. 1. CLE (0.05–0.8 mg/ml) significantly increased DPPH scavenging activity by approximately 45–84% (Fig. 1, p < 0.0001).

Fig. 1

Percentage DPPH scavenging activity of CLE (Values expressed as mean ± SD, *** p < 0.0001 compared to control)

The in vitro cytotoxicity of CLE

The WST-1 assay was used to determine cell viability of THP-1 cells and PBMC’s after treatment with CLE (Fig. 2). At 24 h, CLE (0.2–0.8 mg/ml) dose dependently decreased PBMC viability by 33.25–61.85% (Fig. 2a, p < 0.0001), whereas THP-1 viability was not significantly altered as compared to the control (Fig. 2c, p = 0.0003). At 72 h, CLE decreased both PBMC (Fig. 2b, 34.268–74.547%) and THP-1 (Fig. 2d, czmg/ml respectively as compared to the control (p < 0.0001), suggesting that PBMC’s are more sensitive to CLE treatment than THP-1 cells.

Fig. 2

Cell viability of PBMC (a – 24 h, b – 72 h) and THP-1 (c – 24 h, d – 72 h) cells treated with CLE for 24 and 72 h (Values expressed as mean ± SD, ** p < 0.001, *** p < 0.0001 compared to the control)

The immune suppressive properties of CLE

CLE altered cytokine levels in PBMC’s and THP-1 cells which are shown in Figs. 3 and 4 respectively. The levels of TNF-α, IL-1β, IL-6 and IL-10 produced in LPS stimulated PBMC’s was 309.60, 152.83, 626.33 and 23.55 pg/ml respectively. CLE (0.05–0.2 mg/ml) increased PBMC IL-1β and IL-6 concentrations relative to the control (Fig. 3bc, p < 0.0001). In PBMC’s, TNF-α, IL-1β and IL-6 concentrations were decreased at 0.05–0.8 mg/ml CLE, 0.4–0.8 mg/ml CLE and 0.4 mg/ml CLE respectively as compared to the control (Fig. 3ac, p < 0.0001). The levels of TNF-α, IL-1β, IL-6 and IL-10 produced in LPS stimulated THP-1 cells was 5.96, 25.92, 98.63, and 2.46 pg/ml respectively. TNF-α concentration in THP-1 cells was increased by CLE (0.05, 0.8 mg/ml, Fig. 4a, p < 0.0001) relative to the control. In THP-1 cells, IL-1β and IL-6 concentrations were increased by 0.05 mg/ml CLE whereas decreased by 0.2–0.8 mg/ml CLE as compared to the control (Fig. 4bc, p < 0.0001). Concentration of the anti-inflammatory cytokine, IL-10 was decreased in PBMC’s while increased in THP-1 cells by CLE (0.05–0.8 mg/ml) relative to the control (Figs. 3d and 4d, p < 0.0001).

Fig. 3

Concentration of TNF-α (a), IL-1β (b), IL-6 (c) and IL-10 (d) in CLE treated PBMC’s (Values expressed as mean ± SD, * p < 0.01, *** p < 0.0001, compared to the control)

Fig. 4

Concentration of TNF-α (a), IL-1β (b), IL-6 (c) and IL-10 (d) in CLE treated THP-1 cells (Values expressed as mean ± SD, ** p < 0.001, *** p < 0.0001 compared to the control)

The antioxidant potential of CLE

The endogenous antioxidant activity of CLE was determined by measuring GSH levels in both cell lines (Table 1). At 24 h, GSH levels in PBMC’s were increased by 0.05–0.2 mg/ml CLE but decreased by 0.4–0.8 mg/ml CLE relative to the control (Table 1, p < 0.0001). In THP-1 cells, CLE (0.05–0.8 mg/ml) increased GSH levels as compared to the control (Table 1, 24 h, p < 0.0001). At 24 h, GSH concentrations were increased to a greater extent in THP-1 cells (0.068–3.890 μM) than PBMC’s (0.191–1.746 μM). At 72 h, CLE (0.05–0.8 mg/ml) increased GSH concentrations in PBMC’s and THP-1 cells by 1.13–5.91 μM and 0.12–0.19 μM respectively as compared to the control (Table 1, p < 0.0001). Notably, CLE increased GSH levels to a greater extent in PBMC’s as compared to THP-1 cells at 72 h.

