Characterization of the Cubamyces Menziesii Terpenome

Abstract

Long‐lasting polypore fungi are significant producers of terpene cyclases of high interest for medicinal or biotechnological applications. Following the 1000 Fungal Genomes initiative launched by the Joint Genome Institute, the genome of Cubamyces (C.) menziesii and identified 18 genes encoding sesquiterpene cyclases (STCs) is explored. In a search for robust catalysts suitable for practical applications, the 18 codon‐optimized open reading frames are cloned and overproduced the C. menziesii STCs in Escherichia coli. In ten cases, the catalytically active enzyme is purified and tested with three chemically synthesized linear diphosphates: geranyl diphosphate, farnesyl diphosphate (FDP), and geranylgeranyl diphosphate. Only FDP proved to be a substrate for these 10 enzymes. The product specificity of all these enzymes is determined by (GC‐MS) gas chromatography mass spectrometry and (NMR) nuclear magnetic resonance analysis. Among the 10 enzymes, four produced a predominant compound, four yielded two main compounds, and the remaining two acted as a multiproduct catalysts. This work sheds light on the potential sesquiterpenes involved in the chemical ecology of the polypore C. menziesii and provides evidence for the potential of Polyporales fungi in the identification of new sesquiterpene cyclase activities.

The genome of Cubamyces menziesii reveals 18 putative sesquiterpene cyclase genes. These genes are cloned and expressed in Escherichia coli, yielding 10 active enzymes. Using farnesyl diphosphate as substrate, the enzymes are analyzed, and the products characterized after bioconversion, uncovering diverse sesquiterpene structures. This study highlights the phylogenetic prediction and the ecological and biotechnological relevance of C. menziesii sesquiterpenes.

1. Introduction

Terpenes, the largest family of natural compounds,^{[ 1 ]} have garnered significant interest across various scientific areas, including phytochemistry, synthetic chemistry, biochemistry, and more recently synthetic biology. This interest is mainly due to their remarkable structural diversity,^{[ 2} , ³ , ^{4 ]} which results in a vast array of physicochemical properties (aromas, pigments, and rubber) and biological properties (anticancerous, antibiotic, and antimalarial).^{[ 2} , ³ , ^{4 ]} This structural diversity arises from two types of enzymes: 1) terpene cyclases (TCs) (or synthases); and 2) tailoring enzymes. The first group of enzymes catalyzes, in a single step, the rather chemically challenging cyclization of linear diphosphate precursors (geranyl diphosphate (GPP), farnesyl diphosphate (FDP), geranylgeranyl diphosphate (GGDP), and geranylfarnesyl diphosphate (GFDP)) leading to the formation of (poly)cyclic hydrocarbons or alcohols that possess numerous stereogenic centers with defined stereochemistry.^{[ 2 ]} These hundreds of (poly)cyclic hydrocarbons or alcohols represent a highly diversified collection of carbon backbones that can be further modified by tailoring enzymes. The latter is mainly hydroxylases (such as CYP450 and α‐keto glutarate‐dependent dioxygenases), alcohol dehydrogenases, methyl transferases, glycosyl transferases, and acyl transferases.^{[ 5 ]} The interplay between these two types of enzymes in various organisms—mainly plants, bacteria, and fungi—has resulted in the identification of over 100,000 different terpenes currently described,^{[ 1 ]} with many more yet to be discovered. The knowledge of TCs was primarily derived from enzymes found in plants. However, as genome sequencing has become increasingly accessible to many laboratories, the microbial world has emerged as a treasure trove for such enzymes.^{[ 6 ]} Although the characterization of TCs can be time‐consuming, it is important from a biotechnological point of view. Indeed, several microbial enzymes have been recently characterized after the retrieval of their genes from genome libraries and have been utilized to catalyze the production of terpenes or to synthesize new‐to‐nature compounds from linear diphosphates (Ref. [7] and cited herein).

The interest in microbial TCs is heightened by the challenges associated with expressing plant‐derived TCs in microbial chassis^{[ 8 ]} and the often suboptimal catalytic performance of these plant TCs compared to their microbial counterparts.^{[ 8 ]} Building on the 1000 fungal genomes initiative (Joint Genome Institute, JGI), this study focuses on the characterization of TCs from the basidiomycete fungus Cubamyces menziesii (formerly known as Leiotrametes menziesii). C. menziesii is a white rot fungus that inhabits dead fallen trunks of Paleotropical and Neotropical areas. The genome of C. menziesii contains 18 genes predicted to encode sesquiterpene cyclases (STCs). To the best of our knowledge, no natural terpene products have been reported from this fungus^{[ 9 ]} nor have any TCs been characterized. As a member of the polypore family, C. menziesii produces long‐lasting fruiting bodies, which likely use chemical communications to contend with the antagonistic organisms in its environment. Such a role is often fulfilled in nature by volatile terpenes synthesized by TCs, which are crucial for understanding fungal ecology.^{[ 10} , ¹¹ , ^{12 ]} To characterize these 18 putative STCs from C. menziesii (CmSTCs), we synthesized their coding sequences with codon optimization for expression in Escherichia coli. The resulting His‐tagged enzymes were purified using affinity chromatography. Each enzyme was tested against three chemically synthesized linear diphosphates (GDP, FDP, and GGDP), which are the respective precursors of mono‐, sesqui‐, and diterpenes. The structures of the reaction products were determined by (NMR) and/or (GC‐MS). Out of the 18 putative TCs, 10 were characterized as STCs.

2. Results

2.1. Bioinformatic Analysis of Cubamyces menziesii SesquiTerpene Cyclases (CmSTCs), Conserved Motifs and Phylogenetic Tree Relationships

Following the launch of the “1000 Fungal Genomes” program by the JGI, a wealth of genomic information regarding the physiological and biosynthetic capacities of fungi is now accessible to the scientific community. In previous investigation,^{[ 9 ]} a bioinformatic analysis highlighted the potential of the C. menziesii polypore for terpene production, identifying 18 genes that encoded predicted STCs. All 18 enzymes exhibited (Figure S5, Supporting Information) the classical motifs of TCs,^{[ 13 ]} which are involved in the coordination of Mg²⁺ and the ionization of diphosphate substrates, such as the DD/E/NxxD motif (DDxxQ for CmSTC7) and the NDxxSxxxE motif, as well as the RY motif.^{[ 9 ]} According to prior reports,^{[ 14 ]} it has been suggested that STCs from basidiomycetes can be classified based on their mode of cyclization of FDP or nerolidyl diphosphate (NDP). It has been proposed that fungal STCs from clade I catalyze the 1,10‐cyclization of farnesyl cation, those from clade II catalyze the 1,10‐cyclization of nerolidyl cation, those from clade III catalyze the 1,11‐cyclization of farnesyl cation, and those from clade IV catalyze the 1,6‐ or 1,7‐cyclization of nerolidyl cation. To analyze the distribution of CmSTCs among these four clades, we generated a phylogenetic tree using 32 previously characterized basidiomycetes STCs^{[ 7 ]} and the 18 CmSTCs from C. menziesii (Figure S4, Supporting Information). Out of the 18 CmSTCs, four clustered with STCs from clade I, four clustered in clade II, eight clustered in clade III, and one clustered in clade IV (Figure 5). The clustering of CmSTC6 was poorly supported by bootstrap values, leaving its classification ambiguous (see Figure S2, Supporting Information).

