5Z,9S - ACS Publications - American Chemical Society

Article Cite This: J. Nat. Prod. 2018, 81, 2266−2274

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Identification and Enantiodivergent Synthesis of (5Z,9S)‑Tetradec-5en-9-olide, a Queen-Specific Volatile of the Termite Silvestritermes minutus Aleš Machara,* Jan Křivánek, Klára Dolejšová, Jana Havlíčková, Lucie Bednárová, Robert Hanus, Pavel Majer, and Pavlína Kyjaková*

J. Nat. Prod. 2018.81:2266-2274. Downloaded from pubs.acs.org by UNIV OF TEXAS AT EL PASO on 10/26/18. For personal use only.

Institute of Organic Chemistry and Biochemistry of the CAS, Flemingovo n. 542/2, 166 10, Prague 6, Czech Republic S Supporting Information *

ABSTRACT: The queens of social insects differ from sterile colony members in many aspects of their physiology. Besides adaptations linked with their specialization for reproduction and extended lifespan, the queens also invest in the maintenance of their reproductive dominance by producing exocrine chemicals signaling their presence to the nestmates. The knowledge of the chemistry of queen-specific cues in termites is scarce. In addition to the contact recognition based on cuticular hydrocarbons, long-range signals mediated by volatiles are expected to participate in queen signaling, especially in populous colonies of higher termites (Termitidae). In queens of the higher termite Silvestritermes minutus (Syntermitinae), we have detected a previously undescribed volatile. It is present in important quantities on the body surface and in the headspace, ovaries, and body cavity. MS and GC-FTIR data analyses led us to propose the structure of the compound to be a macrolide 10-pentyl-3,4,5,8,9,10hexahydro-2H-oxecin-2-one. We performed enantiodivergent syntheses of two possible enantiomers starting from enantiopure (S)-glycidyl tosylate. The synthetic sequence involved macrolide-closing metathesis quenched with a ruthenium scavenging agent. The absolute and relative configuration of the compound was assigned to be (5Z,9S)-tetradec-5-en-9-olide. Identification and preparation of the compound allow for investigation of its biological significance.

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When compared to the other major group of social insects, the ants, the knowledge on the chemistry of queen recognition and fertility signals in termites is rather scarce and restricted to only a few empirical reports from the past decade. Several of these studies put forward the role of cuticular hydrocarbons (CHCs) in queen recognition and queen fertility signaling, drawing a strong analogy to the situation in eusocial Hymenoptera, in which the crucial role of CHCs in queen signaling is well-established.3,4 It has been shown in several termite species that individual castes have caste-specific CHC profiles and that reproductive castes (kings and queens) differ from other colony members either qualitatively or quantitatively due to shifted proportions of several CHCs.5,6 In a wellstudied model of lower termites, Cryptotermes secundus (Kalotermitidae), silencing of a queen-specific P450 gene

ueens of social insects often dramatically differ from the sterile colony members in fundamental aspects of their physiology, anatomy, and behavior. Their reproductive monopoly is commonly accompanied by large body size, extreme fecundity, and long lifespan.1 This fully applies to the queens of termites, especially in the most advanced termite group, Termitidae (higher termites). Their queens may reach spectacular sizes due to the gradual growth of their abdomens (physogastry) during sexual maturation and the fecund period of their lives, which may reach up to more than a decade. Besides adaptations directly linked with their specialization for reproduction and extended lifespan, the queens also invest in the maintenance of their reproductive dominance in the colony, most commonly by producing queen-specific exocrine chemicals (fertility signals, queen pheromones, queen recognition pheromones). These affect the behavior and physiology of the nestmates and make them tend the queen and her offspring and forego their own reproduction.2 © 2018 American Chemical Society and American Society of Pharmacognosy

Received: July 30, 2018 Published: October 9, 2018 2266

DOI: 10.1021/acs.jnatprod.8b00632 J. Nat. Prod. 2018, 81, 2266−2274

Journal of Natural Products

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Figure 1. Caste specificity of chemical profiles. GC×GC chromatograms of headspace samples (A−E) of neotenic queens (A), eggs (B), primary king (C), workers (D), and soldiers (E) from one colony of S. minutus; the samples were collected from 2 h of headspace sampling using PDMScoated SPME fiber. GC×GC chromatograms of hexane extracts (F−H) of workers (F), neotenic queens (G), and a primary queen (H) from another colony. Asterisk indicates the queen-specific peak.

volatiles or CHCs, may eventually be explained by the differences in the social complexity and population sizes in different termite lineages. While in small societies (typically Kalotermitidae and Archotermopsidae) most individuals are in frequent contact with reproductives, the populous colonies of socially advanced species (such as many Rhinotermitidae and most higher termites) require long-range signals to spread the information on queen presence to the whole population of complex nests.2,14 The putative role of volatiles in queen signaling was later suggested also in the higher termite Nasutitermes takasagoensis (Termitidae: Nasutitermitinae), in which a queen-specific volatile has been identified as phenylethanol. However, the presumed function could not be verified, in part due to the difficulties in experimental manipulations with advanced termites living in populous colonies.14,15 Yet, these findings prompt the interest in airborne signals of queens in higher termites and their functional significance, possibly ranging from a variety of primary functions to the derived roles in signaling. We have investigated recently the chemical profiles of queens in several species of higher termites (Termitidae), with emphasis on low-molecular-weight secondary metabolites, eventually involved in queen-specific physiological features. These may include signaling of fertility to nestmates and maintenance of reproductive dominance, on one hand, or metabolism related to queen fertility and extended longevity on the other hand. In fertile queens of the South American higher termite Silvestritermes minutus (Termitidae: Syntermitinae), we detected a previously unknown volatile. The studied species is known for its unusual breeding system, called asexual queen succession, in which young colonies are headed by a pair of founding primary reproductives (primary queen and primary king), while in mature colonies the foundress is replaced by multiple neotenic queens that she produces from unfertilized eggs through thelytokous parthenogenesis.16 The reported compound was initially detected in the headspace of fertile queens of both types, while being absent in other castes and life stages, i.e., kings, workers, soldiers, nymphs, and young

