Monoterpenes structure

Monoterpenes structure DEFAULT

Chemical structures of p-menthane monoterpenes with special reference to their effect on seed germination and termite mortality

Journal of Wood Sciencevolume 59, pages 229–237 (2013)Cite this article

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Abstract

A series of p-menthane monoterpenes was investigated to confirm any correlation between their bioactivity (effect on seed germination and termite mortality) and chemical structure. The germination percentages of Brassica rapa seeds at a concentration of 0.1 mg/Petri dish of (+)-pulegone, isopulegol, piperitone, (−)-dihydrocarveol, terpinen-4-ol and (−)-menthol were found to be 21.6, 27.3, 27.3, 29.1, 42.9 and 43.4, respectively. The lethal concentration 50 values of carvacrol, (+)-pulegone, thymol, (−)-menthol and (−)-terpinen-4-ol for termites (Reticulitermes speratus) were 0.34, 0.50, 0.65, 0.92 and 1.26 (mg/Petri dish), respectively. Of all the compounds tested, phenols produced the highest levels of termite mortality, with ketones and alcohols also showing bioactivity. An assessment of the bioactivity revealed that the presence of a phenol group was effective for termite mortality, with a carbonyl group also showing strong bioactivity. The presence of an alcohol or isopropyl group in a ring also contributed to the bioactivity, whereas the presence of an isopropenyl group at the same position, however, exhibited an inhibitory effect on seed germination. In conclusion, the bioactivity of the p-menthane monoterpenoids was dependent upon the presence and position of certain functional groups and the degree of saturation in the functional group of the side chain.

Introduction

Monoterpenes are a class of organic compounds composed of two isoprene units and can be further classified as either acyclic (e.g., myrcene), cyclic (e.g., p-menthanes) or bicyclic (e.g., bornanes, fenchanes, caranes, pinanes and thujanes) depending upon their molecular connectivities. In addition to their basic hydrocarbon forms, they can exist as the corresponding oxygenated compounds, containing aldehyde, alcohol, ketone, ester and ether functionalities [1, 2]. Monoterpenes have been isolated from the essential oils of many higher plants. Essential oils are very complex natural mixtures and, in the majority of cases, contain somewhere in the region 20–60 components at quite different concentrations. Monoterpenes are an important class of compounds in perfumery and flavor industries [3].

Thyme oil contains thymol, carvacrol, p-cymene and γ-terpinene as its major components [3, 4], and is used as a flavoring in the processed food industry, as well as in soaps and detergents [3]. Thyme oil has been shown to exhibit a range of biological activities, including antimicrobial [5, 6], antifungal [7], trypanocidal [8], and antioxidant activities [9].

Peppermint oil contains about 40 % menthol and 20 % menthone, and is used in various medicinal, pharmaceutical, cosmetic and perfumery products as well as in foodstuffs [3, 10]. (−)-Menthol is a particularly important flavoring compound and is used extensively in toothpastes and chewing gums [2, 3, 11]. Peppermint oil is known to possess bioactivities such as antimicrobial [10], and genotoxic activities [12].

p-Menthane monoterpenes are characteristic of thyme and peppermint oils [2] and the bioactivities of both of these oils are possibly related to the functions of p-menthane monoterpenes.

Recently, the essential oils of needles of Pinus thunbergii and Cryptomeria japonica were found to be composed predominantly of p-menthane monoterpenes at 19.8 and 27.1 %, respectively [13]. In addition, the monoterpenes α-terpineol and terpinen-4-ol, both of which possess a p-menthane skeleton, were identified as the bioactive components of the needle essential oils of P.thunbergii and C. japonica, respectively [13]. Of all of the compounds contained within the essential oils, the bioactivity levels of α-terpineol and terpinen-4-ol were found to be particularly pronounced against plant seeds (Raphanus sativus and Brassica rapa) and termites (Reticulitermes speratus) [13]. These findings suggested that the bioactivity of p-menthanes was affected by the presence of an alcoholic hydroxyl group. With this in mind, a fundamental evaluation of the correlation between the bioactivity of p-menthane compounds and their chemical structures would therefore be required to develop a better understanding of this particular series of compounds. There are, however, a few reports on the chemical structure of p-menthane monoterpenes with special reference to their effects on seed germination and termite mortality. Therefore, in the present study, p-menthane monoterpenes have been investigated with the aim of confirming the structure–activity relationship (SAR), in terms of their effect on seed germination and termite mortality, and their chemical structure.

Materials and methods

Test samples

p-Cymene, α-terpinene, (−)-α-phellandrene, terpinolene, thymol, carvacrol, (−)-terpinen-4-ol, trans-sobrerol, (−)-menthol, isopulegol, (−)-carvone, piperitone and (+)-pulegone were purchased from Tokyo Chemical Industry Co. Ltd. (Tokyo, Japan). γ-Terpinene, (+)-limonene and (−)-α-terpineol were purchased from Wako Pure Chemical Industries, Ltd. (Osaka, Japan). p-Menthane, p-cymen-8-ol, 1,8-p-menthadiene-4-ol and (−)-dihydrocarveol were supplied by Nippon Terpene Chemicals, Inc. (Hyogo, Japan). (−)-Carveol was purchased from Sigma-Aldrich Japan Co. LLC. (Tokyo, Japan). The chemical structures of these compounds are shown in Fig. 1.

Chemical structures of the monoterpenoids containing a p-menthane skeleton

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Seeds

In the present study, Raphanus sativus var. sativus and Brassica rapa var. perviridis seeds were used, and were provided by Takii & Co. Ltd. (Kyoto, Japan).

Germination test

The germination test was carried out according to a method described previously in the literature [13]. The effects of the p-menthane compounds on germination were tested on the R. sativus and B. rapa seeds. A sample of each of the compounds (50, 5 and 0.5 mg) was dissolved separately in acetone (10 mL), and a portion (2.0 mL) of the resulting solutions was poured onto a filter paper (90 mm diameter, Advantec No. 2, Toyo Roshi Kaisha, Ltd., Tokyo, Japan) in a Petri dish. In a control Petri dish, acetone (2.0 mL) was only used. The acetone solvent was then removed by evaporation over a period of 20 min in a fume hood. Deionized water (10 mL) was then added to each of the Petri dishes together with 20 seeds of R. sativus or B. rapa. Petri dishes containing concentrations of 10.0, 1.0 and 0.1 (mg/Petri dish) of each of the compounds were prepared in this way. The screening germination test was conducted only at a concentration of 10.0 mg/Petri dish. Following on from the screening test, germination tests were conducted for the active components at concentrations of 1.0 and 0.1 mg/Petri dish. Each of the compound tests and control treatments were performed in triplicate. The Petri dishes were then covered and placed in a growth chamber (Eyelatron FLI-301NH, Tokyo Rikakikai Co. Ltd., Tokyo, Japan) at a temperature of 24 ± 2 °C for 3 days (sequential 10 h photoperiods and 14 h dark periods).

The number of germinated individuals was recorded on a daily basis throughout the experimental period to determine the germination percentage. The germination percentages were calculated according to formula (1) by considering the control as 100 (%):

$$ {\text{Germination}}\;{\text{percent}}\;{\text{of}}\;R. \, sativus\; ( {\text{or}}\;B.\;rapa)\;(\% ) = {\text{number}}\;{\text{of}}\;{\text{germinated}}\;{\text{seeds}}\;{\text{following}}\; 2\;{\text{days (or }}1\;{\text{day)}}\;{\text{of}}\;{\text{the}}\;{\text{tests}}\;{\text{for}}\;{\text{each}}\;{\text{sample/number}}\;{\text{of}}\;{\text{germinated}}\;{\text{seeds}}\;{\text{following}}\; 3\;{\text{days}}\;{\text{of}}\;{\text{the}}\;{\text{control}}\;{\text{tests}} \times 100 $$

(1)

Termites

Colonies of Reticulitermes speratus were collected from the Institute of Wood Technology at the Akita Prefectural University of Japan, in July 2009. The colonies were maintained in a room at 20 ± 2 °C for one and a half years prior to the initiation of the test.

Termiticidal test

The test was conducted according to a method previously described in the literature [13]. Each of the compounds (10.0, 5.0, and 2.5 mg) was dissolved separately in acetone (10 mL) and portion (2.0 mL) of each of the resulting solutions was poured onto a filter paper (90 mm diameter, Advantec No. 2, Toyo Roshi Kaisha, Ltd., Tokyo, Japan) in a Petri dish. In a control Petri dish, acetone (2 mL) was only used. The acetone solvent was then removed by evaporation over a period of 20 min in a fume hood. Deionized water (1.5 mL) was then added to each of the Petri dishes together with 20 active termites (workers). Petri dishes containing concentrations of 2.0, 1.0 and 0.5 (mg/Petri dish) of each compound were prepared in this way. Each compound test and control experiment was conducted in triplicate. The Petri dishes were then covered and placed in an incubator (ICV-450, As One Co., Osaka, Japan) at a temperature of 25 ± 1 °C (dark) for the entire test period (1 h).

The numbers of surviving and dead termites were counted following the tests to determine the percentage mortality values. The percent mortality data were subjected to probit analysis to calculate the lethal concentration required to kill 50 % of the termites (LC50) [14].

Statistical analysis

Test samples were compared using analysis of variance (ANOVA), and the means were separated using a protected Tukey–Kramer test (p < 0.05; Statcel 2, Saitama, Japan) [15]. The mean separation test followed transformation to arcsine square root percent of seed germination and termite mortality. The actual percentages are reported in Table 1 and Figs. 2 and 3.

Full size table

Effect of the p-menthane monoterpenes on the germination of the R. sativus seedsI. ISome of test results were cited in the literature data [13]. Each determination was made with three replicates of twenty seeds. Bars represent standard deviations. IIGermination percent (%) = number of germination seeds of each sample after 2 days of the test/number of germination seeds after 3 days of the tests of control × 100. IIIMeans in the same column with thesame letters are not significantly different. Tukey–Kramer, p ≤ 0.05

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Effect of p-menthane monoterpenes on the germination of B. rapa seedsI. ISome of test results were cited in the literature data [13]. Each determination was made with three replicates of twenty seeds. Bars represent standard deviations. IIGermination percent (%) = number of germination seeds of each sample after 1 day of the test/number of germination seeds after 3 days of the tests of control × 100. IIIMeans in the same column with thesame letters are not significantly different. Tukey–Kramer, p ≤ 0.05. ns Not significant

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Results

Effect of the p-menthane monoterpenes on seed germination

Following days 2 and 3 of the test period, the actual germination percentages of the R. sativus control reached 72.4 and 78.0 %, respectively. Following days 1 and 3 of the test period, the germination percentages of the B.rapa control reached 76.9 and 89.0 %, respectively. The germination percentages of each of the samples at a concentration of 10.0 mg/Petri dish were expressed relative to the percentages of the controls for the R. sativus and B.rapa seeds (Table 1). Of the hydrocarbon type monoterpenes, terpinolene showed inhibitory activity toward the seed germination, whereas almost all of the phenol, alcohol and ketone containing monoterpenes prevented the plant seeds from germinating.

The germination percentages of the active components at concentrations of 1.0 and 0.1 mg/Petri dish were expressed relative to the percentages of the controls for the R. sativus and B. rapa seeds (Figs. 2, 3). With the exception of terpinolene, the germination percentages of the hydrocarbon type monoterpenes for the R. sativus and B. rapa seeds at a concentration of 0.1 mg/Petri dish were almost identical to those of the control. The germination percentage of the R. sativus seeds in the presence of thymol at a concentration of 0.1 mg/Petri dish was 68.9 %, revealing that thymol exhibited a negligible inhibitory effect on the seed germination process. It was recognized that the hydrocarbons and phenols showed only weak or no inhibitory activity against both types of seeds. Of the alcohols at a concentration of 0.1 mg/Petri dish, (−)-terpinen-4-ol, (−)-carveol and (−)-menthol showed inhibitory effects on the R. sativus seeds, with germination percentages of 14.4, 27.0 and 37.8 %, respectively. The percentage of germination in the B. rapa seeds in the presence of isopulegol and (−)-dihydrocarveol were 27.3 and 29.1 %, respectively. The inhibitory effects of these compounds were relatively high compared to those of the other alcohols. The germination percentages of the R. sativus seeds in the compounds containing ketones at a concentration of 0.1 mg/Petri dish were 18.9, 20.0 and 31.1 % for piperitone, (+)-pulegone and (−)-carvone, respectively. The germination percentages of the B. rapa seeds in the presence of (+)-pulegone and piperitone were 21.6 and 27.3 %, respectively. Taken together, these data demonstrate that all of the ketone containing compounds exhibited significant inhibitory effects against both types of seeds.

Termiticidal activity of p-menthane monoterpenes

The LC50 values for each of the samples are shown in Table 2. The results show that the hydrocarbons did not display any significant termiticidal activity, whereas the phenolic compounds demonstrated significantly higher levels of termite mortality than the controls, with carvacrol and thymol being the most active of the p-menthane compounds with LC50 values of 0.34 and 0.65 mg/Petri dish, respectively. Of the alcohols, (−)-menthol and (−)-terpinen-4-ol were also quite active, providing LC50 values of 0.92 and 1.26 mg/Petri dish, respectively. In contrast, however, isopulegol, 1,8-p-menthadiene-4-ol and p-cymen-8-ol showed only weak activities with LC50 values of 3.67, 3.96 and 4.49 mg/Petri dish, respectively. The LC50 values of the ketones containing compounds (+)-pulegone, (−)-carvone and piperitone, were 0.50, 1.62 and 1.87 mg/Petri dish, respectively. The insecticidal activity of (+)-pulegone was greater than that of piperitone and (−)-carvone.

Full size table

Discussion

A previous study on the effects of monoterpenoids on the germination of Lactuca sativa seeds showed that the inhibitory effects of alcohols, phenols and ketones were larger than those of the corresponding hydrocarbons [16]. Alicyclic monoterpene compounds containing a keto group conjugated with a double bond in particular were highlighted as showing the largest inhibitory effects against the germination of L. sativa seeds [17]. We have proposed that the inhibitory effects of terpinen-4-ol and α-terpineol on the germination of R. sativus and B. rapa seeds were greater than those of bornyl acetate, α-pinene and β-pinene [13].

In the present study, it was recognized that hydrocarbons and phenols showed weak to no inhibitory effects against the germination of both types of seeds, whereas alcohols and ketones, with the exception of sobrerol, exhibited stronger inhibitory effects. Although any correlation between the effects and the number of hydroxyl groups present in these compounds cannot be used as a definitive indication of their activities, these data are in good agreement with previous investigations carried out on activity [16, 17]. In particular, the inhibitory effects of alcohols and ketones against the germination of R. sativus seeds were relatively larger than those encountered against B. rapa.

As shown in Fig. 3, the alcohol containing compounds, isopulegol, (−)-dihydrocarveol, (−)-terpinen-4-ol, and (−)-menthol, all exhibited greater inhibitory effects against the germination of B. rapa seeds than any other p-menthane alcohols. The effect of the isopulegol was similar to that of (−)-menthol, indicating that an isopropenyl group at the C-4 position of the ring was almost equivalent to an isopropyl group at the same position in terms of its effect.

The inhibitory effect of (−)-terpinen-4-ol was also higher than that of (−)-α-terpineol, suggesting that the hydroxyl group at the C-4 position of (−)-terpinen-4-ol was contributing to the inhibition of B. rapa germination to a greater extent than the hydroxyl group at the C-8 position of (−)-α-terpineol. There was, however, no difference in the inhibitory effects of isopulegol and (−)-dihydrocarveol, indicating that a hydroxyl group could be well tolerated at either the C-3 or C-2 positions without any adverse impact on the observed levels of activity.

Reynolds observed that the hydrocarbons showed little inhibitory activity on the germination of L. sativa seeds and correlated this observation with their insolubility in water, whereas monoterpenes bearing hydroxyl groups were on the whole more active [17]. In our previous study, we proposed that the increased water solubility of oxygenated monoterpenoids corresponded well with their increased inhibitory effect on seed germination [13].

It is possible that the performance with regard to the inhibitory effect (B. rapa) was influenced by the position of a hydroxyl group, with hydroxyl groups attached to ring carbons providing higher levels of inhibition than those attached to an isopropyl group. Further work is needed for p-menthane compounds containing hydroxyl groups to thoroughly determine the impact of the relationship between the stereochemistry and the inhibitory effect observed in these compounds.

In a previous report from the literature, the termite (Coptotermes formosanus) mortality within Cinnamomum camphora wood was found to depend predominantly on the camphor content in the wood meal [18]. Ohtani et al. [19] identified α-terpinyl acetate and α-terpineol as the termiticidal (C. formosanus) substances in Chamaecyparis obtusa wood. Carvacrol was also reported to possess potent insecticidal and acaricidal activities against a range of agricultural, stored-product and medical arthropod pests, and was the most active as a termiticide (R.speratus) [20].

In the present study, termite mortality was observed for monoterpenoids bearing particular functional groups, with the termite mortality of monoterpenes bearing phenols being greater than that of those bearing alcohols and ketones. Phenols were also more toxic toward termites than p-cymen-8-ol, and it was recognized that the presence of a phenol hydroxyl group on the benzene ring contributed more to the insecticidal activity than any alcoholic hydroxyl group attached to the isopropyl group.

As shown in Table 2, a comparison of the termite mortalities caused by (−)-terpinen-4-ol and (−)-α-terpineol indicated that a hydroxyl group at the C-4 position of the ring induced greater mortality than a hydroxyl group at C-8 position on the isopropyl group. In addition, (−)-carveol and (−)-dihydrocarveol showed higher levels of activity than 1,8-p-menthadien-4-ol and isopulegol, indicating that the termite mortality of the compounds was affected by the position of the hydroxyl group.

The termite mortalities of (−)-terpinen-4-ol and (−)-menthol were significantly higher than those of 1,8-p-menthadiene-4-ol and isopulegol, indicating that the presence of an isopropyl group at the C-4 position of the ring appeared to induce greater mortality than an isopropenyl group at the same position. The termite mortality of (−)-carveol, however, was found to be similar to that of (−)-dihydrocarveol, and the presence of a double bond at the C-1 position of the ring did not lead to decrease in the level of mortality.

The termiticidal activity of (+)-pulegone was greater than the values found for both (−)-carvone and piperitone, indicating that an external double bond at the C-4 position of the saturated six-membered conjugated with a ketone provided a more potent level of activity than the corresponding unsaturated six-membered ring with an α,β-unsaturated ketone with the double portion internal to the ring.

For the valencenoid derivatives, in terms of their repellant activity against termites (C. formosanus), a reduction of the 1,10-double bond (1,10-dihydronootkatone and tetrahydronootkatone) produced compounds that were more repellent than nootkatone [21]. The isopropenyl group probably did not participate in the binding as evidenced by the absence of a significant difference in the repellent activities of nootkatone, isonootkatone and 11,12-dihydronootkatone [21]. In relation to the suppression of the furylfuramide-induced SOS response activity in the umu test using Salmonellatyphimurium, the strength of suppression was related to the saturated six-membered ring, and the inhibition of the suppressive effect by (+)-menthol was stronger than that of isopulegol, indicating that the presence of an isopropenyl group led to a decrease in the strength of the inhibitory effect [22].

In the present study, the presence of a hydroxyl group in the ring of p-menthane alcohols provided a higher level of termite mortality than a hydroxyl group in the corresponding side chain. Furthermore, alcohols containing a saturated side chain showed a higher mortality level than those with an unsaturated side chain. The mortality of the p-menthane alcohols was affected by the position of the hydroxyl group and the saturation fraction of the functional group in the side chain.

In our previous work, the components, which were classified as p-menthane alcohols, provided an increase in water solubility that corresponded well with the associated reduction in the germination rate and LC50 values of the termites [13]. In this study, p-menthane alcohols, phenols and ketones were bioactive against plant seeds and termites. The results suggested that the bioactivities of these compounds were affected by their solubility in water, which affected their mechanism of action. Although sobrerol is one of the p-menthane alcohols, it exhibited a lower level of bioactivity, suggesting that the bioactivities of the compounds were not only affected by their water solubility, but also by their hydrophobicity.

In conclusion, the bioactivity of p-menthane monoterpenoids was dependent upon the presence and the position of certain functional groups and the degree of saturation fraction in the side chain functional group. A phenolic hydroxyl group was effective for termite mortality, whereas the presence of a carbonyl group provided a strong inhibitory effect on seed germination, and promoted termiticidal activity. In particular, this position was relevant for estimating the performance of p-menthane alcohols because the alcoholic hydroxyl group in the ring had been shown to have an inhibitory effect on seed germination and termite mortality. Although an isopropyl group in the ring also provided an inhibitory effect and termiticidal activity, the presence of an isopropenyl group in the ring showed a particularly marked inhibitory effect. Therefore, the selective bioactivity corresponded to the difference in isopropyl and isopropenyl groups.

More work is needed on other compounds containing the p-menthane skeleton to establish any further correlations between the bioactivity and chemical structure of compounds from this particular structural class.

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Acknowledgments

The authors would like to express their gratitude to Nippon Terpene Chemicals Inc., for providing the p-menthane compounds.

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Affiliations

  1. Faculty of Agriculture, Yamagata University, 1-23 Wakaba-machi, Tsuruoka, Yamagata, 997-8555, Japan

    Nobuhiro Sekine

  2. Institute of Wood Technology, Akita Prefectural University, 11-1 Kaieizaka, Noshiro, Akita, 016-0876, Japan

    Nobuhiro Sekine & Sakae Shibutani

Corresponding author

Correspondence to Nobuhiro Sekine.