Figure 5.

Possible structures differentiating the two compounds formed as a result of CmSTC3 catalysis.

2.2. Production, Purification, and Characterization of CmSTCs

To characterize the products of the 18 putative CmSTCs, we cloned their coding sequence after codon optimization for E. coli and overexpressed them in BL21 (DE3) cells using a modified pET22b(+) vector that allows for the addition of a 6His‐Tag at the N‐terminal of the proteins. Initially, we tested overproduction at temperatures of 18, 25, and 37 °C, using isopropyl β‐D‐1‐thiogalactopyranoside (IPTG, 1 mm final concentration) as the inducer (T7 promoter). Following cell lysis, only two proteins were not overproduced (CmSTC13 and CmSTC15). Out of remaining 16 proteins, 12 were produced as an inclusion bodies, while four were produced as soluble proteins (CmSTC1, 3, 5, and 9). To increase the production of soluble proteins, we relied on two strategies: 1) utilizing the E. coli Lemo21(DE3) strain to slow down the rate of protein synthesis (beneficial effect for CmSTC7 and CmSTC8); and 2) changing the position of the 6His‐Tag fusion from N‐terminal to C‐terminal, which improved protein folding and solubility (beneficial effect for CmSTC2, 4, 6, 10, 11, 12, 14, 16, 17, and 18). In total, we successfully produced and tested 16 soluble CmSTCs, using chemically synthesized GDP, FDP, and GGDP on an analytical scale. The reaction products were extracted and analyzed by gas chromatography (GC; see supplementary materials for details on each STC product). Of the sixteen STCs, six did not convert any of the three tested diphosphates (CmSTC4, 6, 10, 12, 14 and 17). On the contrary, the purified CmSTC1, 2, 3, 5, 7, 8, 9, 11, 16, and 18 (see Figure S6, S15, S27, S37, S44, S51, S57, S62, S68, and S75, Supporting Information) were found to be active on FDP and three of them had high product selectivity (CmSTC1, 5, and 9). At this stage, some STCs products displayed chromatograms with identical retention times (see Supporting Information), a peak at 23.0 min (CmSTC2, 3, and 18), a peak at 28.9 min (CmSTC2, 3, and 18), a peak at 39.3 min (CmSTC16 and 18), and a peak at 46.2 min (CmSTC5 and 16). This observation suggested some redundancy in product specificity for these enzymes.

For further characterization, we first focused on STCs with high product selectivity (CmSTC1, 5, and 9), and on those with products of similar retention times (CmSTC2, 3, 16, and 18). To obtain structural information of their products, we conducted the enzymatic reactions at the preparative scale, using FDP and the purified His‐tagged STCs, followed by product extraction, silica gel flash chromatography, and comprehensive NMR experiments, as well as GC‐MS analysis.

2.2.1. CmSTC1

The GC chromatogram (Figure S7, Supporting Information) indicated the production of a major sesquiterpene hydrocarbon, which aligns with the ¹³C NMR experiment (Figure S8, Supporting Information), showing 13 aliphatic carbons and two ethylenic ones. The presence of only one double bond suggested a tricyclic carbon backbone. The distorsionless enhancement polarisation transfert (DEPT) 135 experiment (Figure S9, Supporting Information) demonstrated that both carbons involved in the double bond were quaternary, along with two additional quaternary carbons, while six were primary or tertiary and five were secondary. The ¹H NMR spectra (Figure S10, Supporting Information) revealed that the four methyl groups were located on quaternary carbons, as they each appeared as sharp singlets, indicating that only two tertiary carbons were present in the structure, leading to a molecular formula of C₁₅H₂₄. All these characteristics potentially pointed to Δ⁶‐protoilludene as the main product of CmSTC1.

A careful comparison of ¹H and ¹³C chemical shifts (Table S1, Supporting Information) with literature data^{[ 15 ]} was fully consistent with the proposed structure. This structure was further confirmed by nuclear overhauser effect spectroscopY (NOESY), data not shown) and 2D NMR experiments, including heteronuclear single quantum coherence (HSQC), Figure S11, Supporting Information) and heteronuclear multiple bond correlation (HMBC), Figure S22, Supporting Information), firmly establishing CmSTC1 as a Δ⁶‐protoilludene cyclase. The proposed structure was also validated by GC‐MS analysis (Figure S13, Supporting Information), after comparing the obtained spectrum with literature data.^{[ 14 ]} The previously proposed reaction mechanism^{[ 14} , ^{16 ]} for such an enzyme begins with FDP 1,11‐ring closure, characteristic of clade III fungal sesquiterpene cyclase, as expected from Figure  1 . A small peak was also observed on the chromatogram and was further characterized by GC‐MS analysis as pentalenene (Figure S14, Supporting Information). It is important to note that the formation of both Δ⁶‐protoilludene and pentalenene can be explained by the same cyclization mechanism except at the final steps. In these steps, a dyotropic rearrangement followed by proton loss yields pentalenene from the protoilludyl cation,^{[ 17 ]} while a single proton loss from the same cation leads to Δ⁶‐protoilludene (Figure  2 ).

Figure 1.

Classification of 18 newly identified STCs from C. menziesii. The phylogenetic tree was constructed using the Maximum Likelihood method, with the predicted protein sequences from C. menziesii STCs (CmSTC) and from previously characterized basidiomycete STCs from clade I (blue), clade II (pink), clade III (green), and clade IV (red).

Figure 2.

Proposed mechanism for CmSTC1 (Adapted from permission).^{[ 14} , ^{16 ]}

For further characterization, we conducted kinetic measurements using a malachite green assay,^{[ 18 ]} with chemically synthesized FDP.^{[ 19 ]} The purified CmSTC1 had the following kinetic constants: K _m= 68.2 μm, k _cat= 3.2 s⁻¹, and k _cat/K _m= 0.047 μm ⁻¹ S⁻¹.

2.2.2. CmSTC5

The GC chromatogram and GC retention times (Figure S16, Supporting Information) indicated the production of a major compound, a sesquiterpene alcohol, which aligned with the results of the ¹³C NMR experiment (Figure S17, Supporting Information) showed a peak at 72.7 ppm, characteristic of a C—O bond. Additionally, 12 aliphatic carbons and two ethylenic carbons were identified. The presence of an alcohol and a C=C double bond suggested a bicyclic carbon backbone. The DEPT‐135 experiment (Figure S18, Supporting Information) further revealed the presence of eight primary or tertiary carbons, four secondary carbons, and three quaternary carbons, with one of the two ethylenic carbons being quaternary, as well as two other carbons, while the other ethylenic carbon was identified as tertiary. The ¹H NMR spectra (Figure S19, Supporting Information) showed the presence of one ethylenic proton and of four groups integrating for 3H, two of which appeared as a doublet. These observations suggested the presence of two methyl groups on quaternary carbons, with the other two methyl groups potentially supported by a single tertiary carbon due to identical coupling constants (J = 8 Hz), indicating the presence of an isopropyl group. Considering the presence of one —OH group, four —CH₃ groups, four —CH₂ groups, four —CH groups, one =CH, and one quaternary ethylenic carbon, we proposed the molecular formula C₁₅H₂₆O, which corresponds to a bicyclic alcohol with a trisubstituted double bond. This formula was confirmed by high resolution mass spectrometry (HR‐MS), which yielded an exact mass of 222.1984 Da (theorical 222.1986 Da). A search for NMR data in the literature corresponding to collected information (Table S2, Supporting Information), enabled us to propose δ‐cadinol as the main product formed during CmSTC5 catalysis.^{[ 20 ]} The proposed structure was confirmed by the conjunction of GC‐MS analysis (Figure S25, Supporting Information), comparisons to literature data,^{[ 20 ]} NOESY (Figure S23, Supporting Information) and 2D NMR experiments, including HSQC (Figure S20 and S21, Supporting Information), HMBC (Figure S22, Supporting Information), and correlation spectroscopy (COSY, Figure S24, Supporting Information). The optical rotatory power measurements ([α]²⁵ = +37 at 589 nm) established the stereochemistry as (+)‐δ‐cadinol.^{[ 21 ]}

The previously proposed reaction mechanism for this enzyme,^{[ 20 ]} initiated by FDP 1,10‐ring closure, is characteristic of clade I fungal STCs, as anticipated in Figure 1. In addition to the main peak of δ‐cadinol, a minor peak was identified by GC‐MS analysis and comparison with literature data^{[ 22 ]} as the sesquiterpene hydrocarbon α‐muurolene (Figure S26, Supporting Information). The biosynthesis of α‐muurolene could be explained by a mechanism similar to that of δ‐cadinol, with the exception of the final step, which involves proton loss instead of water addition (Figure  3 ).

Figure 3.

Proposed mechanism for CmSTC5 (Adapted with permission^{[ 20 ]}).

The purified CmSTC5 had the following kinetic constants: K _m= 12.0 μm, k _cat= 0.08 s⁻¹, and k _cat/K _m= 0.006 μm ⁻¹s⁻¹.

2.2.3. CmSTC9

The GC chromatogram (Figure S28, Supporting Information) indicated the production of a sesquiterpene alcohol, which aligns with the ¹³C NMR experiment (Figure S29, Supporting Information) that showed a peak at 80.5 ppm, characteristic of a C—O bond. A 14 aliphatic carbons and no ethylenic carbons were identified, suggesting a tricyclic alcohol structure. Furthermore, the DEPT 135 NMR experiment (Figure S30, Supporting Information) led to the conclusion that the carbon bearing the alcohol functionality was quaternary, along with another aliphatic carbon. Additionally, we observed four secondary carbons and nine primary/tertiary carbons. The ¹H NMR spectra (Figure S31, Supporting Information) revealed the presence of four groups, each integrating for three protons, one relatively unshielded singlet (1.28 ppm) indicating a position on the quaternary carbon bearing the alcohol functionality and three doublets with the same coupling constant (7 Hz). This likely indicates the presence of methyl and/or isopropyl groups, all attached to a tertiary carbon. The remaining five carbons were tertiary leading to a molecular formula of C₁₅H₂₆O. All this structural information was consistent with a structure resembling cubebol. Numerous cubebol epimers are known (1‐epi‐, 4‐epi‐ and 10‐epi‐cubebol) and only the NMR data from 4‐epi‐cubebol^{[ 23 ]} matched ours (Table S3, Supporting Information), ascribing CmSTC9 as a 4‐epi‐cubebol cyclase. This was further confirmed by NOESY (Figure S34, Supporting Information) and 2D NMR experiments, including HSQC (Figure S32, Supporting Information), HMBC (Figure S33, Supporting Information), and COSY (Figure S35, Supporting Information) experiments as well as by GC‐MS (Figure S36, Supporting Information).

The previously proposed reaction mechanism^{[ 13 ]} of this enzyme begins with the NDP 1,10‐ring closure (Figure  4 ), a characteristic feature of clade II fungal STCs, as anticipated in Figure 1.

Figure 4.

Proposed mechanism for CmSTC9 (Adapted with permission^{[ 13 ]}).

The purified CmSTC9 had the following kinetic constants: K _m= 2.1 μm, k _cat= 0.33 s⁻¹, and k _cat/K _m= 0.163 μm ⁻¹ s⁻¹.

2.2.4. CmSTC3

After incubation with FDP, CmSTC3 mainly produced two sesquiterpene hydrocarbons, as indicated by the GC retention times (Figure S38, Supporting Information). The NMR analysis of the purified mixture was facilitated by the ≈2:1 ratio of the two sesquiterpene hydrocarbons. The ¹³C NMR spectra (Figure S39, Supporting Information) revealed only aliphatic and ethylenic carbons. Two shielded ethylenic carbons were observed for the main product, while one was noted for the minor product, indicating the presence of two and one methylene (=CH₂) groups, respectively. The DEPT 135 NMR experiment confirmed this finding (Figure S40, Supporting Information). The aforementioned NMR experiment also indicated that these three methylene groups were connected to three quaternary carbons, and both the main and minor compounds shared a very similar C=CH₂ group (108.3 and 108.4 ppm for the =CH₂ carbon, respectively, and 151.1 and 151.2 ppm for the C= carbon, respectively). In the case of the minor compound, a second double bond between a quaternary and a tertiary carbon (C=CH) was present. For each compound, 11 aliphatic carbons were detected. The DEPT 135 NMR experiment (Figure S40, Supporting Information) indicated the presence of six aliphatic CH₂ groups in the main compound and five in the minor compound, along with two and three methyl groups, respectively. For each compound, we observed two CH groups and one quaternary carbon. The ¹H NMR experiment (Figure S41, Supporting Information) revealed the absence of isopropyl groups, with methyl groups appearing as singlets. Thus, the C₁₅H₂₄ formula was attributed to each compound, corresponding to bicyclic dienes. The main compound contained an additional C=CH₂ double bond and one extra CH₂ group, while the minor compound featured an extra methyl group and a C=CH group, along with very similar chemical shifts for many corresponding carbons in each molecule. This information suggested the existence of two isomers differing solely in the position of a single double bond, as illustrated in Figure  5 .