resulted in the modification of the CHC profile of the queens toward that of the sterile colony members and led to behavioral syndromes indicating reproductive disinhibition of the nestmates. This suggested that CHCs indeed do convey the information on the presence of a functional queen in the colony and are the long-sought fertility signals and queen pheromones.7 Experimental application of a synthetic analogue of juvenile hormone, the major endocrine factor controlling reproduction, has been shown to shift the CHC profiles toward those of fertile reproductives in another lower termite, Zootermopsis nevadensis (Archotermopsidae).8 One specific CHC, heneicosane, has been recently shown to act as a queen and king recognition pheromone, affecting the behavior of the nestmates in the lower termite Reticulitermes f lavipes (Rhinotermitidae).9 However, the data from other studies suggest that the chemical signaling regarding a queen’s (and a king’s) presence and fertility may be more complex and that the set of queenproduced chemical cues is not restricted only to CHC. In several species across the phylogenetic diversity of termites, proteins and peptides exclusive to queens (sometimes also kings) have been found to be secreted by the reproductives.10 While some of them show similarities to antimicrobial peptides,11 the functional significance of proteinaceous secretions has not yet been elucidated. More importantly, a breakthrough discovery dating back to 2010 ascribed the role of queen pheromone to queen-produced volatiles.12 In the lower termite Reticulitermes speratus (Rhinotermitidae), the neotenic queens emit a blend of butyl butyrate and 2methylbutan-1-ol, which inhibits the reproductive potential of sterile colony members. The same compounds have also been found in the chemical profiles of eggs, have been shown to attract the workers, and have demonstrated antifungal properties. This suggests a plausible ancestral role of the compounds in antimicrobial defense, followed by the derived functions of worker attractants and inhibitors of their reproduction.12−14 The apparent controversy in the chemical identity of queen pheromones, identified to be either small 2267



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Figure 2. Mass spectra of the studied compound. (A) EIMS fragmentation of the native compound. (B) High-resolution CIMS fragmentation of the native compound. (C) EIMS fragmentation of the compound after hydrogenation. (D) EIMS fragmentation of DMDS-derivatized compound.

3 ng in the head and thorax, and 126 ± 60 ng in the body cavity of one female (body mass of mature females ranges from 10 to 25 mg of fresh weight). Tentative Identification. The electron ionization MS spectrum of the compound did not match with any reference in the spectral libraries and suggested the ion m/z 224 to be the molecular weight (Figure 2A), which was further confirmed by high-resolution EIMS, indicating the molecular formula of the compound as C14H24O2 (m/z 224.1771; calcd m/z 224.1776). Chemical ionization confirmed the molecular weight due to the presence of the [M + H]+ ion at m/z 225 and showed two dominant peaks at m/z 207 and m/z 189. The first one likely represents a loss of one H2O molecule, and the second corresponds to [M − 36]+, both being the fragmentation features sometimes encountered in macrolide structures (Figure 2B).17 The analysis of neotenic queen extract using gas chromatography (GC)−Fourier transform infrared (FTIR) spectroscopy detected well the studied compound and indicated two diagnostic bands. The first one, 1749 cm−1, signaled the presence of a carbonyl moiety and assigned the two oxygen atoms, proposed in the molecular formula, to an ester group. The second one, 3012 cm−1, indicates, along with a missing vibration in the 960−980 cm−1 region, the presence of a double bond in the Z configuration (Figure S1, Supporting Information).18−20

subfertile queens. Consequently, it has also been found in important quantities inside queen bodies, i.e., in the hemolymph, fat body, and ovaries. Here, we report on the isolation, identification, and enantiodivergent synthesis of the queen-specific compound.

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RESULTS AND DISCUSSION Extraction of the Compound and Its Distribution in Castes and Tissues. The chemical profiles obtained by GC×GC-TOFMS analyses of solvent washes as well as headspace samples of living primary and neotenic queens were dominated by one major peak (RI 1670 on a DB-5 column), whose quantity exceeded dramatically that of all other analytes (Figure 1). The same compound was detected, albeit in lower concentrations, in the headspace and solvent extracts of eggs laid by the queens. In contrast, the corresponding peak was never observed in the chemical profiles of primary kings, immature stages, and sterile castes, i.e., workers, soldiers, larvae, and nymphs. Analysis of solvent washes of neotenic females in different stages of sexual maturation (freshly molted, several days after molt, and mature neotenic queens) revealed a gradual increase in the quantity of the compound along with the maturation of the queens. Important quantities of the same compound were subsequently found in dissected tissues of mature neotenic queens, in amounts of 90 ± 35 ng (mean ± SD, n = 4) in the ovaries, 7 ± 2268