Table 1 Glutathione levels in CLE treated PBMC’s and THP-1 cells

CLE modulates caspase (−8, −9, −3/7) activities and ATP levels

Luminometry assays were used to determine caspase activity and ATP levels in THP-1 cells and PBMC’s after treatment with CLE. The pro-apoptotic effect of CLE in PBMC’s treated for 24 h is shown in Table 2. At 24 h, PBMC caspase-8 activity was increased by 0.05–0.2 mg/ml CLE, whereas decreased by 0.4–0.8 mg/ml CLE as compared to the control (Table 2, p < 0.0001). CLE (0.05–0.8 mg/ml, 24 h) increased PBMC caspase −9 and −3/7 activities relative to the control (Table 2, p < 0.0001). Increased caspase activity led to the initiation and execution of PBMC apoptosis at 24 h. The PBMC ATP levels were increased by 0.4 mg/ml CLE, whereas decreased by 0.05, 0.2 and 0.8 mg/ml CLE (Table 2, p < 0.0001).

Table 2 Modulation of caspase (−8, −9, −3/7) activities and ATP levels in 24 h CLE treated PBMC’s

CLE pro-apoptotic effects in THP-1 cells treated for 24 h is shown in Table 3. At 24 h, CLE (0.05–0.8 mg/ml) increased THP-1 caspase-8 activity as compared to the control (Table 3, p < 0.0001). In THP-1 cells, caspase-9 activity and ATP levels were decreased by 0.05–0.4 mg/ml CLE, whereas increased by 0.8 mg/ml CLE relative to the control (Table 3, 24 h, p < 0.0001). The THP-1 caspase-3/7 activity was decreased by 0.2–0.4 mg/ml CLE, whereas increased by 0.05 and 0.8 mg/ml CLE as compared to the control (Table 3, 24 h, p < 0.0001). THP-1 caspase (−8, −9, −3/7) activities was increased by 0.8 mg/ml CLE, suggesting an increased initiation and execution of THP-1 apoptosis.

Table 3 Modulation of caspase (−8, −9, −3/7) activities and ATP levels in 24 h CLE treated THP-1 cells

The pro-apoptotic effect of CLE in PBMC’s treated for 72 h is shown in Table 4. At 72 h, PBMC caspase-8 activity was increased by 0.4 mg/ml CLE, whereas decreased by 0.05, 0.2, 0.8 mg/ml CLE relative to the control (Table 4, p < 0.0001). CLE (0.05–0.8 mg/ml) decreased PBMC caspase (−9, −3/7) activities and ATP levels as compared to the control (Table 4, 72 h, p < 0.0001). Decreased PBMC caspase activity suggests a decrease in PBMC apoptotic cell death.

Table 4 Modulation of caspase (−8, −9, −3/7) activities and ATP levels in 72 h CLE treated PBMC’s

CLE pro-apoptotic effects in THP-1 cells treated for 72 h is shown in Table 5. At 72 h, THP-1 caspase-8 activity was increased by 0.4 mg/ml CLE whereas decreased by 0.05, 0.2, 0.8 mg/ml CLE relative to the control (Table 5, p < 0.0001). CLE (0.05–0.8 mg/ml) decreased THP-1 caspase (−9, −3/7) activities and ATP levels as compared to the control (Table 5, 72 h, p < 0.0001). Decreased THP-1 caspase activity suggests a decrease in THP-1 apoptotic cell death.

Table 5 Modulation of caspase (−8, −9, −3/7) activities and ATP levels in 72 h CLE treated THP-1 cells


Cancer and cachexia have been associated with increased levels of oxidative stress, pro-inflammatory cytokines and apoptosis [6, 27]. The medicinal plant, C. asiatica possesses anti-inflammatory [29] and anti-tumor activity [35], which can be beneficial in the treatment of cancer cachexia.

Previously, Zainol et al. (2003) reported that C. asiatica possessed antioxidant potential, possibly associated with phenolic compounds [36]. The DPPH assay revealed that CLE has oxidant scavenging potential ranging between 45 and 84% at 0.05–0.8 mg/ml CLE. ROS plays a pivotal role in tumour initiation, inflammation, protein degradation and apoptosis. The significant oxidant scavenging potential of CLE may decrease inflammatory cytokine levels and ROS induced apoptosis.

At 24 h, CLE dose dependently decreased PBMC viability, whereas THP-1 viability remained unchanged. However, at 72 h, CLE significantly decreased both PBMC and THP-1 viability. C. asiatica derived compounds, asiatic acid and asiticoside, were shown to reduce RAW 264.7 cell viability (120 μM, 24 h) by 82% and 71% respectively [37]. Additionally, C. asiatica extracts inhibited breast (MCF-7) and liver (HepG2) cancer cell proliferation [33, 38], indicating our data on CLE cytotoxicity is in agreement with previous studies.

Inflammatory cytokines play an essential role in tumourgenesis and the cachectic syndrome [6]. Previously, Punturee et al. (2004) reported that C. asiatica ethanolic extract modulated/suppressed TNF-α production in mouse macrophages [39]. Our results also show that CLE decreased TNF-α concentration in PBMC’s. Yun et al. (2008) reported that the pre-treatment of RAW264.7 cells with asiatic acid significantly reduced IL-6 production with minimal effects on TNF-α and IL-1β levels [37]. Our findings, however, suggest that CLE modulates pro-inflammatory cytokine levels. In both PBMC’s and THP-1 cells, IL-1β and IL-6 levels were increased by the lower 0.05 mg/ml CLE concentration but decreased at the higher 0.4 mg/ml CLE concentration. Pro-inflammatory cytokines, over a chronic time period, stimulate the production of genotoxic molecules [nitric oxide (NO), ROS] and tumour progression by promoting angiogenesis and metastasis [1, 3]. Previous literature has shown that IL-1 stimulates malignant cell growth and invasiveness [3]. In addition, IL-6 exerts its tumour proliferative and anti-apoptotic potential by targeting genes involved in cell cycle progression and the suppression of apoptosis [3]. The ability of CLE to increase pro-inflammatory cytokines such as IL-1β in PBMC’s may aid in cancerous cell elimination through increased host anti-tumour activity. Conversely, in THP-1 cells, the decrease in IL-6 and IL-1β concentrations by CLE may diminish cytokine induced tumour immunosuppressive activity and cancer progression.

With regard to the cachectic syndrome, TNF-α inhibits the production of LPL and reduces the rate of LPL gene transcription [40,41,42], thereby preventing the formation of new lipid stores while stimulating HSL and increasing lipolysis [43]. In adipose tissue (in vivo), IL-6 decreased LPL activity leading to tissue wasting in cachectic individuals [19]. The potential of CLE (0.4 mg/ml) to decrease IL-6 and IL-1β concentrations in PBMC’s and THP-1 cells suggests a decrease in LPL inhibition and HSL stimulation, thus contributing to lipogenesis maintenance and minimal lipolysis. IL-6 and TNF-α further contribute to cachexia by stimulating muscle catabolism through the activation of the ubiquitin-proteasome pathway [21, 22, 44]. Furthermore, pro-inflammatory cytokines activate NF-κB which regulates the expression of genes involved in the suppression of tumour apoptosis, stimulation of tumour cell cycle progression and enhancement of inflammatory mediators [1, 3]. Taken together, NF-κB promotes tumour progression, invasion, angiogenesis and metastasis [1, 3]. In cachexia, NF-κB activation induces ubiquitin–proteasome pathway activity and suppresses MyoD expression [45], thereby increasing proteolysis and decreasing muscle replenishment [46]. By decreasing IL-6 and IL-1β concentrations in PBMC’s and THP-1 cells, CLE (0.4 mg/ml) may prevent excessive activation of NF-κB and proteasome pathways, ultimately decreasing proteolysis. Taken together, CLE may be able to decrease tissue wasting through the down regulation of pro-inflammatory cytokine levels.

The immunosuppressive and anti-inflammatory cytokine IL-10, inhibits tumour development, tumour progression, modulates apoptosis and suppresses angiogenesis during tumour regression [1, 3]. Additionally, IL-10 inhibits NF-κB activation and subsequently inhibits pro-inflammatory cytokine production (TNF-α, and IL-6) [3]. With regard to tissue wasting, increased IL-10 levels in colon 26- bearing mice was reported to reverse the cachectic syndrome [47]. The decreased PBMC IL-10 concentration may be due to IL-10 combating increased pro-inflammatory cytokine levels (IL-6 and IL-1β). In THP-1 cells, the potential of CLE to increase IL-10 levels will facilitate a decrease in pro-inflammatory cytokine levels, a decrease in malignant cell progression and possibly alleviate the cancer cachectic syndrome.

GSH, a potent antioxidant [48], effectively scavenges ROS both directly and indirectly [49]. In PBMC’s and THP-1 cells, CLE increased GSH concentrations. At 72 h, CLE (0.4 mg/ml) increased GSH levels more significantly in PBMC’s (1.45-fold) than THP-1 cells (1.11-fold). This suggests that CLE induces a higher antioxidant defense in normal PBMC’s than cancerous THP-1 cells at 72 h.

Apoptosis is a tightly regulated process involving a number of check points before an irreversible point is reached [50]. The extrinsic (death receptors) and intrinsic (mitochondria) pathways are the two main apoptotic pathways [26]. Activation of initiator caspases (−8, −9) leads to the activation of execution caspase-3/7 resulting in activation of cytoplasmic endonucleases [26].

Previous studies reported that asiatic acid decreased cell viability, induced apoptosis and DNA fragmentation [51, 52]. In PBMC’s, CLE (0.4–0.8 mg/ml, 24 h) decreased caspase-8 activity. An increase in TNF-α levels initiates the extrinsic apoptotic pathway subsequently activating caspase-8. However, CLE decreased PBMC TNF-α levels which may have contributed to the decreased caspase-8 activity. At 24 h, CLE increased PBMC caspase (−8 (0.05–0.2 mg/ml), −9, −3/7 (0.05–0.8 mg/ml)) activities, suggesting the activation of the extrinsic and intrinsic apoptotic pathways. GSH regulates apoptosis by preventing ROS accumulation [53]. Previous studies have demonstrated that elevated GSH levels have been associated with resistance to apoptosis [54, 55]. In PBMC’s, the decrease in GSH levels and the increase in caspase (−9, −3/7) activities by CLE (0.4–0.8 mg/ml, 24 h) may have increased apoptosis ultimately decreasing PBMC cell viability. In THP-1 cells, CLE (0.05–0.4 mg/ml) increased caspase-8 activity and decreased caspase-9 activity, suggesting initiation of apoptosis through the extrinsic pathway (24 h). In CLE treated THP-1 cells, the decreased caspase-9 activity may have been a consequence of the increased GSH levels. Although extrinsic apoptosis was activated in THP-1 cells, CLE (0.2–0.4 mg/ml) decreased caspase-3/7 activity, indicating that apoptosis was not fully executed (24 h). Interestingly, CLE increased THP-1 caspase (−8, −9, −3/7) activities at 0.8 mg/ml (24 h), suggesting an increased initiation and execution of THP-1 apoptosis.

At 72 h, caspase activities were decreased in both cell lines, suggesting a decreased activation of apoptosis. In PBMC’s and THP-1 cells, the increase in GSH levels and the decrease in caspase (−9, −3/7) activities by CLE (0.05–0.8 mg/ml, 72 h) may have decreased apoptotic cell death. However, PBMC and THP-1 cell viability was deceased at 0.4–0.8 mg/ml CLE and 0.8 mg/ml CLE respectively, suggesting an alternative form of cell death occurred.

Increased caspase-3 and proteasome activity, as well as E3 ubiquitin-conjugating enzyme expression are associated with increased proteolysis [56]. Thus the ability of CLE to down regulate caspase activities in PBMC’s and THP-1 cells may decrease proteolysis and the progression of cancer cachexia.

The cachectic syndrome is characterized by a negative energy balance due to reduced food intake and abnormal metabolism [57]. The inability to ingest/ use nutrients [5] and the negative energy balance present in cachectic patients leads to catalysis of muscle and fat stores for energy production [58]. In PBMC’s, CLE decreased ATP levels, a possible consequence of the decreased cell viability. Cancer cells require high levels of ATP for cellular proliferation [59]. In THP-1 cells, CLE decreased ATP levels which may decrease THP-1 cell proliferation. However in cachexia, a decrease in ATP levels may contribute to tissue wasting.

The potent feeding stimulant neuropeptide Y (NPY) promotes food and energy intake [60]. Increased cytokine (IL-1, IL-6, TNF-α) levels may inhibit NPY signalling leading to decreased food intake and increased energy expenditure [60]. Leptin functions as a suppresser of food intake and stimulator of energy consumption [6]. Pro-inflammatory cytokines may inhibit feeding by mimicking the hypothalamic negative-feedback signalling effect of leptin [61]. Thus, the ability of CLE to decrease pro-inflammatory cytokine levels may increase food intake, decrease energy expenditure and possibly combat the negative energy balance associated with cancer cachexia.


Our results show that CLE increased oxidant scavenging activity and GSH levels, modulated pro-inflammatory cytokine levels and regulated apoptosis and caspase activity in normal PBMC’s and THP-1 cells. CLE may thus be effective in cancer cachexia.



One way analysis of variance


Adenosine triphosphate


Butylated hydroxytoluene

C. asiatica :

Centella asiatica


C: asiatica ethanolic leaf extract


Dimethyl sulphoxide


Deoxyribonucleic acid


2, 2-diphenyl-1 picrylhydrazyl


Enzyme-linked immunosorbant assay


Fatty acids


Foetal calf serum






Hormone sensitive lipase




Lipoprotein lipase






Nuclear factor kappa B


Nitric oxide


neuropeptide Y


Poly (ADP-ribose) polymerase


Peripheral blood mononuclear cells


Phosphate buffered saline


Pen/Strep Amphotericin B


Relative light units


Reactive oxygen species


Room temperature


South Africa




A leukaemic cell line


Tumour necrosis factor-α


4-[3-(4-iodophenyl)-2-(4-nitrophenyl)-2H-5-tetrazolio]-1,3-benzene disulfonate


  1. 1.

    Landskron G, De la Fuente M, Thuwajit P, Thuwajit C, Hermoso MA. Chronic Inflammation and cytokines in the tumor microenvironment. J Immunol Res. 2014;Article ID 149185.:1–19.

  2. 2.

    Grivennikov SI, Greten FR, Karin M. Immunity, inflammation, and cancer. Cell. 2010;140(6):883–99.

  3. 3.

    Lin WW, Karin M. A cytokine-mediated link between innate immunity, inflammation, and cancer. J Clin Invest. 2007;117(5):1175–83.

  4. 4.

    Martignoni ME, Kunze P, Friess H. Cancer cachexia. Mol Cancer. 2003;2(36):1–3.

  5. 5.

    Inui A. Cancer anorexia-cachexia syndrome: current issues in research and management. CA Cancer J Clin. 2002;52(2):72–91.

  6. 6.

    Tisdale MJ. Mechanisms of cancer cachexia. Physiol Rev. 2009;89(2):381–410.

  7. 7.

    Lanza-Jacoby S, Lansey SC, Miller EE, Cleary MP. Sequential changes in the activities of lipoprotien lipase and lipogenic enzymes during tumor growth in rats. Cancer Res. 1984;44(11):5062–7.

  8. 8.

    Vlassara H, Speigel RJ, San Doval D, Cerami A. Reduced plasma lipoprotien lipase activity in patients with malignancy-associated weight loss. Horm Metab Res. 1986;18(10):698–703.

  9. 9.

    Thompson MP, Cooper ST, Parry BR, Tuckey JA. Increased expression of the mRNA for hormone-sensitive lipase in adipose tissue of cancer patients. Biochim Biophys Acta. 1993;1180(3):236–42.

  10. 10.

    Lundholm K, Bennegard K, Eden E, Svaninger G, Emery PW, Rennie MJ. Efflux of 3-methylhistidine from the leg in cancer patients who experience weight loss. Cancer Res. 1982;42(11):4807–11.

  11. 11.

    Lecker SH, Solomon V, Mitch WE, Goldberg AL. Muscle protein breakdown and critical role of the ubiquitin-proteasome pathway in normal and disease states. J Nutr. 1999;129(1):227S–37S.

  12. 12.

    Anker SD, Chua TP, Ponikowski P, Harrington D, Swan JW, Kox WJ, et al. Hormonal changes and catabolic / anabolic imbalance in chronic heart failure and their importance for cardiac cachexia. Circulation. 1997;96(2):526–34.

  13. 13.

    Mehta VB, Hart J, Wewers MD. ATP-stimulated release of interleukin (IL)-1beta and IL-18 requires priming by lipopolysaccharide and is independent of caspase-1 cleavage. J Biol Chem. 2001;276(6):3820–6.

  14. 14.

    Akira S. Toll-like receptor signaling. J Biol Chem. 2003;278(40):38105–8.

  15. 15.

    Asehnoune K, Strassheim D, Mitra S, Kim JY, Abraham E. Involvement of reactive oxygen species in Toll-like receptor 4-dependent activation of NF-kappa B. J Immunol. 2004;172(4):2522–9.

  16. 16.

    Janssen-Heininger YM, Poynter ME, Baeuerle PA. Recent advances towards understanding redox mechanisms in the activation of nuclear factor kappaB. Free Radic Biol Med. 2000;28(9):1317–27.

  17. 17.

    Park HS, Jung HY, Park EY, Kim J, Lee WJ, Bae YS. Cutting edge: direct interaction of TLR4 with NAD(P)H oxidase 4 isozyme is essential for lipopolysaccharide-induced production of reactive oxygen species and activation of NF-kappaB. J Immunol. 2004;173(6):3589–93.

  18. 18.

    Beutler BA, Cerami A. Recombinant interleukin 1 suppresses lipoprotein lipase activity in 3T3-L1 cells. J Immunol. 1985;135(6):3969–71.

  19. 19.

    Greenberg AS, Nordan RP, McIntosh J, Calvo JC, Scow RO, Jablons D. Interleukin 6 reduces lipoprotien lipase activity in adipose tissue of mice in vivo and in 3T3-L1 adipocytes: a possible role for interleukin 6 in cancer cachexia. Cancer Res. 1992;52(15):4113–6.

  20. 20.

    Argil´es JM, Busquets S, Felipe A, L´opez-Soriano FJ. Molecular mechanisms involved in muscle wasting in cancer and ageing: cachexia versus sarcopenia. Int J Biochem Cell Biol. 2005;37(5):1084–104.

  21. 21.

    Garcia-Martinez C, Agell N, Llovera M, López-Soriano FJ, Argilés JM. Tumour necrosis factor-alpha increases the ubiquitinization of rat skeletal muscle proteins. FEBS Lett. 1993;323(3):211–4.

  22. 22.

    Garcia-Martinez C, Llovera M, Agell N, López-Soriano FJ, Argilés JM. Ubiquitin gene expession in skeletal muscle is increased during sepsis: involvement of TNF-alpha but not IL-1. Biochem Biophys Res Comm. 1995;217(3):839–44.

  23. 23.

    Ebisui C, Tsujinaka T, Morimoto T, Kan K, Iijima S, Yano M, et al. Interleukin-6 induces proteolysis by activating intracellular proteases (cathepsins B and L, proteasome) in C2C12 myotubes. Clin Sci. 1995;89(4):431–9.

  24. 24.

    Tisdale MJ. Loss of skeletal muscle in cancer: biochemical mechanisms. Front Biosci. 2001;6:D164–74.

  25. 25.

    Buck M, Chojkier M. Muscle wasting and dedifferentiation induced by oxidative stress in a murine model of cachexia is prevented by inhibitors of nitric oxide synthesis and antioxidants. EMBO J. 1996;15(8):1753–65.

  26. 26.

    Elmore S. Apoptosis: a review of programmed cell death. Toxicol Pathol. 2007;35(4):495–516.

  27. 27.

    Du J, Wang X, Miereles C, Bailey JL, Debigare R, Zheng B, et al. Activation of caspase-3 is an initial step triggering accelerated muscle proteolysis in catabolic conditions. J Clin Invest. 2004;113(1):115–23.

  28. 28.

    Orhan IE. Centella asiatica (L.) Urban: From Traditional Medicine to Modern Medicine with Neuroprotective Potential. Evid. Based Complement. Alternat. Med. 2012;Article ID 946259:1–8.

  29. 29.

    Farnsworth NR, Bunyapraphatsara N. Thai Medicinal plants: recommended for primary Health care system. Medicinal Plant Information Center: Mahidol University, Thailand; 1992.

  30. 30.

    Rao AV, Gurfinkel DM. The bioactivity of saponins: triterpenoid and steroidal glycosides. Drug Metabol Drug Interact. 2000;17(1–4):211–35.

  31. 31.

    Odhav B, Beekrum S, Akula US, Baijnath H. Preliminary assessment of nutritional value of traditional leafy vegetables in KwaZulu-Natal. South Africa J Food Comp Anal. 2007;20(5):430–5.

  32. 32.

    Dasgupta N, De B. Antioxidant activity of some leafy vegetables of India: a comparative study. Food Chem. 2007;101(2):471–4.

  33. 33.

    Babykutty S, Padikkala J, Sathiadevan PP, Vijayakurup V, Azis TK, Srinivas P, et al. Apoptosis induction of Centella asiatica on human breast cancer cells. Afr J Tradit Complement Altern Med. 2008;6(1):9–16.

  34. 34.

    Royal Botanic Gardens, Kew, Garden tMB. The plant list 2010.

  35. 35.

    Babu TD, Kuttan G, Padikkala J. Cytotoxic and anti-tumor properties of certain taxa of Umbelliferae with special reference to Centella asiatica (L.) Urban. J Ethnopharmacol. 1995;48(1):53–7.

  36. 36.

    Zainol MK, Abd-Hamid A, Yusof S, Muse R. Antioxidative activity and total phenolic compounds of leaf, root and petiole of four accessions of Centella asiatica (L.) Urban. Food Chem. 2003;81(4):575–81.

  37. 37.

    Yun KJ, Kim JY, Kim JB, Lee KW, Jeong SY, Park HJ, et al. Inhibition of LPS-induced NO and PGE2 production by asiatic acid via NF-κB inactivation in RAW 264.7 macrophages: possible involvement of the IKK and MAPK pathways. Int Immunopharmacol. 2008;8(3):431–41.

  38. 38.

    Hussin F, Eshkoor SA, Rahmat A, Othman F, Akim A. The Centella asiatica juice effects on DNA damage, apoptosis and gene expression in hepatocellular carcinoma (HCC). BMC Complement Altern Med. 2014;14:1–7.

  39. 39.

    Punturee K, Wild CP, Vinitketkumneun U. Thai medicinal plants modulate nitric oxide and tumor necrosis factor-alpha in J774.2 mouse macrophages. J Ethnopharmacol. 2004;95(2–3):183–9.

  40. 40.

    Cornelius P, Enerback S, Bjursell G, Olivecrona T, Pekala PH. Regulation of lipoprotien lipase mRNA content in 3T3-L1 cells by tumour necrosis factor. Biochem J. 1988;249(3):765–9.

  41. 41.

    Fried SK, Zechner R. Cachectin/tumor necrosis factor decreases human adipose tissue lipoprotien lipase mRNA levels, synthesis and activity. J Lipid Res. 1989;30:1917–23.

  42. 42.

    Zechner R, Newman TC, Sherry B, Cerami A, Breslow JL. Recombinant human cachectin/tumor necrosis factor but not interleukin-1 alpha downregulates lipoprotien lipase gene expression at the transcriptional level in mouse 3T3-L1 adipocytes. Mol Cell Biol. 1988;8(6):2394–401.

  43. 43.

    Elborn JS, Cordon SM, Western PJ, MacDonald IA, Shale DJ. Tumor necrosis factor α, resting energy expenditure and cachexia in cystic fibrosis. Clin Sci (Lond). 1993;85(5):563–8.

  44. 44.

    Llovera M, Garcia-Martinez C, Agell N, López-Soriano FJ, Argilés JM. TNF can directly induce the expression of ubiquitin-dependent proteolytic system in rat soleus muscles. Biochem Biophys Res Commun. 1997;230(2):238–41.

  45. 45.

    Russell ST, Rajani S, Dhadda RS, Tisdale MJ. Mechanism of induction of muscle protein loss by hyperglycaemia. Exp Cell Res. 2009;315(1):16–25.

  46. 46.

    Guttridge DC, Mayo MW, Madrid LV, Wang CY, Baldwin ASJ. NF-κB-induced loss of MyoD messenger RNA: possible role in muscle decay and cachexia. Science. 2000;289(5488):2363–6.

  47. 47.

    Fujiki F, Mukaida N, Hirose K, Ishida H, Harada A, Ohno S, et al. Prevention of adenocarcinoma colon 26-induced cachexia by interleukin 10 gene transfer. Cancer Res. 1997;57(1):94–9.

  48. 48.

    Kehrer JP, Lund LG. Cellular reducing equivalents and oxidative stress. Free Radic Biol Med. 1994;17(1):65–75.

  49. 49.

    Fang YZ, Yang S, Wu G. Free radicals, antioxidants, and nutrition. Nutrition. 2002;18(10):872–9.

  50. 50.

    Kroemer G, Petit P, Zamzami N, Vayssiere JL, Mignotte B. The biochemistry of programmed cell death. FASEB J. 1995;9(13):1277–87.

  51. 51.

    Lee YS, Jin DQ, Kwon EJ, Park SH, Lee E, Jeong TC, et al. Asiatic acid, a triterpene, induces apoptosis through intracellular Ca2+ release and enhanced expression of p53 in HepG2 human hepatoma cells. Cancer Lett. 2002;186(1):83–91.

  52. 52.

    Park BC, Bosire KO, Lee ES, Lee YS, Kim JA. Asitic acid induces apoptosis in SK-MEL-2 human melanoma cells. Cancer Lett. 2005;218(1):81–90.

  53. 53.

    Franco R, Cidlowski JA. Apoptosis and glutathione: beyond an antioxidant. Cell Death Differ. 2009;16(10):1303–14.

  54. 54.

    Friesen C, Kiess Y, Debatin KM. A critical role of glutathione in determining apoptosis sensitivity and resistance in leukemia cells. Cell Death Differ. 2004;11(1):S73–85.

  55. 55.

    Cazanave S, Berson A, Haouzi D, Vadrot N, Fau D, Grodet A, et al. High hepatic glutathione stores alleviate Fas-induced apoptosis in mice. J Hepatol. 2007;46(5):858–68.

  56. 56.

    Wang X, Hu Z, Hu J, Du J, Mitch WE. Insulin resistance accelerates muscle protein degradation: activation of the ubiquitin– proteasome pathway by defects in muscle cell signaling. Endocrinology. 2006;147(9):4160–8.

  57. 57.

    Bluma D, Omlin A, Baracos VE, Solheim TS, Tan BH, Stone P, et al. Cancer cachexia: a systematic literature review of items and domains associated with involuntary weight loss in cancer. Crit Rev Oncol Hematol. 2011;80(1):114–44.

  58. 58.

    Falconer JS, Fearon KC, Plester CE, Ross JA, Carter DC. Cytokines, the acute-phase response, and resting energy expenditure in cachectic patients with pancreatic cancer. Ann Surg. 1994;219(4):325–31.

  59. 59.

    Gorrini C, Harris IS, Mak TW. Modulation of oxidative stress as an anticancer strategy. Nat Rev Drug Discov. 2013;12(12):931–47.

  60. 60.

    Inui A. Cancer anorexia-cachexia syndrome: are neuropeptides the key? Cancer Res. 1999;59(18):4493–501.

  61. 61.

    Aleman MR, Santolaria F, Batista N, de La Vega M, Gonzalez-Reimers E, Milena A, et al. Leptin role in advanced lung cancer. A mediator of the acute phase response or a marker of the status of nutrition? Cytokine. 2002;19(1):21–6.

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We are grateful to the National Research Foundation, the South African Medical Research Council, Department of Science and Technology, Government of India and Manipal University, India for financial support to conduct experimentation. The authors also acknowledge Miss Tarylee Reddy for assistance with statistical analysis of data.


Sources of funding included the National Research Foundation, the South African Medical Research Council and Department of Science and Technology, India and Manipal University, India. The funding sources were not involved in study design, collection of samples, analysis of data, interpretation of data, writing of the report and decision to publish. Scientific out-put is a requirement of the National Research Foundation.

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DBN carried out all experimentation except the luminometry (Caspase, ATP, GSH) assays. DBN analysed and interpreted data, performed statistical analysis, drafted and revised the manuscript. AC and AP carried out luminometry assays and provided intellectual input into the manuscript. VS, KPG and KS gave substantial contributions to conception, design and supervision of the study and revision of the manuscript. All authors read and approved the final manuscript.

Correspondence to Vikash Sewram.

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Naidoo, D.B., Chuturgoon, A.A., Phulukdaree, A. et al. Centella asiatica modulates cancer cachexia associated inflammatory cytokines and cell death in leukaemic THP-1 cells and peripheral blood mononuclear cells (PBMC’s). BMC Complement Altern Med 17, 377 (2017).

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  • Cancer
  • Cachexia
  • Cytokines
  • Apoptosis
  • Centella asiatica