Thus, we searched for bicyclic isomers containing two or three methyl groups, as depicted in Figure 5, and no isopropyl group. The two sesquiterpene hydrocarbons α‐ and β‐selinene were first selected as candidate structures for the products. Published NMR data for both compounds^{[ 24} , ²⁵ , ²⁶ , ^{27 ]} corresponded with our findings, despite our use of a mixture. This analysis established β‐selinene as the main compound and α‐selinene as the minor one.

It is known that the purification of germacrene A on silica gel can lead to α‐ and β‐selinene.^{[ 28 ]} We therefore considered the possibility that the two selinenes might result from such a rearrangement occurring during purification. This assumption was reinforced by a notable feature observed on the GC chromatogram (Figure S38, Supporting Information). Specifically, before the emergence of the second peak, the baseline increased until reaching the second peak and then returned to a basal level. This characteristic feature suggested an on‐column rearrangement. Such a phenomenon has previously been documented in the analysis of sesquiterpenes produced by a plant germacrene A cyclase.^{[ 29 ]} Consequently, we attributed the second peak to germacrene A. Furthermore, a GC‐MS analysis of the mixture confirmed the presence of two major peaks, with the first being β‐elemene (Figure S42, Supporting Information), a known thermal rearrangement product of germacrene A. To our surprise, the second peak (as observed in both GC and GC‐MS) was not the same. In the GC‐MS experiment (Figure S43, Supporting Information), the second peak was unequivocally identified as β‐farnesene, as deduced from comparison of retention time, coinjection and GC‐MS mass spectrum of an authentic standard. However, in the GC experiments, β‐farnesene could be excluded, as the authentic sample did not match the retention time of the second peak (initially attributed to germacrene A). This discrepancy between the two sets of experiments (GC and GC‐MS) could only be explained by differences in the columns used and/or by variations in the temperature program, although the inlet injection temperature was consistent in both cases (250 °C). A third minor peak, appearing just before β‐elemene, was assigned to cis‐β‐elemene based on the GC‐MS experiment. All these results indicated that CmSTC3 was an almost exclusive germacrene A cyclase.

The previously proposed reaction mechanism of this enzyme^{[ 30 ]} begins with the FDP 1,10‐ring closure, a feature characteristic of clade I fungal STCs. Consequently, the proposed mechanism (Figure  6 ) is inconsistent with the phylogenetic tree, which places CmSTC3 in clade III (Figure 1).

Figure 6.

Proposed mechanism for CmSTC3 (Adapted with permission^{[ 30 ]}).

The purified CmSTC3 had the following kinetic constants: K _m= 19.9 μm, k _cat= 2.4 s⁻¹, and k _cat/K _m= 0.121 μm ⁻¹ s⁻¹

2.2.5. CmSTC18

The GC chromatogram (Figure S45, Supporting Information) indicated the production of two minor sesquiterpene hydrocarbons and one main sesquiterpene alcohol. Both hydrocarbons exhibited the same retention times as the two compounds generated by CmSTC3 and were,thus, designated as β‐elemene and germacrene A, respectively, following GC‐MS analysis confirmation. When ¹³C and DEPT 135 NMR experiments (Figure S46 and S47, Supporting Information) were carried out on the mixture of the three products, after extraction but prior to any purification, the peaks of β‐ and α‐selinene were observed, suggesting that CDCl₃ is sufficiently acidic to catalyze the conversion of germacrene A into the two selinene isomers. Prolonged evaporation of the mixture removed the selinenes, yielding an essentially pure alcohol that allowed for an unambiguous attribution of the ¹³C NMR peaks. Four ethylenic peaks were present in the ¹³C NMR spectrum (Figure S48, Supporting Information), three of which were tertiary (=CH) and one was quaternary. The presence of a peak at 73.3 ppm confirmed that this compound is an alcohol and that the hydroxyl group was attached to a quaternary carbon (Figure S48, Supporting Information). Four CH₂ groups were also present, leaving six –CH and –CH₃ groups. The ¹H NMR spectrum (Figure S49, Supporting Information) suggested the presence of four methyl groups (each integrating for 3H), two of which appeared as doublets and two as singlets, the latter being associated with two quaternary carbons. The other two methyl groups could be assigned to an isopropyl group based on the coupling constants. The carbon count was, thus, completed by two tertiary –CH carbons, leading to the molecular formula of C₁₅H₂₆O, indicative of a monocyclic sesquiterpene alcohol with two double bonds. This formula was confirmed by HR‐MS, which yielded an exact mass of 222.1984 Da (theorical mass 222.1986 Da). One of the rare characterized sesquiterpene with this configuration is germacradiene‐4‐ol. The NMR spectra (¹H and ¹³C) were concurrent with those described in the literature.^{[ 31 ]} Furthermore, the GC‐MS analysis (Figure S50, Supporting Information), when compared with literature data,^{[ 31 ]} was consistent with this assignment, establishing CmSTC18 as a germacradiene‐4‐ol cyclase.

The previously proposed reaction mechanism for the production of germacradiene‐4‐ol^{[ 31 ]} begins with the 1,10‐ring closure of FDP, a characteristic feature of clade I fungal STCs, as anticipated in Figure 1. This mechanism also accounts for the formation of germacrene A (Figure  7 ).

Figure 7.

Proposed mechanism for CmSTC18 (Adapted with permission^{[ 31 ]}).

The purified CmSTC18 had the following kinetic constants: K _m= 6.6 μm, k _cat= 0.57 s⁻¹, and k _cat/K _m= 0.085 μm ⁻¹ s⁻¹.

We then focused on the characterization of CmSTC2 and CmSTC16, both of which exhibiting products with retention times identical to those of CmSTC3, CmSTC5, and CmSTC18 after incubation with FDP.

2.2.6. CmSTC2

The GC chromatogram and GC retention times indicated the formation of one main and five minor compounds (Figure S52, Supporting Information). The main compound had a retention time and a GC‐MS mass spectrum (Figure S53, Supporting Information) identical to that of the major compound produced by CmSTC3 (β‐elemene), confirming that CmSTC2 also functioned as a germacrene A cyclase. One of the minor compounds was identified as β‐farnesene (Figure S54, Supporting Information), by comparing its retention time with that of an authentic standard. The other two minor compounds were identified as sesquiterpene hydrocarbons: β‐selinene (Figure S55, Supporting Information) and α‐maalinene (Figure S56, Supporting Information) by GC‐MS analysis, while the identity of the last compound, a sesquiterpene alcohol, remained undetermined. The formation of β‐selinene from germacrene‐A, to the best of our knowledge, has only been described by following silica gel purification. Furthermore, β‐selinene was not detected in the chromatogram of CmSTC3, a quasiexclusive germacrene‐A cyclase. Therefore, we deduced that CmSTC2 was indeed capable of catalyzing the transformation of FDP into β‐selinene as a minor compound. The fourth sesquiterpene hydrocarbon, identified as α‐maalinene by GC‐MS, was potentially a rearranged product of bicyclogermacrene, something reminiscent of the acidic rearrangement of germacrene A into α‐ and β‐selinene (Figure  8 ). However, to the best of our knowledge, such a rearrangement has never been documented during GC experiments, and we conclude that CmSTC2 truly produced α‐maalinene instead of bicyclogermacrene as a minor compound.

Figure 8.

Acid‐catalyzed rearrangement of germacrene A into α‐ and β‐selinene and suggested rearrangement of bicyclogermacrene into α‐maalinene (Adapted with permission^{[ 28 ]}).

CmSTC2 is, thus, a germacrene A cyclase but it is also able of generating β‐farnesene, α‐maalinene, and β‐selinene. The formation of these compounds can be explained by the same central mechanism, diverging as the reaction proceeds (Figure  9 ). The previously proposed reaction mechanism^{[ 30 ]} of this enzyme begins with the 1,10‐ring closure of FDP, a characteristic feature of clade I fungal STCs. Consequently, this proposed mechanism is not consistent with the phylogenetic tree placing CmSTC2 in clade III (Figure 1), a situation similar to that of germacrene A cyclase CmSTC3.

Figure 9.

Proposed mechanism for CmSTC2 (Adapted with permission^{[ 30 ]}).

2.2.7. CmSTC16

The GC chromatogram (Figure S58, Supporting Information) indicated the formation of two sesquiterpene alcohols. The minor compound had the same retention time (confirmed by coinjection, see Figure S59, Supporting Information) and mass spectrum (Figure S61, Supporting Information) as δ‐cadinol (CmSTC5). The main compound had the same retention time (confirmed by coinjection, see Figure S60, Supporting Information) and the same mass spectrum (Figure S61, Supporting Information) as germacradiene‐4‐ol (CmSTC18), establishing CmSTC16 as a germacradiene‐4‐ol cyclase. As seen previously, the reaction mechanism for the formation of both alcohols starts with the 1,10‐ring closure of FDP, which is characteristic of clade I fungal sesquiterpene cyclase, as expected from Figure 1. At the germacradienyl cation step, the direct addition of water generates germacradiene‐4‐ol (Figure 7), whereas in the case of δ‐cadinol, the water molecule is added to the cadinyl cation derived from the germacradienyl cation (Figure 3).

Finally, we completed the study by investigating CmSTC7, 8, and 11.

2.2.8. CmSTC7

The GC chromatogram (Figure S63, Supporting Information) indicated the formation of two sesquiterpene hydrocarbons. The GC‐MS spectrum of the minor compound (Figure S64, Supporting Information) was identical to that of dauca‐4,8‐diene (daucene)^{[ 14 ]} and the spectrum of the main compound (Figure S65, Supporting Information) was identical to that of dauca‐4(11),8‐diene^{[ 14 ]}, indicating that CmSTC7 was a trans‐dauca‐4(11),8‐diene cyclase. NMR experiments (Figure S66 and S67, Supporting Information) on purified dauca‐4(11),8‐diene confirmed the characterization of this enzyme. In addition, the ¹³C NMR data were consistent with those reported in the literature (Table S4, Supporting Information).^{[ 32 ]} Furthermore, the C₁₅H₂₄ formula, indicative of a bicyclic sesquiterpene hydrocarbon with four unsaturations, was confirmed by HR‐MS and the expected exact mass of 204.1878 Da (theorical mass 204.1878 Da). The previously proposed reaction mechanism^{[ 14 ]} of this enzyme (Figure  10 ) starts with the 1,6‐ or 1,7‐ring closure of NDP, which is characteristic of clade IV fungal STCs, as expected from Figure 1. Daucene formation can be explained by following the same scheme but leading, by a hydride shift, to another tertiary carbocation before the final proton loss.

Figure 10.

Proposed mechanism for CmSTC7 (Adapted with permission^{[ 14 ]}).

The purified CmSTC7 had the following kinetic constants: K _m= 3.5 μm, k _cat= 0.055 s⁻¹, and k _cat/K _m= 0.0159 μm ⁻¹ s⁻¹.

2.2.9. CmSTC8

GC chromatogram and GC retention times (Figure S69, Supporting Information) were indicative of the formation of numerous sesquiterpene hydrocarbons as well as some sesquiterpene alcohols. The main compound was identified as β‐copaene by comparison of the GC‐MS spectrum (Figure S70, Supporting Information) with the literature,^{[ 33 ]} establishing that CmSTC8 is a β‐copaene cyclase. The second largest peak was assigned by GC‐MS (Figure S71, Supporting Information) to germacrene D,^{[ 22 ]} and the two minor hydrocarbons (Figure S72 and S73, Supporting Information) were identified as bicyclogermacrene^{[ 33 ]} and δ‐cadinene,^{[ 33 ]} respectively. The largest alcohol peak was assigned (Figure S74, Supporting Information) to 1‐epi‐cubenol.^{[ 33 ]}

The previously proposed reaction mechanism of this enzyme begins with the NDP 1,10‐ring closure,^{[ 12 ]} a characteristic feature of clade II fungal STCs, as anticipated from Figure 1. All the assigned compounds are potentially produced from the last two carbocations, leading to β‐copaene, germacrene D, bicyclogermacrene, or δ‐cadinene by proton loss or to 1‐epi‐cubebol by addition of a water molecule (Figure  11 ).

Figure 11.

Proposed mechanism for CmSTC8 (Adapted with permission^{[ 14 ]}).

The purified CmSTC8 had the following kinetic constants: K _m= 8.6 μm, k _cat= 0.124 s⁻¹, and k _cat/K _m= 0.0144 μm ⁻¹ s⁻¹.

2.2.10. CmSTC11

The GC chromatogram (Figure S76, Supporting Information) indicated the formation of two sesquiterpene hydrocarbons. The minor compound exhibited a retention time and a GC‐MS spectrum identical to that of an authentic β‐farnesene standard (Figure S77, Supporting Information). The main peak was assigned to α‐farnesene by comparison with the mass spectrum (Figure S78, Supporting Information) described in the literature, thus, establishing CmSTC11 as an α‐farnesene synthase. A preparative experiment was conducted to obtain a sufficient amount of material for purification using silica gel chromatography and NMR analysis. The comparison of the ethylenic part of the ¹H NMR spectra (Figure S79, Supporting Information) of the purified mixture arising from CmSTC11 catalysis on FDP, with that of authentic β‐farnesene and the one of α‐farnesene reported in the literature,^{[ 34 ]} confirmed the presence of an ≈2/1 mixture of α/β farnesene isomers. The formation of both farnesene isomers can be easily explained by the formation of the farnesyl cation through the loss of the diphosphate ion, followed by proton loss, leading to a conjugated diene system (Figure  12 ).

Figure 12.

Proposed mechanism for CmSTC11.

3. Discussion

Fungi are recognized as a producers of a vast diversity of terpenes and terpenoid derivatives. In particular, their genomes are rich in genes encoding STCs, which catalyze the cyclization of FDP into sesquiterpene backbones.^{[ 9} , ^{35 ]} It is known that basidiomycete fungi produce a range of sesquiterpenes that are involved in environmental communication, such as attracting beneficial organisms, repelling harmful ones, and facilitating interactions between fungi and between fungi and plants.^{[ 36} , ^{37 ]} However, fewer than 30 basidiomycete species have been thoroughly investigated for their repertoires of STCs, leading to the characterization of ≈150 fungal STCs. Among basidiomycetes, the order Polyporales contains the majority of long‐lived fungi that colonize wood. Polyporales are estimated to be 183 million years old, and we expect that such an extensive evolutionary timeframe could promote an increase in the diversity of STC enzymes.^{[ 38 ]} The identification of 18 genes encoding putative STCs in C. menziesii provides an opportunity to explore in detail the terpenome of this polypore fungus. Furthermore, to the best of our knowledge, no secondary metabolites, including terpenes, have been extracted so far from C. menziesii, and none of the TCs encoded in its genome have been characterized. As a representative of the Polyporaceae family, we aimed to enhance our understanding of the capacity of C. menziesii to generate terpenes for comparative studies among basidiomycete fungi. Out of 18 predicted TCs from C. menziesii, 10 were overproduced and their products were characterized primarily using NMR and GC‐MS. The main compounds produced by CmSTCs were Δ⁶‐protoilludene (CmSTC1), germacrene A (CmSTC2, 3 and 18), δ‐cadinol (CmSTC5 and 16), trans‐dauca‐(11),8‐diene (CmSTC7), β‐copaene and germacrene D (CmSTC8), 4‐epi‐cubebol (CmSTC9), α‐ and β‐farnesene (CmSTC11), and germacrene‐D‐4‐ol (CmSTC16 and 18) (see Table S5, Supporting Information, for a summary). To date, only three studies have investigated the complete sesquiterpenome of fungi from the Polyporaceae family, i.e., in Cerrena unicolor,^{[ 39 ]} Trametes versicolor, ^{[ 40 ]} and Lignosus rhinocerotis. ^{[ 41 ]} Only the Δ⁶‐protoilludene cyclase,^{[ 40 ]} δ‐cadinol cyclase^{[ 39} , ^{41 ]} and trans‐dauca‐4(11),8‐diene cyclase^{[ 40 ]} were commonly found within the C. menziesii terpenome. The protein sequence of CmSTC1 shared 42% identity (Figure S80, Supporting Information) with the Δ⁶‐protoilludene cyclase from T. versicolor.^{[ 40 ]} The protein sequence of CmSTC5 shared 45% and 54% identity (Figure S81, Supporting Information) with the δ‐cadinol cyclases from C. unicolor ^{[ 39 ]} Cun7050 and Cun0716, respectively, and 53% identity with the one from L. rhinoceratis. ^{[ 41 ]} The protein sequence of CmSTC7 shared 54% identity (Figure S82, Supporting Information) with the trans‐dauca‐4(11),8‐diene cyclase^{[ 40 ]} from T. versicolor. Thus, even within the same taxonomic family, fungal STCs with the same product specificity exhibit only ≈40–55% sequence identity. We also compared the sequence of CmSTC1 and CmSTC5 with the characterized homologs from other fungi within the Basidiomycota division. CmSTC1 shared 39–45% identity with 15 characterized Δ⁶‐protoilludene cyclases.^{[ 7 ]} However, this identity dropped to 8% when all the sequences were compared altogether (data not shown). In the case of the δ‐cadinol cyclase CmSTC5, the pairwise alignment with five other characterized δ‐cadinol cyclases showed 42–53% sequence identity, whereas the overall identity between the whole set of sequences dropped to 27% (data not shown). These results indicate a relatively low sequence conservation within the Basidiomycota division for enzymes that catalyze the same reaction but are derived from different fungi within the Basidiomycota division. Consequently, it is challenging to infer the selectivity of such enzymes based solely on sequence identity within the subphylum Basidiomycetes. Notably, the 4‐epi‐cubebol cyclase activity of CmSTC9 has not yet been described in basidiomycete fungi.^{[ 7 ]}

We also compared the cyclization mechanisms inferred from the identification of the reaction products with those predicted by the phylogenetic analysis. We observed that the cyclization mechanisms predicted for clade II and clade IV STCs (nerolidyl cation 1,10‐cyclization and nerolidyl cation 1,6‐ or 1,7‐cyclization, respectively) were consistent with the identified reaction products. In contrast, we found that some of the CmSTCs that clustered in clade III did not catalyze the expected farnesyl cation 1,11‐cyclization. Instead, they catalyzed the farnesyl cation 1,10‐cyclization, which is typical of clade I. This discrepancy can be attributed to the low confidence we observed in the separation of clade I from clade III in our phylogenetic analysis (bootstrap value of 74 out of 100 sampled tree configurations, Supporting Information). This finding underscores the need for caution when predicting the cyclization mechanism of STCs from clades I and III based solely on the protein sequences. Notably, CmSTC6 did not cluster with enzymes from any of the four clades in our phylogenetic analysis. This observation highlights the potential for identifying additional STC clades as more fungal enzymes are studied, and aligns with the possibility for a fifth clade, as suggested by other groups.^{[ 40 ]}

At the molecular level, some CmSTCs were identified as intriguing catalysts. Specifically, the k _cat from CmSTC1 and CmSTC3, which exclusively produced hydrocarbons, were 3.2 and 2.4 s⁻¹, respectively. In contrast, the k _cat values for CmSTCs that generated alcohols were lower (0.08 s⁻¹ for CmSTC5, 0.33 s⁻¹ for CmSTC9, and 0.57 s⁻¹ for CmSTC18). All of these k _cat values compared favorably with those of other STCs, such as the protoilludane cyclase from the basidiomycete Omphalotus olearius (k _cat= 0.22 s⁻¹),^{[ 14 ]} the hirsutene cyclase from Coprinus cinereus (k _cat= 0.67 s⁻¹),^{[ 42 ]} and the improved double mutant of plant amorphadiene cyclase (k _cat= 1.00 s⁻¹).^{[ 43 ]}

4. Conclusion

The 10 out of 18 putative STCs from C. menziesii have been characterized in this study, thereby expanding the known repertoire of STCs from basidiomycete fungi. As more and more terpenomes are characterized, the role of terpenes and terpenoids in the chemical ecology of these organisms will become clearer. Furthermore, the determination of kinetic constants during this study revealed that some of these enzymes hold significant potential for biotechnological applications. Our findings suggest that the cyclization mechanism was conserved among STCs from phylogenetic clades II and IV. In contrast, the phylogenetic distinction between clades I and III is insufficiently robust to determine whether the enzymes in these clades catalyze farnesyl cation 1,10‐cyclization or farnesyl cation 1,11‐cyclization. Our work highlight the significant potential of polypore fungi, such as C. menziesii, for the discovery of new enzymes. We identified a CmSTC that does not cluster within any of the four clades previously described in the literature, and the reaction product of this enzyme remains to be identified. Among the 10 CmSTCs we analyzed, we identified one CmSTC that produced a sesquiterpene (4‐epi‐cubebol) not previously reported in basidiomycetes. The 18 CmSTCs discussed here were identified through classical basic local alignment search tool (BLAST) searches for proteins that shared a sequence similarity with previously characterized enzymes. Recently, 10 additional STCs have been identified from the genome sequence of C. menziesii using hiden markov model (HMM) profiles.^{[ 9 ]} These enzymes are phylogenetically closely related to the 18 enzymes described here, suggesting that the predictions for STC activity are accurate and that the sesquiterpenome of C. menziesii may be wider than anticipated (Figure S3, Supporting Information). Among these 10 additional STCs, eight clustered in clade IV, indicating that sequence similarity searches are less efficient for this clade of enzymes. Characterizing the reaction products of these enzymes will be essential to enhance our understanding of the overall sesquiterpenome of this fungus. This study also paves the way for a more detailed investigation into the role of each STC, and for the potential localization of the genes into biosynthesis gene clusters, within the genome of C. menziesii. Finally, the forthcoming exploration of genomic and postgenomic data, using data mining tools, such as HMM profiles will undoubtedly reveal a broader diversity of STC enzymes in these fungi.

5. Experimental Section

5.1.

5.1.1.

Phylogenetic Analysis

The predicted protein sequences of CmSTCs and 32 previously characterized STCs were aligned using multiple alignment fast fourier transform (MAFFT).^{[ 44 ]} Spurious sequences or poorly aligned regions were removed using trimAl.^{[ 45 ]} A total of 188 residues were utilized for phylogeny analysis using PhyML 3.3.20,220,408,^{[ 46 ]} the LG substitution matrix, a discrete gamma model, and 100 bootstrap samples. All steps were conducted on the Galaxy Europe server. The phylogenetic tree was visualized using Itol,^{[ 47 ]} and the protein accession numbers are presented in Table S6, Supporting Information.

Strain and Plasmid Construction

The coding sequences of the CmSTCs (Figure S1, Supporting Information) were codon‐‐optimized, synthesized, cloned into a pET‐22b(+) vector by Gibson assembly (refer to Supporting Information) and transformed into different E. coli BL21(DE3) strains (see text). The resulting transformants were subsequently spread on LB (Lysogeny Broth) agar plates supplemented with ampicillin (100 μg mL⁻¹).

Construction of CmSTC1‐18 Expression Plasmids by the Gibson Assembly Method

The full‐length plasmid was amplified by inverse polymerase chain reaction (PCR) using the primers “pET22b_6H‐N or C‐term” forward (Fw) and reverse (Rev) (Table S7, Supporting Information) and the full‐length Cmstc genes were PCR‐amplified using a personal collection as template, with the primers “CmSTC_6H‐N or C‐term” forward (Fw) and reverse (Rev), except for Cmstc7, 10 and 16 genes, see below (Table S8, Supporting Information).

The PCR reaction in a total volume of 50 μL (35.5 μL distilled water, 10 μL of Q5 5X buffer (NEB), 1 μL dNTP (10 mm), 1.25 μL of each primer (forward and reverse, each diluted to 20 μm), 0.5 μL of Q5 high‐fidelity DNA Polymerase (NEB)), and 5 ng of DNA template) was performed using the following program: 98 °C, 2 min; (98 °C, 20 s; specific primer Tm°C (calculated on https://tmcalculator.neb.com/#!/main), 30 s; 72 °C, 30 s kb⁻¹) × 30 cycles; and 72 °C, 5 min. Then, 8 μL of the PCR mix was digested for 3 h at 37 °C by DpnI (8 μL PCR mix, 1 μL DpnI, 1 μL 10X Cutsmart buffer (NEB)) to remove the PCR matrix.

Cloning was performed using the Gibson assembly approach^{[ 48 ]} at 50 °C during 1 h, using a molar ratio of 2:1 insert:vector (calculated at https://nebiocalculator.neb.com/#!/ligation), with a homemade enzymatic mix.

The cloned sequence was confirmed by Sanger sequencing (Eurofins Genomics, Cologne, Germany).

Heterologous Expression of C. Menziesii STCs and Purification

One clone from each CmSTCs transformant was picked up from an LB agar plate to inoculate tubes containing 5 mL of LB medium supplemented with ampicillin (100 μg mL⁻¹). After an overnight culture at 37 °C, this preculture was used to inoculate a baffled conical flask filled with one‐fifth (vol/vol) of TB (terrific broth) medium supplemented with ampicillin (100 μg mL⁻¹) at a dilution of 1:100 (vol/vol). The culture was incubated for ≈2 h at 37 °C and 180 rpm. When the culture OD reached a value between 1 and 2, induction was initiated by adding IPTG to a final concentration of 1 mm (0.4 mm IPTG + 2 mm rhamnose for the Lemo21(DE3) strain). Following induction, the culture was incubated at 18 °C for 20 h. The cells were harvested by centrifugation at 4 °C, 6000 g for 20 min and washed with cooled NPI‐10 buffer (50 mm NaH₂PO₄, 300 mm NaCl, 10 mm imidazole, pH 8) and resuspended in the same cooled buffer. After a second centrifugation (4 °C, 10,000 g, 30 min), the cells were frozen in liquid nitrogen and stored at 20 °C until use. For the purification of CmSTCs, the cells were thawed on ice, resuspended in NPI‐10 buffer at a concentration of 200 OD units mL⁻¹, and disrupted at a pressure of 1.3 kBar (cell disintegrator, CellD, TS serie) or by sonication (3 s pulse, 3 s rest, total duration 5 min, current 30%). The resulting mixture was then incubated at 4 °C, 30 rpm for 30 min with the addition of DNAse (DNase I from bovine pancreas, Roche diagnostics Gmbh, and final concentration, 0.5 mg mL⁻¹) and lysozyme (22,770 U mg⁻¹, Euromedex, and final concentration 1 mg mL⁻¹). The total soluble proteins were recovered after centrifugation at 6000 g for 30 min at 4 °C. The CmSTCs of interest were purified by affinity chromatography. The total soluble proteins were loaded onto a 5 mL Ni Histrap column (Cytiva) and eluted using an ÄKTA pure 25 (Cytiva) with a gradient of 20–250 mm imidazole in 50 mm Tris‐HCl, 150 mm NaCl, pH 7.5. Buffer exchange was performed using gel filtration on a PD10 column (Cytiva), with proteins eluted in 50 mm Tris‐HCl, 5 mm MgCl₂, and pH 7.5 buffer. The fractions were analyzed by sodium dodecyl sulfate ‐poly acrylamide gel electrophoresis (SDS‐PAGE) electrophoresis. The enzymes were stored at 4 °C until activity assays were conducted.

The small scale bioconversion assays were conducted using either chemically synthesized (E,E)‐farnesyl diphosphate (FDP), (E,E)‐geranyl diphosphate (GDP), or (E,E)‐geranylgeranyl diphosphate (GGDP) at a final concentration of 5 mm, in 50 mm Tris‐HCl, 5 mm MgCl₂, and pH 7.5, at 37 °C for 24 h with gentle agitation in a final reaction volume of 500 μL. The enzyme concentration varied from 1 to 14 mg mL⁻¹ (final concentration) depending on the enzyme used. For control experiments, either the enzyme or the substrate was omitted. The reaction products were extracted using 300 μL of pentane or ethyl acetate. After 1 min of vortexing and 1 min of centrifugation at 12,500 rpm, the organic phase was transferred to a GC vial for subsequent GC and GC‐MS analysis.

The large‐scale bioconversion assays were conducted on chemically synthesized (E,E)‐farnesyl diphosphate (final concentration: 5 mm), under the same conditions as those used in the small‐scale assays. The reaction products were extracted with pentane or ethyl acetate, dried over anhydrous sodium sulfate, filtered, evaporated, and purified by silica gel flash chromatography. Pentane was used as the eluent for sesquiterpene hydrocarbons, while a mixture of pentane/diethyl ether (8/2) was employed for sesquiterpene alcohols.

Kinetic Constants Determination

The determination of catalytic constants was performed with the green malachite method^{[ 18 ]} with slight modifications. In brief, the assays were monitored in 96‐well plates in a total reaction volume of 180 μL with a Tris‐HCl 50 mm, MES 25 mm, CAPS 25 mm, and MgCl₂ 5 mm buffer at pH 7.8 and 2U of pyrophosphatase from Saccharomyces cerevisiae. Different concentrations of FDP were used: 3.125 to 50 or 75 μm, and the reactions were started with an enzyme solution (0.09–0.49 μm, depending on the protein). Enzymatic reactions were stopped and developed by the addition of 24 μL of the malachite green solution and incubated for 10 min prior to readings at 623 nm on a UV–Vis spectrophotometer (U‐2810 Hitachi High‐Technologies Co., Tokyo, Japan). The catalytic constants (K _m, k _cat, k _cat/K _m) were determined with GraphPad Prism version 10.0.

Identification of C. Menziesii STCs Products

Small‐scale bioconversion assays were first analyzed using GC chromatography with a CLARUS 500 (Perkin Elmer) equipped with an Optima WAX Plus column (Macherey‐Nagel, 0.25 μm thickness, 30 m X 0.25 mm ID). The temperature program was as follows: 60 °C (1 min) followed by a ramp of 3 °C min⁻¹ up to 165 °C (5 min), and a second ramp of 3 °C min⁻¹ up to 220 °C (20 min). The inlet and detector temperatures were maintained at 250 °C. The GC‐MS experiments were performed using a GC‐2010 Plus (Shimadzu) coupled with a GC‐MS QP2010 SE mass detector (Shimadzu). The column was a ZB‐5MS (Zebron, 0.25 μm thickness, 60 m X 0.25 mm ID). Terpene separation was achieved using the following temperature program: 55 °C (4 min), followed by a ramp of 7 °C min⁻¹ up to 190 °C (30 min). The mass detector parameters for electron impact mode were set to 250 °C for the ion source, 200 °C for the MS quadrupole, and 70 eV for electron energy. The data were acquired in scan mode over a range of 50–300 amu. NMR experiments were routinely conducted on a Bruker AV400 nanospectrometer, while in‐depth NMR experiments were conducted on a Bruker AVL600 spectrometer, using CDCl₃ as the solvent in both cases. Optical rotations were measured using a Jasco P‐2000 polarimeter equipped with a sodium lamp (589 nm) and a halogen lamp (578 and 546 nm), in a 10 cm cell that was thermostated at 25 °C with a Peltier‐controlled cell holder.

Conflict of Interest

The authors declare no conflict of interest.

Supporting information

Supplementary Material

Data Availability Statement

The data that support the findings of this study are available in the supplementary material of this article.