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The molecular formula suggested an additional degree of unsaturation to the two listed above, indicating again a possible ring structure of the compound. Therefore, we performed a microhydrogenation of the extract with a Pd/C catalyst.21 The hydrogenated product showed a molecular mass two units higher (m/z 226), suggesting only one CC double bond in the molecule (Figure 2C), thus confirming the presence of the ring. The EIMS spectrum of the native compound further showed a fragment at m/z 153 [M − 71]+, which was also found as an m/z 155 equiv in the hydrogenated compound, suggesting the loss of a pentyl side chain by α-cleavage. The high relative intensity of these two corresponding fragments led us to assign the structure as a 10-membered ring. We concluded that the molecule has a lactone structure with a saturated C5 side chain.17 The proposed structure is supported by the similarities of the MS spectra of the hydrogenated compound (Figure 2C) to that of 10-hexyloxecan-2-one (pentadecan-9-olide), listed in the NIST library (#194473), differing only by the side chain length. To assign the double-bond position in the ring, we performed an iodine-mediated methylthiolation using dimethyl disulfide (DMDS). The MS spectrum of the adduct (Figure 2D) showed highly abundant diagnostic signals at m/z 129 and m/z 147. The adduct of the unsaturated macrolide and dimethyl disulfide undergoes a McLafferty rearrangement involving the hydrogen at the β-position from the ring oxygen.17,22,23 The bond between the two thiomethyl groups cleaves, and the two observed fragments are formed by the cleavage of the ester bond. The “acid part” of the fragment loses H2O, forming the base peak at m/z 129. This indicates the likely presence of the double bond at position 5. Proposed fragmentation of the DMDS adduct is given in Figure S2, together with additional fragmentation support.17,24,25 The presented indices converge at the proposed structure (5Z)tetradec-5-en-9-olide, with unknown configuration at C-9 (Figure 3).

Scheme 1. Retrosynthesis Analysis

macrolide formation, would be performed by ring-closing metathesis under standard conditions.28−30 Because it is difficult to control the E/Z configuration of the double bond during the formation of medium-size rings via RCM,31 such cyclization most likely furnishes both E- and Z-isomers as was described previously.32,33 Nevertheless, the olefin configuration was of minor concern at this stage because we were dealing with a synthesis of a compound whose constitution (arrangements of atoms) had yet to be fully elucidated. The synthesis of the proposed macrolide started from commercially available optically pure (S)-glycidyl tosylate (1), whose epoxide moiety was opened by treatment with allylmagnesium bromide in the presence of a catalytic amount of Li2CuCl4 to yield the desired intermediate 2 (Scheme 2).34 Scheme 2. Synthesis of Macrolides with the (9R)Stereogenic Centera

Figure 3. Proposed structure of the studied compound.

Enantiodivergent Synthesis and Structure Confirmation. To confirm the structure and elucidate the absolute configuration at C-9, we proceeded to enantiodivergent synthesis of the compound. Retrosynthetic analysis is depicted in Scheme 1. Its rationale consisted in a utilization of easily accessible chiral building blocks, which could be converted into the target molecule through a series of steps that do not allow any racemization. A macrolide possessing the oxecin-2-one scaffold was envisaged to arise from connection of two terminal alkenyl moieties by ring-closing metathesis (RCM).23,26,27 The key ester should be accessible from Steglich esterification of a secondary alcohol with hex-5enoic acid. The chiral secondary alcohol was anticipated to derive from commercially available glycidyl tosylate via consecutive epoxide opening with a Grignard reagent followed by nucleophilic substitution of the tosylate with another alkylmagnesium halide. The last synthetic step, 10-membered

a Reagents and conditions: (i) allylmagnesium bromide, Li2CuCl4 (5 mol %), −40 °C, 1 h, 83%; (ii) butylmagnesium bromide, CuCl (3 mol %), rt, 4 h, 78%; (iii) hex-5-enoic acid, DCC, DMAP, rt, 2 h, 73%; (iv) Hoveyda−Grubbs second generation (5 mol %), toluene, 80 °C, 5 h, 41% (ratio E/Z 4:1).

Compound 2 was then involved in a second reaction with Grignard reagent using butylmagnesium bromide in the presence of CuI catalyst, affording secondary alcohol 3 in very good yield. Subsequent Steglich esterification under standard conditions gave the ester 4 in 73% yield. This key intermediate was subjected to macrolide formation mediated by the Hoveyda−Grubbs second-generation catalyst under high dilution in toluene at 80 °C. We were pleased to find that RCM resulted in the formation of a mixture of E (5) and Z (6) 2269



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olefins (ratio 4:1), whose separation from each other was feasible. Although the overall yield of macrolides was rather low (41%), there was enough material to proceed with further investigation. At this moment, we did not perform a rigorous separation of the E- and Z-isomers from each other because we were more interested in the configuration assignment at C-9. Comparison of retention characteristics of the compounds 5 and 6 with the native compound from the queen extract on a chiral-phase GC column did not show a close match, suggesting thus that the native compound is likely the (9S) enantiomer (Figure 4).

Scheme 4. Synthesis of Macrolides with the (9S)Stereogenic Centera

a Reagents and conditions: (i) butylmagnesium bromide, Li2CuCl4 (5 mol %), −40 °C, 94%; (ii) allylmagnesium bromide, CuCl (3 mol %), rt, 4 h, 82%, or allylmagnesium bromide, Li2CuCl4 (5 mol %), 40%; (iii) hex-5-enoic acid, DCC, DMAP, rt, 4 h, 77%; (iv) Hoveyda− Grubbs second generation (2.2 mol %), toluene, 80 °C, 3 h, 61% (ratio E/Z 4:1); (v) 2-morpholinoethyl isocyanide, 30 min then wash with aqueous citric acid (5%).

Figure 4. Chiral-phase GC analysis of neotenic queen extract (A), synthetic (5E,9R)-tetradec-5-en-9-olide 5, (5Z,9R)-tetradec-5-en-9olide 6 (B), (5E,9S)-tetradec-5-en-9-olide 10, and (5Z,9S)-tetradec-5en-9-olide 11 (C). Chromatograms B and C are the result of GC analysis of fractions enriched in the Z-isomers.

bromide in the presence of CuCl or Li2CuCl4 precatalyst provided the alcohol 8 in 82% and 40% yield, respectively. After successful preparation of the key intermediate 9, we focused on the RCM. In this case we decided to perform the metathesis with a few modifications. First, loading of catalyst was reduced to 2 mol %, and argon was allowed to bubble through the reaction mixture for the whole time. Second, the time scale of the RCM reaction was kept as short as possible (approximately 3 h). Regardless of these conditions, the overall yield of both products was still low. We deemed that the critical phase in the preparation of the macrolides is actually the evaporation of the solvent (approximately 1 L of toluene) after the completion of the reaction. During this timedemanding operation, still active metathetic catalyst could negatively affect the yield of the products. Especially, in the end of the evaporation, when the material is very concentrated, the residues of the catalyst can cause product decomposition. To address this problem, we used morpholinoethyl isocyanide, a reaction-quenching agent for olefin metathesis described recently.37 Application of this ruthenium scavenger resulted in the increase of the yield from 41% to 61%. In addition, during the separation of macrolide 11 by column chromatography, we have observed an improved purity of all collected fractions containing either 10 or 11. Eventually, the optically pure macrolide 11 with S configuration at C-9 was separated from the E-isomer 10 by repetitive column chromatography of fractions rich in 11. (5Z,9S)-Tetradec-5-en-9-olide (11) showed a perfect match with the native compound in the chiral-phase GC analysis (Figure 4C), as well as in EI- and CIMS fragmentation and FTIR spectra (Figures S1, S3). Identification of the queen-specific volatile compound in the higher termite S. minutus supports the presumed production of volatile queen fertility signals in species inhabiting complex nests containing large populations, especially the higher termites. Mature colonies of S. minutus contain multiple

We then proceeded to the synthesis of the (9S) enantiomer with the Z double-bond configuration. In principal, we could start from (R)-glycidyl tosylate, which is also commercially available. Instead, we decided to pursue the synthesis from the same building block as for the (9R) enantiomer. An intriguing feature of glycidyl tosylate is the presence of so-called “proenantiotopic” functionalities at carbons C-1 and C-3, making it suitable for enantiodivergent synthesis (Scheme 3). Scheme 3. Enantiodivergent Approach in the Synthesis

Referring to these functionalities as “proenantiotopic” indicates that a controlled sequence of chemical transformations can lead to both enantiomers of the target molecule.35 This feature can be exploited in the design of flexible synthetic routes that allow an “enantiomeric switch” at incipient chemical steps. In other words, an enantiodivergent approach is accomplished by changing the order of steps in the synthetic route without changing the reagents or the conditions for any of the chemical transformations.36 Therefore, a single enantiomer of the starting material could be used for the preparation of both enantiomers of the target molecule. Synthesis of the (9S) enantiomer required 1, which was in this case treated with butylmagnesium bromide at −40 °C, providing compound 7 in an excellent yield of 94% (Scheme 4). Subsequent reaction of tosylate 7 with allylmagnesium 2270



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(7.26 ppm for 1H NMR and 77.0 ppm for 13C NMR spectra). Highresolution mass spectrometry with electrospray ionization (HRMSESI, positive ion) was performed on an LTQ Orbitrap XL (Thermo Fisher Scientific). Origin of Termites. Several entire colonies of Silvestritermes minutus (Termitidae: Syntermitinae) were collected in 2015−2017 in the rainforest at multiple sites along the route to Petit Saut in French Guiana (N 5°02.6620−N 5°07.2020, W 53°03.2950−W 52°57.8780). The species builds small but conspicuous nests from soil material, usually situated at the base of a young palm tree. The living colonies were transported to Prague and kept in glass jars lined with original rainforest soil at high humidity (>85%) and 29 °C in complete darkness. Extraction and Quantification. Headspace extracts were collected for 2 h on a red Supelco solid-phase microextraction (SPME) fiber with poly(dimethylsiloxane) coating (PDMS), inserted through the cap into 2 mL glass vials containing the studied phenotypes (1 queen, 5 workers, 5 soldiers, 50 eggs, 1 primary king). Solvent extracts of the body surface were prepared by extracting coldanesthetized individuals (1 queen, 5 workers, 5 soldiers, 50 eggs, 1 primary king) in a 2 mL glass vial containing 200 μL of distilled nhexane at 4 °C for 10 min. For quantification of the compound in different queen tissues, four S. minutus neotenic queens were cold anesthetized and dissected into four parts, i.e., gonads, head and thorax, abdominal cavity, and digestive tube. Each tissue was homogenized individually and sonicated in glass vials with 100 μL of distilled n-hexane. The supernatant was transferred into another vial. The amount of the compound was estimated by comparison with the added internal standard 1-bromodecane. Chemical Identifications, Microderivatizations. Preliminary chemical analyses and quantifications were carried out using gas chromatography coupled with mass spectrometric detection (GCMS) on a TRACE 1310 gas chromatograph coupled with an ISQ LT mass spectrometer with electron ionization (70 eV) and a quadrupole mass analyzer (Thermo Scientific) equipped with a DB-5 column (30 m, i.d. 0.25 mm, 0.25 μm film thickness). Detailed analyses were performed using a two-dimensional gas chromatograph with time-of-flight mass spectrometric detection (GC×GC-TOFMS; Pegasus 3D, Leco), equipped with a combination of nonpolar ZB-5MS (30 m, i.d. 0.25 mm, 0.25 μm film thickness) and medium-polarity RTX-50 (1.5 m, i.d. 0.1 mm, 0.1 μm film thickness) columns. In both cases, the temperature program was 50 °C (1 min) to 320 °C (10 min) at 8 °C/min. The secondary column of GC×GC-TOFMS was set 10 °C higher. Chiral-phase analyses were carried out using gas chromatography with a flame-ionization detector equipped with a chiral-phase column (30 m, i.d. 0.25 mm, 0.25 μm film thickness) with a permethylated βcyclodextrin phase (HP-CHIRAL-20B, J&W Scientific) enabling separation of chiral compounds. The gas chromatograph (HP-6850, Agilent) inlet was heated to 200 °C and the flame-ionization detector to 220 °C, and hydrogen was used as a carrier gas at a flow rate of 1.5 mL/min. Temperature program of the oven was 70 (1 min) to 100 °C (0 min) at 1 °C/min and then to 200 °C (5 min) at 25 °C/min. FTIR spectra were recorded on a Nicolet 6700FT-IR spectrometer (Thermo Scientific) combined with an Agilent 6850 gas chromatograph. The temperature program on the DB-5 column (30 m, i.d. 0.32 mm, 0.25 μm film thickness) was 40 °C (2 min) to 240 °C (5 min) at 5 °C/min. A liquid-nitrogen-cooled photoconductive mercury− cadmium−telluride (MCT) detector was used with an FTIR resolution of 8 cm−1; the light pipe temperature was 200 °C. For microhydrogenation, 0.5 mg of Pd on activated charcoal (10 wt % loading; Sigma-Aldrich) was added into 50 μL of extract, and a light flow of H2 bubbled through the solution for 10 min. The resulting mixture was filtered through 0.1 g of silica gel in a Pasteur pipet, and a small aliquot analyzed on GC-MS. The position of the double bond was determined by MS fragmentation of queen body extracts treated with DMDS. Fifty microliters of extract was mixed with 50 μL of DMDS and 20 μL of I2 (60 mg/mL in ether) and kept in the dark at 40 °C overnight to allow

thousands of termites, and the nest itself is a structured construction with large foraging territory.16 Thus, also in this species, it is plausible to expect the need for a long-range queen signal. Lactones of various ring sizes and levels of unsaturation and with various side chains occur frequently as semiochemicals in insects, other animals, and microorganisms. They most commonly originate from fatty acids, making them readily available. This, combined with desirable physicochemical properties (e.g., volatility), makes them an ideal candidate for use in communication.38 Volatile emissions produced by the queens may eventually serve several purposes. On one hand, it can be used as a queen recognition pheromone, eliciting the tending behavior in the nestmates, even though this function has been attributed to cuticular hydrocarbons acting in direct contact recognition in R. flavipes.9 To test this hypothesis, we performed a series of experiments targeting the eventual perception of (5Z,9S)tetradec-5-en-9-olide by the colony members. These bioassays included electrophysiological experiments testing the perception of the compound by the brains of individual workers and soldiers, as well as open area tests of attractiveness or preference of the compound by groups of termites using the electroantennographic and behavioral techniques that we described earlier.39 Yet, we did not observe any indications that the compound is perceived and/or directly affects the behavior of the nestmates. Alternatively, and more likely, the compound can play the role of a queen primer pheromone, arresting the sexual maturation of other females under the presence of fertile queens.12,14,15 It is specifically in this context that a long-range queen signal is deemed to be required in large colonies.2,14 In S. minutus, this second function may apply especially during the replacement of the primary queen by its parthenogenetic daughters. Our previous observations have shown that the queen parthenogens only start their sexual maturation in the absence of an active primary queen, due to either her death or senescence. When an active primary queen is present in the colony, the parthenogenetic daughters remain subfertile, most often arrested in the fourth nymphal stage. By contrast, when these nymphs are separated from the queen, they rapidly molt into neotenic queens.16 As noted previously, the investigation of the presumed function of the volatile in reproductive inhibition is a particularly challenging task in higher termites, due to their large colony population, de novo built nests from soil material, and complex life habits, making it difficult to keep separated groups of termites outside their nests.14,15 Nevertheless, as observed in the lower termite R. speratus, the putative ancestral functions of the compound should also be taken into consideration (e.g., antimicrobial or antioxidative properties), given the known wide range of biological activities reported for lactones with medium-sized rings (8−10 membered)40−43 and also because of the relatively large amounts of the compound detected in the queens and its presence in different body tissues rather than in a specific exocrine organ.

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EXPERIMENTAL SECTION

General Experimental Procedures. Optical rotations were measured in CHCl3 using a PerkinElmer 341 polarimeter at the sodium D-line using a cell with a 100 mm path. IR spectra were recorded on PerkinElmer FT-IR or Bruker Alpha-P FT-IR spectrophotometers. 1H and 13C NMR spectra were recorded using Bruker 400 and 600 MHz instruments at ambient temperature. The chemicals shifts are given relative to the chloroform solvent signal 2271



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h, and then hexane (20 mL) was added. The formed suspension was filtered through a pad of Celite, and the filtrate was evaporated to dryness. Column chromatography (hexane/toluene, 1:1) of the residue furnished ester 4 (0.13 g, 73%) as an oil: Rf 0.4 (hexane/ toluene, 1:1); [α]D −2.0 (c 0.34, CHCl3); 1H NMR (400 MHz, CDCl3) δ 5.79 (m, 2H), 5.24−4.74 (m, 4H), 2.30 (t, J = 7.5 Hz, 2H), 2.21−1.97 (m, 4H), 1.73 (m, 2H), 1.69−1.58 (m, 2H), 1.58−1.46 (m, 2H), 1.40−1.16 (m, 6H), 0.98−0.77 (m, 3H); 13C NMR (101 MHz, CDCl3) δ 173.2, 137.9, 137.7, 115.3, 114.8, 73.5, 34.1, 33.8, 33.4, 33.1, 31.7, 29.6, 24.9, 24.2, 22.5, 13.9; HRESIMS m/z 275.1982 [M + Na]+ (calcd for C16H28O2Na 275.1981). Macrolide-Closing Metathesis. A solution of ester 4 (0.35 g; 1.39 mmol) and Hoveyda−Grubbs second-generation catalyst (0.025 g; 0.04 mmol) in toluene (750 mL) was at first bubbled thoroughly with argon for 15 min and then was stirred for 5 h at 80 °C. The reaction mixture was allowed to cool to rt and was concentrated down on a rotavap. GC analysis of the residue determined the ratio of the E/Z products as 4:1. Multiple column chromatography (in straight toluene) led to separation of both isomers from each other. This tedious technique furnished 0.10 g (32%) of E-isomer 5, which eluted first and 0.03 g (9%) of Z-isomer 6. E-isomer 5: oil; 1H NMR (400 MHz, CDCl3) δ 5.60−5.39 (m, 1H, H-6), 5.33−5.18 (m, 1H, H-5), 5.15−4.96 (m, 1H, H-9), 2.42−2.22 (m, 3H, H-2′, H-4′, and H-7′), 2.12−1.87 (m, 4H, H-2″, H-4″, H-7″, and H-3′), 1.81−1.72 (m, 1H, H-3″), 1.65−1.56 (m, 2H, H-8′ and H-8″), 1.47 (dd, J = 11.8, 6.1 Hz, 2H, H-10′ and H-10″), 1.38−1.20 (m, 6H, H-11′, H-11″, H-12′, H12″, H-13′, and H-13″), 0.96−0.79 (t, J = 11.8 Hz, 3H, Me); 13C NMR (101 MHz, CDCl3) δ 176.1 (C-1), 133.5 (C-6), 128.1 (C-5), 74.9 (C-9), 35.8 (C-10), 34.8 (C-2), 33.9 (C-4), 32.9 (C-8), 31.6 (C7 and C-12), 26.0 (C-3), 24.8 (C-11), 22.53 (C-13), 14.0 (C-14); HRESIMS m/z 224.1777 (calcd for C14H24O2 224.1776). Spectroscopic data of the Z-isomer 6 match with data of its enantiomer 11 (see below). (S)-2-Hydroxyhept-1-yl 4-methylbenzenesulfonate (7). To a solution of lithium tetrachlorocuprate (0.1 M solution in THF, 2.2 mL; 0.22 mmol) was added dropwise butylmagnesium bromide (1.0 M solution in diethyl ether, 5.7 mL; 5.7 mmol) at −50 °C. The mixture was stirred for 1 h while the temperature of the mixture ranged from −50 to −40 °C (measured by internal thermometer). The precooled solution of (2S)-glycidyl tosylate (1) (1.0 g; 4.38 mmol) in THF (20 mL) was added dropwise through a dropping funnel over about 1 h at −40 °C (internal temperature). The reaction mixture was stirred for 1 h, then was quenched with aqueous NH4Cl and warmed to rt. The product was extracted with EtOAc (3 × 20 mL), and the combined organic layers were washed with brine and concentrated in vacuo. The residue was purified with column chromatography (cyclohexane/EtOAc, 3:1) to afford 7 as a colorless oil (1.18 g, 94%): Rf 0.4 (cyclohexane/EtOAc, 2:1); [α]D +7.8 (c 0.34, CHCl3); 1H NMR (400 MHz, CDCl3) δ 7.82 (d, J = 8.0 Hz, 2H), 7.38 (d, J = 8.0 Hz, 2H), 4.23−4.02 (m, 1H), 3.96−3.87 (m, 1H), 3.85 (m, 1H), 2.47 (s, 3H), 2.13 (d, J = 4.6 Hz, 1H), 1.43 (m, 2H), 1.28 (m, 4H), 0.89 (t, J = 6.9 Hz, 3H); 13C NMR (101 MHz, CDCl3) δ 145.1, 132.7, 129.9, 127.9, 74.0, 69.5, 32.6, 31.6, 24.9, 22.5, 21.7, 13.9; HRESIMS m/z 309.1130 [M + Na]+ (calcd for C14H22O4NaS 309.1131). (S)-Dec-1-en-5-ol (8). To a solution of CuCl (0.009 g; 0.09 mmol) and 7 (0.53 g; 1.85 mmol) in THF (5 mL) was added allylmagnesium bromide (1 M solution in THF, 4.2 mL; 4.2 mmol) dropwise at rt. The reaction mixture was stirred for 4 h and then was quenched with aqueous NH4Cl. Extraction with EtOAc (3 × 10 mL), washing of the combined organic layers with brine, and concentration in vacuo gave a residue, which was purified by column chromatography (cyclohexane/EtOAc, 4:1) to afford alcohol 8 (0.24 g, 82%) as a colorless oil: [α]D +1.7 (c 0.40, CHCl3); the 1H NMR and 13C NMR data match with data of the above-mentioned enantiomer 3. ((S)-1-Pentylpent-4-enyl) hex-5-enoate (9). To a solution of sec-alcohol 8 (0.75 g; 4.8 mmol), hex-5-enoic acid (0.66 g; 5.8 mmol), and DMAP (0.18 g; 1.44 mmol) in CH2Cl2 (10 mL) was added solid DCC (1.39 g; 6.73 mmol) at rt. The reaction mixture was stirred for 4

the excess iodine to react with sodium thiosulfate. DMDS adducts were extracted with n-hexane and analyzed by GC-MS. Accurate mass measurements were performed on a GCT Premier (GC-TOFMS, Waters) instrument with electron and chemical ionization modes connected to an Agilent GC equipped with a nonpolar ZB-5MS column (Phenomenex; 30 m, i.d. 0.25 mm, 0.25 μm film thickness) using a temperature program of 50 °C (1 min) to 320 °C (10 min) at 8 °C/min. Samples were injected in splitless mode. General Procedures of the Synthesis. All reactions except those in aqueous media were carried out under an argon atmosphere or with the use of standard techniques for exclusion of moisture. Reactions were monitored by thin-layer chromatography on silica gel plates (Merck Kieselgel 60 F254) and visualized with UV and Seebach’s stain. Purification by column chromatography was typically performed using silica gel 60 (Merck), and solvent mixtures and gradients are recorded herein. Standard workup procedures were used for all reactions, and combined organic layers were dried over MgSO4 before evaporation. Chemicals. All commercial chemicals and solvents were reagent grade and used without further purification unless otherwise specified. The following solvents and reagents have been abbreviated: dichloromethane (CH2Cl2), N,N′-dicyclohexylcarbodiimide (DCC), N,N-dimethylaminopyridine (DMAP), tetrahydrofuran (THF), chloroform (CHCl3), ethyl acetate (EtOAc). Prior to use, THF was distilled from sodium/acetophenone and toluene was distilled from sodium. (S)-2-Hydroxyhex-5-en-1-yl 4-methylbenzenesulfonate (2). To a solution of lithium tetrachlorocuprate (0.1 M solution in THF, 6.8 mL; 0.68 mmol) was added dropwise allylmagnesium bromide (1.0 M solution in diethyl ether, 17.9 mL; 17.9 mmol) at −50 °C. The mixture was stirred for 1 h while the temperature of the mixture ranged from −50 to −40 °C (measured by internal thermometer). The precooled solution of (2S)-glycidyl tosylate (1) (3.14 g; 13.75 mmol) in THF (35 mL) was added dropwise through a dropping funnel over about 1 h at −40 °C (internal temperature). The reaction mixture was stirred for 1 h, then was quenched with aqueous NH4Cl and warmed to room temperature (rt). The product was extracted with EtOAc (3 × 20 mL), and the combined organic layers were washed with brine and concentrated in vacuo. The residue was purified with column chromatography (cyclohexane/EtOAc, 4:1) to afford 2 as a colorless oil (3.1 g, 83%). The analytical data were in accordance with those published in the literature.30 Rf 0.6 (cyclohexane/EtOAc, 2:1); 1H NMR (400 MHz, CDCl3) δ 7.83 (d, J = 8.0 Hz, 2H), 7.38 (d, J = 8.0 Hz, 2H), 5.79 (ddt, J = 17.0, 10.2, 6.7 Hz, 1H), 5.15−4.83 (m, 2H), 4.06 (d, J = 7.0 Hz, 1H), 4.00−3.83 (m, 2H), 2.31−2.08 (s, 3H), 1.55 (m, 2H); 13C NMR (101 MHz, CDCl3) δ 145.2, 137.6, 132.7, 130.1, 128.1, 115.5, 73.9, 68.9, 31.8, 29.5, 21.8; HRESIMS m/z 293.0819 [M + Na]+ (calcd for C13H18O4NaS 293.0818). (R)-Dec-1-en-5-ol (3). To a solution of CuCl (0.034 g; 0.344 mmol) and 2 (3.1 g; 11.48 mmol) in THF (30 mL) was added butylmagnesium bromide (2 M solution in THF, 12.6 mL; 25.2 mmol) dropwise at rt. The reaction mixture was stirred for 4 h and then was quenched with aqueous NH4Cl. Extraction with EtOAc (3 × 20 mL), followed by washing of combined organic layers with brine and concentration in vacuo gave a residue that was purified by column chromatography (cyclohexane/EtOAc 4:1) to afford compound 3 (1.40 g, 78%) as a colorless oil: Rf 0.7 (cyclohexane/EtOAc, 2:1); [α]D −1.5 (c 0.41, CHCl3); 1H NMR (400 MHz, CDCl3) δ 5.87 (ddt, J = 17.0, 10.2, 6.7 Hz, 1H), 5.07 (dd, J = 17.0, 1.9 Hz, 1H), 5.04−4.95 (m, 1H), 3.68−3.61 (m, 1H), 2.33−2.09 (m, 2H), 1.69−1.55 (m, 2H), 1.50−1.41 (m, 4H), 1.39−1.27 (m, 4H), 0.99−0.79 (m, 3H); 13 C NMR (101 MHz, CDCl3) δ 138.7, 114.6, 71.4, 37.4, 36.4, 31.9, 30.1, 25.3, 22.6, 14.0; HRESIMS m/z 156.1514 (calcd for C10H20O 156.1514). ((R)-1-Pentylpent-4-enyl) hex-5-enoate (4). To a solution of alcohol 3 (0.11 g; 0.70 mmol), hex-5-enoic acid (0.1 g; 0.85 mmol), and DMAP (0.026 g; 0.21 mmol) in CH2Cl2 (5 mL) was added solid DCC (0.20 g; 1.0 mmol) at rt. The reaction mixture was stirred for 2 2272


Journal of Natural Products h, and then hexane (30 mL) was added. The formed suspension was filtered through a pad of Celite, and the collected filtrate was evaporated to dryness. Column chromatography (hexane/toluene, 1:1) of the residue afforded an oily ester 9 (0.93 g, 77%): [α]D +0.5 (c 0.42, CHCl3); the 1H NMR and 13C NMR data match with data of the above-mentioned enantiomer 4. Macrolide-Closing Metathesis Using a Ru Scavenging Agent. A solution of ester 9 (0.60 g; 2.38 mmol) and Hoveyda− Grubbs second-generation catalyst (0.033 g; 0.052 mmol) in toluene (950 mL) was at first bubbled thoroughly with argon for 15 min and then was stirred for 3 h at 80 °C. The reaction was allowed to cool to rt and then was quenched with the addition of 2-morpholinoethyl isocyanide (0.03 g; 0.02 mmol). The reaction mixture changed color to brownish, and then the mixture was treated with a 5% aqueous solution of citric acid (20 mL). The organic layer was washed with H2O and concentrated in vacuo. GC analysis of the residue determined the ratio of the E/Z products as 4:1. Multiple column chromatography (toluene) led to separation of both isomers from each other. This tedious technique furnished 0.25 g (46%) of the Eisomer 10, which eluted first, and 0.80 g (15%) of the Z-isomer 11. (5E,9S)-Tetradec-5-en-9-olide (10): Rf 0.38 (toluene); [α]D +45.2 (c 0.26, CHCl3); 1H NMR (400 MHz, CDCl3) δ 5.62−5.38 (m, 1H), 5.32−5.14 (m, 1H), 5.09−4.91 (m, 1H), 2.38−2.22 (m, 3H), 2.15− 1.84 (m, 4H), 1.75 (m, 1H), 1.68−1.57 (m, 3H), 1.47 (m, 2H), 1.29 (m, 6H), 0.95−0.81 (m, 3H); 13C NMR (101 MHz, CDCl3) δ 176.1, 133.5, 128.1, 74.9, 35.8, 34.9, 33.9, 32.9, 31.7, 26.0, 24.8, 22.6, 14.0; HRESIMS m/z 224.1776 (calcd for C14H24O2 224.1776); (5Z,9S)Tetradec-5-en-9-olide (11): Rf 0.31 (toluene); [α]D +54.6 (c 0.47, CHCl3); 1H NMR (400 MHz, CDCl3) δ 5.42 (m, 1H, H-6), 5.36 (m, 1H, H-5), 4.85 (m, 1H, H-9), 2.46 (m, 1H, H-7′), 2.44−2.33 (m, 1H, H-2″, H-4′, and H-4″), 2.3−2.15 (m, 3H,), 2.05−1.93 (m, 1H, H-7″), 1.85 (m, 1H, H-10′), 1.81−1.69 (m, 2H, H-3′, and H-3″), 1.66−1.43 (m, 3H, H-8′, H-8″, and H-10″), 1.37−1.26 (m, 6H, H-11′, H-11″, H-12′, H-12″, H-13′, and H-13″), 0.89 (t, J = 6.9 Hz, 3H, CH3); 13C NMR (101 MHz, CDCl3) δ 174.4 (C-1), 129.8 (C-6), 129.6 (C-5), 73.4 (C-9), 35.7 (C-8), 34.9 (C-2), 31.8 (C-12), 31.3 (C-10), 25.9 (C-4), 24.9 (C-3), 24.8 (C-11), 23.2 (C-7), 22.6 (C-13), 14.0 (C14); HRESIMS m/z 224.1776 (calcd for C14H24O2 224.1776).

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ACKNOWLEDGMENTS

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REFERENCES

We are grateful to the Czech Science Foundation (18-21200S), the Institute of Organic Chemistry and Biochemistry, Czech Academy of Sciences (RVO: 61388963), and the Mobility Plus Programme of CAS and FNRS (FNRS-17-02). We thank Dr. A. Jirošová and J. Titzenthalerová for their contributions to this manuscript.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jnatprod.8b00632.

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Article

Fourier transform infrared spectra of the natural compound, synthetic (5Z,9S)-tetradec-5-en-9-olide, and synthetic (5E,9S)-tetradec-5-en-9-olide, proposed EIMS fragmentation of the DMDS adduct of (5Z,9S)-tetradec5-en-9-olide, EIMS fragmentation of the natural compound, synthetic (5Z,9S)-tetradec-5-en-9-olide, and synthetic (5E,9S)-tetradec-5-en-9-olide, and 1H and 13C NMR spectra for compounds 2−5, 7, 11 (PDF)

AUTHOR INFORMATION

Corresponding Authors

*Tel: +420-220-183-397. E-mail: [email protected]. *Tel: +420-220-183-319. E-mail: [email protected]. cz. ORCID

Aleš Machara: 0000-0002-1139-2762 Robert Hanus: 0000-0002-7054-1975 Pavlína Kyjaková: 0000-0003-0081-3717 Notes

The authors declare no competing financial interest. 2273



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(30) Sudina, P. R.; Motati, D. R.; Seema, A. J. Nat. Prod. 2018, 81, 1399−1404. (31) Montgomery, T. P.; Johns, A. M.; Grubbs, R. H. Catalysts 2017, 7, 87−124. (32) Fürstner, A.; Radkowski, K.; Wirtz, C.; Goddard, R.; Lehmann, C. W.; Mynott, R. J. Am. Chem. Soc. 2002, 124, 7061−7069. (33) Katsuta, R.; Masada, N.; Shimodaira, Y.; Ueda, S.; Yäjima, A.; Nukada, T. Tetrahedron 2017, 73, 1733−1739. (34) Li, J.; Zhao, C.; Liu, J.; Du, Y. Tetrahedron 2015, 71, 3885− 3889. (35) Hudlicky, T.; Price, J. D.; Rulin, F.; Tsunoda, T. J. Am. Chem. Soc. 1990, 112, 9439−9440. (36) Hudlicky, T.; Reed, J. W. Chem. Soc. Rev. 2009, 38, 3117−3132. (37) Szczepaniak, G.; Urbaniak, K.; Wierzbicka, C.; Kosiński, K.; Skowerski, K.; Grela, K. ChemSusChem 2015, 8, 4139−4148. (38) Schulz, S.; Hötling, S. Nat. Prod. Rep. 2015, 32, 1042−1066. (39) Jirošová, A.; Sillam-Dussès, D.; Kyjaková, P.; Kalinová, B.; Dolejšová, K.; Jančařík, A.; Majer, P.; Cristaldo, P. F.; Hanus, R. J. Chem. Ecol. 2016, 42, 1070−1081. (40) Dräger, G.; Kirschning, A.; Thiericke, R.; Zerlin, M. Nat. Prod. Rep. 1996, 13, 365−375. (41) Rukachaisirikul, V.; Pramjit, S.; Pakawatchai, C.; Isaka, M.; Supothina, S. J. Nat. