Next Article in Journal
A Robust Fermentation Process for Natural Chocolate-like Flavor Production with Mycetinis scorodonius
Next Article in Special Issue
New Derivatives of 5-Substituted Uracils: Potential Agents with a Wide Spectrum of Biological Activity
Previous Article in Journal
Components of Banisteriopsis caapi, a Plant Used in the Preparation of the Psychoactive Ayahuasca, Induce Anti-Inflammatory Effects in Microglial Cells
Previous Article in Special Issue
Recent Advances in Divergent Synthetic Strategies for Indole-Based Natural Product Libraries
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Molecules
Volume 27
Issue 8
10.3390/molecules27082501
molecules-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Anticancer Activity of Natural and Semi-Synthetic Drimane and Coloratane Sesquiterpenoids

by
Lorenz Beckmann
1,
Uta Sandy Tretbar
2,
Reni Kitte
2 and
Maik Tretbar
1,*
1
Medical Faculty, Institute for Drug Discovery, Leipzig University, Bruederstr. 34, 04103 Leipzig, Germany
2
Department of GMP Process Development/ATMP Design, Fraunhofer Institute for Cell Therapy and Immunology IZI, Perlickstr. 1, 04103 Leipzig, Germany
*
Author to whom correspondence should be addressed.
Molecules 2022, 27(8), 2501; https://doi.org/10.3390/molecules27082501
Submission received: 29 March 2022 / Revised: 9 April 2022 / Accepted: 11 April 2022 / Published: 13 April 2022
(This article belongs to the Special Issue Bioactive Compounds: Design, Synthesis and Biological Evaluation)
Browse Figures
Review Reports Versions Notes

Abstract

:
Drimane and coloratane sesquiterpenoids are present in several plants, microorganisms, and marine life. Because of their cytotoxic activity, these sesquiterpenoids have received increasing attention as a source for new anticancer drugs and pharmacophores. Natural drimanes and coloratanes, as well as their semi-synthetic derivatives, showed promising results against cancer cell lines with in vitro activities in the low micro- and nanomolar range. Despite their high potential as novel anticancer agents, the mode of action and structure–activity relationships of drimanes and coloratanes have not been completely enlightened nor systematically reviewed. Our review aims to give an overview of known structures and derivatizations of this class of sesquiterpenoids, as well as their activity against cancer cells and potential modes-of-action. The cytotoxic activities of about 40 natural and 25 semi-synthetic drimanes and coloratanes are discussed. In addition to that, we give a summary about the clinical significance of drimane and coloratane sesquiterpenoids.

1. Introduction

According to the World Health Organization, cancer is the most common cause of death in the 21st century and one of the world’s major health challenges [1,2,3]. Particularly, the growing world population and increasing life expectancy will lead to a constant increase in cancer cases per year. Therefore, the development of effective anticancer treatments is indispensable. Cancer treatment often includes the application of chemotherapeutic drugs, which have become an essential part of cancer drug therapy [4]. Most chemotherapeutic agents are cytotoxic compounds targeting fast-dividing cells by impairing processes of cell division. In general, cytotoxic anticancer drugs include alkylating agents, platinums, anti-metabolites, topoisomerase I and II inhibitors, tubulin-binding drugs, DNA intercalators, and DNA cleaving agents [5,6]. Despite the broad clinical use of chemotherapeutics, there are several drawbacks such as severe side effects due to the lack of selectivity or development of drug resistance [7,8]. Consequently, the investigation of improved anticancer drugs represents a continuous task in medicinal chemistry.
Natural products have proven to be a versatile source of new drugs and pharmacophores in recent decades [9,10,11]. For example, approved chemotherapeutics—such as paclitaxel, vinblastine, or belotecan—are natural products or contain natural product pharmacophores. Despite the long history of natural products as anticancer drugs, their potential has not been exhausted so far. Among natural products, sesquiterpenoids are a prominent group, which has been shown to exhibit cytotoxic effects against cancer cells [12]. Some sesquiterpenoids—such as artemisinin, thapsigargin, or mipsagargin (G-202)—have already entered clinical trials as novel chemotherapeutic agents, showing that this class of natural products holds great potential in anticancer therapy [13]. In nature, sesquiterpenoids are formed in higher plants and microorganisms often to act as antifeedants, deterrents, or attractants [14]. They are 15-carbon compounds derived from the assembly of three isoprenoid units and can be subdivided into acyclic, monocyclic, bicyclic, and tricyclic derivatives. Some examples of cytotoxic sesquiterpenoids are depicted in Figure 1. In this review, we will focus on bicyclic drimane and coloratane sesquiterpenoids.
The general structure of drimanes is derived from drimenol (4), which was first isolated in 1948 from the stem bark of Drimys winterii (Figure 2) [15]. Drimane sesquiterpenoids are characterized by a trans-decalin core structure and common methyl groups at the C4 and C10 positions [16]. Natural drimanes are often substituted by different oxygen-containing groups—such as aldehyde, hydroxy, acetate, and lactone functionalities—which mainly occur at the C6, C9, C11, and C12 positions [16,17,18,19,20,21]. Rearranged drimanes—also known as coloratanes—are closely related compounds characterized by the 1,2 rearrangement of a C4 methyl group, leading to a terminal double bond at the C4 position.
Sources rich in naturally occurring drimanes and coloratanes are liverworts, ferns and higher plants including Canellaceae, Polygonaceae, and Winteraceae species [22,23]. Due to the pharmacological effects of drimane and coloratanes, some of these plants are commonly used in traditional medicine [24,25,26,27]. Other sources of drimanes and coloratanes are fungi [28,29,30] and marine sponges [31,32,33]. Drimane and coloratane sesquiterpenoids have been shown to exhibit antibacterial [34,35,36], antifungal [37,38], antifeedant [39,40,41], and antimalarial [42,43,44] activity.
This review focuses on the cytotoxic properties of drimane and coloratane sesquiterpenoids against cancer cells. We give an overview of the anticancer activity of natural and semi-synthetic drimanes and coloratanes as well their modes of action. In addition to that, the clinical significance of this class of sesquiterpenoids will be summarized.

2. Anticancer Properties

2.1. Natural Drimanes and Coloratanes

Natural sources provide structural diverse and bioactive drimane and coloratane sesquiterpenoids. In this subsection, cytotoxic drimanes and coloratanes from natural sources are described and some structure–activity relationships are discussed. All values that are relevant for the discussion of the cytotoxic activity are summarized in Table 1 at the end of this subsection.

2.1.1. Plant Sources

In 1980, Mahmoud et al. showed that natural drimanes from Cinnamodendron dinisii—cinnamodial (7), capsicodendrin (8), and cinnamosmolide (9)—exhibit cytotoxic activities against P388 leukemia cells and cells of nasopharynx cancer (Eagle’s 9KB-cells, Figure 3) [45]. It is supposed that the cytotoxic activities of 7 and 8 are similar due to the tendency of the acetal group of 8 to hydrolyze under aqueous conditions to yield 7. Interestingly, no cytotoxicity was observed for drimanes lacking an α,β-unsaturated double bond at the C7,8 position such as 10. Karmahapatra et al. also obtained cinnamodial (7) and capsicodendrin (8) from the species Cinnamosma fragrans and tested their cytotoxic and antiproliferative activity against HL-60 and K562 leukemia cells [46]. Cinnamodial (7) was less cytotoxic than capsicodendrin (8). Despite that, 7 is supposed to be the active species because it was shown that 8 is converted to 7 in DMSO. It is suggested that 8 is a prodrug of 7 and the in situ formation of 7 inside the cell increases its cytotoxic activity.
Nomoto et al. isolated drimane lactones and lactams from Cinnamosma fragrans and evaluated their cytotoxicity against A549 carcinoma cells (Figure 4) [47]. Lactone 11 was more potent than 12 and 13, suggesting that certain hydrophilic substituents could lead to an increase in cytotoxic activity due to improved bioavailability.
Fratoni et al. purified drimanes from the stem bark of Drimys brasiliensis and investigated their cytotoxic potential against leukemia cell lines K562 and Nalm-6 (Figure 5) [48]. The drimanes 14–19 and 2 were active against both cancer cell lines. Especially, the cinnamoyl derivatives 18 and 19 exhibited increased cytotoxic activities. Compound 18 was 10- to 20-fold more active than polygodial (2), showing an increase in cytotoxicity by the conjugated π-substituent. As previously reported, drimanes lacking an α,β-unsaturated double bond at the C7,8 position lead to no or a strongly decreased cytotoxic activity. In further experiments, Fratoni et al. tested derivatives 16, 18, and 19 against 16 human cancer cell lines [49]. Compound 18 was the most active compound against most of the cancer cell lines. For seven cancer cell lines, the IC50 values of 18 are located within 1–2 µM. Interestingly, there are a few exceptions where 19 was more active than 18, suggesting a cell-type dependent susceptibility.
Allouche et al. isolated cinnamoyl derivatives 17, 20–22, and polygodial (2) from the bark and leaves of Zygogynum pancheri and Zygogynum acsmithii and tested their cytotoxic activity against KB adenocarcinoma, HL-60 leukemia, and HCT116 colon cancer cell lines [50]. The cinnamoyl derivatives 17, 20–22 showed increased cytotoxicity compared to polygodial (2) as reported by Fratoni et al. [48].
The glycosylated drimane 23 was isolated by Dong et al. from seeds of Antairis toxicaria (Figure 6) and exhibited a moderate cytotoxic activity towards K562 leukemia and SMMC-7721 HeLa cell lines [51].
Xu et al. isolated several drimanes from the stem bark of Warburgia ugandensis and evaluated their cytotoxicity towards nasopharyngeal cancer cells (KB cells) (Figure 7) [52]. Only the dialdehyde drimanes polygodial (2), warburganal (24), and mukaadial (25) showed a strong cytotoxic activity. Compound 24 was more active than 2 suggesting that the C9 hydroxyl group leads to an increase in cytotoxicity. On the other side, 25 is less cytotoxic than 2 showing that a C6 hydroxylation is disfavored or there is a critical number of hydrophilic groups, leading to a loss of anticancer activity by reduced membrane mobility.
Kitte and Tretbar et al. obtained the sesquiterpenoids polygodial (2), warburganal (24), and muzigadial (26) from the stem bark of Warburgia ugandensis and tested their cytotoxic activity against leukemia (MV4-11 and THP-1), melanoma (Sk-Mel29), and glioblastoma (LN-229) cell lines (Figure 7) [53]. Muzigadial (26) exhibited three-to-five-fold higher activities than polygodial (2). The activity of warburganal (24) was also increased compared to polygodial (2) as previously observed by Xu et al. [52]. The results by Kitte and Tretbar et al. suggest that a terminal double bond at the C4 position of muzigadial (26) increases its cytotoxicity.

2.1.2. Marine Sources

Sakio et al. obtained drimane lactones and acetals 27–33 from Dendrodoris carbunculosa (Figure 8) [54]. Drimanes 27–33 exhibited cytotoxic effects against the lymphoma cell line P388 and adriamycin-, and vincristin-resistant (ADR, VDR) P388 cells with IC50 values of 4–17 µg/mL. Lactone 33 showed increased cytotoxicity compared to 27–32. Simultaneous incubation of P388 cells with drimanes 27–33 and adriamycin or vincristine led to no reversal effects.
Drimane acetals 34–36 were isolated from Perenniporia maackiae by J. Kwon et al. and tested against ACHN renal, HCT-15 colon, MDA-MB-231 breast, NCI-H23 lung, NUGC-3 stomach, and PC-3 prostate cancer cell lines (Figure 9) [55]. Compound 35 was shown to be the most active compound. In general, Kwon et al. observed that only drimane acetals with an (S)-configuration or a keto group at the C6 position proved to be cytotoxic during their investigations.

2.1.3. Fungal Sources

Liu et al. isolated polyene-substituted drimanes 37–39 from the fungus Aspergillus ustus and investigated their impact on a murine lymphoma cell line (L5178Y) on PC-12 and HeLa cells (Figure 10) [56]. Compound 37 was shown to be cytotoxic towards all cancer cell lines, whereas 38 and 39 were less active than 37. In general, L5178 cells were more susceptible towards 37–39 than PC-12 and HeLa cells.
Lu et al. also obtained the polyene-substituted drimanes 37 and 38 and an additional compound 40 from Aspergillus ustus and investigated their cytotoxicity against A549 carcinoma and HL-60 leukemia cell lines (Figure 10) [57]. The highest activities were observed for 37 and 40. Surprisingly, 37 exhibited no cytotoxicity against A549 cells and 40 no activity against HL-60 cells. For 38 only a minor impact against A549- and HL-60 cells was observed. The results by Liu et al. and Lu et al. suggest that hydrophilic and electron-deficient groups at the terminal position of the conjugated π-system (Figure 10, green circle) are beneficial for an increase in cytotoxic activity [56,57].
Liu et al. isolated dihydroxylated polyene-substituted drimane lactones 41–44 from Aspergillus flavus (Figure 11) [58]. All sesquiterpenoids 38–41 displayed IC50 values ranging from 1.4 to 8.3 µM against HeLa, MCF-7 breast cancer, MGC-803, and A549 cancer cells, of which 41 exhibited the strongest cytotoxicity.
W. Feng et al. used Fomes officinalis as a source of drimane sesquiterpenoids (Figure 12) [59]. Only drimane 45 exhibited a moderate cytotoxic activity against HL-60 leukemia, Bel-7402 hepatoma, and KB nasopharyngeal cells.
S. Ngokpol et al. isolated drimane sesquiterpene-conjugated amino acids 46–48 from Talaromyces minioluteus and evaluated their cytotoxicity against the human hepatocellular carcinoma cell line HepG2 (Figure 13) [60]. Compounds 47 and 48 are four-fold more active than 46.
All cytotoxic activities of drimane and coloratane sesquiterpenoids from this subsection are concluded in the following table (Table 1). The compounds are classified by their source and the anticancer activity of each compound refers to certain cell lines.
Table
Table 1. Anticancer activity of natural drimane and coloratane sesquiterpenoids 1.
Table 1. Anticancer activity of natural drimane and coloratane sesquiterpenoids 1.
SourceSpeciesComp.Anticancer Activity 2 Value
(Unit) 3		Cell LinesRef.
PlantsA. toxicaria236.60 a, 7.20 bIC50
(μg/mL)		a: K562,
b: SMMC-7721		[51]
C. dinisii72.2EC50
(μg/mL)		P388
(9: KB)		[45]
82.9
91.2
C. fragrans70.48 a, 0.21 bEC50
(µM)		a: HL-60,
b: K562		[46]
80.20 a, 0.09 b
C. fragrans1144.6IC50
(µM)		A549[47]
1262.0
1369.6
D. brasiliens278.20 a, 83.85 bIC50
(µM)		a: Nalm6,
b: K562		[48] 4
1468.03 a, 16.24 b
1578.67 a, 16.66 b
1623.0 a, 26.10 b
188.18 a, 3.56 b
1912.73 a, 12.98 b
W. ugandensis21.0IC50
(µM)		KB[52]
240.3
255.3
W. ugandensis26.91 a, 2.22 b, 7.81 c, 22.40 dIC50
(µM)		a: MV4-11,
b: THP-1,
c: Sk-Mel29,
d: LN-229		[53]
243.20 a, 1.12 b, 4.99 c, 4.11 d
262.62 a, 0.44 b, 2.46 c, 2.74 d
Z. pancheri,
Z. acsmithii		21 a, 1.2 b, 0.7 cIC50
(µM)		a: KB,
b: HL-60,
c: HCT116		[50]
170.5 a, 0.5 b, 0.2 c
200.4 a, 0.3 b, 0.1 c
210.4 a, 0.3 b, 0.1 c
220.4 a, 0.2 b, 0.1 c
Marine lifeD. carbunculosa2710.5IC50 (µg/mL)P388[54]
2817
2911.5
3015
3110.8
3210
333.2
P. maackiae342.0 a, 1.8 b, 2.3 c, 2.1 d, 2.4 e, 6.0 fGI50
(µM)		a: ACHN,
b: HCT-15, c: MDA-MB-231,
d: NCI-H23,
e: NUGC-3,
f: PC-3		[55]
351.2 a, 1.6 b, 1.5 c, 1.4 d, 1.9 e, 2.0 f
363.6 a, 5.4 b, 4.0 c, 4.3 d, 2.3 e, 4.6 f
FungiA. flavus413.6 a, 1.4 b, 6.8 c, 5.0 dIC50
(µM)		a: HeLa,
b: MCF-7,
c: MGC-803,
d: A549		[58]
422.9 a, 5.7 b, 1.9 c, 2.5 d
438.3 a, 3.1 b, 7.1 c, 6.3 d
444.2 a, 6.6 b, 2.3 c, 7.5 d
A. ustus370.6 a, 7.2 b, 5.9 cEC50
(μg/mL)		a: L5178Y,
b: PC-12,
c: HeLa		[56]
381.9 a, >10 b, 7.5 c
395.3 a, >10 b, >10 c
A. ustus37>100 a, 9.0 bIC50
(μM)		a: A549,
b: HL-60		[57]
3810.5 a, >100 b
4030.0 a, 20.6 b
F. officinalis4551.2 a, 88.7 b, 146.0 cIC50
(µM)		a: HL-60,
b: Bel-7402,
c: KB		[59]
T. minioluteus46193.3IC50
(µM)		HepG2[60]
4750.6
4857.0
1 The comparison of anticancer activities is only possible within the same publication because of varying cell viability assays. 2 The anticancer activity refers to a specific cell line which is indicated by superscripts (a–f) for each value within one species. 3 IC50 is the drug concentration causing 50% inhibition of the desired activity and is primarily applied for a specific target. For whole-cell assays, the use of GI50 and ED50 (or EC50) is recommended. GI50 is the concentration for 50% of maximal cell proliferation, whereas ED50 (or EC50) is the dose/concentration causing 50% of a maximum effect for any measured biological activity [61]. 4 See also [49] for more results.

2.2. Semi-Synthetic Drimane Derivatives

Synthetic modification of natural products provides a valuable strategy for increasing pharmacological properties [9,10,11]. For drimane modification, polygodial (2) turned out to be a suitable target because it can be isolated in sufficient quantities from different sources and possesses a less complex structure with reactive aldehyde groups [62,63]. Montenegro et al. isolated polygodial (2), its epimer 9-epipolygodial (49), and drimenol (4) from Drimys winteri [64]. Based on polygodial (2), three semi-synthetic drimanes were obtained by acetalization and reduction. Modification of drimenol (4) yielded two cytotoxic drimane derivatives. Some examples are given in Figure 14.
All derivatives were evaluated against human prostate cell lines DU-145 and PC-3 as well as the breast cancer cell line MCF-7. An increased cytotoxicity was observed for polygodial (2) and the semi-synthetic derivatives 50, 52, and 53. These data allow a more systematic insight into the structure–activity relationships of drimanes. The electrophilic position at the C7 position and the configuration of the C9 position are critical for cytotoxic activity. Reduction of both aldehyde groups of polygodial (2) leads to complete inactivation of both epimers of 51. Interestingly, acetalization of the C11 aldehyde group of 2 as to 50 has no impact on cytotoxic activity. However, it can be assumed that the acetal group of 50 is converted to 2 under aqueous conditions. Furthermore, 9-epipolygodial (49) showed no cytotoxicity suggesting a critical role in the configuration of the C9 position. Transforming the C7 position of drimenol (4) to an electrophilic keto group (52 and 53) leads to cytotoxic activity compared to 4.
Dasari et al. modified polygodial (2) to increase its antiproliferative activity against A549-, Sk-Mel28, MCF-7-, U373-, and Hs683 cancer cells [65]. 9-epipolygodial (49) was obtained by an epimerization reaction of polygodial (2) (Scheme 1). Typical GI50 values of 49 were at 2–6 μM and are 10-to-20-fold smaller than GI50 values of polygodial (2) Thus, contradictory to Montenegro et al. (Table 2), 9-epipolygodial (49) displayed a high antiproliferative activity [64]. Certainly, the impact of anticancer agents is cell line dependent-but even for the same cancer cell line MCF-7, both groups obtained contrary results [64,65]. The two groups performed different cell viability assays and control culture treatments to measure the cytostatic effects of 49 which might lead to different results. Dasari et al. measured the cell viability as sextuplicate in only one experiment [65]. On the other side, Montenegro et al. performed three independent experiments in triplicates [64]. Discrepancies in the data could therefore be due to the lack of biological replicates. Despite that, 9-epipolygodial (49) also showed antiproliferative effects against the multi-resistant uterus cancer cell lines MES-SA and MES-SA/Dx5, which were measured in two independent experiments. The effects are comparable to the chemotherapeutics paclitaxel and vinblastine.
Based on 9-epipolygodial (49), Wittig derivative 54 was synthesized leading to reduced cytotoxicity compared to 49 (Scheme 2) [65]. By selective reduction of the double bond of polygodial (2), derivative 55 was obtained that is in equilibrium with 56. The reduced derivatives 55/56 still showed some antiproliferative activity suggesting that the presence of a dialdehyde structure causes a part of the cytotoxicity. Indeed, Dasari et al. observed a high reactivity of the dialdehyde compounds with primary amines forming unstable pyrrole adducts [65]. The authors suggested that dialdehyde drimanes might react with the ε-amine group of lysine residues of proteins, leading to the alteration of their functions. However, no protein drimane adducts were identified so far.
Dasari et al. proposed that the reversible formation of bis-acetals might lead to a controlled release of reactive dialdehydes inside the cell and an increased cytotoxicity [65]. Therefore, different bis-acetals were formed from polygodial (2) (Scheme 3). Bis-acetals 57a, b, 62a, b, 63a, b and 58 showed comparable or even enhanced antiproliferative effects compared to polygodial (2). A possible reason is also the enhanced membrane mobility of the bis-acetals because of the less polar structure compared to polygodial (2).
In further works, Dasari et al. focused on the Wittig derivatization of the C12 aldehyde group of polygodial (2) (Scheme 4) [66]. The reactivity of the C12 position is increased compared to the C11 position, enabling the selective modification of 2. Scheme 4 represents the C12 Wittig derivatives synthesized. All derivatives were tested against A549-, Sk-Mel28, MCF-7-, U373-, and Hs683 cancer cells. Best results were obtained for derivative 64 against MCF-7 cancer (GI50 = 7.0 µM) being 10-fold more active than polygodial (2). In general, GI50 values of all Wittig derivatives 64–71 were about 21–42 μM for A549-, Sk-Mel-28-, U373-, and Hs683 cancer cells. Additionally, it was demonstrated that 64 forms an isolable pyrrole product with benzylamine. No Wittig derivatives of 9-epipolygodial (49) were synthesized.
Antiproliferative effects of 9-epipolygodial (49) and the Wittig derivative 64 were investigated more intensively [67,68]. In addition to that, the derivative 73 was obtained by a Horner–Wadsworth–Emmons (HWE) reaction with 72 (Scheme 5). Because of the basic reaction conditions, 73 is formed as a C9 epimer. HWE derivative 73 showed robust inhibition of LNCaP prostate cancer cell growth (IC50 = 5 µM) but does not exhibit significant cytotoxicity towards PC-3, DU-145 prostate cancer cells, and normal prostate cells (RWPE-1). Compound 73 is also antiproliferative towards oral squamous cell carcinoma cell lines Cal27 and HSC3 and HeLa cells. GI50 values were observed between 2 and 8 μM. For H460 lung cancer cells, the antiproliferative effect of 73 is smaller with a GI50 value of 70 μM.
Recently, Maslivetc et al. synthesized ethylene glycol-linked dimers of polygodial (2) (Figure 15) [69]. Antiproliferative effects of 74a and 74b were increased compared to polygodial (2). Against A549-, SkMel-28-, U373-, MCF-7-, Hs683-, and B16F10 cell lines GI50 values between 4 and 14 μM were obtained for 74a. A longer chain length of the polyethylene linker leads to a decreased cytotoxicity. Consequently, derivative 74c shows higher GI50 values than polygodial (2). The increased cytotoxicity of 74a and 74b is explained by the ability to crosslink antitumor targets by pyrrole formation with lysine residues. However, no crosslink adducts were identified by Maslivetc et al. [69].
In general, there are certain modifications of polygodial (2) leading to a loss or an increase in cytotoxicity (Figure 16). It was observed that the reduction of both aldehyde groups extinguishes cytotoxic activity [64,65]. On the other side, the selective reduction of the C7–C8 double bond only diminishes the cytotoxicity without a complete loss [65]. Current results suggest that the presence of an α,β-unsaturated system at the C7,8 position is a key feature for high cytotoxic activity. The conversion of the C9 aldehyde group by Wittig- and HWE reactions maintains a π-conjugation with an electron-deficient carbonyl function and leads to an increase in cytotoxicity [65,67,68]. It was also shown, that the C9 configuration influences the anticancer activity but contradictory results are currently present [64,65]. Additionally, the bis-acetalization and dimerization of polygodial (2) are strategies to increase its cytotoxic effects [65,69].

3. Mode-of-Action–Specific Effects or ‘Just’ Detergents?

Despite the known cytotoxic effect of natural and semi-synthetic sesquiterpenoids, their mode of action is not completely enlightened and different targets are discussed in the literature. The following part aims to give a comparative overview of accomplished results regarding the mode of action of drimane and coloratane sesquiterpenoids. In Figure 17, different modes-of-action are summarized and will be discussed in further detail during this section. It should be mentioned that a compound can act by more than one mechanism, which can change even within the same compound class depending on its structural details. Especially, data derived from experiments using plant extracts must be interpreted carefully because observed biological activity cannot be exclusively depicted to drimane and coloratane sesquiterpenoids.

3.1. Detergent Effects on Mitochondrial Membrane

Quite early, polygodial (2) was described to inhibit the mitochondrial ATP synthesis in fungus and eukaryotic cells [70,71,72]. Because of its amphiphilic nature, polygodial (2) is supposed to act as a non-ionic detergent on the mitochondrial membrane (MM), leading to the distortion of the mitochondrial membrane potential (MMP). The loss of the MMP is known to be a common signal for cells to undergo apoptosis [73]. In this context, Montenegro et al. observed higher mitochondrial membrane permeabilities in cancer cells after treatment with polygodial (2) and two other drimanes [64]. Additionally, the activity of caspase 3—a main executor of apoptosis—was increased. Despite that, these results should be interpreted carefully because the permeabilization of the MM and loss of MMP are part of several pro-apoptotic signaling cascades and are not restricted to detergent effects on the mitochondrial membrane, exclusively [73]. Therefore, it must be determined if the loss of MMP results from the detergent effects of the sesquiterpenoids on the MM or occurs as a downstream event of another apoptotic pathway. Furthermore, the detergent effect on subcellular and cellular membranes should be examined.
The susceptibility of cellular membranes strongly depends on their lipid composition and varies between cell types [74]. Especially, cancer cells show abnormal lipid metabolism resulting in altered membrane compositions [75]. An interesting target for detergents is the lysosome which got increasing attention as an anticancer target for lysosomotropic detergents [76,77,78]. These agents accumulate in the lysosome and lead to the permeabilization of the lysosomal membrane, which is a signal for apoptosis via the lysosomal death pathway [79]. For the lysosomes of chronic lymphocytic and acute myeloid leukemia cells, altered membrane composition and expression profiles were observed, leading to higher susceptibility against detergents [77,80].

3.2. TRPV1 Ion Channel

Polygodial (2), its epimer 49 and warburganal (24) were found to be agonists of the non-selective ion channel TRPV1 (transient receptor potential vanilloid 1), which is expressed on the cell surface of several cancer cell lines as well as on the mitochondrial membrane of different human cells [81,82,83,84,85,86,87,88,89,90,91]. TRPV1 regulates the influx of Ca2+ ions into the cytosol and plays a role in inflammation, proliferation, and pain [92,93]. It is known that Ca2+ signaling is a critical part of the proliferation and apoptosis resistance of cancer cells [94,95]. On the other side, Ca2+ overload can induce apoptosis [96,97]. In the past, different TRPV1 agonists, especially capsaicin, were shown to trigger cell death in TRPV1 expressing cancer cells [98,99,100,101,102,103,104,105]. Based on that, Dasari et al. investigated whether the antitumor effects of different drimanes and derivatives including polygodial (2) and its epimer 49 correlate with TRPV1 activity [65,66]. They showed that only polygodial (2) and some drimanes with minor cytotoxicity act as agonists on the TRPV1 ion channel, but at concentrations much higher than the GI50 values determined. In contrast to previous results, no TRPV1 activity for 9-epipolygodial (49) was observed. 9-epipolygodial (49) shows high antiproliferative effects against several cancer cell lines. In addition to that, no TRPV1 activity was observed for the highly active Wittig derivative 64 and HWE derivative 73. To support their results, Dasari et al. used computer models docking the drimane compounds into the VBS (vanilloid binding site) of TRPV1, where only for polygodial (2) and some inactive compounds energetically relevant docking positions were obtained [65,66]. These results indicate that the anticancer effects of drimanes might be independent of the TRPV1 ion channel.

3.3. TRPA1 Ion Channel

Another potential target of drimanes and coloratanes is the TRPA1 (transient receptor potential ankyrin 1) ion channel. TRPA1 is related to TRPV1 and leads to the influx of Ca2+ ions into the cytosol [106,107]. It is overexpressed in different cancer cell lines and attracts increasing attention in cancer-related processes [108,109]. For several drimanes and coloratane dialdehyde compounds, the activation of TRPA1 was observed [110,111]. TRPA1 is activated by electrophilic pungent electrophiles, such as isothiocyanates, forming reversible covalent bonds to cysteine residues of the ion channel protein [112,113,114]. Especially, cysteine 621 had been suggested to be most critical for electrophile-binding [115]. For polygodial (2), chemical reactions with cysteine residues have been proposed but no defined reaction product could be identified [116,117,118]. Besides cysteine 621, lysine 620 is suggested to be another critical residue for TRPA1 activation. Mathie et al. showed that polygodial (2) reacts rapidly with the amino group of Lys-Nα-Ac while no reaction occurred with Cys-Nα-Ac [111]. In a mutational study, the dialdehydes polygodial (2) and isovelleral retained full activity on a triple cysteine to lysine TRPA1 mutant [110]. These results suggest that TRPA1 activation of drimane dialdehyde compounds is mediated by covalent modification of critical lysine residues within TRPA1. However, the contribution of TRPA1 activation to the anticancer activity of drimane and coloratane sesquiterpenoids needs to be evaluated in greater detail.

3.4. ROS Generation and DNA Damage

DNA damage results in the activation of a variety of downstream pathways, which induce either DNA repair mechanisms or cell cycle arrest and apoptosis [119]. Especially, DNA double-strand breaks (DSBs) are critical events leading to the activation of apoptotic downstream events. Typical sources for DNA damage and DSBs are methylating agents and oxidative stress by reactive oxygen species (ROS). In terms of anticancer treatment, radiation therapy (RT) is closely related to ROS and DNA damage [120]. Due to ionizing radiation (IR), ROS are generated at localized regions in the nucleus leading to DNA lesions and consequent cell death. The generation of ROS is not restricted to IR and can also result from endogenous (e.g., mitochondria) and exogenous sources (e.g., heavy metals, xenobiotics) [121,122,123]. An overload of ROS induces apoptosis and cell cycle arrest in cancer cells indicating the potential of ROS in anticancer therapy [124].
For the measurement of DSBs, the amount of phosphorylated histone 2AX (γH2AX) has proven to be a valuable marker for DSBs [125]. DSBs cause the phosphorylation of neighboring γH2AX which is assumed to be functional for DNA repair by causing the chromatin to be more accessible for DNA repair [126]. However, there is also evidence that γHA2X is required for DNA ladder formation during apoptosis [127]. Another protein involved in DNA repair is poly-(ADP-ribose)-polymerase 1 (PARP1) [128,129]. During apoptosis, PARP1 is cleaved by caspases 3 and 7, leading to the inactivation of PARP1-mediated DNA repair. Therefore, the concentration of c-PARP1 levels is a marker for apoptosis.
For the Wittig derivative 64 and HWE derivative 73, increased concentrations of c-PARP1 and γH2AX were observed after exposure to cancer cells [67,68]. Interestingly, the antiproliferative effects of 64 and 73 can be terminated by the addition of the antioxidant N-acetyl cysteine (NAC), suggesting that ROS play a critical role in the mode of action of drimanes by inducing DNA damage. Increased ROS and γH2AX levels were also observed in HT-29 and HCT116 and A549 cells after incubation with extracts of W. ugandensis [130,131]. In addition to that, extracts of W. ugandensis lead to an increase in c-PARP1 concentrations.
Contradictory to the previous results, Karmahapatra et al. detected no ROS increase in HL-60 leukemia cells after incubation with capsicodendrin (8) [46]. Capsicodendrin (8) forms cinnamodial (7) under aqueous conditions and it was shown that 7 forms covalent bonds with cysteine residues of glutathione—an intracellular thiol-based antioxidant—without altering intracellular ROS concentrations. Despite that, DNA damage still occurred in HL-60 cells after exposure to cinnamodial (7). It is suggested that DNA damage might be caused by the 2-alkenal motif of 7. In a previous study, it was shown that 2-alkenals lead to DNA damage in V79 lung fibroblasts and Caco-2 colorectal carcinoma cells [132].
In general, there are still open questions regarding the generation and contribution of ROS towards the mode-of-action of drimane and coloratane sesquiterpenoids. It needs to be evaluated whether the generation of ROS depends on specific structural details of the sesquiterpenoids and if ROS production is cell-type dependent. Furthermore, the origin of ROS production must be evaluated in further detail.
It is known that ROS can be produced as a response to mitochondrial stress [133]. As mentioned, drimanes and coloratanes might act as detergents on the MM, leading to mitochondrial stress and ROS generation [70,71,72]. Therefore, ROS could be a link between the detergent effects of drimanes and coloratanes on the MM and DNA damage. It is also possible that mitochondrial-induced ROS are generated by Ca2+ influx after activation of TRPV1 or TRPA1. An overload of Ca2+ ions is known to induce ROS release from the mitochondrion. Additionally, it was shown that ROS can trigger Ca2+ release from the endoplasmic reticulum (ER), resulting in a positive feedback loop with accelerated ROS production [134].

3.5. Cell Cycle Arrest

The cell cycle comprises a series of cellular events during the growth and division of cells [135,136]. In general, the cell cycle is divided into different phases (G0, G1-, S-, G2-, and M-phase) that are characterized by the action of specific proteins. Important regulators of the cell cycle are cyclin-dependent kinases (CDKs) that require different cyclins for kinase activity [137].
Zhang et al. observed reduced expression of cyclin D1 and E1 after exposure of A549, HT-29, and HCT 116 cells to the extract of the W. ugandensis [130,131]. Cyclin D1 and E1 play an important role in the progression from the G1 to the S phase [138,139]. The expression of cyclin D1 is tightly regulated by the PI3K/Akt/GSK3β pathway [140]. After incubation of HT-29 cells with W. ugandensis extract, expression levels of PI3K, Akt, p-Akt, and p-GSK3β were changed leading to the suppression of the PI3K/Akt/GSK3β pathway and reduced amounts of cyclin D1 [131]. In addition to that, Zhang et al. measured an increase in P27 in A549 cells [130]. P27 is a negative regulator of CDKs and is ubiquitinylated by S-phase kinase-associated protein 2 (SKP2), reducing its cellular concentration by proteasomal degradation [141,142,143]. Furthermore, SKP2 is negatively regulated by Forkhead-Box-Protein O3 (FOXO3A) [144,145]. An increased expression of FOXO3A was also observed after cell exposure to extracts of W. ugandensis [130]. In future experiments, it would be important to evaluate whether these results are caused by drimane and coloratane sesquiterpenoids or other bioactive compounds from W. ugandensis. In this context, Lohberger et al. have already shown that a dehydrocostus lactone sesquiterpenoid leads to cell cycle arrest in soft tissue sarcoma cells by a decrease in CDK2 levels [146]. Therefore, the impairment of CDK-dependent cell cycle progression could be a more common feature of sesquiterpenoids including also drimanes and coloratanes.

3.6. Inhibition of DNA-Binding by NF-kB and Stat-3

Felix et al. observed reduced expression of survivin—an antiapoptotic protein critical in apoptosis resistance of cancer cells—in Colo-320 cells after the addition of the drimane lactone 75 (Figure 18) [147].
In further experiments, it was shown that 75 inhibits DNA binding of the transcription factors NF-kB and Stat-3 to the survivin promoter. DNA binding of NF-κB and Stat-3 to the CMV promoter was not impaired. Interestingly, structurally related eudasmane sesquiterpenoids and meroterpenoids lead to the attenuation of NF-κB dependent pathways [148,149]. Tang et al. have shown that upstream of the NF-κB activation the phosphorylation of p38 mitogen-activated protein kinases (p38 MAPK) is inhibited by 1,10-seco-eudesmane sesquiterpenoids [148]. In future studies, it should be evaluated if the inhibition of NF-κB contributes to the anticancer activity of drimane and coloratane sesquiterpenoids.

4. Clinical Significance

Around half of the small molecules that have been approved for cancer chemotherapy since 1940 are natural products [150]. All of these compounds were proven to be effective and safe. The efficacy of a compound can only be evaluated if a proper target is identified, which is the main step in drug discovery [151]. Selectivity toward tumor cells preventing off-target effects plays an important role in clinical approval. Although more than 200 compounds derived from natural products are currently in preclinical and clinical investigations [152] only a few of them will be used in clinical routines in the future. The main reason why 95% of small-molecule oncology therapeutics fail in clinical trials is due to a lack of clinical safety and efficacy [153].
Drimane and coloratane sesquiterpenoids were shown to exert promising anticancer activity in vitro. However, no preclinical studies evaluating their in vivo cytotoxic activity nor clinical trials were accomplished until now. In vivo studies are only available for antidiabetic and antimalarial activities of some drimane sesquiterpenoids and plant extracts [44,154]. In these experiments, no unspecific toxicity was observed in rats, indicating a low risk of severe side effects. In addition to that, plant extracts containing drimanes and coloratanes are already used in traditional medicine leading to the conclusion that a clinical application appears feasible [24,25,26,27]. Other sesquiterpenoids, such as sesquiterpene lactones, already made it into clinical trials as chemotherapeutics for their anticancer activity such as artemisinin, showing that sesquiterpenoids are indeed a valuable source for anticancer agents and provide candidates for in vivo studies and clinical trials [13]. We must then ask: why have none of the aforementioned drimanes or coloratanes made it into the clinic yet?
Possible reasons are the lack of detailed knowledge about a specific target or modes-of-action, cancer cell specificity, or insufficient anticancer activities of drimane and coloratane sesquiterpenoids. Until now, IC50 values in the low nanomolar range were only observed for a few drimanes and coloratanes against certain cancer cell lines [46,49,50,52,53]. Therefore, the optimization of the anticancer activity of drimane and coloratane sesquiterpenoids should be addressed in future projects. Especially, new synthetic methods could provide access to novel drimanes and coloratanes with enhanced cytotoxicity and selectivity [155,156,157]. Furthermore, a detailed understanding of the mode-of-action would be beneficial for the rational functionalization of the drimane and coloratane scaffold.

5. Conclusions

In this review, we gave an overview of the cytotoxic activity of drimane and coloratane sesquiterpenoids against cancer cells. Over the past decades, natural drimanes and coloratanes were isolated and their cytotoxic potential was evaluated. Additionally, several semi-synthetic derivatives were obtained from polygodial to increase its cytotoxic activity. It is demonstrated that the drimane and coloratane scaffold holds great potential as a lead structure for anticancer drug development. However, none of the components are used as a therapeutic agent or in a clinical trial so far. Furthermore, several modes-of-action of drimane and coloratane sesquiterpenoids were discussed. Detergent effects on subcellular membranes, DNA damage, and ROS production are commonly observed for drimanes. It was also shown that cancer cell exposure to drimane-containing plant extracts can cause cell cycle arrest. Additionally, dialdehyde drimanes activate the TRPA1 ion channel. The related ion channel TRPV1 was often supposed to be addressed by drimanes, but current data show that TRPV1 only plays a minor role in the cytotoxic action of this class of sesquiterpenoids. However, whether there is a relation between these actions or they are caused independently needs to be further evaluated. Therefore, a detailed analysis of differentially expressed genes using microarrays or next-generation sequencing methods could be key methods to reveal targets or mode-of-action of drimane and coloratane type sesquiterpenoids in anticancer activity. Proper target identification will lay a solid basis for future lead compound discovery and candidate selection for preclinical and clinical development.

Author Contributions

Writing—original draft preparation, L.B.; Writing—review and editing, U.S.T., M.T. and R.K.; Supervision, U.S.T. and M.T.; Project administration, U.S.T. and M.T.; Funding acquisition, M.T. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Not applicable.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. WHO. Available online: https://www.who.int/news-room/fact-sheets/detail/cancer (accessed on 25 March 2022).
  2. Sung, H.; Ferlay, J.; Siegel, R.L.; Laversanne, M.; Soerjomataram, I.; Jemal, A.; Bray, F. Global Cancer Statistics 2020: GLOBOCAN Estimates of Incidence and Mortality Worldwide for 36 Cancers in 185 Countries. CA Cancer J. Clin. 2021, 71, 209–249. [Google Scholar] [CrossRef] [PubMed]
  3. Pillerson, S.; Soto-Perez-de-Cellis, E.; Vignat, J.; Ferlay, J.; Soerjomataram, I.; Bray, F.; Sarfati, D. Estimated global cancer incidence in the oldest adults in 2018 and projections to 2050. Int. J. Cancer. 2021, 148, 601–608. [Google Scholar] [CrossRef] [PubMed]
  4. Chabner, B.A.; Roberts, T.G., Jr. Chemotherapy and the war on cancer. Nat. Rev. 2005, 5, 65–72. [Google Scholar] [CrossRef] [PubMed]
  5. Lind, M.J. Principles of cytotoxic chemotherapy. Medicine 2008, 36, 19–23. [Google Scholar] [CrossRef]
  6. Nussbaumer, S.; Bonnabry, P.; Veuthey, J.-L.; Fleury-Souverain, S. Analysis of anticancer drugs: A review. Talanta 2011, 85, 2265–2289. [Google Scholar] [CrossRef]
  7. Pearce, A.; Haas, M.; Viney, R.; Pearson, S.-A.; Haywood, P.; Brown, C.; Ward, R. Incidence and severity of self-reported chemotherapy side effects in routine care: A prospective cohort study. PLoS ONE 2017, 12, e0184360. [Google Scholar] [CrossRef]
  8. Bukowski, K.; Kciuk, M.; Kontek, R. Mechanisms of Multidrug Resistance in Cancer Chemotherapy. Int. J. Mol. Sci. 2020, 21, 3233. [Google Scholar] [CrossRef]
  9. Cragg, C.M.; Grothaus, P.G.; Newman, D.J. Impact of Natural Products on Developing New Anti-Cancer Agents. Chem. Rev. 2009, 109, 3012–3043. [Google Scholar] [CrossRef]
  10. Lee, K.-H. Discovery and Development of Natural Product-Derived Chemotherapeutic Agents Based on a Medicinal Chemistry Approach. J. Nat. Prod. 2010, 73, 500–516. [Google Scholar] [CrossRef] [Green Version]
  11. Demain, A.L.; Vaishnav, P. Natural products for cancer chemotherapy. Microb. Biotechnol. 2011, 4, 687–699. [Google Scholar] [CrossRef] [Green Version]
  12. Chen, Q.-F.; Liu, Z.-P.; Wang, F.-P. Natural Sesquiterpenoids as Cytotoxic Anticancer Agents. Mini-Rev. Med. Chem. 2011, 11, 1153–1164. [Google Scholar] [CrossRef] [PubMed]
  13. Ghantous, A.; Gali-Muhtasib, H.; Vuorela, H.; Saliba, N.A.; Darwiche, N. What made sesquiterpene lactones reach cancer clinical trials? Drug Discov. Today 2010, 15, 668–678. [Google Scholar] [CrossRef] [PubMed]
  14. Fraga, B.M. Natural sesquiterpenoids. Nat. Prod. Rep. 2013, 30, 1226–1264. [Google Scholar] [CrossRef]
  15. Appel, H.H. Estudio químico de la corteza del árbol Drimys winteri Forst. Scientia 1948, 15, 31–32. [Google Scholar]
  16. Jansen, B.J.M.; de Groot, A. Occurrence, biological activity and synthesis of drimane sesquiterpenoids. Nat. Prod. Rep. 2004, 21, 449–477. [Google Scholar] [CrossRef]
  17. Stork, G.; Burgstahler, A.W. The Stereochemistry of Polyene Cyclization. J. Am. Chem. Soc. 1955, 77, 5068–5077. [Google Scholar] [CrossRef]
  18. Parker, W.; Roberts, J.S. Sesquiterpene Biogenesis. Q. Rev. Chem. Soc. 1967, 21, 331–363. [Google Scholar] [CrossRef]
  19. Dawson, G.W.; Hallahan, D.L.; Mudd, A.; Patel, M.M.; Pickett, J.A.; Wadhams, L.J.; Wallsgrove, R.M. Secondary plant metabolites as targets for genetic modification of crop plants for pest resistance. Pestic. Sci. 1989, 27, 191–201. [Google Scholar] [CrossRef]
  20. Fleck, W.F.; Schlegel, B.; Hoffmann, P.; Ritzau, M.; Heinze, S.; Gräfe, U. Isolation of Isodrimenediol, a Possible Intermediate of Drimane Biosynthesis from Polyporus arcularius. J. Nat. Prod. 1996, 59, 780–781. [Google Scholar] [CrossRef]
  21. Huang, Y.; Hoefgen, S.; Valiante, V. Biosynthesis of Fungal Drimane-Type Sesquiterpene Esters. Angew. Chem. Int. Ed. 2021, 60, 23763–23770. [Google Scholar] [CrossRef]
  22. Ludwiczuk, A.; Gradstein, S.R.; Nagashima, F.; Asakawa, Y. Chemosystematics of Porella (Marchantiophyta, Porellaceae). Nat. Prod. Commun. 2011, 6, 315–321. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  23. Asakawa, Y.; Ludwiczuk, A.; Harinantenaina, L.; Toyota, M.; Nishiki, M.; Bardon, A.; Nii, K. Distribution of Drimane Sesquiterpenoids and Tocopherols in Liverworts, Ferns and Higher Plants: Polygonaceae, Canellaceae and Winteraceae Species. Nat. Prod. Commun. 2012, 7, 685–692. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  24. Choudhary, R.K.; Oh, S.; Lee, J. An ethnomedicinal inventory of knotweeds of Indian Himalaya. J. Med. Plant. Res. 2011, 5, 2095–2103. [Google Scholar] [CrossRef]
  25. Maroyi, A. The genus Warburgia: A review of its traditional uses and pharmacology. Pharm. Biol. 2014, 52, 378–391. [Google Scholar] [CrossRef] [Green Version]
  26. Hug, A.K.K.M.; Jamal, J.A. Ethnobotanical, Phytochemical, Pharmacological, and Toxicological Aspects of Persicaria hydropiper (L.) Delarbre. eCAM 2014, 2014, 782830. [Google Scholar] [CrossRef] [Green Version]
  27. Nasir, A.; Khali, A.A.K.; Bhatti, M.Z.; Rehman, A.U.; Li, J.; Parveen, Z. Review on Pharmacological and Phytochemical Prospects of Traditional Medicinal Plant: Persicaria hydropiper (Smartweed). Curr. Top. Med. Chem. 2021, 21, 1027–1036. [Google Scholar] [CrossRef]
  28. Zhou, H.; Zhu, T.; Cai, S.; Gu, Q.; Li, D. Drimane Sesquiterpenoids from the Mangrove-Derived Fungus Aspergillus ustus. Chem. Pharm. Bull. 2011, 59, 762–766. [Google Scholar] [CrossRef] [Green Version]
  29. Lacey, H.J.; Gilchrist, C.L.M.; Crombie, A.; Kalaitzis, J.A.; Vuong, D.; Rutledge, P.J.; Turner, P.; Pitt, J.I.; Lacey, E.; Chooi, Y.-H.; et al. Nanangenines: Drimane sesquiterpenoids as the dominant metabolite cohort of a novel Australian fungus, Aspergillus nanangensis. Beilstein J. Org. Chem. 2019, 15, 2631–2643. [Google Scholar] [CrossRef] [Green Version]
  30. Ma, M.; Ge, H.; Yi, W.; Wu, B.; Zhang, Z. Bioactive drimane sesquiterpenoids and isocoumarins from the marine-derived fungus Penicillium minioluteum ZZ1657. Tetrahedron Lett. 2020, 61, 151504. [Google Scholar] [CrossRef]
  31. Montagnac, A.; Martin, M.-Z.; Debitus, C.; Païs, M. Drimane Sesquiterpenes from the Sponge Dysideafusca. J. Nat. Prod. 1996, 59, 866–868. [Google Scholar] [CrossRef]
  32. Paul, V.J.; Seo, Y.; Cho, K.W.; Rho, J.R.; Shin, J.; Bergquist, P.R. Sesquiterpenoids of the Drimane Class from a Sponge of the Genus Dysidea. J. Nat. Prod. 1997, 60, 1115–1120. [Google Scholar] [CrossRef] [PubMed]
  33. Schmitz, F.J.; Lakshmi, V.; Powell, D.R.; Van der Helm, D. Arenarol and arenarone: Sesquiterpenoids with rearranged drimane skeletons from the marine sponge Dysidea arenaria. J. Org. Chem. 1984, 49, 241–244. [Google Scholar] [CrossRef]
  34. Rabe, T.; van Staden, J. Isolation of an antibacterial sesquiterpenoid from Warburgia salutaris. J. Ethnopharmacol. 2000, 73, 171–174. [Google Scholar] [CrossRef]
  35. Kubo, I.; Fujita, K.; Lee, S.H.; Ha, T.J. Antibacterial activity of polygodial. Phytother. Res. 2005, 19, 1013–1017. [Google Scholar] [CrossRef] [PubMed]
  36. Neuhaus, G.F.; Loesgen, S. Antibacterial Drimane Sesquiterpenes from Aspergillus ustus. J. Nat. Prod. 2021, 84, 37–45. [Google Scholar] [CrossRef]
  37. Kubo, I.; Himejima, M. Potentiation of antifungal activity of sesquiterpene dialdehydes against Candida albicans and two other fungi. Experientia 1992, 48, 1162–1164. [Google Scholar] [CrossRef]
  38. Edouarzin, E.; Horn, C.; Paudyal, A.; Zhang, C.; Lu, J.; Tong, Z.; Giaever, G.; Nislow, C.; Veerapandian, R.; Hua, D.H.; et al. Broad-spectrum antifungal activities and mechanism of drimane sesquiterpenoids. Microb. Cell 2020, 7, 146–159. [Google Scholar] [CrossRef]
  39. Montenegro, I.; Pino, L.; Werner, E.; Madrid, A.; Espinoza, L.; Moreno, L.; Villena, J.; Cuellar, M. Comparative Study on the Larvicidal Activity of Drimane Sesquiterpenes and Nordrimane Compounds against Drosophila melanogaster til-til. Molecules 2013, 18, 4192–4208. [Google Scholar] [CrossRef]
  40. Moreno-Osorio, L.; Cortés, M.; Armstrong, V.; Bailén, M.; González-Coloma, A. Antifeedant Activity of Some Polygodial Derivatives. Z. Naturforsch. 2008, 63, 215–220. [Google Scholar] [CrossRef]
  41. Inocente, E.A.; Nguyen, B.; Manwill, P.K.; Benatrehina, A.; Kweka, E.; Wu, S.; Cheng, X.; Rakotondraibe, L.H.; Piermarini, P.M. Insecticidal and Antifeedant Activities of Malagasy Medicinal Plant (Cinnamosma sp.) Extracts and Drimane-Type Sesquiterpenes against Aedes aegypti Mosquitoes. Insects 2019, 10, 373. [Google Scholar] [CrossRef] [Green Version]
  42. Claudino, V.D.; da Silva, K.C.; Filho, V.C.; Yunes, R.A.; Monache, F.D.; Giménez, A.; Salamanca, E.; Gutierrez-Yapu, D.; Malheiros, A. Drimanes from Drimys brasiliensis with leishmanicidal and antimalarial activity. Mem. Inst. Oswaldo Cruz 2013, 108, 140–144. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  43. Corrêa, D.S.; Tempone, A.G.; Reimão, J.Q.; Taniwaki, N.N.; Romoff, P.; Fávero, O.A.; Sartorelli, P.; Mecchi, M.C.; Lago, J.H.G. Anti-leishmanial and anti-trypanosomal potential of polygodial isolated from stem barks of Drimys brasiliensis Miers (Winteraceae). Parasitol. Res. 2011, 109, 231–236. [Google Scholar] [CrossRef] [PubMed]
  44. Nyaba, Z.N.; Murambiwa, P.; Opoku, A.R.; Mukaratirwa, S.; Shode, F.O.; Simelane, M.B.C. Isolation, characterization, and biological evaluation of a potent anti-malarial drimane sesquiterpene from Warburgia salutaris stem bark. Malar. J. 2018, 17, 296. [Google Scholar] [CrossRef]
  45. Mahmoud, I.I.; Kinghorn, A.D.; Cordell, G.A.; Farnsworth, N.R. Potential anticancer agents. XVI. Isolation of bicyclofarnesane sesquiterpenoids from Capsicodendron dinisii. J. Nat. Prod. 1980, 43, 365–371. [Google Scholar] [CrossRef] [PubMed]
  46. Karmahapatra, S.; Kientz, C.; Shetty, S.; Yalowich, J.C.; Pakotondraibe, L.H. Capsicodendrin from Cinnamosma fragrans Exhibits Antiproliferative and Cytotoxic Activity in Human Leukemia Cells: Modulation by Glutathione. J. Nat. Prod. 2018, 81, 625–629. [Google Scholar] [CrossRef] [PubMed]
  47. Nomoto, Y.; Harinantenaina, L.; Sugimoto, S.; Matsunami, K.; Otsuka, H. 3,4-seco-24-homo-28-nor-Cycloartane and drimane-type sesquiterpenes and their lactams from the EtOAc-soluble fraction of a leaf extract of Cinnamosma fragrans and their biological activity. J. Nat. Med. 2014, 68, 513–520. [Google Scholar] [CrossRef]
  48. Fratoni, E.; Claudino, V.D.; Yunes, R.A.; Franchi, G.C., Jr.; Nowill, A.E.; Filho, V.C.; Monache, F.D.; Malheiros, A. Further drimane sesquiterpenes from Drimys brasiliensis stem barks with cytotoxic potential. Naunyn-Schmiedeberg’s Arch. Pharmacol. 2016, 389, 791–797. [Google Scholar] [CrossRef]
  49. Fratoni, E.; de Athayde, A.E.; da Silva Machado, M.; Zermiani, T.; Venturi, I.; dos Santos, M.C.; Lobato, F.; Filho, V.C.; Franchi, G.C., Jr.; Nowill, A.E.; et al. Antiproliferative and toxicological properties of drimanes obtained from Drimys brasiliensis stem barks. Biomed. Pharmacother. 2018, 103, 1498–1506. [Google Scholar] [CrossRef]
  50. Allouche, N.; Apel, C.; Martin, M.-T.; Dumontet, V.; Guéritte, F.; Litaudon, M. Cytotoxic sesquiterpenoids from Winteraceae of Caledonian rainforest. Phytochemistry 2009, 70, 546–553. [Google Scholar] [CrossRef]
  51. Dong, W.-H.; Mei, W.-L.; Zhao, Y.-X.; Zeng, Y.-B.; Wang, H.; Dai, H.-F. A new drimane sesquiterpenoid glycoside from the seeds of Antiaris toxicaria. J. Asian Nat. Prod. Res. 2011, 13, 561–565. [Google Scholar] [CrossRef]
  52. Xu, M.; Litaudon, M.; Krief, S.; Martin, M.-T.; Kasenene, J.; Kiremire, B.; Dumontet, V.; Guéritte, F. Ugandenial A, a New Drimane-type Sesquiterpenoid from Warburgia ugandensis. Molecules 2009, 14, 3844–3850. [Google Scholar] [CrossRef] [PubMed]
  53. Kitte, R.; Tretbar, M.; Dluczek, S.; Beckmann, L.; Marquardt, P.; Duenkel, A.; Schubert, A.; Fricke, S.; Tretbar, U.S. Chemical and Cytotoxic Activity of three main Sesquiterpenoids from Warburgia ugandensis. Results Chem. 2021, 3, 100242. [Google Scholar] [CrossRef]
  54. Sakio, Y.; Hirano, Y.J.; Hayashi, M.; Komiyama, K.; Ishibashi, M. Dendocarbins A–N, New Drimane Sesquiterpenes from the Nudibranch Dendrodoris carbunculosa. J. Nat. Prod. 2001, 64, 726–731. [Google Scholar] [CrossRef] [PubMed]
  55. Kwon, J.; Lee, H.; Seo, Y.H.; Yun, J.; Lee, J.; Kwon, H.C.; Guo, Y.; Kang, J.S.; Kim, J.-J.; Lee, D. Cytotoxic Drimane Sesquiterpenoids Isolated from Perenniporia maackiae. J. Nat. Prod. 2018, 81, 1444–1450. [Google Scholar] [CrossRef] [PubMed]
  56. Liu, H.; Edrada-Ebel, R.; Ebel, R.; Wang, Y.; Schulz, B.; Draeger, S.; Müller, W.E.G.; Wray, V.; Lin, W.; Proksch, P. Drimane Sesquiterpenoids from the Fungus Aspergillus ustus Isolated from the Marine Sponge Suberites domuncula. J. Nat. Prod. 2009, 72, 1585–1588. [Google Scholar] [CrossRef]
  57. Lu, Z.; Wang, Y.; Miao, C.; Liu, P.; Hong, K.; Zhu, W. Sesquiterpenoids and Benzofuranoids from the Marine-Derived Fungus Aspergillus ustus 094102. J. Nat. Prod. 2009, 72, 1761–1767. [Google Scholar] [CrossRef]
  58. Liu, Y.-F.; Yue, Y.-F.; Feng, L.-X.; Zhu, H.-J.; Cao, F. Asperienes A–D, Bioactive Sesquiterpenes from the Marine-Derived Fungus Aspergillus flavus. Mar. Drugs 2019, 17, 550. [Google Scholar] [CrossRef] [Green Version]
  59. Feng, W.; Yang, J.-S. A new drimane sesquiterpenoid and a new triterpene lactone from fungus of Fomes officinalis. J. Asian Nat. Prod. Res. 2015, 17, 1065–1072. [Google Scholar] [CrossRef]
  60. Ngokpol, S.; Suwakulsir, W.; Sureran, S.; Lirdprapamongkol, K.; Aree, T.; Wiyakrutta, S.; Mahidol, C.; Ruchirawat, S.; Kittakoop, P. Drimane Sesquiterpene-Conjugated Amino Acids from a Marine Isolate of the Fungus Talaromyces minioluteus (Penicillium Minioluteum). Mar. Drugs 2015, 13, 3567–3580. [Google Scholar] [CrossRef] [Green Version]
  61. Brooks, E.A.; Galarza, S.; Gencoglu, M.F.; Cornelison, R.C.; Munson, J.M.; Peyton, S.R. Applicability of drug response metrics for cancer studies using biomaterials. Philos. Trans. R. Soc. B 2019, 374, 20180226. [Google Scholar] [CrossRef] [Green Version]
  62. Hagendoorn, M.J.M.; Geelen, T.A.M.; van Beek, T.A.; Jamar, D.C.L.; Tetteroo, F.A.A.; van der Plas, L.H.W. Occurrence of polygodial in plant organs and tissue culture of Polygonum hydropiper. Physiol. Plant. 1994, 92, 595–600. [Google Scholar] [CrossRef]
  63. Just, J.; Jordan, T.B.; Paull, B.; Bissember, A.C.; Smith, J.A. Practical isolation of polygodial from Tasmannia lanceolata: A viable scaffold for synthesis. Org. Biomol. Chem. 2015, 13, 11200–11207. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  64. Montenegro, I.; Tomasoni, G.; Bosio, C.; Quinones, N.; Madrid, A.; Carrasco, H.; Olea, A.; Martinez, R.; Cuellar, M.; Villena, J. Study on the Cytotoxic Activity of Drimane Sesquiterpenes and Nordrimane Compounds against Cancer Cell Lines. Molecules 2014, 19, 18993–19006. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  65. Dasari, R.; de Carvalho, A.; Medellin, D.C.; Middleton, K.N.; Hague, F.; Volmar, M.N.M.; Frolova, L.V.; Rossato, M.F.; de la Chapa, J.J.; Dybdal-Hargreaves, N.F.; et al. Synthetic and Biological Studies of Sesquiterpene Polygodial: Activity of 9-Epipolygodial against Drug-Resistant Cancer Cells. ChemMedChem 2015, 10, 2014–2026. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  66. Dasari, R.; de Carvalho, A.; Medellin, D.C.; Middleton, K.N.; Hague, F.; Volmar, M.N.M.; Frolova, L.V.; Rossato, M.F.; de la Chapa, J.J.; Dybdal-Hargreaves, N.F.; et al. Wittig derivatization of sesquiterpenoid polygodial leads to cytostatic agents with activity against drug resistant cancer cells and capable of pyrrolylation of primary amines. Eur. J. Med. Chem. 2015, 103, 226–237. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  67. Dasari, S.; Samy, A.L.P.A.; Narvekar, P.; Dontaraju, V.S.; Dasari, R.; Kornienko, A.; Munirathinam, G. Polygodial analog induces apoptosis in LNCaP prostate cancer cells. Eur. J. Pharm. 2018, 828, 154–162. [Google Scholar] [CrossRef] [PubMed]
  68. De la Chapa, J.; Singha, P.K.; Sallaway, M.; Self, K.; Nasreldin, R.; Dasari, R.; Hart, M.; Kornienko, A.; Just, J.; Smith, J.A.; et al. Novel polygodial analogs P3 and P27: Efficacious therapeutic agents disrupting mitochondrial function in oral squamous cell carcinoma. Int. J. Oncol. 2018, 53, 2627–2636. [Google Scholar] [CrossRef] [Green Version]
  69. Maslivetc, V.; Laguera, B.; Chandra, S.; Dasari, R.; Olivier, W.J.; Smith, J.A.; Bissember, A.C.; Masi, M.; Evidente, A.; Mathieu, V.; et al. Polygodial and Ophiobolin a Analogues for Covalent Crosslinking of Anticancer Targets. Int. J. Mol. Sci. 2021, 22, 11256. [Google Scholar] [CrossRef]
  70. Lunde, C.; Kubo, I. Effect of Polygodial on the Mitochondrial ATPase of Saccharomyces cerevisiae. Antimicrob. Agents Chemother. 2000, 44, 1943–1953. [Google Scholar] [CrossRef] [Green Version]
  71. Kubo, I.; Fujita, K.; Lee, S.H. Antifungal Mechanism of Polygodial. J. Agric. Food Chem. 2001, 49, 1607–1611. [Google Scholar] [CrossRef]
  72. Castelli, M.V.; Lodeyro, A.F.; Malheiros, A.; Zacchino, S.A.S.; Roveri, O.A. Inhibition of the mitochondrial ATP synthesis by polygodial, a naturally occurring dialdehyde unsaturated sesquiterpene. Biochem. Pharmacol. 2005, 70, 82–89. [Google Scholar] [CrossRef] [PubMed]
  73. Kim, R.; Emi, M.; Tanabe, K. Role of mitochondria as the gardens of cell death. Cancer Chemother. Pharmacol. 2006, 57, 545–553. [Google Scholar] [CrossRef] [PubMed]
  74. Casares, D.; Escribá, P.V.; Rosselló, C.A. Membrane Lipid Composition: Effect on Membrane and Organelle Structure, Function and Compartmentalization and Therapeutic Avenues. Int. J. Mol. Sci. 2019, 20, 2167. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  75. Szlasa, W.; Zendran, I.; Zalesińska, A.; Tarek, M.; Kulbacka, J. Lipid composition of the cancer cell membrane. J. Bioenerg. Biomembr. 2020, 52, 321–342. [Google Scholar] [CrossRef]
  76. Ostenfeld, M.S.; Høyer-Hansen, M.; Bastholm, L.; Fehrenbacher, N.; Olsen, O.D.; Groth-Pedersen, L.; Puustinen, P.; Kirkegaard-Sørensen, T.; Nylandsted, J.; Farkas, T.; et al. Anti-cancer agent siramesine is a lysosomotropic detergent that induces cytoprotective autophagosome accumulation. Autophagy 2008, 4, 487–499. [Google Scholar] [CrossRef] [Green Version]
  77. Dielschneider, R.F.; Eisenstat, H.; Mi, S.; Curtis, J.M.; Johnston, J.B.; Gibson, S.B. Lysosomotropic agents selectively target chronic lymphocytic leukemia cells due to altered sphingolipid metabolism. Leukemia 2016, 30, 1290–1300. [Google Scholar] [CrossRef]
  78. Rafiq, S.; McKenna, S.L.; Muller, S.; Tschan, M.P.; Humbert, M. Lysosomes in acute myeloid leukemia: Potential therapeutic targets? Leukemia 2021, 35, 2759–2770. [Google Scholar] [CrossRef]
  79. Boya, P.; Kroemer, G. Lysosomal membrane permeabilization in cell death. Oncogene 2008, 27, 6434–6451. [Google Scholar] [CrossRef] [Green Version]
  80. Sukhai, M.A.; Giaever, G.; Schimmer, A.D. Lysosomal disruption preferentially targets acute myeloid leukemia cells and progenitors. J. Clin. Investig. 2013, 123, 315–328. [Google Scholar] [CrossRef]
  81. Szallas, A.; Jonassohn, M.; Ács, G.; Bíró, T.; Ács, P.; Blumberg, P.M.; Sterner, O. The stimulation of capsaicin-sensitive neurones in a vanilloid receptor-mediated fashion by pungent terpenoids possessing an unsaturated 1,4-dialdehyde moiety. Br. J. Pharmacol. 1996, 119, 283–290. [Google Scholar] [CrossRef] [Green Version]
  82. André, E.; Camp, B.; Trevisani, M.; Ferreira, J.; Malheiros, Â.; Yunes, R.A.; Calixto, J.B.; Geppetti, P. Pharmacological characterisation of the plant sesquiterpenes polygodial and drimanial as vanilloid receptor agonists. Biochem. Pharmacol. 2006, 71, 1248–1254. [Google Scholar] [CrossRef] [PubMed]
  83. Hartel, M.; di Mola, F.F.; Selvaggi, F.; Mascetta, G.; Wente, M.N.; Felix, K.; Giese, N.A.; Hinz, U.; di Sebastiano, P.; Büchler, M.W.; et al. Vanilloids in pancreatic cancer: Potential for chemotherapy and pain management. Gut 2006, 55, 519–528. [Google Scholar] [CrossRef] [Green Version]
  84. Marincsák, R.; Tóth, B.I.; Czifra, G.; Márton, I.; Rédl, P.; Tar, I.; Tóth, L.; Kovács, L.; Bíró, T. Increased expression of TRPV1 in squamous cell carcinoma of the human tongue. Oral Dis. 2009, 15, 328–335. [Google Scholar] [CrossRef]
  85. Czifra, G.; Varga, A.; Nyeste, K.; Marincsák, R.; Tóth, B.I.; Kovács, I.; Kovács, L.; Bíró, T. Increased expressions of cannabinoid receptor-1 and transient receptor potential vanilloid-1 in human prostate carcinoma. J. Cancer Res. Clin. Oncol. 2009, 135, 507–514. [Google Scholar] [CrossRef]
  86. Weber, L.V.; Al-Refal, K.; Wölk, G.; Bonatz, G.; Altmüller, J.; Becker, C.; Gisselmann, G.; Hatt, H. Expression and functionality of TRPV1 in breast cancer cells. Breast Cancer 2016, 8, 243–252. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  87. Faussone-Pellegrini, M.S.; Taddei, A.; Bizzoco, E.; Lazzeri, M.; Vannuchi, M.G.; Bechi, P. Distribution of the vanilloid (capsaicin) receptor type 1 in the human stomach. Histochem. Cell Biol. 2005, 124, 61–68. [Google Scholar] [CrossRef]
  88. Sun, Z.; Han, J.; Zhao, W.; Zhang, Y.; Wang, S.; Ye, L.; Liu, T.; Zheng, L. TRPV1 Activation Exacerbates Hypoxia/Reoxygenation-Induced Apoptosis in H9C2 Cells via Calcium Overload and Mitochondrial Dysfunction. Int. J. Mol. Sci. 2014, 15, 18362–18380. [Google Scholar] [CrossRef]
  89. Miyake, T.; Shirakawa, H.; Nakagawa, T.; Kaneko, S. Activation of mitochondrial transient receptor potential vanilloid 1 channel contributes to microglial migration. Glia 2015, 63, 1870–1882. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  90. Hurt, C.M.; Lu, Y.; Stary, C.M.; Piplani, H.; Small, B.A.; Urban, T.J.; Qvit, N.; Gross, G.J.; Mochly-Rosen, D.; Gross, E.R. Transient Receptor Potential Vanilloid 1 Regulates Mitochondrial Membrane Potential and Myocardial Reperfusion Injury. J. Am. Heart Assoc. 2016, 5, e003774. [Google Scholar] [CrossRef] [PubMed]
  91. Otto, M.; Bucher, C.; Liu, W.; Müller, M.; Schmidt, T.; Kardell, M.; Driessen, M.N.; Roissaint, J.; Gross, E.R.; Wagner, N.-M. 12(S)-HETE mediates diabetes-induced endothelial dysfunction by activating intracellular endothelial cell TRPV1. J. Clin. Investig. 2020, 130, 4999–5010. [Google Scholar] [CrossRef]
  92. Jardin, I.; López, J.J.; Diez, R.; Sánchez-Collado, J.; Cantonero, C.; Albarrán, L.; Woodard, G.E.; Redondo, P.C.; Salido, G.M.; Smani, T.; et al. TRPs in Pain Sensation. Front. Physiol. 2017, 8, 392. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  93. Bujak, J.K.; Kosmala, D.; Szopa, I.M.; Majchrzak, K.; Bednarczyk, P. Inflammation, Cancer and Immunity—Implication of TRPV1 Channel. Front. Oncol. 2019, 9, 1087. [Google Scholar] [CrossRef] [PubMed]
  94. Roderick, H.L.; Cook, S.J. Ca2+ signalling checkpoints in cancer: Remodelling Ca2+ for cancer cell proliferation and survival. Nat. Rev. 2008, 8, 361–375. [Google Scholar] [CrossRef] [Green Version]
  95. Wu, L.; Lian, W.; Zhao, L. Calcium signaling in cancer progression and therapy. FEBS J. 2021, 288, 6187–6205. [Google Scholar] [CrossRef]
  96. Rizzuto, R.; Pinton, P.; Ferrari, D.; Chami, M.; Szabadkai, G.; Magalhães, P.J.; Di Virgilio, F.; Pozzan, T. Calcium and apoptosis: Facts and hypotheses. Oncogene 2003, 22, 8619–8627. [Google Scholar] [CrossRef] [Green Version]
  97. Hajnóczky, G.; Davies, E.; Madesh, M. Calcium signaling and apoptosis. Biochem. Biophys. Res. Commun. 2003, 304, 445–454. [Google Scholar] [CrossRef]
  98. Jambina, E.; Alonso, R.; Alcalde, M.; Rodríguez, M.d.C.; Serrano, A.; Martínez, C.; García-Sancho, J.; Izquierdo, M. Calcium Influx through Receptor-operated Channel Induces Mitochondria-triggered Paraptotic Cell Death. J. Biol. Chem. 2003, 278, 14134–14145. [Google Scholar] [CrossRef] [Green Version]
  99. Shirakawa, H.; Yamaoka, T.; Sanpei, K.; Sasaoka, H.; Nakagawa, T.; Kaneko, S. TRPV1 stimulation triggers apoptotic cell death of rat cortical neurons. Biochem. Biophys. Res. Commun. 2008, 377, 1211–1215. [Google Scholar] [CrossRef]
  100. Gil, Y.-G.; Kang, M.-K. Capsaicin induces apoptosis and terminal differentiation in human glioma A172 cells. Life Sci. 2008, 82, 997–1003. [Google Scholar] [CrossRef]
  101. Ip, S.W.; Lan, S.-H.; Lu, H.-F.; Huang, A.-C.; Yang, J.-S.; Lin, J.-P.; Huang, H.-Y.; Lien, J.-C.; Ho, C.-C.; Chiu, C.-F.; et al. Capsaicin mediates apoptosis in human nasopharyngeal carcinoma NPC-TW 039 cells through mitochondrial depolarization and endoplasmic reticulum stress. Hum. Exp. Toxicol. 2012, 31, 539–549. [Google Scholar] [CrossRef]
  102. Krizanova, O.; Steliarova, I.; Csaderova, L.; Pastorek, M.; Hudecova, S. Capsaicin induces apoptosis in PC12 cells through ER stress. Oncol. Rep. 2014, 31, 581–588. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  103. Hou, N.; He, X.; Yang, Y.; Fu, J.; Zhang, W.; Guo, Z.; Hu, Y.; Liang, L.; Xie, W.; Xiong, H.; et al. TRPV1 Induced Apoptosis of Colorectal Cancer Cells by Activating Calcineurin-NFAT2-p53 Signaling Pathway. Biomed. Res. Int. 2019, 2019, 6712536. [Google Scholar] [CrossRef] [PubMed]
  104. Gosh, A.K.; Basu, S. Fas-associated factor 1 is a negative regulator in capsaicin induced cancer cell apoptosis. Cancer Lett. 2010, 287, 142–149. [Google Scholar] [CrossRef] [PubMed]
  105. Punzo, F.; Manzo, F.; Tortora, C.; Pota, E.; D’Angelo, V.; Bellini, G.; di Paolo, A.; Verace, F.; Casale, F.; Rossi, F. Effects of CB2 and TRPV1 receptors’ stimulation in pediatric acute T-lymphoblastic leukemia. Oncotarget 2018, 9, 21244–21258. [Google Scholar] [CrossRef] [Green Version]
  106. Meents, J.E.; Ciotu, C.I.; Fischer, M.J.M. TRPA1: A molecular view. J. Neurophysiol. 2019, 121, 427–443. [Google Scholar] [CrossRef]
  107. Talavera, K.; Startek, J.B.; Alvarez-Collazo, J.; Boonen, B.; Alpizar, Y.A.; Sanchez, A.; Naert, R.; Nilius, B. Mammalian Transient Receptor Potential TRPA1 Channels: From Structure to Disease. Physiol. Rev. 2020, 100, 725–803. [Google Scholar] [CrossRef]
  108. Schaefer, E.A.M.; Stohr, S.; Meister, M.; Aigner, A.; Gudermann, T.; Buech, T.R.H. Stimulation of the chemosensory TRPA1 cation channel by volatile toxic substances promotes cell survival of small cell lung cancer cells. Biochem. Pharmacol. 2013, 85, 426–438. [Google Scholar] [CrossRef]
  109. Takahashi, N.; Chen, H.-Y.; Harris, I.S.; Stover, D.G.; Selfors, L.M.; Bronson, R.T.; Deraedt, T.; Cichowski, K.; Welm, A.L.; Mori, Y.; et al. Cancer Cells Co-opt the Neuronal Redox-Sensing Channel TRPA1 to Promote Oxidative-Stress Tolerance. Cancer Cell 2018, 33, 985–1003. [Google Scholar] [CrossRef] [Green Version]
  110. Escalera, J.; von Hehn, C.A.; Bessac, B.F.; Sivula, M.; Jordt, S.-E. TRPA1 Mediates the Noxious Effects of Natural Sesquiterpene Deterrents J. Biol. Chem. 2008, 283, 24136–24144. [Google Scholar] [CrossRef] [Green Version]
  111. Mathie, K.; Lainer, J.; Spreng, S.; Dawid, C.; Andersson, D.A.; Bevan, S.; Hofmann, T. Structure–Pungency Relationships and TRP Channel Activation of Drimane Sesquiterpenes in Tasmanian Pepper (Tasmannia lanceolata). J. Agric. Food Chem. 2017, 65, 5700–5712. [Google Scholar] [CrossRef] [Green Version]
  112. Hinman, A.; Chuang, H.-H.; Bautista, D.M.; Julius, D. TRP channel activation by reversible covalent modification. Proc. Natl. Acad. Sci. USA 2006, 103, 19564–19568. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  113. Macpherson, L.J.; Dubin, A.E.; Evans, M.J.; Marr, F.; Schultz, P.G.; Cravatt, B.F.; Patapoutian, A. Noxious compounds activate TRPA1 ion channels through covalent modification of cysteines. Nature 2007, 445, 541–545. [Google Scholar] [CrossRef] [PubMed]
  114. Takahashi, N.; Mizuno, Y.; Kozai, D.; Yamamoto, S.; Kiyonaka, S.; Shibata, T.; Uchida, K.; Mori, Y. Molecular characterization of TRPA1 channel activation by cysteine-reactive inflammatory mediators. Channels 2008, 2, 287–298. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  115. Bahia, P.K.; Parks, T.A.; Stanford, K.R.; Mitchell, D.A.; Karma, S.; Stevens, S.M., Jr.; Taylor-Clark, T.E. The exceptionally high reactivity of Cys 621 is critical for electrophilic activation of the sensory nerve ion channel TRPA1. J. Gen. Physiol. 2016, 147, 451–465. [Google Scholar] [CrossRef] [Green Version]
  116. Taniguchi, M.; Adachi, T.; Haraguchi, H.; Oi, S.; Kubo, I. Physiological Activity of Warburganal and Its Reactivity with Sulfhydryl Groups. J. Biochem. 1983, 94, 149–154. [Google Scholar] [CrossRef]
  117. Jonassohn, M.; Davidsson, R.; Kahnberg, P.; Sterner, O. The reactivity of drimane unsaturated dialdehydes towards nucleophiles. Tetrahedron 1997, 53, 237–244. [Google Scholar] [CrossRef]
  118. Lam, P.Y.-S.; Frazier, J.L. Model study on the mode of action of muzigadial antifeedant. Tetrahedron Lett. 1987, 28, 5477–5480. [Google Scholar] [CrossRef]
  119. Norbury, C.J.; Zhivotovsky, B. DNA damage-induced apoptosis. Oncogene 2004, 23, 2797–2808. [Google Scholar] [CrossRef] [Green Version]
  120. Kim, W.; Lee, S.; Seo, D.; Kim, D.; Kim, K.; Kim, E.; Kang, J.; Seong, K.M.; Youn, H.; Youn, B. Cellular Stress Responses in Radiotherapy. Cells 2019, 8, 1105. [Google Scholar] [CrossRef] [Green Version]
  121. Ercal, N.; Gurer-Orhan, H.; Aykin-Burns, N. Toxic Metals and Oxidative Stress Part I: Mechanisms Involved in Metal-induced Oxidative Damage. Curr. Top. Med. Chem. 2001, 1, 529–539. [Google Scholar] [CrossRef] [PubMed]
  122. Indo, H.P.; Davidson, M.; Yen, H.-C.; Suenaga, S.; Tomita, K.; Nishii, T.; Higuchi, M.; Koga, Y.; Ozawa, T.; Majima, H.J. Evidence of ROS generation by mitochondria in cells with impaired electron transport chain and mitochondrial DNA damage. Mitochondrion 2007, 7, 106–118. [Google Scholar] [CrossRef] [PubMed]
  123. Henkler, F.; Brinkmann, J.; Luch, A. The Role of Oxidative Stress in Carcinogenesis Induced by Metals and Xenobiotics. Cancers 2010, 2, 376–396. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  124. Perillo, B.; Donato, M.D.; Pezone, A.; Zazzo, E.D.; Giovannelli, P.; Galasso, G.; Castoria, G.; Migliaccio, A. ROS in cancer therapy: The bright side of the moon. Exp. Mol. Med. 2020, 52, 192–203. [Google Scholar] [CrossRef] [PubMed]
  125. Mah, L.-J.; El-Osta, A.; Karagiannis, T.C. γH2AX: A sensitive molecular marker of DNA damage and repair. Leukemia 2010, 24, 679–686. [Google Scholar] [CrossRef] [Green Version]
  126. Rogakou, E.P.; Pilch, D.R.; Orr, A.H.; Ivanova, V.S.; Bonner, W.M. DNA Double-stranded Breaks Induce Histone H2AX Phosphorylation on Serine 139. J. Biol. Chem. 1998, 273, 5858–5868. [Google Scholar] [CrossRef] [Green Version]
  127. Lu, C.; Zhu, F.; Cho, Y.-Y.; Tang, F.; Zykova, T.; Ma, W.-y.; Bode, A.M.; Dong, Z. Cell Apoptosis: Requirement of H2AX in DNA Ladder Formation, but Not for the Activation of Caspase-3. Mol. Cell. 2006, 23, 121–132. [Google Scholar] [CrossRef] [Green Version]
  128. Bouchard, V.J.; Rouleau, M.; Poirier, G.G. PARP-1, a determinant of cell survival in response to DNA damage. Exp. Hematol. 2003, 31, 446–454. [Google Scholar] [CrossRef]
  129. Soldani, C.; Scovassi, A.I. Poly(ADP-ribose) polymerase-1 cleavage during apoptosis: An update. Apoptosis 2002, 7, 321–328. [Google Scholar] [CrossRef]
  130. Zhang, Y.-L.; Chen, G.-L.; Liu, Y.; Zhuang, X.-C.; Guo, M.-Q. Stimulation of ROS Generation by Extract of Warburgia ugandensis Leading to G0/G1 Cell Cycle Arrest and Antiproliferation in A549 Cells. Antioxidants 2021, 10, 1559. [Google Scholar] [CrossRef]
  131. Zhang, Y.; Chen, G.; Zhuang, X.; Guo, M. Inhibition of Growth of Colon Tumors and Proliferation of HT-29 Cells by Warburgia ugandensis Extract through Mediating G0/G1 Cell Cycle Arrest, Cell Apoptosis, and Intracellular ROS Generation. Oxid. Med. Cell. Longev. 2021, 2021, 8807676. [Google Scholar] [CrossRef]
  132. Thielen, S. Charakterisierung DNA-Schädigender Wirkungen Mittels Comet Assay und hPRT-Genmutations-Assay: Ein Vergleich Zwischen α,β-Ungesättigten Carbonylverbindungen, Epoxiden und N-Nitroso-Verbindungen. Ph.D. Thesis, TU Kaiserslautern, Kaiserslautern, Germany, 13 December 2007. [Google Scholar]
  133. Redza-Dutordoir, M.; Averill-Bates, D.A. Activation of apoptosis signalling pathways by reactive oxygen species. Biochim. Biophys. Acta Mol. Cell Res. 2016, 1863, 2977–2992. [Google Scholar] [CrossRef] [PubMed]
  134. Zeeshan, H.M.A.; Lee, G.H.; Kim, H.-R.; Chae, H.-J. Endoplasmic Reticulum Stress and Associated ROS. Int. J. Mol. Sci. 2016, 17, 327. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  135. Schäfer, K.A. The Cell Cycle: A Review. Vet. Pathol. 1998, 35, 461–478. [Google Scholar] [CrossRef] [PubMed]
  136. Vermeulen, K.; Van Bockstaele, D.R.; Berneman, Z.N. The cell cycle: A review of regulation, deregulation and therapeutic targets in cancer. Cell Prolif. 2003, 36, 131–149. [Google Scholar] [CrossRef]
  137. Malumbres, M. Cyclin-dependent kinases. Genome Biol. 2014, 15, 122. [Google Scholar] [CrossRef] [Green Version]
  138. Alao, J.P. The regulation of cyclin D1 degradation: Roles in cancer development and the potential for therapeutic invention. Mol. Cancer 2007, 6, 24. [Google Scholar] [CrossRef] [Green Version]
  139. Caldon, C.E.; Musgrove, E.A. Distinct and redundant functions of cyclin E1 and cyclin E2 in development and cancer. Cell Div. 2010, 5, 2. [Google Scholar] [CrossRef] [Green Version]
  140. Yang, W.; Zhang, Y.; Li, Y.; Wu, Z.; Zhu, D. Myostatin Induces Cyclin D1 Degradation to Cause Cell Cycle Arrest through a Phosphatidylinositol 3-Kinase/AKT/GSK-3β Pathway and Is Antagonized by Insulin-like Growth Factor 1. J. Biol. Chem. 2007, 282, 3799–3808. [Google Scholar] [CrossRef] [Green Version]
  141. Chu, I.M.; Hengst, L.; Slingerland, J.M. The Cdk inhibitor p27 in human cancer: Prognostic potential and relevance to anticancer therapy. Nat. Rev. Cancer 2008, 8, 253–267. [Google Scholar] [CrossRef]
  142. Gstaiger, M.; Jordan, R.; Lim, M.; Catzavelos, C.; Mestan, J.; Slingerland, J.; Krek, W. Skp2 is oncogenic and overexpressed in human cancers. Proc. Natl. Acad. Sci. USA 2001, 98, 5043–5048. [Google Scholar] [CrossRef] [Green Version]
  143. Zhong, L.; Georgia, S.; Tschen, S.; Nakayama, K.; Bhushan, A. Essential role of Skp2-mediated p27 degradation in growth and adaptive expansion of pancreatic β cells. J. Clin. Investig. 2007, 117, 2869–2876. [Google Scholar] [CrossRef] [Green Version]
  144. Duan, Y.; Pan, J.; Chen, J.; Zhu, D.; Wang, J.; Sun, X.; Chen, L.; Wu, L. Soluble Egg Antigens of Schistosoma japonicum Induce Senescence of Activated Hepatic Stellate Cells by Activation of the FoxO3a/SKP2/P27 Pathway. PLoS Negl. Trop. Dis. 2016, 10, e0005268. [Google Scholar] [CrossRef]
  145. Li, F.; Dong, X.; Lin, P.; Jiang, J. Regulation of Akt/FoxO3a/Skp2 Axis Is Critically Involved in Berberine-Induced Cell Cycle Arrest in Hepatocellular Carcinoma Cells. Int. J. Mol. Sci. 2018, 19, 327. [Google Scholar] [CrossRef] [Green Version]
  146. Lohberger, B.; Rinner, B.; Stuendl, N.; Kaltenegger, H.; Steinecker-Frohnwieser, B.; Bernhart, E.; Rad, E.B.; Weinberg, A.M.; Leithner, A.; Bauer, R.; et al. Sesquiterpene Lactones Downregulate G2/M Cell Cycle Regulator Proteins and Affect the Invasive Potential of Human Soft Tissue Sarcoma Cells. PLoS ONE 2013, 8, e66300. [Google Scholar] [CrossRef] [Green Version]
  147. Felix, S.; Sandjo, L.P.; Opatz, T.; Erkel, G. SF002-96-1, a new drimane sesquiterpene lactone from an Aspergillus species, inhibits survivin expression. Beilstein J. Org. Chem. 2013, 9, 2866–2876. [Google Scholar] [CrossRef] [Green Version]
  148. Tang, J.-J.; Wang, M.-R.; Dong, S.; Huang, L.-F.; He, Q.-R.; Gao, J.-M. 1,10-Seco-Eudesmane sesquiterpenoids as a new type of anti-neuroinflammatory agents by suppressing TLR4/NF-κB/MAPK pathways. Eur. J. Med. Chem. 2021, 224, 113713. [Google Scholar] [CrossRef]
  149. Appendino, G.; Maxia, L.; Bascope, M.; Houghton, P.J.; Sanchez-Duffhues, G.; Muñoz Sterner, O. A Meroterpenoid NF-κB Inhibitor and Drimane Sesquiterpenoids from Asafetida. J. Nat. Prod. 2006, 69, 1101–1104. [Google Scholar] [CrossRef]
  150. Newman, D.J.; Cragg, G.M. Natural Products as Sources of New Drugs from 1981 to 2014. J. Nat. Prod. 2016, 79, 629–661. [Google Scholar] [CrossRef] [Green Version]
  151. Hughes, J.P.; Rees, S.; Kalindjian, S.B.; Philpott, K.L. Principles of early drug discovery. Br. J. Pharmacol. 2011, 162, 1239–1249. [Google Scholar] [CrossRef] [Green Version]
  152. Harvey, A.L. Natural products in drug discovery. Drug Discov. Today 2008, 13, 894–901. [Google Scholar] [CrossRef]
  153. Westhouse, R.A. Safety Assessment Considerations and Strategies for Targeted Small Molecule Cancer Therapeutics in Drug Discovery. Toxicol. Pathol. 2010, 38, 165–168. [Google Scholar] [CrossRef] [PubMed]
  154. Belhadj, S.; Keskes, H.; Apel, C.; Roussi, F.; Litaudon, M.; Hentati, O.; Allouche, N. Anti-diabetic and lipid-lowering effects of drimane sesquiterpenoids isolated from Zygogynum pancheri. Chem.-Biol. Interact. 2020, 330, 109167. [Google Scholar] [CrossRef] [PubMed]
  155. Zhao, Y.J.; Li, B.; Tan, L.-S.S.; Shen, Z.-L.; Loh, T.-P. Enantioselective Cationic Polyene Cyclization vs Enantioselective Intramolecular Carbonyl−Ene Reaction. J. Am. Chem. Soc. 2010, 132, 10242–10244. [Google Scholar] [CrossRef] [PubMed]
  156. Schafroth, M.A.; Sarlah, D.; Krautwald, S.; Carreira, E.M. Iridium-Catalyzed Enantioselective Polyene Cyclization. J. Am. Chem. Soc. 2012, 134, 20276–20278. [Google Scholar] [CrossRef]
  157. Arnold, A.M.; Pöthig, A.; Drees, M.; Gulder, T. NXS, Morpholine, and HFIP: The Ideal Combination for Biomimetic Haliranium-Induced Polyene Cyclizations. J. Am. Chem. Soc. 2018, 140, 4344–4353. [Google Scholar] [CrossRef]
Molecules 27 02501 g001 550
Figure 1. Examples for cytotoxic sesquiterpenoids: dehydrovomifol (1), polygodial (2), and lemnalol (3) [12].
Figure 1. Examples for cytotoxic sesquiterpenoids: dehydrovomifol (1), polygodial (2), and lemnalol (3) [12].
Molecules 27 02501 g001
Molecules 27 02501 g002 550
Figure 2. Scaffold of drimane and coloratane sesquiterpenoids.
Figure 2. Scaffold of drimane and coloratane sesquiterpenoids.
Molecules 27 02501 g002
Molecules 27 02501 g003 550
Figure 3. Cytotoxic drimanes against P388- and nasopharynx cancer cells (Eagle’s 9KB-cells) [45,46].
Figure 3. Cytotoxic drimanes against P388- and nasopharynx cancer cells (Eagle’s 9KB-cells) [45,46].
Molecules 27 02501 g003
Molecules 27 02501 g004 550
Figure 4. Cytotoxic drimanes from Cinnamosma fragrans against A549 carcinoma cells [47].
Figure 4. Cytotoxic drimanes from Cinnamosma fragrans against A549 carcinoma cells [47].
Molecules 27 02501 g004
Molecules 27 02501 g005 550
Figure 5. Cytotoxic drimanes from Drimys brasiliensis, Zygogynum pancheri, and Zygogynum acsmithii [48,49,50].
Figure 5. Cytotoxic drimanes from Drimys brasiliensis, Zygogynum pancheri, and Zygogynum acsmithii [48,49,50].
Molecules 27 02501 g005
Molecules 27 02501 g006 550
Figure 6. Glycosylated drimane 23 from seeds of Antairis toxicaria [51].
Figure 6. Glycosylated drimane 23 from seeds of Antairis toxicaria [51].
Molecules 27 02501 g006
Molecules 27 02501 g007 550
Figure 7. Cytotoxic drimanes and coloratanes from Warburgia ugandensis against KB, MV4-11, THP-1 leukemia, Sk-Mel29 melanoma, and LN-229 glioblastoma cells [52,53].
Figure 7. Cytotoxic drimanes and coloratanes from Warburgia ugandensis against KB, MV4-11, THP-1 leukemia, Sk-Mel29 melanoma, and LN-229 glioblastoma cells [52,53].
Molecules 27 02501 g007
Molecules 27 02501 g008 550
Figure 8. Drimane lactones from Dendrodoris carbunculosa [54].
Figure 8. Drimane lactones from Dendrodoris carbunculosa [54].
Molecules 27 02501 g008
Molecules 27 02501 g009 550
Figure 9. Cytotoxic drimane acetals from Perenniporia maackiae [55].
Figure 9. Cytotoxic drimane acetals from Perenniporia maackiae [55].
Molecules 27 02501 g009
Molecules 27 02501 g010 550
Figure 10. Cytotoxic drimanes from Aspergillus ustus against L5178Y-, PC-12-, A549-, HL-60-, and HeLa cells [56,57].
Figure 10. Cytotoxic drimanes from Aspergillus ustus against L5178Y-, PC-12-, A549-, HL-60-, and HeLa cells [56,57].
Molecules 27 02501 g010
Molecules 27 02501 g011 550
Figure 11. Cytotoxic drimanes from Aspergillus flavus against HeLa, MCF-7, MGC-803, and A549 cells [58].
Figure 11. Cytotoxic drimanes from Aspergillus flavus against HeLa, MCF-7, MGC-803, and A549 cells [58].
Molecules 27 02501 g011
Molecules 27 02501 g012 550
Figure 12. Cytotoxic drimane 45 from Fomes officinalis against HL-60, Bel-7402, and KB cells [59].
Figure 12. Cytotoxic drimane 45 from Fomes officinalis against HL-60, Bel-7402, and KB cells [59].
Molecules 27 02501 g012
Molecules 27 02501 g013 550
Figure 13. Cytotoxic drimane sesquiterpene-conjugated amino acids from Talaromyces minioluteus [60].
Figure 13. Cytotoxic drimane sesquiterpene-conjugated amino acids from Talaromyces minioluteus [60].
Molecules 27 02501 g013
Molecules 27 02501 g014 550
Figure 14. Cytotoxic and inactive drimanes from Montenegro et al. [64].
Figure 14. Cytotoxic and inactive drimanes from Montenegro et al. [64].
Molecules 27 02501 g014
Molecules 27 02501 sch001 550
Scheme 1. Epimerization reaction of polygodial (2) [65].
Scheme 1. Epimerization reaction of polygodial (2) [65].
Molecules 27 02501 sch001
Molecules 27 02501 sch002 550
Scheme 2. (a) Wittig derivatization of 9-epipolygodial (49). (b) Chemoselective reduction of polygodial (2) [65].
Scheme 2. (a) Wittig derivatization of 9-epipolygodial (49). (b) Chemoselective reduction of polygodial (2) [65].
Molecules 27 02501 sch002
Molecules 27 02501 sch003 550
Scheme 3. Polygodial derived bis-acetals 57–63 [65].
Scheme 3. Polygodial derived bis-acetals 57–63 [65].
Molecules 27 02501 sch003
Molecules 27 02501 sch004 550
Scheme 4. Wittig derivatization of polygodial (2) [66].
Scheme 4. Wittig derivatization of polygodial (2) [66].
Molecules 27 02501 sch004
Molecules 27 02501 sch005 550
Scheme 5. HWE reaction of polygodial (2) with phosphonate 72 [67,68].
Scheme 5. HWE reaction of polygodial (2) with phosphonate 72 [67,68].
Molecules 27 02501 sch005
Molecules 27 02501 g015 550
Figure 15. Polyethylene-linked polygodial dimers 74a–c [69].
Figure 15. Polyethylene-linked polygodial dimers 74a–c [69].
Molecules 27 02501 g015
Molecules 27 02501 g016 550
Figure 16. General trends in cytotoxicity for semi-synthetic derivatives from polygodial (2). Derivatizations leading to an increase in cytotoxicity are depicted in red and functional groups, leading to a decrease are shown in blue. Other functionalizations with a minor/moderate impact on cytotoxic activity are depicted in purple.
Figure 16. General trends in cytotoxicity for semi-synthetic derivatives from polygodial (2). Derivatizations leading to an increase in cytotoxicity are depicted in red and functional groups, leading to a decrease are shown in blue. Other functionalizations with a minor/moderate impact on cytotoxic activity are depicted in purple.
Molecules 27 02501 g016
Molecules 27 02501 g017 550
Figure 17. Schematic overview of potential modes-of-action of drimanes, coloratanes, and drimane-containing plant extracts.
Figure 17. Schematic overview of potential modes-of-action of drimanes, coloratanes, and drimane-containing plant extracts.
Molecules 27 02501 g017
Molecules 27 02501 g018 550
Figure 18. Drimane lactone 75 inhibits NF-κB and Stat-3 to the promoter of the survivin gene [147].
Figure 18. Drimane lactone 75 inhibits NF-κB and Stat-3 to the promoter of the survivin gene [147].
Molecules 27 02501 g018
Table
Table 2. Cytotoxicity of drimanes from Montenegro et al.; IC50 values against DU-145, PC-, and MCF-7 cancer cells are given [64].
Table 2. Cytotoxicity of drimanes from Montenegro et al.; IC50 values against DU-145, PC-, and MCF-7 cancer cells are given [64].
Comp.DU-145
(μM)		PC-3
(μM)		MCF-7
(μM)
2 171.4 ± 8.589.2 ± 6.893.7 ± 9.1
4 1>200>200>200
49 1>200>200>200
50 270.6 ± 5.965.4 ± 5.597.1 ± 7.2
51 3>200>200>200
52 493.5 ± 6.797.5 ± 10.4>200
53 5>20090.2 ± 8.888.4 ± 7.1
1 Obtained from Drimys winteri. 2 Starting material: polygodial (2), p-TosOH, ethylene glycol. 3 Starting material: polygodial (2), NaBH4 (excess), MeOH, r.t. or KOH, MeOH (50%), NaBH4 (excess). 4 Starting material: drimenol (4) (i) PCC, CH2Cl2, r.t., 2 h (ii) m-CPBA, CH2Cl2, r.t., 1.5 h 5 Starting material: drimenol (4), (i) Pb(IV)-acetate, benzol, reflux (ii) m-CPBA, CH2Cl2, r.t. (iii) MeONa, MeOH, rt (iv) PDC, CH2Cl2, r.t.
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Beckmann, L.; Tretbar, U.S.; Kitte, R.; Tretbar, M. Anticancer Activity of Natural and Semi-Synthetic Drimane and Coloratane Sesquiterpenoids. Molecules 2022, 27, 2501. https://doi.org/10.3390/molecules27082501

AMA Style

Beckmann L, Tretbar US, Kitte R, Tretbar M. Anticancer Activity of Natural and Semi-Synthetic Drimane and Coloratane Sesquiterpenoids. Molecules. 2022; 27(8):2501. https://doi.org/10.3390/molecules27082501

Chicago/Turabian Style

Beckmann, Lorenz, Uta Sandy Tretbar, Reni Kitte, and Maik Tretbar. 2022. "Anticancer Activity of Natural and Semi-Synthetic Drimane and Coloratane Sesquiterpenoids" Molecules 27, no. 8: 2501. https://doi.org/10.3390/molecules27082501

Article Metrics

Back to TopTop