The cyclic peptide mallotumide A inhibits colon and breast cancer cell growth and motility by targeting cellular respiration and lipogenesis

Laowittawat, Chayanee; Sawektreeratana, Natthapat; Katewongsa, Kanlaya; Payomhom, Pattaree; Hongthong, Sakchai; Reutrakul, Vichai; Kuhakarn, Chutima; Jitrapakdee, Sarawut

doi:10.1038/s41598-025-24547-x

Download PDF

Article
Open access
Published: 19 November 2025

The cyclic peptide mallotumide A inhibits colon and breast cancer cell growth and motility by targeting cellular respiration and lipogenesis

Chayanee Laowittawat¹,
Natthapat Sawektreeratana²,
Kanlaya Katewongsa³,
Pattaree Payomhom³,
Sakchai Hongthong⁴,
Vichai Reutrakul²,
Chutima Kuhakarn² &
…
Sarawut Jitrapakdee^1,3

Scientific Reports volume 15, Article number: 40774 (2025) Cite this article

1869 Accesses
1 Citations
Metrics details

Subjects

Abstract

We have recently isolated, and determined the structure of a cycloheptapeptide, Mallotumide A from the Mallotus spodocarpus root extract. Here we reported the anti-cancer activity of Mallotumide A in highly invasive colon cancer, HCT116 and triple-negative breast cancer, MDA-MB-231 cell lines. Mallotumide A, at concentrations of 1 nM and 10 nM, completely inhibited the clonogenic growth, migration, and invasion of HCT116 and MDA-MB-231 cells, respectively. While the compound interfered with cell cycle progression without inducing apoptosis, exposure to 10 nM Mallotumide A for 48 h reduced the expression of two key lipogenic enzymes, ACC1 and FASN, by approximately 50% in both cell lines. The downregulation of ACC1 and FASN was accompanied by a 50% reduction in intracellular triglyceride levels while the cholesterol levels remained unaffected. Mallotumide A also moderately decreased AMP-activated protein kinase (AMPK) and ATP levels. Extracellular flux analysis revealed that acute exposure of both cancer cell lines to 1 nM and 10 nM Mallotumide A for 24 h markedly lowered the oxygen consumption rate. This was accompanied by reductions in basal and ATP-linked respiration, maximal respiration, and mitochondrial spare respiratory capacity. Mallotumide A also decreased the extracellular acidification rate, affecting both basal glycolysis and the glycolytic reserve. These findings suggest that the anti-cancer effects of Mallotumide A are associated with disruptions in cellular energy metabolism and the de novo lipogenesis pathway in cancer cells. This study underscores the potential of Mallotumide A as a novel anti-cancer agent.

Tumor metabolome remolded by low dose mitochondrial uncoupler elicites robust CD8⁺ T cell response

Article Open access 01 July 2025

Prognosis prediction and risk stratification of breast cancer patients based on a mitochondria-related gene signature

Article Open access 03 February 2024

Triple negative breast cancer migration is modified by mitochondrial metabolism alteration induced by natural extracts of C. spinosa and P. alliacea

Article Open access 31 August 2024

Introduction

Cyclic peptides are circularized via an amide bond formed between the first and the last amino acids to produce a cyclic structure. Several cyclic peptides are found in natural sources including fungi, bacteria, marine organisms, and plants as part of the defense molecules against pathogen invasion^1,2. Due to the absence of N- and C-termini, the cyclic peptides are more stable and resistant to proteases. The cyclic peptides also have high structural diversity attributed to different amino acid compositions, shapes, and conformations. Furthermore, some cyclic peptides possess a relatively large surface area, allowing them to interact with the target molecules with high affinity and specificity^3,4. Because of their rigid structures, unique conformations, novel cyclic peptides with various biological activities such as anti-bacterial, anti-inflammation, and anti-cancer have been screened from natural products, aiming for their uses as the therapeutic application. Significant efforts have been made to generate cyclic peptides through genetic and chemical methods, with the goal of producing molecules that possess desired biological activities. Due to their high affinity and specificity, cyclic peptides with anti-cancer properties have garnered significant attention as therapeutic agents. Currently, there are more than 100 cyclic peptides that have been discovered from nature; however, only 40 have been approved as therapeutic peptides^5,6. Although many cyclic peptides exhibit anti-cancer activity, only a few demonstrate efficacy at micromolar concentrations, and none have been reported to exhibit activity at the nanomolar level⁷. For example, RA-V, a plant-derived cyclopeptide, possessed an anti-cancer activity by inducing apoptosis at 1 µM concentration⁸. RA-XII, a natural cyclopeptide isolated from Rubia yunnanensis, possessed an anti-proliferative activity by inducing apoptosis in HepG2 cell at 1.34 µM concentration⁹.

Our research group has recently isolated Mallotus spodocarpus plant-derived cyclic peptides, namely Mallotumides A-C. The Mallotumide A is a cycloheptapeptide comprising the non-canonical amino acids, cyclo-[(2R,3S)-D-allo-Ile¹]-[(2R,3S)-D-allo-Ile²]-[(2S)-L-Me-Ala¹]-[(2S,3R)-L-threo-Me-FAS]-[(2S,4S,5S)-DMCG]-[(2S)-Me-Ala²]-[(2R,3S)-D-threo-FAS] (FAS = β-(3-furyl)-O-acetylserine, DMCG = α-(2,3-dimethylcyclopropyl)glycine) with a molecular weight of 926.08 Da (C₄₆H₆₇N₇O₁₃)¹⁰. Its chemical structure is shown in Fig. 1A. Mallotumide A exhibits dose- and time-dependent cytotoxicity against hypopharyngeal carcinoma (FaDu) and keratinocyte (HaCaT) cells with IC₅₀ of 70 nM and 350 nM, respectively following 24 h exposure¹⁰. In this work, we investigated the anti-cancer activity of this compound in the highly invasive colon and triple negative breast cancer cell lines, namely HCT116 and MDA-MB-231, respectively. Mallotumide A inhibits cell proliferation, clonogenic growth, migration, invasion, lipogenic enzyme expression, and triglyceride accumulation at nanomolar concentrations. Furthermore, it interferes with cellular bioenergetics by lowering glycolysis and mitochondrial respiration.

Results

Mallotumide A inhibits growth, migration and invasion of HCT116, MDA-MB-231 cells at nanomolar range

The Mallotumide A used in this study exhibited a purity greater than 98%, as determined by HPLC analysis (supplementary Fig. 1). To investigate the anti-cancer activity of Mallotumide A, its cytotoxicity was evaluated in the highly invasive colon cancer (HCT116) and triple-negative breast cancer (MDA-MB-231) cell lines. Cells were treated with various concentrations of Mallotumide A for 24 and 48 h, and the IC₅₀ values were determined. Growth inhibition curves for HCT116 and MDA-MB-231 cells are shown in Fig. 1B,C, respectively. Overall, Mallotumide A induced dose-dependent cytotoxicity in both cell lines at both time points. Mallotumide A at lower concentrations (0.0001 nM–0.1 nM) gradually inhibited growth of HCT116 cells while 1 nM concentration sharply inhibited cell growth to 30% and 20% following 24 h and 48 h, respectively. Mallotumide A at the highest concentration (10 nM) inhibited the growth of cells by 80% and 95% following 24 h and 48 h treatment. Unlike HCT116 cell lines, low concentrations (0.0001–10.0001 nM) of Mallotumide A gradually inhibited the growth of MDA-MB-231 cells with the remaining cell viability around 50% and 25% following 24 h and 48 h exposure, respectively. However, growth inhibition reached a plateau at the maximal inhibition of 65% and 85% when higher concentrations of Mallotumide A (100 nM and 1,000 nM) were exposed to the cells. According to the growth inhibition curves, IC₅₀ of Mallotumide A for the HCT116 cell line following 24 h and 48 h exposure was approximately 0.44 ± 0.13 nM and 0.13 ± 0.06 nM, respectively while that for the MDA-MB-231 cell line at 24 h and 48 h was approximately 6.05 ± 4.69 nM and 0.19 ± 0.11 nM, respectively. In contrast, the IC₅₀ of 5-fluorouracil (5-FU), the chemotherapeutic drug was within the micromolar range, which is > 100,000 higher than that of Mallotumide A both in HCT116 (Fig. 1D) and MDA-MB-231 (Fig. 1E) cells.

Microscopic examination of HCT116 cells exposed to 0.1 nM, 1 nM, and 10 nM of Mallotumide A for 24 h revealed that their morphology was similar to that of the control cells, while those being exposed to 100 nM Mallotumide A were rounded up and did not grow to form a monolayer (Fig. 1F). At 48 h, HCT116 cells exposed to 0.1 nM, 1 nM and 10 nM of Mallotumide A still showed the similar morphology as the control cells; however, cells exposed to 100 nM Mallotumide A become swelling, suggesting the cellular damage (Fig. 1F). MDA-MB-231 cells exposed to 0.1 nM and 1 nM of Mallotumide A showed similar cell shape as the control cells following 24 h. However, when exposed to 10 nM, MDA-MB-231 cells became spindle-shaped. Most cells started to lose the contact with adjacent cells while most cells were shrunk and much less confluence when being treated with 100 nM concentration (Fig. 1G). Following 48 h exposure, cells exposed to 10 nM had much less confluence than 24 h while significant portion of cells being exposed to 100 nM concentrations started to detach and disintegrate. Mallotumide A’s inhibitory effect on both cancer cell lines suggested that this compound may trigger cell death via apoptosis. However, surprisingly, measurements of caspase-3/7 activity in both HCT116 and MDA-MB-231 cells treated with Mallotumide A showed a decrease (Fig. 2A,B, respectively), contrary to the expected induction seen with 25 µM 5-FU treatment. Furthermore, western blot analysis of poly(ADP-ribose) polymerase) (PARP), another apoptosis marker, revealed no apparent change in its cleavage in either cell line (Fig. 2C,D, respectively). This data suggests that the compound’s cytotoxicity in these cells may be attributed to a non-apoptotic mechanism. The inhibitory effect of Mallotumide A on cell growth, without inducing apoptosis, suggests that this compound may disrupt cell cycle progression. Figure 2E,F show the cell cycle profiles of HCT116 and MDA-MB-231 cells treated with 1 nM and 10 nM Mallotumide A. The result showed that 1 nM Mallotumide A caused a modest but significant decrease in the G0/G1 population of HCT116 cells, along with an increase in the G2/M phase. At 10 nM, there was a 40–50% reduction in both G0/G1 and S phases, accompanied by a threefold increase in the G2/M population, indicating G2/M cell cycle arrest (Fig. 2G). A similar, albeit less pronounced, trend was observed in MDA-MB-231 cells (Fig. 2H). These results suggest that Mallotumide A inhibits cell proliferation partly by inducing G2/M arrest without triggering apoptosis.

Next, we examined the clonogenic assay which measures the capacity of single cells to form cancer colonies in vitro. Both HCT116 and MDA-MB-231 cells were treated with indicated concentrations of Mallotumide A for 10 days. As shown in Fig. 3A, Mallotumide A inhibited clonogenic growth in a dose-dependent manner with 1 nM concentration completely inhibited both cell lines to form the colony. Both HCT116 and MDA-MB-231 cells exhibited a dose-dependent reduction in surviving fraction—the proportion of cells retaining the ability to form colonies following treatment—upon exposure to increasing concentrations of Mallotumide A (Fig. 3A, left and right panels, respectively). Since Mallotumide A at 100 nM caused morphological changes and cell death in both cell lines, subsequent experiments were conducted using concentrations of 0.1 nM, 1 nM, and 10 nM. Cell migration and invasion are key characteristics of cancer cells. To assess the effect of Mallotamide A on these properties, we evaluated its ability to inhibit these processes. Both cell lines were treated with Mallotamide A for 48 h before being tested for migration through the extracellular matrix. Mallotumide A inhibited cell migration in HCT116 cells at a concentration of 0.1 nM, resulting in a 50% decrease in migration. A concentration of 1 nM nearly completely inhibited migration. For MDA-MB-231 cells, a concentration of Mallotumide A that was 10 times higher than that used for HCT116 cells was required to achieve the same level of inhibition (Fig. 3B). A similar range of Mallotumide A concentration was required to inhibit invasion of HCT116. In contrast, up to 10 nM was required to entirely inhibit the invasion of MDA-MB-231 cells (Fig. 3C). The inhibitory effect of Mallotumide A on cell invasion was accompanied by the reduced expression of matrix metalloproteinase-1 (MMP1), a major enzyme responsible for invasion through the extracellular matrix (Fig. 3D). In general, the HCT116 cell line was more sensitive to Mallotumide A than MDA-MB-231 cells. These data indicated that Mallotumide A exerts anti-cancer activity by inhibiting clonogenic growth, migration, and invasion in both cell lines.

Mallotumide A down-regulates expression of lipogenic enzyme

Lipid is an essential structural component of the cell, and we investigate whether Mallotumide A disrupts lipid synthesis in cancer cells, potentially contributing to growth inhibition. To assess this, we measured the expression of key enzymes involved in de novo lipogenesis—acetyl-CoA carboxylase-1 (ACC1) and fatty acid synthase (FASN)—as well as cholesterol synthesis, specifically HMG-CoA reductase (HMGCR). Additionally, we evaluated the levels of adipocyte triglyceride lipase (ATGL), an enzyme responsible for regulating triglyceride hydrolysis, and pyruvate carboxylase (PC), a gluconeogenic enzyme, using Western blot analysis.

Figure 4A,B showed the representative western blots of various lipogenic enzymes and their regulatory proteins of HCT116 and MDA-MB-231 cells treated with different concentrations of Mallotumide A, respectively. In HCT116 cells, 1 nM Mallotumide A suppressed ACC1 expression by approximately 50%, while 10 nM further reduced its expression by 65% (Fig. 4C). Western blot analysis of the phosphorylated form of ACC1 revealed that Mallotumide A increased ACC1 phosphorylation only at 10 nM (Fig. 4D,E), suggesting that ACC1 is inactivated at higher concentrations of the compound. Mallotumide A at both 1 and 10 nM also downregulated FASN expression to a similar extent as ACC1 (Fig. 4F). In contrast, Mallotumide A did not affect the expression of PC (Fig. 4G), an anaplerotic enzyme, or HMGCR, a key regulatory enzyme in cholesterol biosynthesis (Fig. 4H). At 1 and 10 nM, the compound moderately reduced ATGL expression by 25% and 40%, respectively (Fig. 4I). We also examined the expression of AMP-activated protein kinase (AMPK), an energy-sensing kinase, and its phosphorylated form (pAMPK). Mallotumide A at 10 nM caused a slight reduction in total AMPK levels (Fig. 4J). Although phosphorylation of AMPK was also reduced (Fig. 4K), the ratio of phosphorylated AMPK to total AMPK remained unchanged (Fig. 4L).

A similar pattern was observed in MDA-MB-231 cells, where Mallotumide A affected ACC1 expression (Fig. 4N), phosphorylated ACC1 (Fig. 4O), and the pACC1/ACC1 ratio (Fig. 4P), FASN (Fig. 4Q) but did not affect PC (Fig. 4R) and HMGCR expression (Fig. 4S). However, the inhibitory effect on ATGL expression was evident only at 10 nM (Fig. 4T). Similar to the findings in HCT116 cells, Mallotumide A downregulated total AMPK without affecting its phosphorylated form in MDA-MB-231 cells (Fig. 4U–W). We also found that sterol regulatory binding protein 1c (SREBP1c), a common transcriptional activator of both ACC1 and FASN, was reduced by 75% in HCT116 and by 40% in MDA-MB-231 cell lines (Fig. 4M,X, respectively).

As Mallotumide A down-regulated the expression of key lipogenic enzymes, we assessed whether the level of cellular triglyceride was decreased in parallel with that of ACC1 and FASN. HCT116 cells treated with 1 nM and 10 nM Mallotumide A showed 40% and 50% reduction of triglyceride, respectively (Fig. 5A). For MDA-MB-231 cells, only the cells exposed to 10 nM concentration displayed 50% reduction of triglyceride (Fig. 5B). The slight change of cholesterol was consistent with the unnoticeable changes of HMGCR expression in both HCT116 and MDA-MB-231 cells (Fig. 5C,D, respectively). These data suggested that Mallotamide A inhibited the growth of both HCT116 and MDA-MB-231 cells at least partially through the reduction of triglyceride through the down-regulation of key metabolic enzymes in lipid metabolism.

Mallotumaide A perturbs bioenergetic status of cancer cells by diminishing glycolysis, mitochondrial respiration and ATP levels

Because cellular respiration is required to support growth, cell motility, and lipid biosynthesis¹¹, the perturbation of glycolytic and mitochondrial metabolism can affect the above biological processes. Therefore, we assessed the cellular metabolism of Mallotumide A -treated cells using the Seahorse extracellular flux analyzer. The extracellular acidification (ECAR) and oxygen consumption rates (OCR), reflecting glycolytic and mitochondrial oxidative phosphorylation activities, respectively, were measured in both HCT116 and MDA-MB-231 that were acutely exposed to various concentrations of Mallotumide A for 24 h. Figure 6A,B showed the ECAR profiles of HCT116 and MDA-MB-231 cell lines treated with 0.1 nM, 1 nM, and 10 nM after injecting glucose, oligomycin, and 2-DG, respectively. Mallotumide A at 0.1 nM concentration did not affect HCT116 and MDA-MB-231 cell glycolysis. At higher concentrations (1 nM and 10 nM), Mallotumide A decreased glycolysis in HCT116 cells by 30% and 60%, respectively (Fig. 6C). At the same time, they inhibited glycolysis in MDA-MB-231 cells by 40% and 50%, respectively (Fig. 6D). The inhibitory effect on glycolysis by Mallotumide A at 1 nM and 10 nM concentrations was accompanied by the decreased glycolysis capacity (Fig. 6E,F) and glycolytic reserve (Fig. 6G,H) with a mild effect on non-glycolysis acidification (Fig. 6I,J).

Mallotumide A also reduced OCR in both HCT116 and MDA-MB-231 cell lines, as shown in Fig. 7A,B, respectively. Mallotumide A at 0.1 nM did not affect basal respiration of both HCT116 and MDA-MB-231 cell lines (Fig. 7C,I). While higher concentrations of Mallotumide A (1 nM and 10 nM) lowered basal respiration to the similar extent i.e., 40%−50% in HCT116 cells, only 10 nM concentration lowered basal respiration by 40% in MDA-MB-231 cells (Fig. 7C,I). A similar inhibitory effect of Mallotumide A at higher concentrations on ATP-linked respiration to the similar extent as the basal respiration was also observed in both cell lines (Fig. 7D,J). However, only Mallotumide A at concentration of 10 nM markedly decreased the maximal respiration to 30–40% (Fig. 7E,K), and mitochondrial spare respiratory capacity to 10%−20% in both cell lines (Fig. 7F,L) without the effect on non-mitochondrial respiration (Fig. 7G,M). The highest concentration of Mallotumide A decreased proton leak to 40% in HCT116 but not MDA-MB-231 cells (Fig. 7H,N). In general, Mallotumide A at 1 nM and 10 nM inhibited glycolytic activity of both HCT116 and MDA-MB-231 cells, while a higher concentration of Mallotumide A (10 nM) was required to inhibit basal and ATP-linked respiration in MDA-MB-231 cells to the similar extent as that observed in HCT116 cells by 1 nM concentration.

As the ATP-linked respiration was decreased, we sought to confirm whether the level of ATP in Mallotumide A-treated cells was also affected. As shown in Fig. 8A,B, Mallotumide A caused a moderate reduction in ATP levels in both cell lines, with a maximal decrease of approximately 40–50% compared to control cells.

Discussion

Accumulating data indicate that cyclic peptides have been exploited as therapeutic agents, such as anti-microbial, anti-cancer, anti-inflammation agents or hormonal drugs, and against cancer⁴. Our previous report showed that M. spodocarpus root extract contains several phenolic compounds, and cycloheptapeptides, namely Mallotumides A, B, and C. All cyclo-heptapeptides showed most potent cytotoxicity against six cancer cells with IC₅₀ between 0.60 and 4.02 nM¹⁰. Here, we exploited the anti-cancer activity of Mallotumide A and investigated its inhibitory effect on cellular metabolism in highly invasive breast and colon cancer cells.

Mallotumide A exerted a potent cytotoxic effect in a dose-dependent manner for HCT116 and MDA-MB-231 cell lines with IC₅₀ of 0.1 nM following 48 h. The IC₅₀ against HCT116 and MDA-MB-231 cells was 20-fold lower than our previously reported IC₅₀ for FaDu hypopharyngeal cancer and cholangiocarcinoma cell lines¹⁰. However, a 10-fold higher than IC₅₀ concentration was required to inhibit the clonogenic growth of both cell lines. The sub-nanomolar IC₅₀ for Mallotumide A is significantly lower than most cyclic peptides whose IC₅₀ is in the micromolar range. There have been only five cyclic peptides whose IC₅₀ falls below 10 nM range including Apratoxin A (0.5 nM)¹², Nannocystin A (2.6 nM)¹³, Coibamide A (2.8 nM)¹⁴, Vitilevuamide (6 nM)¹⁵ and IB-01212 (10 nM)¹⁶, however the molecular mechanism by which these peptides kill the cancer cells are mostly unknown. While some cyclic peptides exert cytotoxicity by inducing apoptosis, Mallotumide A does not appear to trigger apoptosis at concentrations up to 20 nM. This compound also disrupted cell cycle progression, resulting in a significant G2/M arrest in both HCT116 and MDA-MB-231 cell lines. This disruption, while a key contributor to the compound’s inhibitory effect on cell growth, was not sufficient to induce apoptosis. It is therefore plausible that this compound induces alternative, non-apoptotic cell death pathways. Several anticancer agents are known to promote cell death through non-apoptotic mechanisms such as ferroptosis, necroptosis, or autophagic cell death¹⁷. These pathways may explain the caspase-independent cytotoxic effects observed with Mallotumide A in HCT116 and MDA-MB-231 cells.

Mallotumide A also exerted its anti-cancer activity by inhibiting both migration and invasion. While Mallotumide A at 1 nM concentration completely inhibited migration and invasion of HCT116 cells, a 10-fold higher concentration of this compound was required to achieve the same degree of inhibition for MDA-MB-231 cells. This is consistent with the growth inhibition curve of MDA-MB-231 cells, in which a 10-fold higher concentration of the compound was required to achieve the same response as that of the HCT116 cells. The higher dose requirement for MDA-MB-231cells is likely attributed to this cell line’s highly tumorigenic and metastatic properties¹⁸. The inhibitory effect of Mallotumide A on invasion was partly mediated through the reduced MMP-1 expression. MMP-1 is a crucial matrix metalloproteinase that plays a key role in degrading the extracellular matrix during invasion^19,20,21. Similar to Mallotumide A, cyclopeptide RA-V was reported to inhibit migration and invasion of MCF-7 and MDA-MB-231 cells through down-regulation of MMP-1 expression. Nevertheless, only 1 nM of Mallotumide A was required to inhibit MMP-1 expression, while it required up to 25 nM for RA-V to achieve the same response²².

De novo lipogenesis is a crucial biosynthetic pathway that provides cellular structure and energy storage for cancer cells^23,24. Inhibiting lipogenesis by targeting key enzymes, including ACC1 and FASN, is one of the strategies to inhibit cancer growth^25,26,27. Unlike the inhibitory effect on growth, migration, and invasion at IC₅₀ concentration, the inhibitory effect on the expression of both ACC1 and FASN was only observed above its IC₅₀ (1–10 nM), where this effect on ACC1 expression was more pronounced in HCT116 cells. Nevertheless, the down-regulation of both ACC1 and FASN by Mallotumide A at above IC₅₀ concentration was accompanied by the reduced triglyceride levels in both cell lines to a similar extent. This result suggests that the > 50% reduced growth of HCT116 and MDA-MB-231 cells may be secondary to the depletion of lipids, which serve as energy storage and cellular structure. This result is in line with several previous studies that pharmacological inhibition or genetic suppression of ACC-1^28,29,30 and FASN^31,32 impaired the growth of several cancer models³³. Compared with other published cyclic peptides, Wang et al. reported that cyclic peptide RA-XII inhibited expression of FASN, stearoyl-CoA desaturase, and sterol regulatory element-binding transcription factor 1 transcription factor in the HCT116 cell line accompanied by reduced triglyceride level³⁴. However, this inhibitory effect required RA-XII up to 10 µM concentration to achieve a 50% reduction of FASN³⁴, while only 10 nM Mallotumide A was found to suppress the expression of ACC1 and FASN by 65–70%. Although affecting triglyceride contents, Mallotumide A did not affect the expression level of HMGCR, a key enzyme of cholesterol biosynthesis and cholesterol.

ATGL is the key enzyme that hydrolyzes triglycerides to free fatty acids, which are further used as a fuel for energy production^35,36. Recent studies showed that colorectal cancer up-regulates ATGL expression to increase energy production during rapid proliferation and metastasis^37,38. It is possible that the decreased expression of ATGL is a secondary effect of reduced lipogenesis. ATGL is transcriptionally regulated by the nuclear receptor PPARγ^39,40, which requires endogenous ligands such as polyunsaturated fatty acids (PUFAs) and their derivatives for full activation⁴¹. Therefore, the inhibition of lipogenesis through the down-regulation of ACC1 and FASN could potentially reduce PUFA levels, thereby attenuating PPARγ’s transcriptional activity, and consequently decreasing ATGL expression.

Mallotumide A’s inhibitory effect on metabolic enzyme expression is not random as it did not affect PC, an anaplerotic enzyme that support cancer growth⁴². Thus, Mallotumide A explicitly suppressed de novo lipogenesis enzymes rather than cholesterol synthesis. Although, Mallotumide A also interferes the AMPK, the energy-sensing kinase in the cells, only a high dose of Mallotumide A produced a mild effect on total AMPK without affecting its phosphorylation state. Despite ACC1 phosphorylation being a target of AMPK⁴³, only saw a noticeable change in ACC1 phosphorylation at 10 nM. The lack of correlation between the reduced total AMPK and reduced ACC1 phosphorylation in Mallotumide A-treated cells is not unexpected because AMPK also regulates non-metabolic processes, enabling cancer cells to proliferate and survive under anoikis-resistance conditions^44,45,46. Therefore, mild reduction of AMPK level by Mallotumide A is partly attributed to growth restriction and impaired cell migration and invasion.

Mallotumide A perturbs mitochondrial respiration profiles by lowering the basal- and ATP-linked respiration by approximately 50%, which was consistent with a mild reduction of cellular ATP. Not only affecting basal and ATP-linked respiration, this compound also reduced the maximal respiration and spare respiratory capacity which indicate the mitochondrial flexibility in response to the extra energy requirement during acute cellular stress⁴⁷. The inhibitory effect of Mallotumide A on maximum respiration and spare respiratory capacity suggests that this compound may induce mitochondria stress, lowering the oxidation capacity^48,49.

In addition to mitochondrial oxidative phosphorylation, glycolysis is a crucial metabolic pathway for ATP production in cells, particularly in rapidly proliferating cancer cells that require substantial energy. Several anticancer drugs target key glycolytic enzymes, such as hexokinase 2 (HK2)⁵⁰, LDH^51,52 and monocarboxylic acid transporters (MCT)⁵³, effectively reducing glycolysis and inducing cell death. Our results clearly demonstrate that Mallotumide A significantly reduces glycolytic activity in both HCT116 and MDA-MB-231 cell lines by moderately inhibiting glycolytic capacity and glycolytic reserve. Interestingly, HCT116 was more sensitive to Mallotumide A than MDA-MB-231. HCT116 cells had more decreases in glycolysis and mitochondrial stress parameters than MDA-MB-231, especially the proton leak in mitochondrial stress parameters, which only decreased in HCT116. Proton leak can be a sign of mitochondrial damage or can be used as a mechanism to regulate mitochondrial ATP production. Therefore, the combined results may be why HCT116 cells exposed to Mallotumide A-concentrations can exhibit better anti-cancer effects than MDA-MB-231.

While we provide clear evidence that Mallotumide A inhibits cancer cell growth, motility, and cellular metabolism, several limitations should be acknowledged. First, although the compound reduced the expression of key lipogenic enzymes (ACC1 and FASN), their upstream regulator SREBP1c, and triglyceride levels, it remains uncertain whether Mallotumide A directly inhibits de novo lipogenesis. The rate of de novo lipogenesis was not directly measured, limiting our ability to confirm this mechanism. Second, the extent to which the observed suppression of lipogenic enzymes, reduction in triglyceride levels, and altered cellular respiration are direct effects of Mallotumide A remains unclear, as the molecular target of the compound has not yet been identified. Although Mallotumide A exhibited greater potency in cancer cell lines (IC₅₀ ~2 nM) compared to non-transformed HaCaT cells (IC₅₀ ~20 nM), indicating a potential therapeutic window¹⁰, we did not evaluate its effects on the invasive behavior of non-transformed cells. As a result, the selectivity of Mallotumide A toward malignant versus non-malignant cell invasion remains to be determined. Further investigation is warranted to address this gap.

Conclusion

Mallotumide A, a novel plant-derived cyclic peptide, exhibits potent anti-cancer activity in highly invasive colon and breast cancer cell lines. It suppresses cancer cell proliferation, motility, induces cell cycle arrest without inducing apoptosis and disrupts cellular bioenergetics—including glycolysis, oxidative phosphorylation, and ATP production. It also downregulates key enzymes involved in de novo lipogenesis, such as ACC and FASN. These findings highlight the potential of Mallotumide A as a novel anti-cancer agent.

Methods

Reagents and cell lines

Mallotumide A was isolated from M. spodocarpus roots as described previously¹⁰. The purity and identity were determined by HPLC analysis and spectroscopic data, respectively. A stock solution of Mallotumide A was prepared and then diluted with DMSO to obtain the appropriate concentrations. The colon cancer cell line, HCT116 (ATCC: CCL-247) and breast cancer cell lines, MDA-MB-231 (ATCC: CRM-HTB-26) were cultured in McCoy’s 5 A medium, supplemented with 10% fetal bovine serum (FBS) and 1% (w/v) penicillin-streptomycin or in Dulbecco’s Modified Eagle’s Medium (high glucose) medium, supplemented with 10% FBS and 1% penicillin-streptomycin, respectively. All cell lines were maintained at 37 °C in an incubator with a humidified atmosphere of 95% air and 5% CO₂. The confluent cells were sub-cultured every 2–3 days. The effects of Mallotumide A on various parameters were evaluated in comparison to a vehicle control group, in which cells were treated with 0.1% DMSO.

Cytotoxicity assay

Cell cytotoxicity was determined by MTT (3-(4,5-dimethylthiazol-2-yl)−2,5 diphenyl tetrazolium bromide) assay (AppliChem, Darmstadt, Germany). Cells were seeded at the density of 5 × 10³ cells/well in a 96-well plate and maintained at 37 °C for 24 h in an incubator with a humidified atmosphere of 95% air and 5% CO₂. The cells were then treated with various concentrations of Mallotumide A and incubated at 37 °C for 24 or 48 h. After incubation, 1 mg/mL MTT solution was added to each well before incubation at 37 °C for 1 h. The MTT solution was then removed, followed by adding 150 µL of DMSO to dissolve the formazan crystals. Finally, the absorbance was measured at 540 nm using a microplate reader (Multiskan SkyHigh, Thermo Scientific).

Clonogenic assay

Five hundred cells were seeded in 35-mm culture dish and maintained in a humidified incubator at 37 °C for 24 h. After the incubation, cells were treated with various concentrations of Mallotumide A and incubated at 37 °C for 10 days. Clonogenic survival following Mallotumide A treatment was quantitated by calculating the surviving fraction for each experimental condition. Briefly, after 10 days of growth, colonies (defined as cell clusters of at least 50 cells) were fixed with 100% (v/v) methanol for 20 min and stained with 0.5% (w/v) crystal violet in 25% (v/v) methanol for 1 h. Colonies were then counted from four random fields under a light microscope and averaged. The plating efficiency (PE) of untreated control cells was first determined by dividing the number of colonies formed by the number of cells initially seeded in the control wells. The surviving fraction for each drug-treated group was subsequently calculated by dividing the number of colonies formed in the treated wells by the product of the initial number of cells seeded for that treatment and the plating efficiency of the untreated control. All data were derived from at least three independent biological replicates.

Caspase3/7 assay

HCT116 and MDA-MB-231 cells were seeded at a density of 1 × 10⁴ cells/well in Corning™ 96-well white plates and allowed to adhere overnight. Cells were treated with Mallotumide A (0.1, 1, or 10 nM) for 48 h. As a positive control, cells were treated with fluorouracil (25 µM for HCT116 or 100 µM for MDA-MB-231), while the vehicle control group received 0.1% DMSO. Following treatment, caspase3/7 activity was determined using the Caspase-Glo^® 3/7 Assay (G8091, Promega, Madison, WI, USA). 100 µL of Caspase-Glo^® 3/7 reagent was added directly to each well, and the plate was gently shaken for 30 s to lyse the cells. After incubation at room temperature for 2 h in the dark. The luminescence readouts were recorded using Multimode Plate Reader. To account for variations in cell number, parallel plates were seeded simultaneously in clear 96-well plates under identical treatment conditions. Total protein was extracted from these parallel plates and quantified using the BCA protein assay (Thermo Scientific, Massachusetts, United States). Caspase3/7 activity were normalized to the corresponding protein content and expressed relative to the vehicle-treated control.

Cell cycle analysis

HCT116 cells were seeded at a density of 1 × 10⁶ cells in T25 flasks, while MDA-MB-231 cells were seeded in T75 flasks, and allowed to attach overnight. Cells were then treated with Mallotumide A (1 or 10 nM) for 24 h, with 0.1% DMSO serving as the vehicle control. Following treatment, both adherent and floating cells were harvested, washed with PBS, and fixed by slowly adding 70% ice-cold ethanol. Fixed cells were stored at 4 °C for 30 min, washed with PBS, and incubated with RNase A (final concentration 80 µg/mL; Thermo Fisher Scientific, MA, USA) to remove RNA. The cells were subsequently stained with propidium iodide (final concentration 4 µg/mL) for DNA content analysis. Flow cytometric analysis was performed on a FACSCalibur instrument (BD Biosciences, NJ, USA) using CellQuest software, and the distribution of cells in G0/G1, S, and G2/M phases was determined.

Migration and invasion assays

3 × 10⁵ cells were seeded in a 6-well plate and cultured for 24 h. The cells were then pre-treated with Mallotumide A for 48 h at 37 °C. Following 48 h treatment, only adherence cells were trypsinized and resuspended in a serum-free medium. 1 × 10⁵ cells were plated onto matrix-free transwell for the migration assay and Matrigel-coated transwell (0.2 mg/mL) for the invasion assay. The lower chamber was replaced with a complete medium containing 10% FBS. The trans-well was incubated for 24 h at 37 °C. The non-migrated or non-invaded cells were removed and washed with 1x PBS three times. The migrated and invaded cells were fixed with 4% (v/v) paraformaldehyde for 20 min before staining with 0.5% (w/v) crystal violet in 25% (v/v) methanol for 1 h in the dark at room temperature. In the final step, cells were washed twice with tap water and then randomly counted from 3 fields under a light microscope.

Measurement of Glycolysis and mitochondrial oxygen consumption with seahorse XF24 analyzer

The extracellular acidification rate (ECAR) and the oxygen consumption rate (OCR) of cells treated with Mallotumide A were measured using a Seahorse XFe24 Analyzer (Agilent Technologies). HCT116 or MDA-MB-231 cells were seeded at the density of 3 × 10⁴ cells/well in XF24 cell culture plate with complete medium (McCoy’s 5 A medium supplemented with 10% FBS (HCT116) or DMEM high glucose medium and 10% FBS (MDA-MB-231) and maintained in a humidified incubator at 37 °C for 24 h. The cells were then maintained in the complete growth medium containing 0.1 nM, 1 nM and 10 nM of Mallotumide A at 37 °C for 24 h. Following 24 h, the medium containing Mallotumide A was replaced with DMEM supplemented with 2 mM glutamine and 5 mM HEPES without glucose, L-glutamine, phenol red, sodium pyruvate and sodium bicarbonate) and maintained at 37 °C in non-CO₂ incubator for 1 h. The glycolytic activity of cells was measured using the glycolysis stress test kit (Agilent Technologies,). The ECAR profiles were measured following the serial injections of glucose, oligomycin and 2-deoxyglucose to final concentration 10 mM, 1 µM, and 50 mM, respectively to the medium. The ECAR was recorded at time intervals and the glycolysis rate, glycolysis capacity, glycolysis reserve, and non-glycolytic acidification were analyzed using the Agilent Seahorse Wave software (Agilent Technologies). The ECAR was normalized with protein concentration of cells which was determined using the BCA protein assay (Thermo Scientific, Massachusetts, United States).

The cells were similarly prepared for the OCR measurement using a mitochondrial stress test kit. Following 24 h, the medium containing Mallotumide A was replaced with DMEM supplemented with 2 mM glutamine, 1 mM pyruvate, 10 mM glucose and 5 mM HEPES without phenol red and sodium bicarbonate. The OCR profiles were measured following the serial injections of oligomycin, FCCP and rotenone/antimycin A to final concentrations of 1.5 mM, 1 mM, and 0.5 mM, respectively, to the cells. The OCR was recorded at time intervals and the basal respiration, ATP-linked respiration, proton leak, maximal respiration, and spare respiratory capacity were analyzed with the Agilent Seahorse Wave software (Agilent Technologies). The OCR was normalized with number of cells following staining with Hoechst 33,342 (Abcam, Cambridge, UK).

All Seahorse assays were conducted in three independent experiments. For each experiment, two technical replicate wells were used per treatment. The normalized ECAR and OCR values from the technical replicates were averaged, and the standard deviation (SD) was calculated. The mean values and SDs from the three independent experiments are presented in the final result.

Adenosine triphosphate (ATP) measurement

The cellular ATP was measured using a luminescent ATP detection assay kit (Abcam, Cambridge, UK). Cells were seeded at a density of 1.2 × 10⁴ cells/well in a complete medium containing 10% FBS in a 96-well white plate for 24 h at 37 °C. The cells were then treated with a complete medium containing 10% FBS and Mallotumide A at 0.1 nM, 1 nM, and 10 nM, and incubated at 37 °C for 48 h. Cell lysis was performed by adding 50 µL of detergent solution to each well and mixed for 5 min. 50 µL of substrate solution was added to the cell lysate and incubated for 1 h in dark. The luminescence readouts were recorded using Multimode Plate Reader.

Protein extraction and Western Immunoblot analysis

3 × 10⁵ cells were seeded in 60-mm dish and cultured for 24 h before treatment with various concentrations of Mallotumide A for 48 h at 37 °C. Protein was extracted from cells with RIPA lysis buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 1 mM EDTA, 0.25% sodium deoxycholate, 1% NP-40, 1 mM DTT and 1x protease inhibitor cocktail (Roche)). The protein concentration was determined using the Bradford’s reagent (Bio-Rad, Foster city, CA, USA). 20 µg of proteins were subjected to 10% discontinuous polyacrylamide gel electrophoresis with 1x glycine running buffer (25 mM Tris-HCl pH 8.3, 193 mM glycine and 0.1% (w/v) SDS) under reducing conditions. The proteins were transferred to the polyvinylidene fluoride (PVDF) membranes using Trans-Blot^® Turbo™ System Tour (Bio-Rad, Foster city, CA, USA) with 1x transfer buffer (25 mM Tris-HCl, 192 mM glycine, 0.1% (w/v) SDS and 20% (v/v) methanol). The membranes were incubated in a blocking buffer containing 5% (w/v) non-fat dry milk or 5% (w/v) bovine serum albumin (BSA) in 0.1% Tween 20 in Tris-buffered saline (TBS-T) (20 mM Tris and 150 mM sodium chloride) at room temperature for 1 h. After blocking, PVDF membranes were incubated with the following primary antibodies: 1:1,000 dilutions of anti-Phospho-AMPKα (Thr172) (Cell Signaling, Danvers, MA, USA), anti-AMPK (Cell Signaling, Danvers, MA, USA), anti-ACC1 (Proteintech, Rosemont, USA), anti-FASN (Proteintech, Rosemont, USA), anti-PC⁵⁴, anti-HMGCR (Proteintech, Rosemont, USA), anti-ATGL (Proteintech, Rosemont, USA) and anti-β-actin (Proteintech, Rosemont, USA) antibodies at 4° C, overnight. The excess antibody was washed with TBS-T before incubation with either anti-rabbit or anti-mouse IgG conjugated with horseradish peroxidase HRP for 1 h, at room temperature. The immunoreactive bands were detected using the chemiluminescent HRP substrate (Merck Millipore, Massachusetts, USA), and the images were captured using G: Box Chemi XT4 Chemiluminescence or Fluorescence Imaging System (SYNGENE, Cambridge, UK).

Measurements of triglyceride and cholesterol

Cells were seeded at a density of 1.2 × 10⁴ cells/well for 24 h at 37 °C before treating with various concentrations of Mallotumide A for 48 h. Cell viability was determined prior to measuring triglyceride using CellTiter-Flour fluorescence assay. The fluorescence intensity was recorded using Tecan Spark 10 M Multimode Plate Reader (TECAN, Männedorf, Switzerland) with filter 380–400 nmEx/505Em. Triglyceride level was measured after the CellTiter-Flour fluorescence assay reagent was removed. In brief, 50 µL of glycerol lysis solution with lipase were added to each well, and the plate was gently mixed for 30 s, and incubated at 37 °C for 30 min. Then, 50 µL of glycerol detection reagent was added to cell lysate and further incubated at room temperature for 1 h. The luminescence signals were recorded using Tecan Spark 10 M Multimode Plate Reader. The fluorescence readout was normalized with the cell viability as described earlier.

For cholesterol measurement, the cells were prepared as described for the triglyceride measurement. After removal of CellTiter-Flour fluorescence assay reagent, cholesterol was measured using Cholesterol/Cholesterol Ester-Glo™ assay kit. 50 µL of cholesterol lysis solution was added to each well, and the plate was gently mixed for 30 s and incubated at 37 °C for 30 min. Then, 50 µL of cholesterol detection solution with esterase was added to cell lysate, and further incubated at room temperature for 1 h. The luminescence signals were recorded using Tecan Spark 10 M Multimode Plate Reader. The fluorescence readout was normalized with the cell viability as described earlier.

Statistical analysis

All experiments were performed independently at least three times (n = 3), representing biological replicates conducted on separate days with independently prepared samples. For assays conducted using multi-well plates, statistical analysis was performed by first averaging the two technical replicate wells for each treatment, and their standard deviation was calculated. The resulting values from all biological replicates were then used to calculate an overall mean and standard deviation for that particular assay. Statistical analysis was determined using one-way ANOVA for comparison between control and Mallotumide A-treated groups (GraphPad Prism, GraphPad Software Inc., San Diego, USA). P < 0.05 is considered a significant difference.

Data availability

All data generated or analyzed during this study are included in this published article and its supplementary information. The data that support the findings of this study are available on request from the corresponding authors (S.J. and C.K.).

References

Craik, D. J. Circling the enemy: Cyclic proteins in plant defence. Trends Plant. Sci. 14, 328–335. https://doi.org/10.1016/j.tplants.2009.03.003 (2009).
Article CAS PubMed Google Scholar
Jan-Christoph Westermann, D. J. & Craik,. Plant peptide toxins from nonmarine environments. Elsevier Sci. 63, 257–285 (2010).
Google Scholar
Choi, J. S. & Joo, S. H. Recent trends in Cyclic peptides as therapeutic agents and biochemical tools. Biomol. Ther. (Seoul). 28, 18–24. https://doi.org/10.4062/biomolther.2019.082 (2020).
Article CAS PubMed Google Scholar
Ramadhani, D., Maharani, R., Gazzali, A. M. & Muchtaridi, M. Cyclic peptides for the treatment of cancers: a review. Molecules https://doi.org/10.3390/molecules27144428 (2022).
Article PubMed PubMed Central Google Scholar
Zhang, H. & Chen, S. Cyclic peptide drugs approved in the last two decades (2001–2021). RSC Chem. Biol. 3, 18–31. https://doi.org/10.1039/d1cb00154j (2022).
Article CAS PubMed Google Scholar
Liu, H., Bai, L. & Jiang, X. Recent progress on total synthesis of cyclic peptides. Tetrahedron Lett. https://doi.org/10.1016/j.tetlet.2024.155314 (2024).
Article PubMed PubMed Central Google Scholar
Zhang, J. N., Xia, Y. X. & Zhang, H. J. Natural cyclopeptides as anticancer agents in the last 20 years. Int. J. Mol. Sci. https://doi.org/10.3390/ijms22083973 (2021).
Article PubMed PubMed Central Google Scholar
Fang, X. Y. et al. Plant cyclopeptide RA-V kills human breast cancer cells by inducing mitochondria-mediated apoptosis through blocking PDK1-AKT interaction. Toxicol. Appl. Pharmacol. 267, 95–103. https://doi.org/10.1016/j.taap.2012.12.010 (2013).
Article CAS PubMed Google Scholar
Song, L. et al. Natural cyclopeptide RA-XII, a new autophagy inhibitor, suppresses protective autophagy for enhancing apoptosis through AMPK/mTOR/P70S6K pathways in HepG2 cells. Molecules https://doi.org/10.3390/molecules22111934 (2017).
Article PubMed PubMed Central Google Scholar
Sawektreeratana, N. et al. Potent cytotoxic cycloheptapeptides from the roots of Mallotus Spodocarpus. Org. Lett. 25, 8183–8187. https://doi.org/10.1021/acs.orglett.3c02641 (2023).
Article CAS PubMed Google Scholar
Kim, S. Y. Cancer energy metabolism: shutting power off cancer factory. Biomol. Ther. (Seoul). 26, 39–44. https://doi.org/10.4062/biomolther.2017.184 (2018).
Article CAS PubMed Google Scholar
Kazemi, S. et al. Targeting of HER/ErbB family proteins using broad spectrum Sect. 61 inhibitors Coibamide A and apratoxin A. Biochem. Pharmacol. 183, 114317. https://doi.org/10.1016/j.bcp.2020.114317 (2021).
Article CAS PubMed Google Scholar
Krastel, P. et al. Nannocystin A: an elongation factor 1 inhibitor from Myxobacteria with differential Anti-Cancer properties. Angew Chem. Int. Ed. Engl. 54, 10149–10154. https://doi.org/10.1002/anie.201505069 (2015).
Article CAS PubMed Google Scholar
Medina, R. A. et al. Coibamide A, a potent antiproliferative Cyclic depsipeptide from the Panamanian marine Cyanobacterium leptolyngbya Sp. J. Am. Chem. Soc. 130, 6324–6325. https://doi.org/10.1021/ja801383f (2008).
Article ADS CAS PubMed PubMed Central Google Scholar
Edlera, M. C., Fernandezb, A. M., Lassotac, P., Irelandb, C. M. & Barrowsa, L. R. Inhibition of tubulin polymerization by vitilevuamide, a bicyclic marine peptide, at a site distinct from colchicine, the Vinca alkaloids, and Dolastatin 10. Biochem. Pharmacol. 63, 707–715 (2002).
Article Google Scholar
Luis, J. et al. IB-01212, a New Cytotoxic Cyclodepsipeptide Isolated from the Marine Fungus Clonostachys sp. ESNA-A009. J. Organ. Chem. 71, 3335–3338 (2006).
Lambrecht, R. et al. Drug-induced oxidative stress actively prevents caspase activation and hepatocyte apoptosis. Cell. Death Dis. 15, 659. https://doi.org/10.1038/s41419-024-06998-8 (2024).
Article CAS PubMed PubMed Central Google Scholar
Zhang, B., Shetti, D., Fan, C. & Wei, K. miR-29b-3p promotes progression of MDA-MB-231 triple-negative breast cancer cells through downregulating TRAF3. Biol. Res. 52, 38. https://doi.org/10.1186/s40659-019-0245-4 (2019).
Article CAS PubMed PubMed Central Google Scholar
Kessenbrock, K., Plaks, V. & Werb, Z. Matrix metalloproteinases: regulators of the tumor microenvironment. Cell 141, 52–67. https://doi.org/10.1016/j.cell.2010.03.015 (2010).
Article CAS PubMed PubMed Central Google Scholar
Wang, K. et al. Knockdown of MMP–1 inhibits the progression of colorectal cancer by suppressing the PI3K/Akt/c–myc signaling pathway and EMT. Oncol. Rep. 43, 1103–1112. https://doi.org/10.3892/or.2020.7490 (2020).
Article CAS PubMed PubMed Central Google Scholar
Hui Liu, Y. et al. Price1, the role of MMP-1 in breast cancer growth and metastasis to the brain in a xenograft model. BioMed. Cent. 12, 583 (2012).
Google Scholar
Leung, H. W. et al. Cyclopeptide RA-V inhibits cell adhesion and invasion in both Estrogen receptor positive and negative breast cancer cells via PI3K/AKT and NF-kappaB signaling pathways. Biochim. Biophys. Acta. 1853, 1827–1840. https://doi.org/10.1016/j.bbamcr.2015.04.020 (2015).
Article CAS PubMed Google Scholar
Germain, N. et al. Lipid metabolism and resistance to anticancer treatment. Biology (Basel) https://doi.org/10.3390/biology9120474 (2020).
Article PubMed Google Scholar
Phokrai, P. et al. Suppressed de Novo lipogenesis by plasma membrane citrate transporter inhibitor promotes apoptosis in HepG2 cells. FEBS Open. Bio. 8, 986–1000. https://doi.org/10.1002/2211-5463.12435 (2018).
Article CAS PubMed PubMed Central Google Scholar
Wang, Y. et al. Acetyl-CoA carboxylases and diseases. Front. Oncol. 12, 836058. https://doi.org/10.3389/fonc.2022.836058 (2022).
Article CAS PubMed PubMed Central Google Scholar
Corbet, C. & Feron, O. Emerging roles of lipid metabolism in cancer progression. Curr. Opin. Clin. Nutr. Metab. Care. 20, 254–260. https://doi.org/10.1097/MCO.0000000000000381 (2017).
Article CAS PubMed Google Scholar
Fhu, C. W. & Ali, A. Fatty acid synthase: An emerging target in cancer. Molecules https://doi.org/10.3390/molecules25173935 (2020).
Article PubMed PubMed Central Google Scholar
Liu, S. et al. Acetyl-CoA carboxylase 1 depletion suppresses de novo fatty acid synthesis and mitochondrial beta-oxidation in castration-resistant prostate cancer cells. J. Biol. Chem. https://doi.org/10.1016/j.jbc.2022.102720 (2022).
Article PubMed PubMed Central Google Scholar
Ma, J. et al. Aldo-keto reductase family 1 B10 affects fatty acid synthesis by regulating the stability of acetyl-CoA carboxylase-alpha in breast cancer cells. J. Biol. Chem. 283, 3418–3423. https://doi.org/10.1074/jbc.M707650200 (2008).
Article CAS PubMed Google Scholar
Beckers, A. et al. Chemical Inhibition of acetyl-CoA carboxylase induces growth arrest and cytotoxicity selectively in cancer cells. Cancer Res. 67, 8180–8187. https://doi.org/10.1158/0008-5472.CAN-07-0389 (2007).
Article CAS PubMed Google Scholar
Khan, A., Aljarbou, A. N., Khan, S. & Khan, M. A. Her-2-directed systemic delivery of fatty acid synthase (FASN) SiRNA with novel liposomal carrier systems in the breast cancer mouse model. J. Drug Target. 30, 634–645. https://doi.org/10.1080/1061186X.2022.2038613 (2022).
Article CAS PubMed Google Scholar
Du, Q. et al. FASN promotes lymph node metastasis in cervical cancer via cholesterol reprogramming and lymphangiogenesis. Cell. Death Dis. 13, 488. https://doi.org/10.1038/s41419-022-04926-2 (2022).
Article CAS PubMed PubMed Central Google Scholar
Jones, J. E. et al. Inhibition of Acetyl-CoA carboxylase 1 (ACC1) and 2 (ACC2) reduces proliferation and de Novo lipogenesis of EGFRvIII human glioblastoma cells. PLoS One. 12, e0169566. https://doi.org/10.1371/journal.pone.0169566 (2017).
Article ADS CAS PubMed PubMed Central Google Scholar
Wang, Y. et al. Inhibition of fatty acid synthesis arrests colorectal neoplasm growth and metastasis: Anti-cancer therapeutical effects of natural cyclopeptide RA-XII. Biochem. Biophys. Res. Commun. 512, 819–824. https://doi.org/10.1016/j.bbrc.2019.03.088 (2019).
Article ADS CAS PubMed Google Scholar
Vegliante, R., Di Leo, L., Ciccarone, F. & Ciriolo, M. R. Hints on ATGL implications in cancer: beyond bioenergetic clues. Cell. Death Dis. 9, 316. https://doi.org/10.1038/s41419-018-0345-z (2018).
Article CAS PubMed PubMed Central Google Scholar
Castelli, S., De Falco, P., Ciccarone, F., Desideri, E. & Ciriolo, M. R. Lipid catabolism and ROS in cancer: A bidirectional liaison. Cancers (Basel) https://doi.org/10.3390/cancers13215484 (2021).
Article PubMed Google Scholar
Yin, H. et al. Adipose triglyceride lipase promotes the proliferation of colorectal cancer cells via enhancing the lipolytic pathway. J. Cell. Mol. Med. 25, 3963–3975. https://doi.org/10.1111/jcmm.16349 (2021).
Article CAS PubMed PubMed Central Google Scholar
Xie, H. et al. Adipose triglyceride lipase activity regulates cancer cell proliferation via AMP-kinase and mTOR signaling. Biochim. Biophys. Acta Mol. Cell. Biol. Lipids. 1865, 158737. https://doi.org/10.1016/j.bbalip.2020.158737 (2020).
Article CAS PubMed PubMed Central Google Scholar
Kershaw, E. E. et al. PPARgamma regulates adipose triglyceride lipase in adipocytes in vitro and in vivo. Am. J. Physiol. Endocrinol. Metab. 293, E1736–1745. https://doi.org/10.1152/ajpendo.00122.2007 (2007).
Article CAS PubMed Google Scholar
Kim, J. Y., Tillison, K., Lee, J. H., Rearick, D. A. & Smas, C. M. The adipose tissue triglyceride lipase ATGL/PNPLA2 is downregulated by insulin and TNF-alpha in 3T3-L1 adipocytes and is a target for transactivation by PPARgamma. Am. J. Physiol. Endocrinol. Metab. 291, E115–127. https://doi.org/10.1152/ajpendo.00317.2005 (2006).
Article CAS PubMed Google Scholar
Toshimasa Itoh, L. et al. Structural basis for the activation of PPARγ by oxidized fatty acids. Nat. Struct. Mol. Biol. 15, 9 (2008).
Google Scholar
Kiesel, V. A. et al. Pyruvate carboxylase and cancer progression. Cancer Metab. 9, 20. https://doi.org/10.1186/s40170-021-00256-7 (2021).
Article PubMed PubMed Central Google Scholar
Zordoky, B. N. et al. AMPK-dependent inhibitory phosphorylation of ACC is not essential for maintaining myocardial fatty acid oxidation. Circ. Res. 115, 518–524. https://doi.org/10.1161/CIRCRESAHA.115.304538 (2014).
Article CAS PubMed Google Scholar
Laderoute, K. R. et al. 5’-AMP-activated protein kinase (AMPK) supports the growth of aggressive experimental human breast cancer tumors. J. Biol. Chem. 289, 22850–22864. https://doi.org/10.1074/jbc.M114.576371 (2014).
Article CAS PubMed PubMed Central Google Scholar
Rios, M. et al. AMPK activation by oncogenesis is required to maintain cancer cell proliferation in astrocytic tumors. Cancer Res. 73, 2628–2638. https://doi.org/10.1158/0008-5472.CAN-12-0861 (2013).
Article CAS PubMed Google Scholar
Sravanth, K. et al. Rangarajan1, identification of a novel AMPK-PEA15 axis in the anoikis-resistant growth of mammary cells. Breast Cancer Res. (2014).
Marchetti, P., Fovez, Q., Germain, N., Khamari, R. & Kluza, J. Mitochondrial spare respiratory capacity: Mechanisms, regulation, and significance in non-transformed and cancer cells. FASEB J. 34, 13106–13124. https://doi.org/10.1096/fj.202000767R (2020).
Article CAS PubMed Google Scholar
De Oliveira, M. P. & Liesa, M. The role of mitochondrial fat oxidation in cancer cell proliferation and survival. Cells https://doi.org/10.3390/cells9122600 (2020).
Article PubMed PubMed Central Google Scholar
Martinez-Reyes, I. et al. TCA cycle and mitochondrial membrane potential are necessary for diverse biological functions. Mol. Cell. 61, 199–209. https://doi.org/10.1016/j.molcel.2015.12.002 (2016).
Article CAS PubMed Google Scholar
DeWaal, D. et al. Hexokinase-2 depletion inhibits Glycolysis and induces oxidative phosphorylation in hepatocellular carcinoma and sensitizes to Metformin. Nat. Commun. 9, 446. https://doi.org/10.1038/s41467-017-02733-4 (2018).
Article ADS CAS PubMed PubMed Central Google Scholar
Xie, H. et al. Targeting lactate dehydrogenase–a inhibits tumorigenesis and tumor progression in mouse models of lung cancer and impacts tumor-initiating cells. Cell. Metab. 19, 795–809. https://doi.org/10.1016/j.cmet.2014.03.003 (2014).
Article CAS PubMed PubMed Central Google Scholar
Verma, S. et al. Pharmacologic LDH inhibition redirects intratumoral glucose uptake and improves antitumor immunity in solid tumor models. J. Clin. Invest. https://doi.org/10.1172/JCI177606 (2024).
Article PubMed PubMed Central Google Scholar
Feng, Q. et al. Severely polarized extracellular acidity around tumour cells. Nat. Biomedical Eng. 8, 787–799. https://doi.org/10.1038/s41551-024-01178-7 (2024).
Article CAS Google Scholar
Rohde, M., Lim, F. & Wallace, J. C. Electron microscopic localization of pyruvate carboxylase in rat liver and Saccharomyces cerevisiae by Immunogold procedures. Arch. Biochem. Biophys. 290, 197–201 (1991).
Article CAS PubMed Google Scholar

Download references

Acknowledgements

This work was financially supported by the NSRF through the Program Management Unit for Human Resources & Institutional Development, Research and Innovation (PMU-B) grant no. B16F640120 and the Center of Excellence for Innovation in Chemistry (PERCH-CIC), Ministry of Higher Education, Science, Research and Innovation, and Mahidol University. Chayanee Laowittawat was supported by the Science Achievement PhD scholarship program.

Author information

Authors and Affiliations

Graduate Program in Molecular Medicine, Faculty of Science, Mahidol University, Rama VI Rd, Bangkok, 10400, Thailand
Chayanee Laowittawat & Sarawut Jitrapakdee
Department of Chemistry, Center of Excellence for Innovation in Chemistry (PERCH-CIC), Faculty of Science, Mahidol University, Rama VI Rd, Bangkok, 10400, Thailand
Natthapat Sawektreeratana, Vichai Reutrakul & Chutima Kuhakarn
Department of Biochemistry, Faculty of Science, Mahidol University, Rama VI Rd, Bangkok, 10400, Thailand
Kanlaya Katewongsa, Pattaree Payomhom & Sarawut Jitrapakdee
Division of Chemistry, Faculty of Science and Technology, Rajabhat Rajanagarindra University, Chachoengsao, 24000, Thailand
Sakchai Hongthong

Authors

Chayanee Laowittawat
View author publications
Search author on:PubMed Google Scholar
Natthapat Sawektreeratana
View author publications
Search author on:PubMed Google Scholar
Kanlaya Katewongsa
View author publications
Search author on:PubMed Google Scholar
Pattaree Payomhom
View author publications
Search author on:PubMed Google Scholar
Sakchai Hongthong
View author publications
Search author on:PubMed Google Scholar
Vichai Reutrakul
View author publications
Search author on:PubMed Google Scholar
Chutima Kuhakarn
View author publications
Search author on:PubMed Google Scholar
Sarawut Jitrapakdee
View author publications
Search author on:PubMed Google Scholar

Contributions

C.L. wrote the original draft of the manuscript. S.J., C.K., V.R. reviewed and edited the manuscript. C.L., N.S., P.P. conducted the experiments. N.S., and S.H. provided some resources and methodologies. K.K., C.K. and S.J. supervised the experiments. S.J. an C.K. conceptualized the project. All authors have seen the final manuscript.

Corresponding authors

Correspondence to Chutima Kuhakarn or Sarawut Jitrapakdee.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary Information

Below is the link to the electronic supplementary material.

Supplementary Material 1

Supplementary Material 2

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if you modified the licensed material. You do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by-nc-nd/4.0/.

Reprints and permissions

About this article

Cite this article

Laowittawat, C., Sawektreeratana, N., Katewongsa, K. et al. The cyclic peptide mallotumide A inhibits colon and breast cancer cell growth and motility by targeting cellular respiration and lipogenesis. Sci Rep 15, 40774 (2025). https://doi.org/10.1038/s41598-025-24547-x

Download citation

Received: 04 February 2025
Accepted: 14 October 2025
Published: 19 November 2025
Version of record: 19 November 2025
DOI: https://doi.org/10.1038/s41598-025-24547-x