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Sekine, N., Shibutani, S. Chemical structures of p-menthane monoterpenes with special reference to their effect on seed germination and termite mortality. J Wood Sci59, 229–237 (2013). https://doi.org/10.1007/s10086-013-1327-5

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Keywords

  • Monoterpenoid
  • p-Menthane
  • Termite
  • Seed
Sours: https://jwoodscience.springeropen.com/articles/10.1007/s10086-013-1327-5

Cyclic and Linear Monoterpenes in Phospholipid Membranes: Phase Behavior, Bilayer Structure, and Molecular Dynamics

Abstract Image

Monoterpenes are abundant in essential oils extracted from plants. These relatively small and hydrophobic molecules have shown important biological functions, including antimicrobial activity and membrane penetration enhancement. The interaction between the monoterpenes and lipid bilayers is considered important to the understanding of the biological functions of monoterpenes. In this study, we investigated the effect of cyclic and linear monoterpenes on the structure and dynamics of lipids in model membranes. We have studied the ternary system 1,2-dimyristoyl-sn-glycero-3-phosphocholine–monoterpene–water as a model with a focus on dehydrated conditions. By combining complementary techniques, including differential scanning calorimetry, solid-state nuclear magnetic resonance, and small- and wide-angle X-ray scattering, bilayer structure, phase transitions, and lipid molecular dynamics were investigated at different water contents. Monoterpenes cause pronounced melting point depression and phase segregation in lipid bilayers, and the extent of these effects depends on the hydration conditions. The addition of a small amount of thymol to the fluid bilayer (volume fraction of 0.03 in the bilayer) leads to an increased order in the acyl chain close to the bilayer interface. The findings are discussed in relation to biological systems and lipid formulations.

Sours: https://pubs.acs.org/doi/10.1021/acs.langmuir.5b00856
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Abstract

The plant kingdom supports an extraordinary chemical diversity, with terpenoids representing a particularly diversified class of secondary (or specialized) metabolites. Volatile and semi-volatile terpenoids in the C10–C20 range are often formed in specialized cell types and secretory structures. In the angiosperm lineage, glandular trichomes play an important role in enabling the biosynthesis and storage (or in some cases secretion) of functionalized terpenoids. The ‘decoration’ of a terpenoid scaffold with functional groups changes its physical and chemical properties, and can therefore affect the perception of a specific metabolite by other organisms. Because of the ecological implications (e.g. plant–herbivore interactions) and commercial relevance (e.g. volatiles used in the flavor and fragrance industries), terpenoid functionalization has been researched extensively. Recent successes in the cloning and functional evaluation of genes as well as the structural and biochemical characterization of enzyme catalysts have laid the foundation for an improved understanding of how pathways toward functionalized monoterpenes may have evolved. In this review, we will focus on an up-to-date account of functionalization reactions present in glandular trichomes.

Carbonyl reductase, cytochrome P450, dehydrogenase, double bond reductase, glandular trichome, hydroxylase, oxidoreductase

Introduction

Terpenoids form the largest class of secondary (or specialized) metabolites in plants, with >50000 known structures according to the Dictionary of Natural Products (http://dnp.chemnetbase.com, last accessed 29 December 2018). This enormous diversity is due to (i) the modular assembly of building blocks into a multitude of (mostly) hydrocarbons ranging from C5 (e.g. isoprene) to large polymers with molecular weights approaching 1000000 mass units (e.g. latex); (ii) rearrangement and elimination reactions modifying these backbones; and (iii) functionalization and conjugation reactions that generate additional structural complexity. In many cases, terpenoids are synthesized and accumulated in specialized cell types (e.g. root cork cells) or anatomical structures [e.g. glandular trichomes (GTs), laticifers, or resin ducts]. A comprehensive coverage of biosynthetic reactions and the relevance of tissue-level pathway localization would doubtlessly fill a whole book. The reader is therefore referred to other reviews for special topics in terpenoid biosynthesis. For the same reasons, we will not be able to cover the biosynthesis of terpenophenolics (e.g. humulene in hops or tetrahydrocannabinol in cannabis) and iridoids (e.g. nepetalactone in catnip). This review article is focused on discussing structure–function relationships that determine the functionalization of monoterpene volatiles synthesized in GTs, which contribute substantially to the chemical diversity in the angiosperm lineage. The article first introduces metabolites, enzymes, and biosynthetic pathways, generally organized by substructures of characteristic monoterpenes. This is followed by a more detailed discussion of biocatalyst properties, arranged by enzyme family. Finally, we discuss the implications of newer findings on our understanding of the evolution of pathways toward monoterpenes. While the examples for this review article are drawn from the literature on GTs, the conclusions are equally relevant to enzymes involved in the biosynthesis of other secondary/specialized metabolites.

Enzyme classes involved in monoterpene biosynthesis

The first committed step in monoterpene biosynthesis is usually catalyzed by a monoterpene synthase, which results in the formation of a parent carbon skeleton from a prenyl diphosphate. These enzymes have been classified by the Enzyme Commission as ‘carbon-oxygen lyases acting on phosphates’ (EC 4.2.3.-) or ‘intramolecular lyases’ (EC 5.5.1.-) (Table 1). Alternatively, prenyl diphosphates can be acted upon by a diphosphoric ester hydrolase (EC 3.1.7.-), thereby generating an acyclic monoterpene alcohol (note that in some cases, the enzymes act on cyclic terpene diphosphates). Hydroxyl groups are often added through the action of cytochrome P450-dependent monooxygenases (CYPs). The Enzyme Commission differentiates ‘oxidoreductases acting on paired donors (with NADH or NADPH serving as one donor)’ (EC 1.14.13.-) or ‘oxidoreductases acting on paired donors (with reduced flavin or flavoprotein serving as one donor)’ (EC 1.14.14.-) (Table 1). Further modifications are carried out by various classes of oxidoreductases, including members of the short-chain dehydrogenase/reductase (SDR) family with NAD+ as preferred acceptor (acting on CH–OH functional groups; EC 1.1.1.-), the SDR family with NADPH as preferred cofactor [these can act on carbonyls (EC 1.1.1.-) or CH=CH double bonds (EC 1.3.1.-)], and the medium-chain dehydrogenase/reductase (MDR) family [which act on CH-OH groups (EC 1.1.1.-) or CH=CH double bonds (EC 1.3.1.-)] (Table 1). In the following paragraphs, we will first provide a more detailed description of the pathways of monoterpene biosynthesis and then discuss the structure–function relationships of the diverse classes of enzymes involved in monoterpene functionalization.

Table 1.

Classification of enzymes involved in the biosynthesis of functionalized monoterpenes in glandular trichomes

Enzyme commission classification . Description . 
1.1.1.- Oxidoreductases acting on the CH–OH group of donors 
1.3.1.- Oxidoreductases acting on the CH–CH group of donors 
1.14.13.- Oxidoreductases acting on paired donors, with incorporation or reduction of molecular oxygen 
(NADH or NADPH as one donor, and incorporation of one atom of oxygen) 
1.14.14.- Oxidoreductases acting on paired donors, with incorporation or reduction of molecular oxygen 
(reduced flavin or flavoprotein as one donor, and incorporation of one atom of oxygen) 
3.1.7.- Diphosphoric monoester hydrolases 
4.2.3.- Carbon-oxygen lyases acting on phosphates 
5.5.1.- Intramolecular lyases 
Enzyme commission classification . Description . 
1.1.1.- Oxidoreductases acting on the CH–OH group of donors 
1.3.1.- Oxidoreductases acting on the CH–CH group of donors 
1.14.13.- Oxidoreductases acting on paired donors, with incorporation or reduction of molecular oxygen 
(NADH or NADPH as one donor, and incorporation of one atom of oxygen) 
1.14.14.- Oxidoreductases acting on paired donors, with incorporation or reduction of molecular oxygen 
(reduced flavin or flavoprotein as one donor, and incorporation of one atom of oxygen) 
3.1.7.- Diphosphoric monoester hydrolases 
4.2.3.- Carbon-oxygen lyases acting on phosphates 
5.5.1.- Intramolecular lyases 

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Table 1.

Classification of enzymes involved in the biosynthesis of functionalized monoterpenes in glandular trichomes

Enzyme commission classification . Description . 
1.1.1.- Oxidoreductases acting on the CH–OH group of donors 
1.3.1.- Oxidoreductases acting on the CH–CH group of donors 
1.14.13.- Oxidoreductases acting on paired donors, with incorporation or reduction of molecular oxygen 
(NADH or NADPH as one donor, and incorporation of one atom of oxygen) 
1.14.14.- Oxidoreductases acting on paired donors, with incorporation or reduction of molecular oxygen 
(reduced flavin or flavoprotein as one donor, and incorporation of one atom of oxygen) 
3.1.7.- Diphosphoric monoester hydrolases 
4.2.3.- Carbon-oxygen lyases acting on phosphates 
5.5.1.- Intramolecular lyases 
Enzyme commission classification . Description . 
1.1.1.- Oxidoreductases acting on the CH–OH group of donors 
1.3.1.- Oxidoreductases acting on the CH–CH group of donors 
1.14.13.- Oxidoreductases acting on paired donors, with incorporation or reduction of molecular oxygen 
(NADH or NADPH as one donor, and incorporation of one atom of oxygen) 
1.14.14.- Oxidoreductases acting on paired donors, with incorporation or reduction of molecular oxygen 
(reduced flavin or flavoprotein as one donor, and incorporation of one atom of oxygen) 
3.1.7.- Diphosphoric monoester hydrolases 
4.2.3.- Carbon-oxygen lyases acting on phosphates 
5.5.1.- Intramolecular lyases 

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Monoterpene biosynthesis in glandular trichomes

p-Menthane monoterpenes are common, functionalized volatile constituents of glandular trichomes (Lange, 2015). The relevant biosynthetic pathways were first elucidated in members of the Lamiaceae. The first committed step in the genus Mentha is the cyclization of geranyl diphosphate to (–)-limonene, which is catalyzed by (–)-limonene synthase (EC 4.2.3.16) (Lange, 2015). At this stage, the pathway diverges, with hydroxylation at C3 in Mentha×piperita (peppermint) [catalyzed by (–)-limonene 3-hydroxylase, EC 1.14.13.47], at C6 in Mentha spicata (spearmint) [catalyzed by (–)-limonene 6-hydroxylase, EC 1.14.14.51], or at C7 in Perilla frutescens (perilla) (catalyzed by perillyl alcohol/perillyl aldehyde synthase, EC 1.14.14.52) (Lange, 2015) (Fig. 1). These enzymes are categorized as CYPs (Table 1). The perilla enzyme catalyzes a two-step oxidation (at C7) to yield (–)-perillyl aldehyde (Fujiwara and Ito, 2017). In spearmint, the oxidation of the secondary alcohol at C6 is catalyzed by (–)-carveol dehydrogenase (EC 1.1.1.243) (Ringer et al., 2005), leading to the formation of (–)-carvone, the principal constituent of the essential oil. In peppermint glandular trichomes, a more complex mixture of p-menthane monoterpenes is generated along a branched pathway that involves (–)-trans-isopiperitenol dehydrogenase (EC 1.1.1.223), (–)-trans-isopiperitenone reductase (EC 1.3.1.82), an as yet unidentified isomerase, (+)-pulegone reductase (EC 1.3.1.81), (+)-menthofuran synthase (EC 1.14.13.104), (–)-menthone:(–)-menthol reductase (EC 1.1.1.207), and (–)-menthone:(+)-neomenthol reductase (EC 1.1.1.208) (Lange, 2015) (Fig. 1). Pathways of the opposite stereochemistry have been described to occur in glandular trichomes of Carum carvi (caraway) [accumulating (+)-carvone] and Schizonepeta tenuifolia (Japanese catnip) [where (–)-pulegone, (+)-menthone, and (–)-isomenthone are the most abundant terpenoids] (Bouwmeester et al., 1998, 1999; Liu et al., 2018) (Fig. 1).

Fig. 1.

Outline of monoterpenoid biosynthesis in plant GTs. Abbreviations and symbols: AAT, alcohol acetyltransferase; IPI, isopulegone isomerase; ISPD, isopiperitenol dehydrogenase; ISPR, isopiperitenone reductase; L3H, limonene 3-hydroxylase; NeDH, nerol dehydrogenase; P, phosphate; PulR, pulegone reductase. Enzyme Commission numbers are given in blue font (right next to reaction arrows). Example species that accumulate certain monoterpenoids are listed in red font.

Fig. 1.

Outline of monoterpenoid biosynthesis in plant GTs. Abbreviations and symbols: AAT, alcohol acetyltransferase; IPI, isopulegone isomerase; ISPD, isopiperitenol dehydrogenase; ISPR, isopiperitenone reductase; L3H, limonene 3-hydroxylase; NeDH, nerol dehydrogenase; P, phosphate; PulR, pulegone reductase. Enzyme Commission numbers are given in blue font (right next to reaction arrows). Example species that accumulate certain monoterpenoids are listed in red font.

Different enantiomeric series of bornane monoterpenes are accumulated in glandular trichomes of Salvia officinalis (culinary sage) and Tanacetum vulgare (tansy). The first committed biosynthetic step in each of these species is catalyzed by a stereospecific terpene synthase [(+)-bornyl diphosphate synthase (EC 5.5.1.8) and (–)-bornyl diphosphate synthase (EC 5.5.1.22) in sage and tansy, respectively] (Croteau and Shaskus, 1985; Wise et al., 2001) (Table 1; Fig. 1). The diphosphate moiety is hydrolyzed by a monoterpenyl diphosphatase (EC 3.1.7.3) in sage (Croteau and Karp, 1979), while no information is available for the reaction in tansy. (+)-Borneol dehydrogenase (EC 1.1.1.198) and (–)-borneol dehydrogenase (EC 1.1.1.227) then catalyze oxidation reactions to produce (+)-camphor (sage) and (–)-camphor (tansy) (Croteau et al., 1978; Croteau and Felton, 1980) (Fig. 1).

Acyclic p-menthane monoterpenoids can be formed from several prenyl diphosphate precursors. Geraniol is produced from geranyl diphosphate by geranyl diphosphate diphosphatase (EC 3.1.7.11) and further oxidized to geranial by geraniol dehydrogenase (EC 1.1.1.183) in GTs of Perilla citriodora (Makino) Nakai (Sato-Masumoto and Ito, 2014b) (Fig. 1). In Persicaria minor (Huds.) Opiz (small smartweed), nerol is formed from neryl diphosphate by the concerted action of monoterpenyl diphosphatase (EC 3.1.7.3) and nerol dehydrogenase (PmNeDH) (Tan et al., 2018). GTs of lavandin (Lavandula×intermedia) accumulate monoterpenyl acetates, which are formed from lavadulyl diphosphate-derived monoterpenols through catalysis by acetyltransferases (LiAAT3 and LiAAT4) (Sarker and Mahmoud, 2015) (Fig. 1). The dehydrogenases mentioned here will be discussed in more detail below, while no mechanistic information is available for monoterpene acetyltransferases (which will therefore not be discussed any further).

Members of the cytochrome P450 family (CYPs)

CYPs form a vast superfamily of hemoproteins (containing Fe-heme as the prosthetic group) that occur across all kingdoms of life, but not in all species (Escherichia coli is a prominent example of a species seemingly lacking CYPs). Plant genomes contain expansive CYP gene families (often with more than a couple of hundred members in a single species) and their gene products constitute the largest class of enzymes with roles in metabolism (Nelson and Werck-Reichhart, 2011). The current nomenclature, which is based on sequence identity and not necessarily biological function, includes the term ‘CYP’, followed by a number (family), then a letter (subfamily), and another number (isoform) (e.g. CYP71D13) (exceptions have been made for some plant CYP subfamilies with extremely large clades) (Nelson and Werck-Reichhart, 2011). While CYPs can catalyze a wide array of reactions, they typically are involved in the activation and heterolytic cleavage of molecular oxygen, with insertion of one of its atoms into a non-activated C–H bond and reduction of the other to form water (monooxygenase function). In plants, CYPs can be localized to the membranes of the endoplasmic reticulum (ER), plastids, or mitochondria. Among the CYPs characterized thus far, only those associated with the ER play roles in specialized metabolism. Conserved features of microsomal CYPs include an N-terminal segment with hydrophobic amino acids (membrane anchor), a hinge region with a cluster of basic residues and a cluster of Pro/Gly residues, and a globular domain that contains several conserved motifs whose residues have functions in the binding of the substrate and prosthetic group [for details on this topic, the reader is referred to specialty reviews (Poulos, 2014)]. The heme-bound oxygen within CYPs is activated by the successive transfer of two electrons from NADPH via the two flavin cofactors of a membrane-bound NADPH-cytochrome P450 oxidoreductase (CPR; systematic name: NADPH-hemoprotein reductase, EC 1.6.2.4) (Fig. 2A). Plant genomes contain small families of CPRs (up to three paralogs; status: June 2018). The domains responsible for the CYP–CPR interaction are conserved so that CPRs from different species or even kingdoms are often able to complement each other functionally (Jensen and Møller, 2010).

Fig. 2.

Regiospecificity of CYPs involved in monoterpene hydroxylation in plant GTs. (A) Simplified scheme outlining electron transfer from cytochrome P450 reductase to CYPs. (B) Structure of (–)-limonene substrate with arrows pointing to positions where different CYPs catalyze hydroxylation reactions. (C–E) Limonene hydroxylase structures, based on structural alignments using CYP101 as a template. Stick diagrams of amino acid residues (color palette: blue for C–C bonds, red for C–O bonds; dark blue for C–N bonds; yellow for C–S bonds; and white for X–H bonds), the heme prosthetic group (pink), the heme iron (orange sphere), the oxygen to be inserted into a C–H bond (red sphere), and the substrate (green) relevant for catalysis in CYP71D15 (C), CYP71D18 (D), and CYP71AT146 (E).

Fig. 2.

Regiospecificity of CYPs involved in monoterpene hydroxylation in plant GTs. (A) Simplified scheme outlining electron transfer from cytochrome P450 reductase to CYPs. (B) Structure of (–)-limonene substrate with arrows pointing to positions where different CYPs catalyze hydroxylation reactions. (C–E) Limonene hydroxylase structures, based on structural alignments using CYP101 as a template. Stick diagrams of amino acid residues (color palette: blue for C–C bonds, red for C–O bonds; dark blue for C–N bonds; yellow for C–S bonds; and white for X–H bonds), the heme prosthetic group (pink), the heme iron (orange sphere), the oxygen to be inserted into a C–H bond (red sphere), and the substrate (green) relevant for catalysis in CYP71D15 (C), CYP71D18 (D), and CYP71AT146 (E).

All CYPs known to be involved in the biosynthesis of terpenoids in GTs belong to the CYP71 clan and have similar sizes of between 54 kDa and 58 kDa (the only exception is CYP706B1, which has a calculated size of ~60 kDa) (Table 2). An important question in CYP enzymology pertains to specificity. Based on currently available information, CYPs acting on monoterpene scaffolds in GTs tend to be fairly specific (with Km values in the lower micomolar range), with significantly lower affinity for alternative substrates (Karp et al., 1990; Bouwmeester et al., 1998, 1999; Wüst et al., 2001).

Table 2.

Enzymes involved in the functionalization of monoterpenes in glandular trichomes

Enzyme commission number . Enzymatic function . Gene/enzyme identifier . Species . Size (Da) (predicted) . Specificity (S, substrate) (P, product) . Km (µM) (substrate) . References . 
Cytochrome P450-dependent monooxygenases
1.14.13.47 (–)-Limonene 3-hydroxylase, CYP71D13 Mentha×piperita56601 S=high; P=high 18 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
(S)-Limonene 3-monooxygenase CYP71D15 Mentha×piperita56532 S=high; P=high 18 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
1.14.14.51 (–)-Limonene 6-hydroxylase, CYP71D18 Mentha spicata56149 S=multi; P=multi 20 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
(old: 1.14.13.48) (S)-Limonene 6-monooxygenase 
1.14.14.53 (+)-Limonene 6-hydroxylase, Not cloned yet Carum carvi (annual) – S=multi; P=high 11.4 [(+)-limonene] Bouwmeester et al. (1998)
(old: 1.14.13.80) (R)-Limonene 6-monooxygenase Not cloned yet Carum carvi (biannual) – S=multi; P=high 14.9 [(+)-limonene] Bouwmeester et al. (1999)
1.14.14.52 (–)-Limonene 7-hydroxylase, CYP71D174 Perilla frutescens(Truncated) S=high; P=multi – Mau et al. (2010)
(old: 1.14.13.49) (S)-Limonene 7-monooxygenase 
(–)-Perillyl alcohol/(–)-perillyl aldehyde synthase CYP71AT146 Perilla frutescens56879 S=high; P=high – Fujiwara and Ito (2017)
(S)-Perillyl alcohol/(S)-perillyl aldehyde synthase (two-step oxidation) 
1.14.13.104 (+)-Menthofuran synthase CYP71A32 Mentha×piperita55360 S=high; P=high – Bertea et al. (2001)
Short-chain dehydrogenase/reductase (SDR) family with NAD+as preferred cofactor
1.1.1.198 or1.1.1.227 Borneol dehydrogenase LiBDH Lavandula x intermedia27270 High (no other alcohols)(de/ee not reported) 53 [borneol] Sarker et al. (2012)
1.1.1.198 (+)-Borneol dehydrogenase Not cloned yet Salvia officinalisNA low (multiple alcohols) 30 [(+)-borneol] Croteau et al. (1978)
1.1.1.227 (–)-Borneol dehydrogenase Not cloned yet Tanacetum vulgareNA Low (multiple alcohols) Not reported Dehal and Croteau (1987)
1.1.1.228 (+)-Sabinol dehydrogenase Not cloned yet Salvia officinalisNA Low (multiple alcohols) Not reported Dehal and Croteau (1987)
n.a. endo-Fenchol dehydrogenase Not cloned yet Foeniculum volgareNA Low (multiple alcohols) Not reported Croteau and Felton (1980)
1.1.1.323 (+)-Thujan-3-ol dehydrogenase Not cloned yet Tanacetum vulgareNA Low (multiple alcohols) Not reported Croteau and Felton (1980)
1.1.1.223 (–)-trans-Isopiperitenol dehydrogenase/ ISPD Mentha x piperita27191 Low regioselectivity (C3 and C6) 72 [(–)-trans-Isopiperitenol] Kjonaas et al. (1985); Ringer et al. (2005)
1.1.1.243 (–)-trans-Carveol dehydrogenase (low activity with other alcohols) 1.8 [(–)-trans-carveol] 
NA (–)-Isopiperitenol dehydrogenase PcPKISPD1 Perilla citriodora (PK-type) 29510 Low (deNot reported Sato-Masumoto and Ito (2014a
NA (–)-cis-Isopiperitenol dehydrogenase PcPTISPD2 Perilla citriodora (PT-type) 29580 High (deNot reported 
1.1.1.275 (+)-trans-Carveol dehydrogenase Not cloned yet Carum carviNA Low (carveol isomers) Not reported Bouwmeester et al. (1998)
NA Alcohol dehydrogenase ADH2 Artemisia annua28130 Low (multiple alcohols) 0.1–8.8 Polichuk et al. (2010)
Short-chain dehydrogenase/reductase (SDR) family with NADPH as preferred cofactor
1.3.1.82 (–)-trans-Isopiperitenone reductase ISPR Mentha×piperita34410 Low (α,β-unsaturated cyclic enones) 1.0 [(–)-isopiperitenone] Ringer et al. (2003); Lygidakis et al. (2016)
1.1.1.207 (–)-Menthol dehydrogenase or MMR Mentha×piperita34070 High (cyclic ketones) 3 [(–)-menthone] Davis et al. (2005)
(–)-Menthone:(–)-menthol reductase Low (de41 [(+)-isomenthone] 
1.1.1.208 (+)-Neomenthol dehydrogenase or MNR Mentha×piperita35722 High (cyclic ketones) 674 [(–)-menthone] Davis et al. (2005); Lygidakis et al. (2016)
(–)-Menthone:(+)-neomenthol reductase Low (de>1000 [(+)-isomenthone] 
Medium-chain dehydrogenase/reductase (MDR) family
1.1.1.183 Geraniol dehydrogenase PcGeDH Perilla citriodora38910 Low (multiple alcohols) Not reported Sato-Masumoto and Ito (2014b
CAD1 Ocimum basilicum38768 Low (multiple alcohols) 72 (geraniol) (reversible) Iijima et al. (2006)
GEDH1 Ocimum basilicum39044 Low (multiple alcohols) 30 (geraniol) (reversible) Iijima et al. (2006)
37 (nerol) (reversible) 
NA Nerol dehydrogenase PmNeDH Persicaria minor39300 High (low activity with other 165 (nerol) Tan et al. (2018)
terpenoid alcohols) 446 (geraniol) 
1.3.1.81 (+)-Pulegone reductase PulR Mentha×piperita37915 High (no activity with other 2.3 [(+)-pulegone] Ringer et al. (2003)
unsataturated carbonyl metabolites) Rios-Estepa et al. (2008)
Other
NA Terpenoid oxidoreductase (SDR familyRed1 Artemisia annua34250 Low (various carbonyl- 7.1 [(–)-menthone] Rydén et al. (2010a, b
containing metabolites) 24 (dihydroartemisinic aldehyde) 
Enzyme commission number . Enzymatic function . Gene/enzyme identifier . Species . Size (Da) (predicted) . Specificity (S, substrate) (P, product) . Km (µM) (substrate) . References . 
Cytochrome P450-dependent monooxygenases
1.14.13.47 (–)-Limonene 3-hydroxylase, CYP71D13 Mentha×piperita56601 S=high; P=high 18 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
(S)-Limonene 3-monooxygenase CYP71D15 Mentha×piperita56532 S=high; P=high 18 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
1.14.14.51 (–)-Limonene 6-hydroxylase, CYP71D18 Mentha spicata56149 S=multi; P=multi 20 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
(old: 1.14.13.48) (S)-Limonene 6-monooxygenase 
1.14.14.53 (+)-Limonene 6-hydroxylase, Not cloned yet Carum carvi (annual) – S=multi; P=high 11.4 [(+)-limonene] Bouwmeester et al. (1998)
(old: 1.14.13.80) (R)-Limonene 6-monooxygenase Not cloned yet Carum carvi (biannual) – S=multi; P=high 14.9 [(+)-limonene] Bouwmeester et al. (1999)
1.14.14.52 (–)-Limonene 7-hydroxylase, CYP71D174 Perilla frutescens(Truncated) S=high; P=multi – Mau et al. (2010)
(old: 1.14.13.49) (S)-Limonene 7-monooxygenase 
(–)-Perillyl alcohol/(–)-perillyl aldehyde synthase CYP71AT146 Perilla frutescens56879 S=high; P=high – Fujiwara and Ito (2017)
(S)-Perillyl alcohol/(S)-perillyl aldehyde synthase (two-step oxidation) 
1.14.13.104 (+)-Menthofuran synthase CYP71A32 Mentha×piperita55360 S=high; P=high – Bertea et al. (2001)
Short-chain dehydrogenase/reductase (SDR) family with NAD+as preferred cofactor
1.1.1.198 or1.1.1.227 Borneol dehydrogenase LiBDH Lavandula x intermedia27270 High (no other alcohols)(de/ee not reported) 53 [borneol] Sarker et al. (2012)
1.1.1.198 (+)-Borneol dehydrogenase Not cloned yet Salvia officinalisNA low (multiple alcohols) 30 [(+)-borneol] Croteau et al. (1978)
1.1.1.227 (–)-Borneol dehydrogenase Not cloned yet Tanacetum vulgareNA Low (multiple alcohols) Not reported Dehal and Croteau (1987)
1.1.1.228 (+)-Sabinol dehydrogenase Not cloned yet Salvia officinalisNA Low (multiple alcohols) Not reported Dehal and Croteau (1987)
n.a. endo-Fenchol dehydrogenase Not cloned yet Foeniculum volgareNA Low (multiple alcohols) Not reported Croteau and Felton (1980)
1.1.1.323 (+)-Thujan-3-ol dehydrogenase Not cloned yet Tanacetum vulgareNA Low (multiple alcohols) Not reported Croteau and Felton (1980)
1.1.1.223 (–)-trans-Isopiperitenol dehydrogenase/ ISPD Mentha x piperita27191 Low regioselectivity (C3 and C6) 72 [(–)-trans-Isopiperitenol] Kjonaas et al. (1985); Ringer et al. (2005)
1.1.1.243 (–)-trans-Carveol dehydrogenase (low activity with other alcohols) 1.8 [(–)-trans-carveol] 
NA (–)-Isopiperitenol dehydrogenase PcPKISPD1 Perilla citriodora (PK-type) 29510 Low (deNot reported Sato-Masumoto and Ito (2014a
NA (–)-cis-Isopiperitenol dehydrogenase PcPTISPD2 Perilla citriodora (PT-type) 29580 High (deNot reported 
1.1.1.275 (+)-trans-Carveol dehydrogenase Not cloned yet Carum carviNA Low (carveol isomers) Not reported Bouwmeester et al. (1998)
NA Alcohol dehydrogenase ADH2 Artemisia annua28130 Low (multiple alcohols) 0.1–8.8 Polichuk et al. (2010)
Short-chain dehydrogenase/reductase (SDR) family with NADPH as preferred cofactor
1.3.1.82 (–)-trans-Isopiperitenone reductase ISPR Mentha×piperita34410 Low (α,β-unsaturated cyclic enones) 1.0 [(–)-isopiperitenone] Ringer et al. (2003); Lygidakis et al. (2016)
1.1.1.207 (–)-Menthol dehydrogenase or MMR Mentha×piperita34070 High (cyclic ketones) 3 [(–)-menthone] Davis et al. (2005)
(–)-Menthone:(–)-menthol reductase Low (de41 [(+)-isomenthone] 
1.1.1.208 (+)-Neomenthol dehydrogenase or MNR Mentha×piperita35722 High (cyclic ketones) 674 [(–)-menthone] Davis et al. (2005); Lygidakis et al. (2016)
(–)-Menthone:(+)-neomenthol reductase Low (de>1000 [(+)-isomenthone] 
Medium-chain dehydrogenase/reductase (MDR) family
1.1.1.183 Geraniol dehydrogenase PcGeDH Perilla citriodora38910 Low (multiple alcohols) Not reported Sato-Masumoto and Ito (2014b
CAD1 Ocimum basilicum38768 Low (multiple alcohols) 72 (geraniol) (reversible) Iijima et al. (2006)
GEDH1 Ocimum basilicum39044 Low (multiple alcohols) 30 (geraniol) (reversible) Iijima et al. (2006)
37 (nerol) (reversible) 
NA Nerol dehydrogenase PmNeDH Persicaria minor39300 High (low activity with other 165 (nerol) Tan et al. (2018)
terpenoid alcohols) 446 (geraniol) 
1.3.1.81 (+)-Pulegone reductase PulR Mentha×piperita37915 High (no activity with other 2.3 [(+)-pulegone] Ringer et al. (2003)
unsataturated carbonyl metabolites) Rios-Estepa et al. (2008)
Other
NA Terpenoid oxidoreductase (SDR familyRed1 Artemisia annua34250 Low (various carbonyl- 7.1 [(–)-menthone] Rydén et al. (2010a, b
containing metabolites) 24 (dihydroartemisinic aldehyde) 

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Table 2.

Enzymes involved in the functionalization of monoterpenes in glandular trichomes

Enzyme commission number . Enzymatic function . Gene/enzyme identifier . Species . Size (Da) (predicted) . Specificity (S, substrate) (P, product) . Km (µM) (substrate) . References . 
Cytochrome P450-dependent monooxygenases
1.14.13.47 (–)-Limonene 3-hydroxylase, CYP71D13 Mentha×piperita56601 S=high; P=high 18 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
(S)-Limonene 3-monooxygenase CYP71D15 Mentha×piperita56532 S=high; P=high 18 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
1.14.14.51 (–)-Limonene 6-hydroxylase, CYP71D18 Mentha spicata56149 S=multi; P=multi 20 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
(old: 1.14.13.48) (S)-Limonene 6-monooxygenase 
1.14.14.53 (+)-Limonene 6-hydroxylase, Not cloned yet Carum carvi (annual) – S=multi; P=high 11.4 [(+)-limonene] Bouwmeester et al. (1998)
(old: 1.14.13.80) (R)-Limonene 6-monooxygenase Not cloned yet Carum carvi (biannual) – S=multi; P=high 14.9 [(+)-limonene] Bouwmeester et al. (1999)
1.14.14.52 (–)-Limonene 7-hydroxylase, CYP71D174 Perilla frutescens(Truncated) S=high; P=multi – Mau et al. (2010)
(old: 1.14.13.49) (S)-Limonene 7-monooxygenase 
(–)-Perillyl alcohol/(–)-perillyl aldehyde synthase CYP71AT146 Perilla frutescens56879 S=high; P=high – Fujiwara and Ito (2017)
(S)-Perillyl alcohol/(S)-perillyl aldehyde synthase (two-step oxidation) 
1.14.13.104 (+)-Menthofuran synthase CYP71A32 Mentha×piperita55360 S=high; P=high – Bertea et al. (2001)
Short-chain dehydrogenase/reductase (SDR) family with NAD+as preferred cofactor
1.1.1.198 or1.1.1.227 Borneol dehydrogenase LiBDH Lavandula x intermedia27270 High (no other alcohols)(de/ee not reported) 53 [borneol] Sarker et al. (2012)
1.1.1.198 (+)-Borneol dehydrogenase Not cloned yet Salvia officinalisNA low (multiple alcohols) 30 [(+)-borneol] Croteau et al. (1978)
1.1.1.227 (–)-Borneol dehydrogenase Not cloned yet Tanacetum vulgareNA Low (multiple alcohols) Not reported Dehal and Croteau (1987)
1.1.1.228 (+)-Sabinol dehydrogenase Not cloned yet Salvia officinalisNA Low (multiple alcohols) Not reported Dehal and Croteau (1987)
n.a. endo-Fenchol dehydrogenase Not cloned yet Foeniculum volgareNA Low (multiple alcohols) Not reported Croteau and Felton (1980)
1.1.1.323 (+)-Thujan-3-ol dehydrogenase Not cloned yet Tanacetum vulgareNA Low (multiple alcohols) Not reported Croteau and Felton (1980)
1.1.1.223 (–)-trans-Isopiperitenol dehydrogenase/ ISPD Mentha x piperita27191 Low regioselectivity (C3 and C6) 72 [(–)-trans-Isopiperitenol] Kjonaas et al. (1985); Ringer et al. (2005)
1.1.1.243 (–)-trans-Carveol dehydrogenase (low activity with other alcohols) 1.8 [(–)-trans-carveol] 
NA (–)-Isopiperitenol dehydrogenase PcPKISPD1 Perilla citriodora (PK-type) 29510 Low (deNot reported Sato-Masumoto and Ito (2014a
NA (–)-cis-Isopiperitenol dehydrogenase PcPTISPD2 Perilla citriodora (PT-type) 29580 High (deNot reported 
1.1.1.275 (+)-trans-Carveol dehydrogenase Not cloned yet Carum carviNA Low (carveol isomers) Not reported Bouwmeester et al. (1998)
NA Alcohol dehydrogenase ADH2 Artemisia annua28130 Low (multiple alcohols) 0.1–8.8 Polichuk et al. (2010)
Short-chain dehydrogenase/reductase (SDR) family with NADPH as preferred cofactor
1.3.1.82 (–)-trans-Isopiperitenone reductase ISPR Mentha×piperita34410 Low (α,β-unsaturated cyclic enones) 1.0 [(–)-isopiperitenone] Ringer et al. (2003); Lygidakis et al. (2016)
1.1.1.207 (–)-Menthol dehydrogenase or MMR Mentha×piperita34070 High (cyclic ketones) 3 [(–)-menthone] Davis et al. (2005)
(–)-Menthone:(–)-menthol reductase Low (de41 [(+)-isomenthone] 
1.1.1.208 (+)-Neomenthol dehydrogenase or MNR Mentha×piperita35722 High (cyclic ketones) 674 [(–)-menthone] Davis et al. (2005); Lygidakis et al. (2016)
(–)-Menthone:(+)-neomenthol reductase Low (de>1000 [(+)-isomenthone] 
Medium-chain dehydrogenase/reductase (MDR) family
1.1.1.183 Geraniol dehydrogenase PcGeDH Perilla citriodora38910 Low (multiple alcohols) Not reported Sato-Masumoto and Ito (2014b
CAD1 Ocimum basilicum38768 Low (multiple alcohols) 72 (geraniol) (reversible) Iijima et al. (2006)
GEDH1 Ocimum basilicum39044 Low (multiple alcohols) 30 (geraniol) (reversible) Iijima et al. (2006)
37 (nerol) (reversible) 
NA Nerol dehydrogenase PmNeDH Persicaria minor39300 High (low activity with other 165 (nerol) Tan et al. (2018)
terpenoid alcohols) 446 (geraniol) 
1.3.1.81 (+)-Pulegone reductase PulR Mentha×piperita37915 High (no activity with other 2.3 [(+)-pulegone] Ringer et al. (2003)
unsataturated carbonyl metabolites) Rios-Estepa et al. (2008)
Other
NA Terpenoid oxidoreductase (SDR familyRed1 Artemisia annua34250 Low (various carbonyl- 7.1 [(–)-menthone] Rydén et al. (2010a, b
containing metabolites) 24 (dihydroartemisinic aldehyde) 
Enzyme commission number . Enzymatic function . Gene/enzyme identifier . Species . Size (Da) (predicted) . Specificity (S, substrate) (P, product) . Km (µM) (substrate) . References . 
Cytochrome P450-dependent monooxygenases
1.14.13.47 (–)-Limonene 3-hydroxylase, CYP71D13 Mentha×piperita56601 S=high; P=high 18 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
(S)-Limonene 3-monooxygenase CYP71D15 Mentha×piperita56532 S=high; P=high 18 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
1.14.14.51 (–)-Limonene 6-hydroxylase, CYP71D18 Mentha spicata56149 S=multi; P=multi 20 [(–)-limonene] Karp et al. (1990); Lupien et al. (1999)
(old: 1.14.13.48) (S)-Limonene 6-monooxygenase 
1.14.14.53 (+)-Limonene 6-hydroxylase, Not cloned yet Carum carvi (annual) – S=multi; P=high 11.4 [(+)-limonene] Bouwmeester et al. (1998)
(old: 1.14.13.80) (R)-Limonene 6-monooxygenase Not cloned yet Carum carvi (biannual) – S=multi; P=high 14.9 [(+)-limonene] Bouwmeester et al. (1999)
1.14.14.52 (–)-Limonene 7-hydroxylase, CYP71D174 Perilla frutescens(Truncated) S=high; P=multi – Mau et al. (2010)
(old: 1.14.13.49) (S)-Limonene 7-monooxygenase 
(–)-Perillyl alcohol/(–)-perillyl aldehyde synthase CYP71AT146 Perilla frutescens56879 S=high; P=high – Fujiwara and Ito (2017)
(S)-Perillyl alcohol/(S)-perillyl aldehyde synthase (two-step oxidation) 
1.14.13.104 (+)-Menthofuran synthase CYP71A32 Mentha×piperita55360 S=high; P=high – Bertea et al. (2001)
Short-chain dehydrogenase/reductase (SDR) family with NAD+as preferred cofactor
1.1.1.198 or1.1.1.227 Borneol dehydrogenase LiBDH Lavandula x intermedia27270 High (no other alcohols)(de/ee not reported) 53 [borneol] Sarker et al. (2012)
1.1.1.198 (+)-Borneol dehydrogenase Not cloned yet Salvia officinalisNA low (multiple alcohols) 30 [(+)-borneol] Croteau et al. (1978)
1.1.1.227 (–)-Borneol dehydrogenase Not cloned yet Tanacetum vulgareNA Low (multiple alcohols) Not reported Dehal and Croteau (1987)
1.1.1.228 (+)-Sabinol dehydrogenase Not cloned yet Salvia officinalisNA Low (multiple alcohols) Not reported Dehal and Croteau (1987)
n.a. endo-Fenchol dehydrogenase Not cloned yet Foeniculum volgareNA Low (multiple alcohols) Not reported Croteau and Felton (1980)
1.1.1.323 (+)-Thujan-3-ol dehydrogenase Not cloned yet Tanacetum vulgareNA Low (multiple alcohols) Not reported Croteau and Felton (1980)
1.1.1.223 (–)-trans-Isopiperitenol dehydrogenase/ ISPD Mentha x piperita27191 Low regioselectivity (C3 and C6) 72 [(–)-trans-Isopiperitenol] Kjonaas et al. (1985); Ringer et al. (2005)
1.1.1.243 (–)-trans-Carveol dehydrogenase (low activity with other alcohols) 1.8 [(–)-trans-carveol] 
NA (–)-Isopiperitenol dehydrogenase PcPKISPD1 Perilla citriodora (PK-type) 29510 Low (deNot reported Sato-Masumoto and Ito (2014a
NA (–)-cis-Isopiperitenol dehydrogenase PcPTISPD2 Perilla citriodora (PT-type) 29580 High (deNot reported 
1.1.1.275 (+)-trans-Carveol dehydrogenase Not cloned yet Carum carviNA Low (carveol isomers) Not reported Bouwmeester et al. (1998)
NA Alcohol dehydrogenase ADH2 Artemisia annua28130 Low (multiple alcohols) 0.1–8.8 Polichuk et al. (2010)
Short-chain dehydrogenase/reductase (SDR) family with NADPH as preferred cofactor
1.3.1.82 (–)-trans-Isopiperitenone reductase ISPR Mentha×piperita34410 Low (α,β-unsaturated cyclic enones) 1.0 [(–)-isopiperitenone] Ringer et al. (2003); Lygidakis et al. (2016)
1.1.1.207 (–)-Menthol dehydrogenase or MMR Mentha×piperita34070 High (cyclic ketones) 3 [(–)-menthone] Davis et al. (2005)
(–)-Menthone:(–)-menthol reductase Low (de41 [(+)-isomenthone] 
1.1.1.208 (+)-Neomenthol dehydrogenase or MNR Mentha×piperita35722 High (cyclic ketones) 674 [(–)-menthone] Davis et al. (2005); Lygidakis et al. (2016)
(–)-Menthone:(+)-neomenthol reductase Low (de>1000 [(+)-isomenthone] 
Medium-chain dehydrogenase/reductase (MDR) family
1.1.1.183 Geraniol dehydrogenase PcGeDH Perilla citriodora38910 Low (multiple alcohols) Not reported Sato-Masumoto and Ito (2014b
CAD1 Ocimum basilicum38768 Low (multiple alcohols) 72 (geraniol) (reversible) Iijima et al. (2006)
GEDH1 Ocimum basilicum39044 Low (multiple alcohols) 30 (geraniol) (reversible) Iijima et al. (2006)
37 (nerol) (reversible) 
NA Nerol dehydrogenase PmNeDH Persicaria minor39300 High (low activity with other 165 (nerol) Tan et al. (2018)
terpenoid alcohols) 446 (geraniol) 
1.3.1.81 (+)-Pulegone reductase PulR Mentha×piperita37915 High (no activity with other 2.3 [(+)-pulegone] Ringer et al. (2003)
unsataturated carbonyl metabolites) Rios-Estepa et al. (2008)
Other
NA Terpenoid oxidoreductase (SDR familyRed1 Artemisia annua34250 Low (various carbonyl- 7.1 [(–)-menthone] Rydén et al. (2010a, b
containing metabolites) 24 (dihydroartemisinic aldehyde) 

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In the context of specificity, a more complete discussion of structure–function relationships for monoterpene hydroxylases acting on (–)-limonene would seem particularly relevant. CYP71D13 and CYP71D15 (both of peppermint) hydroxylate at C3, CYP71D18 of spearmint at C6, CYP71AT146 of perilla at C7, and CYP71D174 of perilla at all three positions (Karp et al., 1990; Mau et al., 2010; Fujiwara and Ito, 2017) (Fig. 2B). Ideally, for structure–function analyses, one would compare crystal structures for these enzymes, but progress with obtaining such data for membrane-bound CYPs has generally been extremely slow (with no success for the CYPs of interest here). As an alternative, structural alignment can be considered, an approach that takes advantage of the fact that, despite the limited sequence identity of CYPs across species, there appears to be a common overall topology and three-dimensional fold, thereby enabling the inference of three-dimensional structure by superposition of the atomic co-ordinate set based on a known structure. The well-characterized CYP101 (from Pseudomonas putida) (Poulos et al., 1987), which hydroxylates the monoterpene camphor, can serve as a template for modeling structures of CYPs involved in the biosynthesis of functionalized terpenes. A structural model of CYP71D15 indicated the placement of the heme prosthetic group close to the residues that constitute the highly conserved heme-binding motif (P429-F430-X-X-G-X-R-X-C437-X-G) (Fig. 2C). The active site was predicted, based on binding energies calculated for various docking scenarios, to constrain the available space in favor of an orientation of the (–)-limonene substrate in which the C3–H bond is situated very close (distance <2 Å) to the heme-bound oxygen (visualized in Fig. 2C as ‘compound I’ intermediate [iron(IV) oxo species with an additional oxidizing equivalent delocalized over the porphyrin and thiolate ligands]). The same docking approach with CYP71D18 indicated favorably low binding energies for (–)-limonene in an orientation that places the C6–H bond in close proximity to the iron-oxo group (Fig. 2D). Finally, structural modeling with CYP71AT146 provided evidence for low binding energies for a (–)-limonene orientation where the C7–H bond is pointing toward the iron-oxo group of the heme complex (Fig. 2E). In summary, in silico docking experiments indicate that the spatial and electronic constraints of the active site of these CYPs favor a certain orientation of the substrate in relation to the reactive iron-oxo porphyrin prosthetic group, which then determines the regiospecificity of the catalyzed hydroxylation reaction. Thus far, there has only been one published report that explored the involvement of specific amino acid residues in controlling regiospecificity. A single amino acid substitution (F363I) in CYP81D18 led to a complete conversion of a 6-hydroxylase to a 3-hydroxylase activity, whereas the reciprocal I364F modification of CYP71D15 (attempting to convert a 3-hydroxylase into a 6-hydroxylase) resulted in an inactive enzyme (despite proper folding) (Schalk and Croteau, 2000). These observations illustrate that there is much to be learned about the determinants of regiospecificity in CYPs that act on monoterpene substrates, with homology modeling presenting a valuable tool to guide such endeavors.

Members of the short-chain dehydrogenase/reductase (SDR) superfamily

Short-chain dehydrogenases/reductases (SDRs) are a superfamily of nicotinamide cofactor-dependent enzymes with significant sequence and functional divergence but surprisingly high structural homology, with a characteristic Rossman βαβ fold (Vidal et al., 2018). The electrostatic environment created by the residues of the cofactor-binding motif determines the preference for NAD+/NADH or NADP+/NADPH, and these structure–function relationships are discussed in more detail below.

Members of the SDR superfamily preferring an NAD+ cofactor

Conserved characteristics of SDRs that employ NAD+ as the preferred cofactor are the presence of motif I (TGXXXGXG; important for coenzyme binding), motif II (NAG), the active site residues with motif III (YXXXK), and a conserved aspartate residue that is indicative of the preference for NAD+ over NADP+ (negatively charged residues favor the NAD+/NADH pair) (Moummou et al., 2012) (Fig. 3A). Ringer et al. (2005) suggested that N118, S147, Y160, and K164 may constitute a catalytic tetrad in peppermint (–)-trans-isopiperitenol dehydrogenase/(–)-trans-carveol dehydrogenase (MpISPD). Members of this enzyme class with demonstrated roles in monoterpene biosynthesis in GTs have comparable molecular weights (27–30 kDa) (Table 2). Predictions of subcellular localization based on sequence properties tend to generate conflicting results. Interestingly, MpISPD was demonstrated experimentally to localize to mitochondria of GTs (Turner and Croteau, 2004). It is currently unknown if the mitochondrial SDR localization is a unique feature of peppermint GTs or a more common phenomenon in the biosynthesis of functionalized monoterpenoids. In this context, it should be noted that other oxidoreductases of the monoterpenoid pathway in peppermint GTs (discussed in more detail below) localize to the cytosol (Turner and Croteau, 2004; Turner et al., 2012).

Fig. 3.

Members of the SDR superfamily of dehydrogenases/reductases with preference for an NAD+ cofactor have important roles in oxidizing terpenoid alcohols in GTs. (A) Sequence features. (B) Stereospecificity of various isopiperitenol dehydrogenase isoforms (for details see text). (C) Ribbon diagram of carveol dehydrogenase from Mycobacterium (beige color) overlayed with the predicted structure (based on homology modeling) of peppermint isopiperitenol dehydrogenase (teal color). A stick diagram is used to depict the NAD+ cofactor (centrally located) and a potential ligand [(4S)-2-methyl-2,4-pentanediol; in purple color].

Fig. 3.

Members of the SDR superfamily of dehydrogenases/reductases with preference for an NAD+ cofactor have important roles in oxidizing terpenoid alcohols in GTs. (A) Sequence features. (B) Stereospecificity of various isopiperitenol dehydrogenase isoforms (for details see text). (C) Ribbon diagram of carveol dehydrogenase from Mycobacterium (beige color) overlayed with the predicted structure (based on homology modeling) of peppermint isopiperitenol dehydrogenase (teal color). A stick diagram is used to depict the NAD+ cofactor (centrally located) and a potential ligand [(4S)-2-methyl-2,4-pentanediol; in purple color].

The substrate specificity of monoterpenoid pathway-related SDRs varies considerably. Borneol dehydrogenase from Lavandula×intermedia (lavandin) converted borneol to camphor (with a Km in the low micromolar range) and did not accept other monoterpene alcohols as substrates (specificity for borneol diastereomers was not tested) (Sarker et al., 2012). The affinity of MpISPD was very high for C6-hydroxylated (–)-trans-carveol (Km=1.8 µM) and also quite high for C3-hydroxylated (–)-trans-isopiperitenol (Km=72 µM). The reaction velocity for these reactions, however, was rather slow [kcat of 0.02 s−1 and 0.002 s−1 for (–)-trans-carveol and (–)-trans-isopiperitenol, respectively] (Table 2). Other monoterpene alcohols tested at saturation, with NAD+ as cofactor, exhibited even lower relative velocities (Kjonaas et al., 1985; Ringer et al., 2005). One ISPD isoform of Perilla citriodora (PcPKISPD1) was reported to accept both (–)-isopiperitenol diastereomers, whereas another (PcPTISPD2) was specific for the conversion of (–)-cis-isopiperitenol (no kinetic values reported) (Sato-Masumoto and Ito, 2014b). The reactions catalyzed by these enzymes, based on in vitro assays, were in some cases reversible [reduction of (–)-trans-isopiperitenone with NADH as cofactor], with PcPKISPD1 being an example, whereas others (MpISPD and PcPTISPD2) did not function measurably in reverse mode as reductases (Ringer et al., 2005; Sato-Masumoto and Ito, 2014a) (Fig. 3B). (+)-trans-Carveol dehydrogenase of caraway accepted several carveol isomers as substrates (no data for other monoterpene alcohols were reported) (Bouwmeester et al., 1998) (Table 2). Partially purified fractions with SDR activity from various aromatic plants (Salvia officinalis, Tanacetum vulgare, and Foeniculum vulgare) were active with multiple monoterpene alcohols (Croteau et al., 1978; Croteau and Felton, 1980; Dehal and Croteau, 1987), but these data sets (obtained with crude preparations containing enzyme mixtures) need to be interpreted with caution. A dehydrogenase from sweet wormwood (ADH2), which was produced recombinantly in a microbial host and subsequently purified, converted several monoterpene alcohols to the corresponding carbonyl metabolites (Table 2) (Polichuk et al., 2010).

Crystal structures of NAD+-dependent SDRs with relevance for monoterpenoid functionalization in plants are not yet available. However, some microbes have been demonstrated to be capable of oxidizing monoterpene alcohols, and crystal structures, at ~2 Å resolution, of seven enzymes with carveol dehydrogenase activity from Mycobacterium species (all co-crystallized with a nicotinamide cofactor) were reported recently (Haft et al., 2017). The structural information on Mycobacterium carveol dehydrogenases was employed in homology modeling to predict the structure of MpISPD bound to NAD+ (Fig. 5C). However, these enzyme conformations did not accommodate the docking of relevant substrates, such as (–)-trans-isopiperitenol or (–)-trans-carveol, with reasonable binding energies, primarily due to insufficient space close to where a hydride would need to be transferred to the nicotinamide moiety of the cofactor. In this context, it is important to note that in only one case was a second ligand [(4S)-2-methyl-2,4-pentanediol] co-crystallized with a Mycobacterium carveol dehydrogenase (in addition to the cofactor), and, based on the reported crystal structure, this molecule (which is not a natural substrate) was not present in the active site of the enzyme but rather in an area closer to the enzyme surface (at >6 Å distance from C4 of NAD+, which is where the hydride transfer occurs) (Fig. 3C). It is conceivable, but at this point mere speculation, that an induced fit model of substrate binding (Plapp, 2010) would be appropriate to explain these observations. However, to enable the testing of such a hypothesis, an ISPD crystal structure in which both the cofactor and a relevant substrate analog are present inside the active site would be highly desirable.

Members of the SDR family preferring an NADPH cofactor

Three SDRs that prefer NADPH as a cofactor were cloned from peppermint GTs and functionally characterized as recombinant proteins. One of these acts as a double bond reductase [(–)-trans-isopiperitenone reductase (MpISPR)], while the other two function as carbonyl reductases [(–)-menthone:(–)-menthol reductase (MpMMR) and (–)-menthone:(+)-neomenthol reductase (MpMNR)] (Ringer et al., 2003; Davis et al., 2005) (Fig. 1). These reductases share significant sequence identity (73% for MpMMR/MpMNR, 66% for MpMNR/MpISPR, and 64% for MpMMR/MpISPR), but are only distantly related to MpISPD, a dehydrogenase mentioned above (12–13% identity). The sequences of conserved motifs I and II are identical across the three reductases, as are the residues required for NADPH binding [N15, K16, R37, and R41 (numbering of MpISPR)] (Fig. 4A)]. Interestingly, motif III is different in MpISPR (ERVSK) from that of MpMMR and MpMNR (YKVSK). Of particular note is residue E238 (MpISPR numbering), which corresponds to a highly conserved tyrosine (Y244 in MpMNR) in other reductases of the same class (Ringer et al., 2003) (Fig. 4A). The implications of this amino acid difference are discussed in more detail below. The molecular weights are comparable among these reductases (34–36 kDa), which are larger than those of the dehydrogenases in the previous section. A carbonyl reductase (termed Red1) with considerable sequence identity to the mint reductases was cloned from Artemisia annua (Rydén et al., 2010a, b) but, because of its relatively low specificity (accepts a range of carbonyl-containing monoterpenes and sesquiterpenes), will not be discussed in detail here.

Fig. 4.

Members of the SDR superfamily of dehydrogenases/reductases with preference for NADPH as cofactor can be involved in double bond and carbonyl reduction reactions in GTs. (A) Sequence features. (B) Ribbon diagram of ISPR. (C) Stick diagram of amino acid residues (color palette: blue for C–C bonds, red for C–O bonds; dark blue for C–N bonds; and white for X–H bonds), the cofactor (pink color for C–C bonds), and the substrate (green) relevant for catalysis within ISPR. (D) Ribbon diagram of MNR. (E) Stick diagram of amino acid residues, cofactor, and substrate (color palette as above) relevant for catalysis within MNR. Proposed electron transfers for reductions are indicated by gray arrows.

Fig. 4.

Members of the SDR superfamily of dehydrogenases/reductases with preference for NADPH as cofactor can be involved in double bond and carbonyl reduction reactions in GTs. (A) Sequence features. (B) Ribbon diagram of ISPR. (C) Stick diagram of amino acid residues (color palette: blue for C–C bonds, red for C–O bonds; dark blue for C–N bonds; and white for X–H bonds), the cofactor (pink color for C–C bonds), and the substrate (green) relevant for catalysis within ISPR. (D) Ribbon diagram of MNR. (E) Stick diagram of amino acid residues, cofactor, and substrate (color palette as above) relevant for catalysis within MNR. Proposed electron transfers for reductions are indicated by gray arrows.

MpISPR has the lowest Km value (1 µM) for its natural substrate [(–)-trans-isopiperitenone] (Ringer et al., 2003) but also accepts other α,β-unsaturated cyclic enones for double bond reductions (Lygidakis et al., 2016). The reverse activity [oxidation of (+)-cis-isopulegone with NADP+ as cofactor] was essentially undetectable in in vitro assays (Ringer et al., 2003). In peppermint GTs, MpMMR converts (–)-menthone and (+)-isomenthone into (–)-menthol and (+)-neoisomenthol, respectively (Ringer et al., 2005). MpMNR uses the same substrates as MpMMR [converting them to (+)-neomenthol and (+)-isomenthol, respectively], but with significantly lower affinity [Km values for (–)-menthone and (+)-isomenthone are 674 µM and >1000 µM, respectively]. Both enzymes accept the optical antipode [(+)-menthone] of their primary substrate (Ringer et al., 2005). However, conversions with other, structurally related, cyclic ketones are very slow or undetectable, and there is no measurable double bond reduction activity (Lygidakis et al., 2016). The oxidation of menthol diastereomers with NADP+ as cofactor was mostly undetectable, with the exception of the conversion of (+)-neomenthol to (–)-menthone by MpMNR, but the Km value for this reaction was extremely high (millimolar range) (Davis et al., 2005).

Crystal structures for peppermint MpISPR and MpMNR, in combination with the nicotinamide cofactor, were solved recently (Lygidakis et al., 2016) and have provided the framework to further our understanding of the mechanistic differences that underlie keto reduction and double bond reduction, respectively, in fairly recently diverged SDRs. Based on in silico docking experiments that explore the orientation of the ligand with the lowest calculated binding energy, Cβ of (–)-trans-isopiperitenone is positioned very close (at a distance of ~3 Å) to the 4-pro-S hydride to be transferred from NADPH, thereby initiating a proposed cascade (Lygidakis et al., 2016) of proton transfers involving the carboxyl group of E238 (with a stabilizing role for the hydroxyl group of S182), an active site water molecule (H2O-1), the ribose moiety of NADPH, the amine group of L242, and a second active site water molecule (H2O-2) in MpISPR (Fig. 4B, C). In this proposed mechanism, E238, a residue that is different from the conserved tyrosine found in the equivalent position in keto reductases (Y244 in MpMNR), would serve to stabilize, by hydrogen bonding with its carboxyl group, the enolate anion intermediate forming at C2–C3, which thereby enables a double bond reduction. To complete the double bond reduction, E238 has been proposed to abstract a proton from the enolate intermediate, while the enolate double bond accepts a proton from water, thereby reforming the carbonyl group and leaving a reduced bond at C1–C2 (Lygidakis et al., 2016).

In docking experiments with MpMNR, the substrate [(–)-menthol] is predicted to assume an orientation that places the C3 carbonyl group (rather than a double bond as in MpISPR) close to the 4-pro-S hydride to be transferred from the NADPH cofactor (orientation with lowest binding energy depicted in Fig. 4D). Both (–)-menthone (with S-configuration at C-4) and (+)-isomenthone (with R-configuration at C-4) (Fig. 1) can be accommodated as substrates. It has been proposed that the hydroxyl group of the conserved Y244 residue might be part of a proton transfer relay that also involves the ribose moiety of NADPH, L248, and active site water molecule(s) in MpMNR (Lygidakis et al., 2016) (Fig. 4E). The mechanistically most relevant difference of MpISPR and MpMNR active site residues (E238 in the former, Y244 in the latter) has been further investigated by mutational exchange. The E238Y variant of MpISPR was demonstrated to be capable of catalyzing carbonyl reduction reactions with several ligands, including the MpMNR substrates (–)-menthone and (+)-isomenthone, but the yields were very low (with loss of stereoselectivity), even in 24 h incubations. A double bond reduction activity of the reciprocal Y244E variant of MpMNR was not detectable with various tested substrates (Lygidakis et al., 2016). These results indicate that, while highly intriguing, there is still much to learn about the determinants of carbonyl versus double bond reduction activities in SDRs.

Members of the medium-chain dehydrogenase/reductase (MDR) superfamily preferring an NADP+ cofactor

Zn2+-dependent alcohol dehydrogenases of the medium-chain dehydrogenase/reductase (MDR) superfamily have prominent roles in the oxidation of geraniol and nerol, which are cis/trans isomeric monoterpene alcohols found in GTs of several plant lineages. Functionally characterized members of this class include geraniol dehydrogenases of the Lamiaceae (e.g. PcGeDH from Perilla citriodora (Sato-Masumoto and Ito, 2014b) and GEDH1 from Ocimum basilicum (Iijima et al., 2006)] and nerol dehydrogenase of Persicaria minor (PmNeDH) (Tan et al., 2018). These enzymes share a characteristic sequence motif close to the N-terminus (GHEXXGXXXXXGXXV; residues 72–86 in PmNeDH) (Fig. 5A). Other conserved motifs [GXGXXG (residues 192–197 in PmNeDH) and STSXXK (residues 215–220 in PmNeDH)] play important roles in the binding of the cofactor (NADP+).

Fig. 5.

Members of the Zn2+-containing MDR superfamily of dehydrogenases/reductases with demonstrated roles in monoterpene alcohol oxidation in GTs. (A) Sequence features. Ribbon diagrams of NeDH from Persicaria minor, with nerol (B) or geraniol (C) docked as a substrate (both in green). The NADP+ cofactor is depicted with pink C–C bonds.

Fig. 5.

Members of the Zn2+-containing MDR superfamily of dehydrogenases/reductases with demonstrated roles in monoterpene alcohol oxidation in GTs. (A) Sequence features. Ribbon diagrams of NeDH from Persicaria minor, with nerol (B) or geraniol (C) docked as a substrate (both in green). The NADP+ cofactor is depicted with pink C–C bonds.
Sours: https://academic.oup.com/jxb/article/70/4/1095/5280677
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Sours: https://www.eurekaselect.com/178506/article

Structure monoterpenes

Regular Monoterpenes and Sesquiterpenes (Essential Oils)

Natural Products pp 2973-3008 | Cite as

  • Remigius ChizzolaEmail author
Reference work entry

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Abstract

Monoterpenes resulting from the combination of two and sesquiterpenes from three branched, unsaturated C5 units (isoprene) represent a large class of natural products with a wide range of biological activities. They include unsaturated hydrocarbons and their oxidation products as alcohol, aldehydes, ketones, and rarely ethers. Volatile compounds are constituents of essential oils that are accumulated by numerous plants in special structures in the tissue. Besides the conspicuous aromatic properties, a wide range of biological activities has been documented allowing a wide field of applications. In an ecological context, mono- and sesquiterpenes play an important role in the relations between organisms, for example, as attractants of pollinators or deterrents of herbivores. On a large-scale, monoterpene emissions from vegetation in nature can have ecosystem-wide influences. The further investigation and documentation of this high biodiversity and its sustainable use remains a promising task. This requires the further development of analytical and production techniques and the exact definition and characterization of the plant sources.

Keywords

Aroma compounds Biological activity Essential oil Monoterpene Sesquiterpene 

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Copyright information

© Springer-Verlag Berlin Heidelberg 2013

Authors and Affiliations

  1. 1.Institute of Animal Nutrition and Functional Plant CompundsUniversity of Veterinary Medicine ViennaViennaAustria
Sours: https://link.springer.com/10.1007/978-3-642-22144-6_130
Recognizing Terpenes

Standardised comparison of limonene-derived monoterpenes identifies structural determinants of anti-inflammatory activity

Abstract

Mint species are widely used in traditional and conventional medicine as topical analgesics for osteoarthritic pain and for disorders of the gastrointestinal and respiratory tracts which are all associated with chronic inflammation. To identify the structural determinants of anti-inflammatory activity and potency which are required for chemical optimization towards development of new anti-inflammatory drugs, a selected group of monoterpenes especially abundant in mint species was screened by measuring bacterial lipopolysacharide (LPS)-induced nitric oxide (NO) production in murine macrophages. Nine compounds significantly decreased LPS-induced NO production by more than 30%. IC50 values were calculated showing that the order of potency is: (S)-(+)-carvone > (R)-(−)-carvone > (+)-dihydrocarveol > (S)-8-hydroxycarvotanacetone > (R)-8-hydroxycarvotanacetone > (+)-dihydrocarvone > (−)-carveol > (−)-dihydrocarveol > (S)-(-)-pulegone. Considering the carbon numbering relative to the common precursor, limonene, the presence of an oxygenated group at C6 conjugated to a double bond at C1 and an isopropenyl group and S configuration at C4 are the major chemical features relevant for activity and potency. The most potent compound, (S)-(+)-carvone, significantly decreased the expression of NOS2 and IL-1β in macrophages and in a cell model of osteoarthritis using primary human chondrocytes. (S)-(+)-carvone may be efficient in halting inflammation-related diseases, like osteoarthritis.

Introduction

Inflammation is an orchestrated physiological response elicited by exogenous inducers such as infectious agents, allergens, irritants and toxic compounds, as well as by endogenous triggers released from stressed or damaged tissues/cells1. Although aiming at restoring homeostasis, inflammation has the potential to cause tissue damage and perpetuate itself2. Likewise, inflammation has been reported as an important component associated with most chronic human diseases, such as rheumatic3, metabolic and neurodegenerative diseases and cancer2,4. Due to the increased incidence of these diseases in relation with population aging and the lack of efficacy and adverse side effects of currently available anti-inflammatory drugs more directed to acute inflammation, new therapeutic agents are needed to contend chronic inflammation-associated diseases5,6,7.

Natural products are increasingly used for their anti-inflammatory properties and as sources of new anti-inflammatory compounds5,6. Among the species most widely used, those of the family Lamiaceae, genus Mentha L., commonly designated as mint species, are widely used in traditional8 and conventional medicine, especially as essential oils. These are well-known for anti-inflammatory, antimicrobial, carminative, antispasmodic and analgesic properties. Among several chemical classes identified in mint essential oils, monoterpenes belonging to the limonene synthase pathway, such as menthol, menthone, pulegone and carvone, are especially abundant8. Some components of this group of monoterpenes have been reported to possess anti-inflammatory activity9 that may justify, at least in part, the beneficial effects attributed to mint species by traditional and conventional medicine10,11. However, mint species exhibit many different chemotypes with significant diversity in qualitative and quantitative chemical composition11,12 that causes substantial variability, although poorly characterized, in terms of pharmacological activity of distinct plants and their essential oils. Besides differences related to distinct chemotypes, disparities in the experimental design, namely concerning the range of concentrations tested and the cell and animal models and inflammatory stimuli used, also make comparisons or prediction of the efficacy and potency of different plants, their essential oils and individual compounds impossible. This heterogeneity also makes it impossible to identify the structural determinants of activity, that is the structure-activity relationship (SAR) of this class of natural compounds.

The chemical optimization of an active compound requires that knowledge and is essential to improve its physicochemical properties and/or increase its potency and safety, thus yielding a suitable lead. This is especially important for monoterpenes whose volatility is a major drawback significantly limiting their use as active ingredients for the large scale production of medicines. Hence, elucidating the SAR is essential to guide the chemical modification of these compounds, namely to lower their vapour pressure at room temperature, without compromising pharmacological activity and/or increasing toxicity, and therefore to enable their progression towards new therapeutic agents13. Further, such knowledge is also essential to explain the different anti-inflammatory properties and potency of distinct mint chemotypes and their essential oils and can be used to predict the therapeutic potential of a given product based on its chemical composition.

Thus, the purpose of this study was to assess, under standardized conditions, the anti-inflammatory activity of a selected group of monoterpenes belonging to the limonene synthase pathway that are abundant in mint species (Fig. 1a) and to compare the potency of the active ones by determining their half-maximal inhibitory concentrations (IC50). These data were then correlated with structural features to identify chemical determinants of activity and potency useful to enable chemical optimization of the active compounds.

Structures of the monoterpenes tested. (a) Selected commercially available limonene-derived monoterpenes found in Mentha spp. (b) non-limonene-derived monoterpenes and (c) semi-synthetic limonene-derived monoterpenes were used to elucidate the role of specific chemical features. Stereochemistry of each chiral centre is indicated only where enantiomerically pure compounds were used. The numbering system employed here is based on compound 1.

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For this, the ability of the test compounds to inhibit the production of nitric oxide (NO), a potent and destructive inflammatory mediator14,15,16, induced by bacterial lipopolysaccharide (LPS) in the mouse macrophage cell line, Raw 264.7, was used as a well-established primary screening assay for the identification of small molecules with anti-inflammatory activity17,18. Then and to further confirm their anti-inflammatory activity, we determined the ability of the two most potent compounds to inhibit the expression of NO synthase 2 (NOS2), the enzyme that produces large amounts of NO in response to inflammatory stimuli15,19, and interleukin-1β (IL-1β), two critical inflammatory mediators strongly associated with several acute and chronic human inflammatory diseases3,16,20.

Finally, the most potent compound identified in macrophages, S-(+)-carvone (4), was tested in primary human chondrocyte cultures treated with the pro-inflammatory and catabolic cytokine, IL-1β, as a widely used cell model of osteoarthritis (OA)21. This is the most common musculoskeletal disease, causing pain and loss of mobility and quality of life to millions of people worldwide22. While no curative therapies are yet available23,24 essential oils from Mentha spicata, which is especially abundant in S-(+)-carvone (4), are broadly used to decrease osteoarthritic pain25. Therefore, we hypothesized that such analgesic effect can be, at least in part, secondary to the anti-inflammatory properties of that compound. To test this hypothesis and evaluate its potential as an anti-osteoarthritic drug, we determined whether S-(+)-carvone (4) is also effective in reducing inflammatory responses in human chondrocytes.

Results

Nine out of twenty-one test compounds inhibit LPS-induced NO production in Raw 264.7 macrophages

Commercially available compounds with substituents in specific positions of the p-menthane skeleton in limonene were selected for the primary screening assay (Fig. 1a). We also tested four unrelated natural compounds (Fig. 1b), as well as two semi-synthetic carvone derivatives (Fig. 1c), to further elucidate the relevance to anti-inflammatory activity of specific chemical features of the limonene-derived compounds tested.

Various concentrations of the test compounds, in the absence or presence of LPS, were first evaluated for cytotoxicity using the resazurin reduction assay26 (Figs. S1 and S2). Concentrations above 400 µg/mL (approximately 2600 µM on average) were found not to be completely miscible in aqueous solution, even in the presence of 0.1% DMSO, and so this was the maximal concentration tested. Cytotoxicity was defined, according to the standard for cytotoxicity assessment, ISO 10993-527, as the highest concentration that did not decrease cell viability by more than 30% relative to cells treated with LPS alone. Non-cytotoxic concentrations of each compound were then selected for the screening assay and subsequent studies.

To confirm the quality of the screening assay, we used Bay 11–7082, a selective IκB Kinase inhibitor that abrogates NF-κB activation28 and the expression of its target genes, including NOS229, as a pharmacological control. Pre-treatment with 5 μM Bay 11-7082 decreased NO production to 48.1 ± 5.2% relative to cells treated with LPS alone, as expected.

At non-cytotoxic concentrations, none of the compounds tested affected basal NO production when added to macrophage cultures in the absence of LPS (data not shown) and eight (1, 2, 10, 11, 12, 14, 15 and 16) also had no effect on LPS-induced NO production (Table 1). Thirteen compounds were found to elicit a statistically significant decrease of LPS-induced NO production at the highest concentration tested, but of these, four (3, 17, 18 and 19) had only a negligible effect of less than 20%. The other nine compounds (4, 5, 6, 7, 8, 9, 13, 20 and 21) elicited a robust inhibition greater than 30% (Table 1) and, therefore, were selected for further studies.

Full size table

Thus and to compare the active compounds in terms of potency, the respective concentration required to inhibit NO production by 50% (IC50) was determined by testing further non-cytotoxic concentrations. Since the maximal inhibition achieved with the highest non-cytotoxic concentration of (S)-(−)-pulegone (13) tested, did not exceed 36%, the IC50 for this compound was not determined. Results in Table 2 show that the order of potency of the remaining 8 active compounds is (S)-(+)-carvone (4)> (R)-(−)-carvone (5) ≫ (+)-dihydrocarveol (8)> (S)-8-hydroxycarvotanacetone (20)> (R)-8-hydroxycarvotanacetone (21)> (+)-dihydrocarvone (7)> (−)-carveol (6)> (−)-dihydrocarveol (9).

Full size table

Identification of chemical features relevant for activity by correlation with potency

Having determined the order of potency of the active compounds, we then correlated those results with structural features of all compounds tested to identify the relevant structural determinants of anti-inflammatory activity. For this, we defined a carbon numbering system applicable to all compounds tested, since their different functional groups and application of IUPAC rules would lead to different numbering of the same carbon atoms. Thus, IUPAC rules were used to define carbon numbering for limonene (Fig. 1) and the resulting numbering sequence was applied to all test compounds without considering their specific substituents. Besides limonene-derived compounds, four other natural monoterpenes, β-myrcene (16), p-cymene (17), carvacrol (18) and thymol (19) (Fig. 1b), were tested mainly to assess the relevance of the rigidity or flexibility of the molecule for anti-inflammatory activity. Additionally, two carvone derivatives (20 and 21, Fig. 1c) were synthesized and tested to assess the relevance of the isopropenyl group at C4.

A functional oxygenated group, either a carbonyl or a hydroxyl group, at C6 is present in all active compounds (4-9, 20 and 21) and absent (1 and 2) or present at other positions (3, 1012, 14 and 15) in all inactive or only slightly active (below 20% inhibition at the maximal non-cytotoxic concentration tested) compounds with the exception of (S)-(−)-pulegone (13) which bears a carbonyl group at C3 and showed weak activity.

Another important feature for activity seems to be the presence of an α,β double bond at C1, since its absence is the only difference between (+)-dihydrocarvone (7) and the much more potent carvone enantiomers (4 and 5). Moreover, the conjugation of this double bond to the carbonyl group at C6 also seems relevant for activity since the two most potent compounds, (S)-(+)-carvone (4) and (R)-(−)-carvone (5), present this feature which is also present in their derivatives (20 and 21), but not in the other less potent compounds. Nonetheless, (R)-(+)-pulegone (12) and (S)-(−)-pulegone (13) which have no or little activity, also have an α, β double bond conjugated to a carbonyl group, but involving the carbonyl group at C3 and the double bond at C4. Thus, the localization of the conjugated double bond and carbonyl group seems especially relevant for activity. Nevertheless, while (R)-(+)-pulegone (12) is inactive, its S enantiomer (13) showed weak activity, indicating that the stereochemistry can be relevant for activity.

Then and to elucidate the relevance of the isopropenyl group at C4, present in six active compounds (4, 5, 6, 7, 8 and 9), but also in five inactive ones (1, 2, 3, 10 and 11), we synthesized derivatives of the two most potent compounds, the carvone enantiomers, where that group was replaced by a 2-hydroxyisopropanyl group. The 8-hydroxycarvotanacetone enantiomers (20 and 21) synthesized showed significant activity, although much lower than the respective parent compounds (4 and 5) (Table 2), thus confirming the relevance of the isopropenyl group at C4 for anti-inflammatory activity. Nonetheless, the isopropenyl group per se is not sufficient for activity, as compounds with such group, but lacking the oxygenated group at C6 (1, 2, 3, 10 and 11) are inactive.

The results also show that (S)-(+)-carvone (4) and its derivative (20) are slightly more potent than their respective R isomers (5 and 21), from which they differ only in terms of stereochemistry at the chiral C4 atom. The third most potent compound used, (+)-dihydrocarveol (8), is a mixture of isomers of which the most abundant, (1 S,2 S,5 S)-dihydrocarveol, presents the S configuration at all its chiral centres, including C5 that corresponds to C4 of limonene, while its isomer, (−)-dihydrocarveol (9), four times less potent, is also a mixture of isomers, the most abundant of which, (1 R,2 R,5 R)-dihydrocarveol, presents the R configuration (detailed composition and purity of each test compound in Supplementary Table S1). Moreover, (−)-carveol (6) and (+)-dihydrocarvone (7) also have additional chiral centres at C6 or C1 and the products used are mixtures of S and R isomers at those positions, but in both, the most abundant is the isomer presenting the R configuration at the carbon atom corresponding to C4 (detailed composition in Supplementary Table S1). Similarly, (R)-(+)-pulegone (12) is inactive, while its S enantiomer (13) shows weak activity. Taken together, these results suggest that the S configuration, especially at C4, is more favourable for activity.

Finally, we tested four monoterpenes (1619) unrelated to the limonene synthase pathway (Fig. 1b), but presenting various degrees of rigidity to elucidate the relevance of this feature for activity. Neither the more rigid (1719), nor the more flexible (16) of these four compounds showed any activity.

Table 3 summarizes the chemical features found relevant for anti-inflammatory activity.

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The carvone enantiomers inhibit LPS-induced NOS2 and IL-1β expression in macrophages

To further confirm the anti-inflammatory properties of the two most potent compounds, the expression of NOS2 and IL-1β were evaluated at the mRNA and protein levels. Macrophage treatment with 1 µg/mL LPS significantly increased NOS2 mRNA (Fig. 2a) and protein (Fig. 2b) levels which, as expected, were decreased by Bay 11-7082. The carvone enantiomers, (4) or (5), also significantly reduced LPS-induced NOS2 mRNA (Fig. 2a) and protein levels (Fig. 2b), as well as IL-1β mRNA levels (Fig. 3a). Upon transcription, this mRNA is translated into a precursor protein (pro-IL-1β) that undergoes partial hydrolysis by a proteolytic complex, the inflammasome, being converted into the mature IL-1β protein which is then secreted30. In agreement with the decrease in IL-1β mRNA levels, (S)-(+)-carvone (4) and (R)-(−)-carvone (5) significantly decreased the levels of both pro-IL-1β (Fig. 3b) and mature IL-1β secreted into the cell culture medium (Fig. 3c), relative to treatment with LPS alone.

(S)-(+)-carvone (4) and (R)-(−)-carvone (5) decrease LPS-induced Nos2 mRNA (a) and protein (b) levels in the Raw 264.7 cell line. Macrophage cultures were treated with 1 µg/mL LPS, for 6 h (a) or 18 h (b), following pre-treatment for 1 h with 666 µM of each test compound (a) or with the concentrations indicated in (b). As a positive control, the cells were similarly treated with the selective NF-κB inhibitor, Bay 11-7082, 5 μM. Control cells (Ctrl) were treated with the vehicle (0.1% DMSO) in the absence of LPS. Each column represents the mean ± SEM of four independent experiments. The blots shown are representative of, at least, three independent experiments and are cropped for clarity and conciseness. The corresponding full-length blots are presented in Fig. S3. **p < 0.01, ***p < 0.001 and ****p < 0.0001 relative to LPS-treated cells. ##p < 0.01 relative to the Ctrl. §p < 0.05, §§§p < 0.001 and §§§§p < 0.0001 between the conditions indicated. MW: molecular weight marker.

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(S)-(+)-carvone (4) and (R)-(−)-carvone (5) decrease LPS-induced IL-1β mRNA (a) and protein (b,c) levels in the Raw 264.7 cell line. Macrophage cultures were treated with 1 µg/mL LPS, for 6 h (a) or 18 h (b,c), following pre-treatment for 1 h with 666 µM of each test compound (a and c) or with the concentrations indicated in (b). Control cells (Ctrl) were treated with vehicle (0.1% DMSO) in the absence of LPS. Each column represents the mean ± SEM of, at least, three independent experiments. The blots shown are representative of, at least, three independent experiments and are cropped for clarity and conciseness. The corresponding full-length blots are presented in Fig. S3. *p < 0.05, **p < 0.01 and ****p < 0.0001 relative to LPS-treated cells. ##p < 0.01, ###p < 0.001 and ####p < 0.0001 relative to the Ctrl. §p < 0.05 between the conditions indicated. MW: molecular weight marker.

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To further characterize the mechanism of action of the carvone enantiomers (4 and 5), we determined whether they are also effective when added to the cells after the inflammatory stimulus and the mechanism involved, that is, whether they act by modifying NOS2 protein levels and/or its enzyme activity. For this, we treated macrophages with LPS for 8 h to induce NOS2 expression and protein synthesis. Then, the cells were washed to remove LPS and new medium with the carvone enantiomers (4 and 5) was added to the respective wells for 18 h. The results in Fig. 4a show that treatment with either compound decreased NO production by approximately 30% while NOS2 protein levels were reduced by approximately 60% (Fig. 4b). This suggests that the decrease in NO production is secondary to the decrease in NOS2 protein and not to a direct inhibition of the enzyme activity.

Effects of (S)-(+)-carvone (4) and (R)-(−)-carvone (5) on NO production and NOS2 protein levels pre-induced by treatment with LPS. In panels (a,b), macrophage cultures were treated with 1 μg/mL LPS, for 8 h to induce NOS2 expression. Then, the medium was changed to remove LPS and the cells were treated for another 18 h with 666 µM of each test compound or the vehicle (0.1% DMSO). Controls were set up by leaving the cells untreated for 8 h followed by addition of vehicle for 18 h. In panel c, cells were pre-treated with LPS or left untreated for 8 h and immediately processed for protein extraction or treated with or without LPS for 8 h and then further incubated with vehicle for another 18 h. Each column represents the mean ± SEM of, at least, three independent experiments. The blots shown are representative of, at least, three independent experiments and are cropped for clarity and conciseness. The corresponding full-length blots are presented in Fig. S3. ***p < 0.001 and ****p < 0.0001 relative to LPS, 8 h + vehicle, 18 h. ###p < 0.001 and ####p < 0.0001 relative to untreated cells, 8 h + DMSO, 18 h. Φp < 0.05 relative to LPS, 8 h. N/A: not applicable.

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To determine whether this decrease was due to inhibition of NOS2 protein synthesis, still occurring after removal of LPS, or to induction of its degradation, we evaluated its protein levels 8 h after treatment with LPS. The results obtained show that NOS2 protein levels still increased after removal of LPS (Fig. 4c), indicating that NOS2 protein synthesis continues even after removal of the inducing stimulus. This indicates that the decrease in NO production observed in Fig. 4a is not due to inhibition of NOS2 enzyme activity, but rather to inhibition of its synthesis.

(S)-(+)-carvone (4) inhibits inflammatory responses induced by IL-1β in human chondrocytes

First, the highest concentrations of (S)-(+)-carvone (4) not toxic to murine macrophages were tested in human chondrocytes and found not to affect cell viability relative to cells treated with IL-1β alone (Fig. 5a). At the same concentrations, (S)-(+)-carvone (4) significantly decreased IL-1β-induced NOS2 protein levels (Fig. 5b) and NO production (Fig. 5c) in human chondrocytes. 5 µM Bay 11-7082 significantly decreased both parameters (Fig. 5b,c), confirming the quality of the model system. Finally, (S)-(+)-carvone (4) also significantly decreased pro-IL-1β protein levels in a concentration-dependent manner (Fig. 5d).

(S)-(+)-carvone (4) does not affect cell viability (a) and decreases IL-1β-induced NOS2 protein levels (b) and NO production (c) as well as pro-IL-1β protein levels (d) in human chondrocytes. The cells were treated with 10 ng/mL IL-1β for 24 h (a, b, c and d), following pre-treatment for 1 h with 666 or 1331 µM of (S)-(+)-carvone (4). As a positive control, the cells were similarly treated with 5 μM Bay 11-7082 (b and c). Control cells (Ctrl) were treated with the vehicle (0.1% DMSO) in the absence of IL-1β. Each column represents the mean ± SEM of, at least, three independent experiments. The blots shown are representative of, at least, three independent experiments and are cropped for clarity and conciseness. The blots shown in Fig. 5b were vertically sliced before the last condition (Bay 11-7082) to exclude a condition not relevant for the present study. The corresponding full-length blots are presented in Fig. S3. ***p < 0.001 and ****p < 0.0001 relative to IL-1β-treated cells. ##p < 0.01, ###p < 0.001 and ####p < 0.0001 relative to the Ctrl. §§§p < 0.001 and §§§§p < 0.0001 between the conditions indicated.

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Discussion

To the best of our knowledge and among the nine active compounds, (S)-(+)-carvone (4), (+)-dihydrocarvone (7), (+)- (8) and (−)-dihydrocarveol (9), (S)-(−)-pulegone (13) and the carvone derivatives (20 and 21), were never reported to have anti-inflammatory effects, while carvone, either as the racemic mixture31 or the (R)-(−) enantiomer (5)3,32,33 and (−)-carveol (6)33, were recently reported to inhibit some effects correlated with inflammation.

Among the compounds that showed no activity, the lack of inhibition of LPS-induced NO production by the limonene enantiomers (1 and 2), (R)-(+)-pulegone (12), (−)-menthone (14), (−)-menthol (15), β-myrcene (16), p-cymene (17), carvacrol (18) and thymol (19) contrasts with other reports that suggest anti-inflammatory activity for these compounds34,35,36,37,38,39,40,41,42. At least in part, these discrepancies can be due to the use of distinct models, namely cell lines, animal models and endpoints analysed, as well as to different concentrations tested. Significant anti-inflammatory effects were recently reported for limonene (racemic mixture, up to 200 μg/mL) and β-myrcene (up to 50 μg/mL) in human chondrocytes37, as well as in the Raw 264.7 cell line41,42, but at concentrations much higher than those used in the current study. These discrepancies can be due in part to the use of distinct methods to assess cell viability, namely the lactate dehydrogenase (LDH) release assay, based on the integrity of the plasma membrane43, used in the studies by Kim et al. (2013) and Yoon et al. (2010), versus the resazurin or the MTT reduction assays dependent on the integrity and activity of mitochondria44, used in our studies. Nonetheless, since cytotoxicity to human chondrocytes and murine macrophages was evaluated by similar methods, the much lower cytotoxic concentrations observed in mouse macrophages indicate that these cells are more sensitive to cytotoxicity induced by those monoterpenes, suggesting that their effects are species and/or cell type-specific, which further highlights the importance of standardized side-by-side comparisons of different compounds.

Having performed such a standardized comparison, some chemical features were identified as relevant for activity and potency (Table 3). Among those, the presence of a functional oxygenated group at C6 appears as the major determinant for activity while the conjugation of the carbonyl group at that position with the α,β double bond at C1, present in the carvone enantiomers (4 and 5) and their derivatives (20 and 21), was found to further increase potency. Such a conjugation provides a Michael acceptor site, that is, an electrophilic site due to the presence of an electron-withdrawing group in close proximity to a double bond, which is a chemical feature relevant for interaction with biomolecules, namely proteins45,46, and thus likely relevant for interaction of these molecules with their target. However, the pulegone enantiomers (12 and 13) which have no or little activity, also have an α,β double bond conjugated to a carbonyl group, but involving the carbonyl group at C3 and the double bond of the isopropylidene group at C4. In this case, the proximity of the methyl group at C10 to the carbonyl group at C3 can cause a sterical hindrance impairing the interaction with the molecular target46 and thus impeding activity.

The lower potency of the carvone derivatives (20 and 21) relative to the parent compounds (4 and 5) can only be due to the replacement of the isopropenyl at C4 by a 2-hydroxyisopropanyl group. The presence of the hydroxyl group can provide an additional hydrogen binding site, thus creating a new site for chemical interactions that may negatively impact anti-inflammatory activity. Furthermore, this hydroxyl group also increases the volume of the molecule at that region which can hinder access to the pharmacological target and thus decrease the activity of those compounds. Replacement of the isopropenyl group at C4 can also explain why these two compounds are less potent than the third most potent compound, (+)-dihydrocarveol (8), which despite bearing a hydroxyl group at C6, retains the isopropenyl group at C4.

Another relevant feature for activity must be chirality as, otherwise, all the enantiomer pairs tested should have similar activities which is not the case. Moreover, 3 of the active compounds (6-8) have additional chiral centres at C1 and/or C6. Interestingly, the order of potency of the nine active compounds is closely related to the S configuration of the chiral atom at C4, although other chemical features, namely the presence of a functional oxygenated group at C6, the double bond at C1 and the isopropenyl group at C4, seem more relevant (Table 3). Since the S or R configurations significantly affect the 3D conformation of a molecule, the S conformation at C4 by conferring some planarity to that region of the molecules is likely relevant for access to and interaction with the target.

Finally, we tested four monoterpenes (Fig. 1b) unrelated to the limonene synthase pathway, but presenting various degrees of rigidity to elucidate the relevance of this feature for activity. None of these four compounds showed activity, including carvacrol (18) even though it bears a hydroxyl group at C6. Unlike the nine active compounds, including those bearing a hydroxyl group at the same position (6, 8 and 9), the cyclohexane ring in carvacrol (18) is aromatic suggesting that its rigidity impairs the interaction with the target, leading to almost no activity. Likewise, thymol (19) which differs from carvacrol only in the position of the hydroxyl group, and p-cymene (17) which has no functional groups, also showed no activity. On the other hand, β-myrcene (16), an aliphatic compound representing a flexible structure, also showed no activity, suggesting that too flexible (16) or too rigid (17, 18 and 19) structures are unfavourable for activity.

In summary, the results obtained indicate that higher potency is conferred by 1) the carbonyl group at C6, rather than the hydroxyl group, 2) the presence of a Michael centre resulting from the conjugation of an α,β double bond at C1 to a carbonyl group at C6, 3) an isopropenyl group at C4 or, at least, the absence of hydrogen binding sites and bulky groups at that position, and 4) the S configuration, especially at C4. These findings can be useful to predict the anti-inflammatory activity of distinct mint species and their chemotypes once their composition in limonene-derived monoterpenes is known.

As found for inhibition of NO production, the two most potent compounds found, the carvone enantiomers, also significantly decreased the mRNA and protein levels of NOS2 (Figs. 2 and 4) and IL-1β (Fig. 3), further strengthening their anti-inflammatory activity and suggesting that they act at the transcriptional or pre-transcriptional levels. Moreover, (S)-(+)-carvone (4) was found to have similar anti-inflammatory effects in human chondrocytes. This indicates that (S)-(+)-carvone (4) is not only effective in inhibiting LPS-induced inflammatory responses in macrophages, but also efficiently inhibits the responses induced by a distinct inflammatory and catabolic stimulus in a different cell type. The anti-inflammatory effects of (S)-(+)-carvone (4) in human chondrocytes are especially relevant because inflammatory cytokines, like IL-1β, drive joint destruction by inducing the expression of catabolic enzymes47 and also contribute to OA pain48, namely by inducing the expression of nerve growth factor by synovial macrophages49. Moreover, unlike other monoterpenes, e.g. limonene, (S)-(+)-carvone (4) is effective as an anti-inflammatory agent both in macrophages and human chondrocytes at similar concentrations. Given the relevance of both cell types and the inflammatory stimuli used to OA pathophysiology23,47, (S)-(+)-carvone (4) may be efficient in halting joint destruction in OA and also contribute to reduce pain. Future studies will aim at further elucidating its mechanism of action and evaluating its anti-osteoarthritic properties in vivo.

Methods

Test compounds

Test compounds 117 and 19 were from Sigma-Aldrich Co. (St Louis, MO, USA). Thymol (18) was from British Drug Houses. Compounds 20 and 21 were synthesized at our laboratory, as described below. Details about purity and isomer composition are provided in Supplementary Table S1.

Chemical synthesis of compounds 20 and 21

The synthetic procedure was adapted from Buechi and Wueest (1979).

Synthesis of (S)-5-(2-hydroxypropan-2-yl)-2-methylcyclohex-2-en-1-one [(S)-8-hydroxycarvotanacetone, 20]

1 mL of 50% aqueous sulphuric acid was slowly added to 150 mg (1 mmol) of (S)-(+)-carvone (4) at 0 °C. The mixture was stirred for 24 h at 0 °C. After extraction with 2 mL of hexane-ether (3:1), the aqueous layer was extracted with diethyl ether (6 × 2 mL) for 24 h. The ether solution was washed with brine containing sodium bicarbonate, dried over anhydrous sodium sulphate and evaporated under reduced pressure. The remaining aqueous layer was extracted with ethyl acetate (3 × 2 mL) for 12 h. The organic phases were washed with brine containing sodium bicarbonate, dried over anhydrous sodium sulphate and evaporated under reduced pressure. The combined crude extract was purified by flash chromatography (hexane: ethyl acetate 1:1) to give compound 20 (80 mg, 48% yield) as a viscous liquid. Purity (GC-MS): 99.7%. 1H NMR (400 MHz, CDCl3) δ: 6.77 (m, 1 H), 2.64-2.42 (m, 1 H), 2.28-2.21 (m, 1 H), 2.28-2.21 (m, 2 H), 1.78 (s, 3 H), 1.24 (s, 3 H), 1.23 (s, 3 H). 13C NMR (101 MHz, CDCl3) δ: 200.35 (C=O), 145.15 (CH), 135.24 (CH), 71.64 (C-OH), 46.05 (CH), 39.61 (CH2), 27.31 (CH3), 27.25(CH2), 27.02(CH3), 15.61(CH3). IR (ATR) cm-1: 3424, 2973, 2925, 2891, 1656, 1381, 1367, 1143, 1111, 1059, 928, 903,813, 711, 678. MS m/z: 28.1, 43.0, 59.1, 95.0, 110.1, 135.1, 150.1, 168.1.

Synthesis of (R)-5-(2-hydroxypropan-2-yl)-2-methylcyclohex-2-en-1-one [(R)-8-hydroxycarvotanacetone, 21]

Compound 21 was synthesized as described for compound 20 using as starting material (R)-(−)-carvone (5). The combined crude was purified by flash chromatography (hexane: ethyl acetate 1:1) to give compound 21 (78 mg, 48% yield) as a viscous liquid. Purity (GC-MS): 99.5%. 1H NMR (400 MHz, CDCl3) δ: 6.77 (m, 1 H), 2.64-2.42 (m, 1 H), 2.28-2.21 (m, 1 H), 2.28-2.21 (m, 2 H), 1.78 (s, 3 H), 1.24 (s, 3 H), 1.23 (s, 3 H). 13C NMR (101 MHz, CDCl3) δ: 200.35 (C=O), 145.15 (CH), 135.24 (CH), 71.64 (C-OH), 46.05 (CH), 39.61 (CH2), 27.31 (CH3), 27.25(CH2), 27.02(CH3), 15.61(CH3). IR (ATR) cm-1: 3424, 2973, 2925, 2891, 1656, 1381, 1367, 1143, 1111, 1059, 928, 903,813, 711, 678. MS m/z: 28.1, 43.0, 59.1, 95.0, 110.1, 135.1, 150.1, 168.1.

Cell culture and treatment

Macrophages

The mouse macrophage cell line, Raw 264.7 (ATCC No. TIB-71), was cultured in DMEM supplemented with 10% non-heat inactivated foetal bovine serum (FBS), 100 U/mL penicillin and 100 µg/mL streptomycin. Raw 264.7 cells were plated at a density of 3 × 105 cells/mL and left to stabilize for up to 24 h.

Human chondrocytes

Human knee cartilage was collected within 24 h of death from the distal femoral condyles of multi-organ donors (48–77 years old, mean = 65, n = 7) at the Bone and Tissue Bank of the University and Hospital Centre of Coimbra (CHUC). Only waste tissue resulting from the preparation of bone tissue for cryopreservation was used. All procedures were approved by the Ethics Committee of CHUC (protocol approval number 8654/DC), which follows the Declaration of Helsinki and Oviedo Convention and the Portuguese legislation for organ donation.

Chondrocytes were isolated by enzymatic digestion from cartilage samples as previously described3. Briefly, cartilage shavings underwent sequential digestion with Pronase (Roche, Indianapolis, IN, USA) and collagenase A (Roche, Indianapolis, IN, USA). To avoid chondrocyte dedifferentiation, non-proliferating monolayer cultures were setup by plating 1× 106 chondrocytes/mL in HAM: F12 medium containing 3% antibiotics and 5% FBS and allowed to recover for 24 h at 37 °C in a humidified atmosphere with 5% CO2. Prior to any treatments, chondrocytes were serum-starved overnight and thereafter maintained in culture medium without FBS.

Cell treatments

Test compounds and Bay 11-7082 (Calbiochem, San Diego, CA, USA), used as a pharmacological control, were dissolved in dimethyl sulfoxide (DMSO; Sigma-Aldrich Co) so that its final concentration in the culture medium did not exceed 0.1% (v/v). This vehicle was used as control. Lipopolysaccharides from Escherichia coli 026:B6 (LPS; Sigma-Aldrich Co.) were dissolved in phosphate buffered saline (PBS). Recombinant human interleukin-1β (IL-1β; Peprotech, Rocky Hill, NJ, USA) was dissolved in PBS containing 0.1% Bovine Serum Albumin. Test compounds or the vehicle were added to macrophage cell cultures or human chondrocytes 1 h before the pro-inflammatory stimulus, 1 µg/mL LPS or 10 ng/mL IL-1β respectively, and maintained for the rest of the experimental period, except for experiments in Fig. 4 (details in the Results section and figure legend). The concentrations of each compound and the experimental treatment periods are indicated in figure legends.

Selection of non-cytotoxic concentrations by the resazurin reduction assay

Resazurin is a redox dye used as an indicator of cellular metabolic activity for various applications, namely cell viability, proliferation and toxicity. The assay is based on the intracellular reduction of the non-fluorescent resazurin to resorufin (a fluorescent and pink coloured compound) by mitochondrial or microsomal enzymes that use NADH or NADPH as electron sources. Since only metabolically active cells can reduce the dye, the increase in fluorescence or absorbance is directly proportional to the number of viable cells26,50.

To select non-cytotoxic concentrations of the test compounds, the resazurin solution was added to each well to a final concentration of 50 µM, 90 min before the end of the treatment period indicated in the figure legends. Then, absorbances at 570 nm and 620 nm (reference wavelength) were read in a Biotek Synergy HT plate reader (Biotek, Winooski, VT, USA).

Nitric oxide production

NO production was measured as the amount of nitrite accumulated in the culture supernatants using the Griess reaction which is based in the reaction of nitrite with sulfanilamide under acidic conditions, yielding a diazonium ion that couples to N-(1-napthtyl)ethylenediamine dihydrochloride to form a water-soluble red-violet azo dye that absorbs at 550 nm51. Briefly, equal volumes of culture supernatants and reagents [equal volumes of 1% (w/v) sulphanilamide in 5% (v/v) phosphoric acid and 0.1% (w/v) N-(1-napthtyl)ethylenediamine dihydrochloride] were mixed and incubated for 10 min, at room temperature, in the dark. The concentration of nitrite accumulated in the culture supernatants was calculated by interpolation of the absorbance of each sample, read in Biotek Synergy HT plate reader (Biotek), in a standard curve of sodium nitrite.

Total RNA extraction and quantitative real-time PCR (qRT-PCR)

Total RNA extraction and qRT-PCR were performed as described before52. Briefly, total RNA was extracted using the NZYol (NZYTECH, Lisbon, Portugal) and quantified in a NanoDrop ND-1000 spectrophotometer at 260 nm. RNA purity was assessed by analysis of 260/230 and 260/280 absorption ratios. The cDNA was reverse-transcribed using NZY First Strand cDNA Synthesis Kit (NZYTECH), beginning with 2 µg of total RNA. qRT-PCR was performed, in duplicate for each sample, using NZYSpeedy qPCR Green Master Mix (2×) (NZYTECH) on CFX96 Real-Time PCR Detection System (Bio-Rad, Hercules, CA, USA).

The efficiency of the amplification reaction for each gene was calculated using a standard curve of a series of diluted cDNA samples and the specificity of the amplification products was assessed by analysing the melting curve generated in the process.

Gene expression changes were analysed using the built-in CFX Manager software which enables the analysis of the results by the Pfaffl method, a variation of the ΔΔCT method corrected for gene-specific efficiencies53. The results were normalized using Hprt1 as housekeeping gene. This gene was experimentally determined with Genex software using NormFinder and geNorm algorithms (MultiD Analyses AB, Göteberg, Sweden) as the most stable for the treatment conditions used. Specific sets of primers for Nos2, Il1b and Hprt1 (Table 4) were designed using Beacon Designer software version 8 (Premier Biosoft International, Palo Alto, CA, USA).

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Western blotting

Total cell extracts were prepared and western blot was performed as described before54. Briefly, total (25 µg for Raw 264.7 cell line and 20 µg for human chondrocytes) proteins were separated by SDS-PAGE under reducing conditions and electrotransferred onto PVDF membranes. These were probed overnight at 4 °C or for 2 h at room temperature with rabbit polyclonal antibody against IL-1β (dilution 1:500; sc-7884, Santa Cruz Biotechnology, INC., Texas, USA) or mouse monoclonal antibody NOS2 (dilution 1:500; MAB9502, R&D Systems, Minneapolis, MN, USA) and then with anti-rabbit or anti-mouse alkaline phosphatase-conjugated secondary antibodies (dilution 1:20000; GE Healthcare, Chalfont St. Giles, UK) for 1 h at room temperature. Immune complexes were detected with Enhanced ChemiFluorescence reagent (GE Healthcare) in the imaging system Thyphoon FLA 9000 (GE Healthcare). The membranes were reprobed with a mouse monoclonal anti-β-Tubulin I antibody (Sigma-Aldrich Co.), diluted at 1:20000, as a loading control, for 1 h at room temperature. Image analysis was performed with TotalLab TL120 software (Nonlinear Dynamics Ltd).

Measurement of secreted IL-1β

The concentration of IL-1β in the culture supernatants was measured using the Mouse IL-1β ELISA kit (ThermoScientific, Rockford, USA), following the manufacturer’s instructions.

Statistical Analysis

Results are presented as means ± SEM. Statistical analysis using GraphPad Prism version 6.0 (GraphPad Software, San Diego, CA, USA). Normal distribution of the data was evaluated with the D’Agostino & Pearson omnibus, the Shapiro-Wilk and the Kolmogorov-Smirnov tests. In cases where the number of samples is too small, we assumed the data follow a normal distribution, as this was verified in all cases where the sample number was larger, including analysis of the same analyte under different experimental treatments. Statistical analysis was performed by one-way ANOVA with the Dunnett post-test for comparison to a control group and the Tukey post-test for multiple comparisons, except in Fig. 4c where the unpaired t-test was used to compare a specific condition with its respective control. Results were considered statistically significant at p < 0.05.

Data availability

The datasets generated and analysed during the current study are available from the corresponding author on reasonable request.

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Sours: https://www.nature.com/articles/s41598-020-64032-1

Now discussing:

Structural basis for promiscuous action of monoterpenes on TRP channels

Abstract

Monoterpenes are major constituents of plant-derived essential oils and have long been widely used for therapeutic and cosmetic applications. The monoterpenes menthol and camphor are agonists or antagonists for several TRP channels such as TRPM8, TRPV1, TRPV3 and TRPA1. However, which regions within TRPV1 and TRPV3 confer sensitivity to monoterpenes or other synthesized chemicals such as 2-APB are unclear. In this study we identified conserved arginine and glycine residues in the linker between S4 and S5 that are related to the action of these chemicals and validated these findings in molecular dynamics simulations. The involvement of these amino acids differed between TRPV3 and TRPV1 for chemical-induced and heat-evoked activation. These findings provide the basis for characterization of physiological function and biophysical properties of ion channels.

Introduction

Organisms use sensors such as transient receptor potential (TRP) ion channels to adapt to environmental changes. Members of the TRP ion channel family play important roles as polymodal sensors to detect and respond to changes in temperature, pH, voltage, osmolarity, and exogenous molecules involved in taste, smell, and pheromone responses1. Channels in this family include TRP vanilloid 1 (TRPV1), TRP vanilloid 3 (TRPV3), and TRP melastatin 8 (TRPM8) that are all crucial for sensing temperature and natural compounds2,3. TRPV1 is physiologically important for thermal and chemical nociception in sensory neurons and is activated by capsaicin, toxins, pH, and temperature in the noxious range (>42 °C). The third member of the TRPV subfamily, TRPV3, is also a heat sensor. TRPV3 is expressed in various tissues and organs, but the most intensive expression is in skin epithelial cells. TRPV3 is believed to be responsible for sensation of warm temperature ranging from 35 to 39 °C4, but has also been reported to be initially activated by noxious heat (>50 °C) and sensitized to activation in response to warm temperature5. Chemical agonists of TRPV3 include spice extracts that contain monoterpenes such as menthol, camphor, and synthetic agents (including 2-aminoethoxy diphenylborate, 2-APB) and unsaturated fatty acids6. Whereas many TRPV subfamily members act as heat sensors, the TRPM8 channel is known to be involved in sensation induced by cool temperatures (<22–27 °C) and menthol3,7. TRMP8 is highly expressed in sensory neurons of trigeminal and dorsal root ganglia, and in the prostate epithelium8. PIP2 regulates TRM8 activity and PIP2 depletion can desensitize the channel9,10,11,12,13.

The six TRP subfamilies (TRPC, TRPV, TRPM, TRPA, TRPP, and TRPML) all have a tetrameric assembly consisting of six transmembrane secondary structures (S1–S6) with intracellular N and C termini. Each subunit has a re-entrant loop with a pore loop (P) comprising 1–2 small pore helices (H1/H2) that is located between S5 and S6 (S5–P–S6). The S1–S4 and S5–P–S6 are connected by the S4–S5 linker. The structures of several TRP channels were recently extensively clarified by cryo-EM single-particle analysis14,15,16,17. The combination of cryo-EM microscopy with nanodisc technology allowed the efficient determination of the location of annular and regulatory lipids9,15,18. Lipids in cellular membranes are important regulators that can functionally modulate many TRP channels by direct or specific interactions, especially in the case of phosphatidylinositides (PIP2) and cholesterol. Gao et al.15 reported that a binding pocket in the S4–S5 linker region for vanilloid ligands is shared with that for PIP2. Tightly bound PIP2 works as a co-factor that stabilizes TRPV1 in its resting state by serving as a competitive vanilloid antagonist and a negative allosteric modulator14,15. In a recent study, Singh et al. observed two non-protein densities, presumably representing lipids, in each subunit of the TRPV3 tetramer. The pockets for lipids are located in the intracellular half of S1–S4 domains and C terminus of the TRP domain, which are analogous to the pocket for putative lipids in other TRPV channels16. Lipid location and function were also clarified in recent studies of the TRPM8 structure9,18. Yin et al. (2019) reported that for TRPM8 PIP2 can effectively control conformational transitions associated with gating and enhance the potency of agonist binding9,17.

Menthol, a natural non-reactive cooling compound that has three asymmetric carbon atoms and a molecular formula of C10H20O, is widely used in oral hygiene products, cosmetics, pesticides, and pharmaceuticals, and is also used as a flavoring agent in foods. The major form of menthol found in nature is (−)-menthol (l-menthol), which is frequently studied since it has better cooling properties than other isomers19,20. Menthol is a well-known TRPM8 activator and is required for cool thermosensation in vivo3,7. Menthol can also activate TRPV3 and has bimodal effects on mouse TRPA1 (refs. 21,22). Moreover, several amino acids were shown to be involved in the menthol binding in mouse TRPM8 (Y745, R842, and Y1005)23 and human TRPA1 (S873 and T874)22. We previously reported that menthol inhibits capsaicin-activated human TRPV1 activity24, indicating that it shows promiscuous actions toward several TRP channels. However, no consensus has been reached about menthol-binding sites, especially for TRPV3.

Other structurally related monoterpenes such as camphor, carvacrol, and 1,8-cineol can also regulate several TRP channels in different ways. For example, camphor activates mammalian TRPV1 and TRPV3, but inhibits rat TRPA1 (ref. 21). Interestingly, some studies indicated that these natural compounds can insert into biological membranes where they can cause significant alterations in numerous physico-chemical characteristics of lipid bilayers25,26,27,28. A central question arises about whether the effects of these small and hydrophobic compounds are mediated directly, via direct interaction with ion channels integrated in the membrane, and/or indirectly through alterations in the physico-chemical characteristics of lipid membranes that have indirect effects on channel function.

Here we have characterized the pharmacological effects of menthol and other structurally related monoterpenes on several TRP channels including TRPV1, TRPV3, and TRPM8. We show an agonistic effect of menthol on TRPV1. We also identified several amino acids in the S4–S5 linker that are involved in the agonistic effect of menthol on rat TRPV1 (R557K and G563S) and mouse TRPV3 (R567K and G573S).

Results

Confirmation of mutations that specifically affect menthol responses in TRPM8

We made histidine substitutions at Y745 in the S1 transmembrane segment and R842 in the S4 transmembrane segment of mouse TRPM8 (mTRPM8; Supplementary Figs. 1 and  2a); these amino acids are putative binding sites for menthol on TRPM8 (refs. 10,22). We then used a whole-cell patch-clamp method to test the agonistic effects of menthol (Supplementary Fig. 2b) on these mutants expressed in HEK293T cells. Consistent with previous reports, menthol (500 μM)-evoked currents for mTRPM8 Y745H and R842H mutants were much smaller than those of wild type (WT) mTRPM8 (Supplementary Fig. 2c, d), while their cold sensitivity was preserved.

Involvement of S4–S5 region residues R567 and G573 in agonist responses by mTRPV3

We hypothesized that amino acids associated with the effects of menthol on TRPV3 and TRPM8 would be similar and found three candidate residues, tyrosine (Y) in S1 and arginine (R) and glycine (G) in the S4–S5 linker (Fig. 1a and Supplementary Fig. 1). In this study, we focused mainly on the shared residues in the S4–S5 linker domain since the glycine residue in TRPM8 corresponding to G573 in mTRPV3 is not known to be part of the menthol-binding site and the S4–S5 linker is known to have broad involvement in several functions of thermosensitive TRP channels. We found that Y448 in mTRPV3 is not involved in menthol-evoked mTRPV3 activation (Supplementary Fig. 3a). Camphor and 2-APB sensitivities were not changed for mTRPV3-Y448H (Supplementary Fig. 3b, c). However, unlike WT mTRPV3, menthol did not activate mTRPV3-G573S, whereas heat-evoked activation was still observed (Fig. 1b, c), indicating that G573 is involved in menthol-evoked activation of mTRPV3. This mutant also lost sensitivity to camphor and 2-APB, but was activated by heat (Fig. 1d–i), consistent with a previous report29. The importance of glycine in the S4–S5 linker domain in chemical activation is consistent with the recent result showing heat-evoked activation of TRPV3 wherein temperature-mediated opening of TRPV3 is induced by conformational changes in the extracellular region of the pore domain that then promote channel opening with involvement of the S4–S5 linker domain30.

a Location of Y448 (S1), R567, and G573S (S4–S5 linker) in mouse TRPV3 (mTRPV3). b Representative current traces for HEK293T cells expressing WT or mTRPV3-G573S in response to menthol (3 mM) followed by heat stimulation. c Comparison of menthol-activated current densities in HEK293T cells expressing WT or mTRPV3-G573S at ±60 mV (WT: 86.4 ± 12.7 pA/pF at +60 and 26.2 ± 4.3 pA/pF at −60 mV, n = 16; G573S: 14.0 ± 1.9 pA/pF at +60 mV and 6.9 ± 1.5 pA/pF at −60mV, n = 5). d Structure of camphor. e Representative current traces for WT or mTRPV3-G573S in response to camphor (10 mM) followed by heat stimulation. f Comparison of camphor-activated current densities in HEK293T cells expressing WT or mTRPV3-G573S at ±60 mV (WT: 466.0 ± 43.5 pA/pF at +60 mV and 410.8 ± 29.6 pA/pF at −60mV, n = 26; G573S: 18.6 ± 6.7 pA/pF at +60 mV and 9.6 ± 3.9 pA/pF, n = 5). g Structure of 2-APB. h Representative current traces for WT or mTRPV3-G573S in response to 2-APB (300 μM) followed by heat stimulation. i Comparison of 2-APB-activated current densities in HEK293T cells expressing WT or mTRPV3-G573S at ±60 mV (WT: 361.5 ± 48.8 pA/pF at +60 mV and 187.7 ± 25.3 pA/pF at −60mV, n = 12; G573S: 10.7 ± 1.5 pA/pF at +60 mV and 5.1 ± 1.0 pA/pF, n = 5). Insets in b, e, h indicate current–voltage curves obtained at the different time points (1, 2, 3) shown in each trace. Holding potentials were −60 mV with ramp-pulses (−100 to +100 mV, 300 ms) applied every 3 s. Data represent means ± SEM. Statistical analysis was performed by two-sample t-test, ***p < 0.001.

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Next, we examined the possible involvement of R567 in menthol-evoked activation of TRPV3. Menthol-activated currents were very small in mTRPV3-R567K (Fig. 2a, b), whereas the magnitude of camphor-evoked currents was similar between WT and mTRPV3-R567K (Fig. 2c), which is in contrast to the phenotype seen for mTRPV3-G573S. Similar results were obtained when the application sequence was changed (Supplementary Fig. 4a, b). The sensitivity of WT and mTRPV3-R567K to 2-APB and camphor was also similar (Supplementary Fig. 4c, d). These data indicated that G573 and R567 are differently involved in the chemical responses of mTRPV3 wherein G573 is involved in responses to menthol, camphor, and 2-APB, while R567 is involved in menthol, but not in camphor and 2-APB sensitivity.

a Representative current traces for HEK293T cells expressing wild type (WT) or R567K mutant stimulated with menthol (3 mM) followed by camphor (10 mM). b Comparison of the normalized current densities (Imenthol/Icamphor) at ±60 mV (WT: 0.16 ± 0.03 at +60 mV and 0.08 ± 0.02 at −60 mV, n = 13; R567K: 0.07 ± 0.02 at +60 mV and 0.02 ± 0.01 at −60mV, n = 11). c Comparison of camphor-activated current densities for HEK293T cells expressing WT or mTRPV3-R567K at ±60 mV (WT: 736.5 ± 198.8 pA/pF at +60 mV and 633.5 ± 153.6 pA/pF at −60 mV, n = 6; R567K: 634.9 ± 68.2 pA/pF at +60 mV and 613.6 ± 35.7 pA/pF at −60 mV, n = 9). Holding potentials were −60 mV with ramp-pulses (−100 to +100 mV, 300 ms) every 3 s. Data represent means ± SEM. Statistical analysis was performed by two-sample t-test, *p < 0.05.

Full size image

Given the critical location of G573 and R567 at the S4–S5 linker and S4, mutations in this region could produce structural changes that affect channel gating. Structural changes at these sites could also contribute to the observed effect of a ligand beyond that seen for ligand binding alone. To examine this possibility, we analyzed mTRPV3-F569H that carries a mutation that lies in close proximity to the tip of the S4–S5 linker. We found that current sizes in response to menthol and camphor were similar between WT and F569H (Supplementary Fig. 5), indicating that F569 is not involved in the activation of mTRPV3 by monoterpenes.

Involvement of S4/S4–S5 region residues R557 and G563 in the agonistic effect of menthol on TRPV1

Naturally occurring TRP ligands show some degree of promiscuity. Notably, menthol is not only a TRPM8 agonist, but can also activate mammalian TRPV3 (refs. 10,31) and TRPA1 (ref. 32), suggesting that the mechanism associated with the response to menthol is conserved across several TRP channels. Indeed, we previously reported that hTRPV1 currents activated by capsaicin were inhibited by menthol with IC50 values of 1.2 ± 0.2 mM24, whereas mTRPM8 activation by menthol has EC50 values ranging from 4 to 80 μM3. Therefore, we first attempted to examine the agonistic effect of high concentrations of menthol on TRPV1 since both the tyrosine in S1 and arginine and glycine in the S4/S4–S5 linker of rTRPV1 are conserved (Fig. 3a). We did not examine Y441 since capsaicin sensitivity was altered in rTRPV1-Y441H (Supplementary Fig. 6b) and the importance of this aromatic cluster is underscored by previous studies demonstrating that replacement of Y441 by small non-aromatic residues resulted in non-functional channels33,34.

a Location of residues Y441 (S1), R557 (bottom of S4), and G563 (S4–S5 linker) in rat TRPV1 (rTRPV1). b Representative current traces for HEK293T cells expressing WT, G563S, or R557K rTRPV1 in response to menthol (3 mM) followed by capsaicin (CAP, 1 μM) stimulation. c Comparison of normalized current densities (Imenthol/ICAP) for WT, rTRPV1-G563S, or rTRPV1-R557K expressed by HEK293T cells at +60 mV (WT: 0.08 ± 0.01, n = 11; G563S: 0.03 ± 0.003, n = 10; R557K: 0.01 ± 0.002, n = 9). d Comparison of CAP-activated currents densities for WT, rTRPV1-G563S, or rTRPV1-R557K expressed in HEK293T cells at ±60 mV (WT: 547 ± 26.7 pA/pF at +60 mV and 442 ± 29.2 pA/pF at −60 mV, n = 22; G563S: 521 ± 3.7 pA/pF at +60 mV and 388.1 ± 26.4 pA/pF at −60 mV, n = 14; R557K: 633.5 ± 3.3 pA/pF at +60 mV and 476.4 ± 33.8 pA/pF at −60 mV, n = 11). Holding potentials were −60 mV with ramp-pulses (−100 to +100 mV, 300 ms) applied every 3 s. Data represent means ± SEM. Statistical analysis was performed using two-sample t-test, ***p < 0.001.

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On the other hand, we unexpectedly observed rTRPV1 activation by 3 mM menthol (Fig. 3b), although we could not determine EC50 values of menthol-evoked activation for rTRPV1 since solutions with high menthol concentrations could not be prepared. As expected, rTRPV1-G563S and rTRPV1-R557K lost menthol-evoked activation, although capsaicin (1 μM) sensitivity was not affected (Fig. 3b–d and Supplementary Fig. 6a). However, reduction of capsaicin-evoked currents after washout occurred very slowly for the two mutants compared to WT-rTRPV1. A comparison of the current densities for WT, rTRPV1-G563S and rTRPV1-R557K 30 s after capsaicin washout showed that the two mutants had significantly larger current densities than WT (Supplementary Fig. 6c), indicating the involvement of these two residues in channel gating. We next examined the effects of camphor and 2-APB since their effects on mTRPV3 differed between G573 and R567 (Figs. 1 and 2). Interestingly, neither camphor nor 2-APB activated rTRPV1-G563S or rTRPV1-R557K (Fig. 4).

a Representative current traces for HEK293T cells expressing WT, rTRPV1-G563S, or rTRPV1-R557K stimulated with camphor (10 mM) followed by CAP (1 μM). b Representative current traces for HEK293T cells expressing WT, rTRPV1-G563S, or rTRPV1-R557K stimulated with 2-APB (300 μM) followed by CAP (1 μM). c Comparison of the normalized current densities (Icamphor/ICAP) for WT, rTRPV1-G563S, or rTRPV1-R557K expressed in HEK293T cells at ±60 mV (Icamphor/ICAP for WT: 0.63 ± 0.07 at +60 mV and 0.28 ± 0.06 at −60 mV, n = 14; G563S: 0.03 ± 0.01 at +60 mV and 0.02 ± 0.01 at −60 mV, n = 8; R557K: 0.03 ± 0.01 at +60 mV and 0.02 ± 0.01 at −60 mV, n = 5). d Comparison of normalized current densities (I2-APB/ICAP) in HEK293T cells expressing WT, rTRPV1-G563S, or rTRPV1-R557K at ±60 mV (I2-APB/ICAP for WT: 0.84 ± 0.06 at +60 mV and 0.91 ± 0.08 at −60 mV, n = 13; G563S: 0.03 ± 0.004 at +60 mV and 0.02 ± 0.004 at −60 mV, n = 8; R557K: 0.02 ± 0.001 at +60 mV and 0.02 ± 0.002 at −60mV, n = 5). Holding potentials were −60 mV with ramp-pulses (−100 to +100 mV, 300 ms) applied every 3 s. Data represent means ± SEM. Statistical analysis was performed using two-sample t-test, ***p < 0.001.

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Importance of positive charge at R567 for activation of mTRPV3 by camphor

The above data indicated that the involvement of glycine and arginine in monoterpene-evoked activation differs between mTRPV3 and rTRPV1. Due to the structural similarity of menthol and camphor as monoterpenes, we explored whether the charge at R567 is important for the camphor action by comparing the response of WT, R567K, R567H, R567A and R567F to camphor. We first confirmed that R567F was not sensitive to menthol (Supplementary Fig. 7). Moreover, since the response to 2-APB was not affected for the R567 mutants, we compared the ratio of camphor-activated currents to 2-APB-activated currents. Camphor-evoked mTRPV3 activation was markedly reduced for R567A and R567F (Fig. 5), confirming the importance of the positive charge of R567 for activation of mTRPV3.

a Representative current traces for HEK293T cells expressing WT, mTRPV3-R567A, or mTRPV3-R567F in response to 2-APB (300 μM) treatment followed by camphor (10 mM) stimulation. b Comparison of normalized current densities (Icamphor/I2-APB) in HEK293T cells expressing WT, mTRPV3-R567K, mTRPV3-R567H, mTRPV3-R567A, or mTRPV3-R567F at ±60 mV (WT: 1.93 ± 0.25 at +60 mV and 2.79 ± 0.48 at −60 mV, n = 16; R567K: 1.82 ± 0.22 at +60 mV and 1.95 ± 0.34 at −60 mV, n = 11; R567H: 1.77 ± 0.17 at +60 mV and 1.92 ± 0.28 at −60 mV, n = 12; R567A: 0.21 ± 0.02 at +60 mV and 0.25 ± 0.03 at −60 mV, n = 9; R567F: 0.13 ± 0.01 at −60 mV and 0.18 ± 0.03 at −60 mV, n = 12). Holding potentials were −60 mV with ramp-pulses (−100 to +100 mV, 300 ms) applied every 3 s. Data represent means ± SEM. Statistical analysis was performed by one-way ANOVA, *p < 0.05 (R567A vs. R567K, R567H at −60 mV), **p < 0.01 (R567F vs. R567K, R567H at −60 mV) and ***p < 0.001 (R567A vs. WT, R567K, R567H at +60 mV; R567F vs. WT, R567K, R567H at +60 mV; R567A vs. WT at −60 mV; R567F vs. WT at −60 mV).

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Involvement of phospholipids in monoterpene-induced activation of rTRPV1, but not mTRPV3

Recent structures of TRP channels have been instrumental in understanding how diverse ligands, including 2-APB, menthol, and others act on these channels and suggested that the pocket that binds these ligands also contains native lipid molecules. To understand the effect of specific lipid molecules on camphor-evoked currents, we depleted the specific lipid types from the membrane and then examined the modulation of channel functions by ligands using two chemicals, the specific PI4 kinase inhibitor phenylarsine oxide (PAO)35 and wortmannin, which inhibits both PI3 and PI4 kinases with different concentration dependence36,37. We observed no difference in camphor-activated mTRPV3 currents in the presence and absence of either PAO (100 μM) or wortmannin (10 μM), whereas for rTRPV1 camphor-activated currents were significantly larger in the presence of PAO, but not wortmannin (Supplementary Fig. 8). Consistent with a previous report, PAO itself activated rTRPV1 (Supplementary Fig. 8b)38. Both chemicals are known to reduce PIP2 production, indicating that PIP2 has a larger role in activation of TRPV1 than TRPV3.

Different role for the S4–S5 linker region in heat-evoked activation of TRPV1 and TRPV3

Notably, temperature sensitivity is a hallmark of TRPV1 and TRPV3 channel function4,39,40. Mutagenesis or deletion studies indicated several regions important for heat-evoked activation of TRP channels. The heat-sensitive regions of TRP channels are located at the N-terminal ankyrin repeat domains in TRPA1 (ref. 41), a membrane-proximal N-terminal segment in TRPV1, TRPV2, and TRPV3 (refs. 5,42), the C terminus of TRPV1 (refs. 43,44,45), and the pore domain of TRPV1 and TRPV3 (refs. 46,47,48,49.). Residues that could be involved in temperature sensing may be scattered throughout the receptors rather than a defined temperature-sensing domain50. We thus considered whether glycine and arginine, which are involved in monoterpene-evoked activation, are also involved in heat-evoked activation. We first examined temperature-evoked activation of mTRPV3-G573S and mTRPV3-R567K in HEK293T cells by heating bath solution from 25 °C to 50−53 °C. Similar to WT mTRPV3, large inward currents were observed for both mTRPV3-G573S and mTRPV3-R567K without changes in temperature thresholds (Fig. 6a, b and Supplementary Fig. 9). However, activation of mTRPV3-G573S and mTRPV3-R567K by heat stimulus appeared more rapid than for WT. Therefore, we compared the activation time course of the heat-evoked currents. The times between the current rise and the peak were shorter for mTRPV3-G573S and mTRPV3-R567K than for WT, although only the difference between WT and mTRPV3-R567K was statistically significant (Fig. 6b). These data suggested that these residues are involved in the heat-evoked activation of mTRPV3. On the other hand, rTRPV1-G563S and rTRPV1-R557K did not exhibit heat-evoked activation (Fig. 6c, d), whereas capsaicin sensitivity was not affected (Fig. 3d and Supplementary Fig. 6a). These data indicated that glycine and arginine in S4–S5 of mTRPV3 and rTRPV1 have a different involvement in heat-evoked activation, and the finding that G563 and R557 are involved in heat-evoked activation of rTRPV1 is consistent with a previous report34.

a Representative current (upper) and temperature (lower) traces for HEK293T cells expressing WT, mTRPV3-G573S, or mTRPV3-R567K exposed to heat up to 53 °C. b Comparison of heat-evoked current densities (left) (WT: 183.5 ± 26.1 pA/pF; G573S: 183.6 ± 49.5 pA/pF; R567K: 211.9 ± 44.1 pA/pF), temperature thresholds (middle) (WT: 50.3 ± 0.6 °C, n = 14; G573S: 50.2 ± 1.5 °C, n = 5; R567K: 51.2 ± 0.4 °C), n = 5, and time to current peak (WT: 58.2 ± 5.2 s, n = 14; G573S: 44.2 ± 9.4 s, n = 5; R567K: 28 ± 2.8 s, n = 5) in HEK293T cells expressing WT, mTRPV3-G573S, or mTRPV3-R567K at −60 mV. c Representative current (upper) and temperature (lower) traces for HEK293T cells expressing WT, rTRPV1-G563S, or rTRPV1-R557K exposed to heat up to 45 °C. d Comparison of heat-evoked current densities for HEK293T cells expressing WT, rTRPV1-G563S, or rTRPV1-R557K at −60 mV. Holding potentials were −60 mV (WT: 40.8 ± 10.1 pA/pF, n = 5; G563S: 6.5 ± 0.6 pA/pF, n = 6; R557K: 4.5 ± 0.6 pA/pF, n = 6). Data represent means ± SEM. Statistical analysis was performed by one-way ANOVA, *p < 0.05, and **p < 0.01.

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Stable binding of menthol and camphor by mouse TRPV3

To examine the structural features of the interaction of TRPV3 with menthol and camphor, we performed MD simulations (Fig. 7a–k). The oxygen atoms of menthol and camphor forms hydrogen bonds with amino groups (−NH2) of R567 (Fig. 7b, c, f, g) that promote stable binding of menthol and camphor to this residue in mTRPV3. On the other hand, these hydrogen bonds cannot form for the R567F mutant (Fig. 7d, e), since the phenylalanine does not have an amino group. Thus, menthol and camphor are not fixed in the R567F mutant and are sometimes distant from F567. To observe this phenomenon more clearly, we calculated a time series for the distances between menthol/camphor and R567/F567 (Fig. 7h, i). The distance was calculated as the shortest distance between the heavy atoms (except hydrogen) of menthol/camphor and those of R567/F567. This distance fluctuated slightly around 3 Å for WT and G573S, but ranged between 3 and more than 10 Å for R567F mutant. These results show that both menthol and camphor stably bind to R567 via hydrogen bonding in WT and G573S, but in the R567F mutant the binding states fluctuate. We also calculated the distances between menthol/camphor and G573/S573, as shown in Fig. 7j, k. In the R567F mutant, the distance from G573 to the monoterpenes also fluctuated in the wide range as in the distance from F567. In WT and G573S, the fluctuation of the distance from G573/S573 was much smaller than that in R567F, but slightly more than the distance from R567. This fact indicates that the monoterpenes are bound to R567, but not to G573/S573. In Fig. 7j, the distance in G573S fluctuated less than that in WT. This is because the mutation of G573 to serine caused the steric overlap with W692 and the binding pocket became smaller to eliminate this overlap, meaning that G573 is not involved in ligand binding, but affects the size of the ligand-binding pocket.

Representative snapshots of a the whole WT TRPV3 system, b menthol in WT TRPV3, c camphor in WT TRPV3, d menthol in mTRPV3-G573/R567F, e camphor in mTRPV3-G573/R567F, f menthol and mTRPV3-G573S/R567, g camphor and mTRPV3-G573S/R567. Black frame in a represents the area we focus in bg. Pink highlights indicate hydrogen bonds between monoterpenes and R567. Time series of distance between h menthol and R567/F567 and i camphor and R567/F567. Time series of distance between j menthol and G573/S573 and k camphor and G573/S573.

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Discussion

To date, little is known about how different monoterpenes are sensed by TRPV3 and the sites at which they act are also unclear. Investigation of the actions of monoterpene agonists on TRPV3 will provide clues to clarify the structural basis of activation, which could lead to increased understanding of the role of TRPV3 in physiological function and in disease. We can also exploit the promiscuous actions of monoterpenes toward several TRP channels as an important research tool to characterize the physiological function and biophysical properties of ion channels. Several monoterpene compounds such as menthol and camphor, and the synthetic small, hydrophobic compound 2-APB all activate TRPV1 and TRPV3, even though these two channels share only about 39% amino acid sequence identity21,51,52. In the present work, we identified the putative shared amino acids that are involved in menthol, camphor, and 2-APB-evoked activation in mTRPV3 and rTRPV1 and these residues were further confirmed in MD simulations. Our results showed that the conserved glycine and arginine residues in the S4–S5 linker of mTRPV3 (G573, R567) and rTRPV1 (G563, R557) are required for menthol- and camphor-evoked activation (Supplementary Table 2). Although we did not examine 2-APB binding to the glycine and arginine in TRPV3 in MD simulations, 2-APB actions involving these two residues are consistent with previous reports16,51,53.

As previously reported, several amino acids function as binding sites for menthol in mTRPM8 (Y745 (S1), R842 (S4), and Y1005 (TRP domain))10,23. Among these, Y745 and R842 are conserved in TRPV1 and TRPV3. Y745, R842, and Y1005 are proximal in the TRPM8 structure, but do not lie close to one another in the TRPV1 and TRPV3 structures (Supplementary Fig. 1). PIP2, a positive modulator of the TRPM8 channel, can bind to the pocket formed by the pre-S1 elbow, S1 of one subunit, and S5 in the neighboring subunit9. On the other hand, it was shown that menthol has a bimodal action on mTRPA1 via a mechanism that involves S876 and T877 in S4–S5 linker/S5 region3. These results suggest that the binding pocket and mechanistic action of menthol toward TRPM8 would be distinguishable from that for other TRP channels, and that the S4–S5 linker/S5 region is critical for the action of menthol in TRPV1, TRPV3, and TRPA1, but not TRPM8.

In this study we revealed the activation of rTRPV1 by menthol, which had characteristics similar to that seen for camphor-mediated activation of rTRPV1. Xu et al.21 showed that camphor has strong desensitizing activity of rTRPV1, and suggested that the combination of desensitization of rTRPV1 and inhibition of rTRPA1 could provide a new explanation for the pain-relieving properties of menthol. These actions could be parallel to the analgesic property of menthol, since high concentrations of menthol inhibit both mammalian TRPA1 and TRPV1 (refs. 23,26). Moreover, total outcomes in terms of whole-body sensation would be very interesting to examine. As we previously reported, menthol activates TRPM8, TRPV3, TRPA1, and TRPV1 and inhibits TRPV1 and TRPA1 (at high concentrations for the mouse clone). We think that total outcomes would be determined by which channel has the most prominent activation. Menthol activates TRPM8 most effectively at relatively low concentrations, explaining why exposure to menthol typically induces a cooling sensation.

In our study we found some differences in the actions of menthol, camphor, and 2-APB between rTRPV1 and mTRPV3. The single point mutations G563S and R557K in rTRPV1 eliminated sensitivity to menthol, camphor, and 2-APB (Figs. 3 and 4), whereas the G573S and R567K mutations exhibited different responses from those seen for mTRPV3. G573S lost sensitivity to all three compounds, and the sensitivity of R567 to menthol was affected, but that for 2-APB was not. Although the structures of camphor and menthol are similar, the positive charge at R567 was necessary for activation of mTRPV3 by camphor (Fig. 5). These results could be explained in part by the smaller size of camphor relative to menthol that requires the positive charge to promote an effective fit in the binding pocket formed by G573 and R567.

Remarkably, the G573S mutation of human TRPV3 corresponding to mouse TRPV3-G573S in our study is associated with Olmsted syndrome, which is characterized by bilateral mutilating palmoplantar keratoderma and periodic keratotic plaques accompanied by severe itching54,55. This mutation in mTRPV3 was reported to show large basal currents in the absence of agonists and no responses to 2-APB and camphor in either HEK293 cells or Xenopus oocytes29,56. Meanwhile, in this study basal currents before stimulation were not larger than those for WT-expressing HEK293T cells. We also did not see 2-APB and camphor sensitivity for this mutant (Fig. 1) or differences in heat sensitivity between mTRPV3-WT and G573S (Fig. 6). The reason for the apparent difference in the basal channel activities between this study and previous studies is unclear. However, the cryo-EM-based structure indicates that G573 in the S4–S5 linker lies at the junction with the lower part of S4, S5, the TRP domain, and the lower part of S6 (ref. 57). This structural feature raises the possibility that mTRPV3-G573 is involved in channel gating in concert with W692, which is in the vicinity of G573 and reported to cause large basal currents29,56. However, the mechanisms responsible for this high basal activity remain unknown.

The mutants G563S and R557K essentially lost the heat sensitivity of rTRPV1, while G573S and R567K exhibited similar heat-evoked responses in WT mTPRV3 with a small difference in activation kinetics, indicating that the involvement of these two corresponding residues in heat sensitivity differs between mTRPV3 and rTRPV1 (Fig. 6). No clear evidence for the mechanisms of heat-evoked activation of TRP channels is available, although multiple studies involving mutant analyses or cryo-EM studies showed that several regions are involved in the heat sensitivity of TRPV1 and TRPV3 (refs. 32,43,44,45,46,58). Our results indicated that the activation mechanisms of heat and chemical stimulation of TRPV3 differ, but could have somehow converged in the sensitivity of TRPV1 to some chemicals such as monoterpenes, suggesting that mechanisms for heat sensitivity of TRPV3 and TRPV1 are different. These results could clarify the mechanisms of heat-evoked activation of thermosensitive TRP channels. Singh et al. structurally identified two conformational steps for the heat-evoked activation of TRPV3: a strongly temperature-dependent first step (sensitization) and a weakly temperature-dependent second step (channel opening)59. In both steps, heat stimuli induce a conformational change in the ankyrin repeat domain that consequently promotes a structural change in the transmembrane domain. Moreover, in both steps TRPV3 shows strong gating-associated changes in annular lipid occupancy wherein the first density is sandwiched between the extracellular half of S4 in one subunit and the pore domain of the adjacent subunit. The second density lies in a pocket formed by the intracellular half of the S1–S4 domain and the C-terminal portion of the TRP domain59. Interestingly, a recent structure involving lipid nanodisc-reconstitution of TRPV3 reveals a lipid–protein interaction in TRPV3 and a bound lipid stabilized the selectivity filter of the pore in the narrow state since the lipid displacement in this region might be involved in the structural transitions from close-π to open state during TRPV3 activation60. Although the type and exact role of these putative lipids in thermo-sensitization was unclear due to the limited resolution of the structures, the authors highlighted an important role of lipids in heat sensing by TRPV3 through its interactions with the surrounding membrane lipids.

Several studies suggested that TRPV1 is intrinsically heat sensitive61, and that manipulating the cholesterol composition in native membranes likely does not play a critical role in temperature sensing62. However, the structure of TRPV1 in nanodiscs revealed the presence of a phosphatidylinositol lipid between the S4 of one subunit and S5 and S6 of an adjacent subunit. This finding suggested that heat may open the channel through displacement of resident phosphatidylinositides in this S4–S5 linker region. In this earlier study, R557 of rTRPV1 binds to PIP2, which is consistent with our results. In addition, Wen et al.63 observed in an MD simulation heat-activated conformation changes in several key domains including the S4–S5 linker of TRPV1, indicating the importance of this region for heat-evoked activation. They also found that heat-evoked expansion of the TRPV1 pore is greater at 72 °C than 60 °C, and such expansion was detected in the open structure of TRPV1, thus supporting a putative role for the S4–S5 linker in TRPV1 gating. Although the residues found in this study substantially reduced the heat-evoked activation of rTRPV1, whether they contribute directly to temperature sensing or have other roles in allosteric coupling remains unclear.

Given that both G573 and R567 may be involved in the binding of monoterpenes in mTRPV3, we performed MD simulations focusing on these residues (Fig. 7). We confirmed that the tight binding of monoterpenes to R567 was lost in the R567F mutant. Interestingly, however, we saw no significant changes in the distance between glycine or serine of mTRPV3 and monoterpenes even though monoterpene-induced currents were substantially reduced for G573S (Fig. 1). These data suggest that G573 is not involved in ligand binding, but rather in channel gating. On the other hand, the amplitudes and temperature thresholds for heat-evoked currents were not different between WT and the mutants (G573S and R567K), although the time to peak of heat-evoked R567K currents was significantly shorter compared with WT (Fig. 6), suggesting that R567 is involved in both heat-evoked channel gating and ligand binding. Glycine at 573 in mTRPV3 is well-conserved in TRP channels, suggesting the importance of this residue in channel function. Indeed, in rTRPV1 the corresponding residue G563 was found to be involved in multiple functions including channel activation by small hydrophobic compounds, heat-evoked activation, and desensitization of capsaicin-activated currents34. Taken together, we concluded that G573 is involved in mTRPV3 gating while R567 is involved in both ligand binding and heat-evoked channel activation. The role of G573 in channel gating could be invoked downstream of ligand binding at R567. This possibility would be consistent with a report that G573 interacts with residues in the TRP box that modulates channel gating64.

The S4–S5 linker mediates gating motions in many ion channels with six transmembrane domains65. For voltage-gated potassium (Kv) and sodium (Nav) channels, conformational changes in S4 are coupled to movement of the S4–S5 linker that promotes opening of the activation gate. Several studies indicated the importance of the S4–S5 linker in ligand binding of TRPV1. Gao et al. suggested that vanilloid agonists function by displacing the resident phosphatidylinositol lipid in the pocket formed by the S4 and S4–S5 linker. Using Rosetta-based molecular docking, Yang et al.66 observed a common structural mechanism underlying vallinoid-evoked activation of TRPV1 where the ligand serves as molecular “glue” that bridges the S4–S5 linker to the S1–S4 domain to induce channel opening. In addition, the authors found that, upon binding, capsaicin initiates a conformational wave that propagates through the S4–S5 linker toward the S6 bundle to open the restriction site in the selectivity filter, consequently inducing conformational rearrangements of the selectivity filter that eventually lead to channel opening67. Since there are putative lipids located in the S4–S5 interface in TRPV1 and TRPV3 (refs. 15,16,57,60), we hypothesize that monoterpenes displace the lipid in the S4–S5 interface pocket and in turn displace the S4–S5 linker away from the central axis to mediate S5 and S6 movement, leading to channel activation. However, the exact mechanism may differ between TRPV1 and TRPV3 since there are some differences in these channels such as the kind of lipid, the regulation of lipid, and the properties of the S6 helix related to channel gating15,16,53,60,68. However, whether there is a rearrangement of the S1–S4 bundle upon ligand binding that contributes to channel opening is unclear, particularly since monoterpenes are generally smaller than vanilloids. Therefore, mimicking the action of vanilloid in several other channels would be difficult66,69.

Menthol and camphor, as mentioned above, are small and hydrophobic (Log p value of 3.4 and 2.74 (refs. 28,70), respectively). Several previous studies showed that the physico-chemical characteristics of lipid bilayer membranes such as membrane fluidity and thickness can be modified by menthol and other related monoterpenes (refs. 27,28,71). TRPM8 requires PIP2 for activation and can be activated by PIP2 alone9,10,11,12,13. Interestingly, beyond the positive effects of PIP2 in menthol-evoked activation of TRPM8 and TRPV3 (ref. 72), previous studies also reported the location of putative lipids in several regions including the S4–S5 linker in TRPV1, TRPV3, and TRPV6 (refs. 15,16,57). For example, the pocket at the S4–S5 interface that binds to an activating lipid in TRPV6 (the density observed in this pocket may represent the natural lipid agonist in TRPV6-binding sites), and the exchange between phosphoinositides and activator/inhibitor in this vanilloid-binding pocket, could allosterically regulate TRPV1. Indeed, 2-APB is proposed to induce TRPV3 activation and inhibit TRPV6 by modulating interactions with bound lipids16. Thus, we suggest that monoterpenes could modulate the activity of several TRP channels including TRPM8, TRPV1, and TRPV3 by manipulating interactions with bound lipids, particularly since monoterpenes can act on several channels and the binding pocket locations overlapped with those for lipids. Regarding the involvement of PIP2 in channel activation by monoterpenes, PAO increased camphor-evoked rTRPV1 currents, but not mTRPV3 (Supplementary Fig. 8). This result supports the involvement of PIP2 in TRPV1 activity61. On the other hand, the negative results for mTRPV3 suggest a less important role for PIP2 in TRPV3 activity, despite a report showing that TRPV3 activity is potentiated by PIP2 with PAO72. Even with these data, we cannot exclude the possibility that other lipids are involved in modulating TRPV3 and TRPV1 activity. In any case, activation or inhibition of TRP channel activity by monoterpenes and the intensity with which these agents act could depend on the regulatory roles of lipids for TRP channel gating and on what kinds of lipid, such as PIP2 and cholesterol, bind to the channels. However, we cannot rule out that both changes in the physico-chemical properties of the lipid bilayer and direct binding of hydrophobic compounds to ion channels explain the actions of menthol and other monoterpenes on the functional properties of TRP channels and other ion channels.

Methods

Construction of mutant mouse TRPV1, TRPV3, or TRPM8

TRPV3 or TRPM8 point mutants were constructed using a PrimeSTAR mutagenesis Basal kit according to the manufacturer’s recommendations (Takara Bio Inc., Shiga, Japan). TRPV1 point mutants were constructed using PrimeSTAR GXL DNA polymerase (Takara Bio Inc., Shiga, Japan). Point mutations were introduced by PCR using rat TRPV1-pcDNA3, mouse TRPV3-pcDNA3, and mouse TRPM8-pcDNA5/FRT as templates with oligonucleotide primers (Supplementary Table 1) containing the intended mutations. The amplified PCR products were transformed into Escherichia coli pcDNA3 vectors containing rat TRPV1 or mouse TRPV3, or pcDNA5/FRT vectors containing mouse TRPM8 that were then purified using standard procedures. The entire rat TRPV1, mouse TRPV3, and mouse TRPM8 coding sequences were determined to confirm that only the intended mutations were introduced.

Cell culture

Human embryonic kidney‐derived 293T (HEK293T) cells were maintained at 37 °C and 5% CO2 in Dulbecco’s modified Eagle’s Medium (WAKO Pure Chemical Industries, Osaka, Japan) containing 10% fetal bovine serum (Biowest SAS, Nuaillé, France), 100 units ml−1 penicillin (Invitrogen Corp.), 100 μg ml−1 streptomycin (Invitrogen Corp.), and 2 mm GlutaMAX (Invitrogen Corp.). For patch‐clamp recordings, 1 μg mouse TRPM8 in pcDNA5/FRT, rat TRPV1 or mouse TRPV3 in pcDNA3 vector and 0.1 μg pGreen Lantern 1 cDNA were transfected to HEK293T cells cultured in 35 mm dishes using Lipofectamine Plus Reagent (Invitrogen Corp.). After incubating for 3–4 h, the cells were reseeded on coverslips and further incubated at 33 °C for mouse TRPV3 or 37 °C for rat TRPV1 or mouse TRPM8 in 5% CO2. Patch‐clamp recordings were performed 1 day after transfection.

Chemicals

l-menthol, capsaicin, 2-aminoethoxydiphenyl borate, phenylarsine oxide (PAO), and wortmannin were purchased from Sigma-Aldrich (St. Louis, MO, USE) and camphor was purchased from WAKO chemicals (Tokyo, Japan). 2-APB, PAO, and wortmannin were dissolved in DMSO to make 1 M, 300 mM, and 20 mM stock solutions. Camphor and l-menthol were dissolved in ethanol to make 3 M stock solutions that were diluted to the desired final concentration with bath solution.

Electrophysiology

For whole‐cell experiments, the experimental solutions were bath solution: 140 mM NaCl, 5 mM KCl, 2 mM MgCl2, 5 mM EGTA, 10 mM HEPES, and 10 mM glucose at pH 7.4 adjusted with NaOH; pipette solution: 140 mM CsCl, 5 mM EGTA, and 10 mM HEPES at pH 7.4 adjusted with CsOH. Data from whole‐cell voltage‐clamp recordings were acquired at 10 kHz throughout the experiments and filtered at 5 kHz for analysis (Axon 200B amplifier with pCLAMP software; Axon Instruments, Foster City, CA, USA). The membrane potential was clamped at −60 mV. Series resistance and membrane capacitance were compensated.

All experiments were performed at 25 °C unless otherwise stated. Heat stimulation was induced by increasing the bath temperature using a pre‐heated solution warmed in an inline heater (1 °C s−1, with a maximum of 55 °C). The temperature was monitored using a thermocouple (TC‐344; Warner Instruments, Hamden, CT, USA) placed within 100 μm of the patch‐clamped cell. The heat stimulation was stopped upon confirming that rat TRPV1 or mouse TRPV3 currents were desensitized or inactivated. Temperature profiles and Arrhenius plots for the data from whole‐cell voltage‐clamp recordings were calculated using Origin 8.5 software (OriginLab, Northampton, MA, USA). The absolute current values were plotted on a log scale against the reciprocal of the absolute temperature (T) (Arrhenius plot), and the temperature threshold for channel activation was determined by the temperature that caused a change in the slope. For current density analysis of TRPV3 channels, the peak currents induced by heat or chemical stimulation were measured and presented as pA pF−1.

Molecular dynamics simulation

To confirm stable binding of monoterpenes, we performed molecular dynamics (MD) simulations of TRPV3 with a l-menthol molecule and a camphor molecule. The PDB structure (ID: 6DVW) was used as the initial structure for TRPV3. Because the N-terminal (residues 1–114) and C-terminal (residues 759–791) residues are absent in the PDB structure, the N terminus (residue 115) and C terminus (residue 758) were blocked with an acetyl group and a N-methyl group, respectively. Docking conformations of the l-menthol and camphor molecules with TRPV3 were obtained using the docking program AutoDock73. The N- and C-terminal residues (up to residue 391 and after residue 720) were removed in our MD simulations. We also performed MD simulations for the TRPV3 mutant R567F with both l-menthol and camphor. The initial conformation of the R567F mutant was prepared by replacing the arginine residue (R567) with a phenylalanine residue. In addition, MD simulations for G573S were performed with the monoterpenes. The initial conformation was prepared as follows. Residue 573 was replaced from a glycine residue to a serine residue, as with R567F. Because the serine residue was sterically overlapped with W692, potential energy minimization was carried out to remove the overlap. After the minimization, the docking program AutoDock was utilized again to obtain docking conformations of the monoterpenes with G573S.

The electrostatic charges of atoms in the l-menthol and camphor molecules were determined using restrained electrostatic potential fits71. Quantum chemical calculations were performed using the Gaussian16 program74. Structure optimization and electrostatic potential calculations were carried out using the Hartree–Fock level with a 6-31G(d) basis set. For parameters other than atomic electrostatic charges, General Amber force field parameters75 were used.

The MD simulations were performed using the Generalized-Ensemble Molecular Biophysics program developed by one of the authors (H.O.). This program has been used for several proteins and peptides76,77. The AMBER parm14SB force field78 was used for TRPV3. A cubic unit cell having a side length of 200 Å with periodic boundary conditions was used. The electrostatic potential was calculated with the particle-mesh Ewald method79. The cut-off distance for the Lennard–Jones potential was 12 Å. The Nosé–Hoover thermostat80,81 was used to control the temperature at 310 K and a multiple-time-step method82 was used. The time step was set as ∆t = 0.5 fs and ∆t = 2.0 fs for bonding and non-bonding interactions, respectively. To maintain the atomic structure of TRPV3 in a vacuum, the N, Cα, and C atoms of residues involved in α-helix structures were restrained with a harmonic potential83. Each MD simulation was performed for 100 ns.

Statistics and reproducibility

Data for the patch-clamp experiments were obtained from at least three independent transfections. Data are presented as the mean ± SEM. Statistical analysis was performed with Origin 8.5 software (OriginLab, Haverhill, MA, USA). Significant changes were identified using a two-sample t-test, or one-way ANOVA followed by a Bonferroni post hoc test with p < 0.05 considered as statistically significant (p values: *<0.05, **<0.01, ***<0.001). EC50 values was determined using Origin 8.5 software.

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability

All data and materials used in the analysis are available in the main text, Figures and Supplementary Figures and Tables. EC50 values are available in the end of Supplementary Data 1 and 2 for rat TRPV1 and mouse TRPV3, respectively. The demonstrations of MD simulations are available in Supplementary Movies 1 and 2. Primer sequences and a summary of mutant responses are available in Supplementary Tables 1 and 2, respectively.

Other data or information that support the findings of this study are available from the corresponding author M.T ([email protected]) upon request.

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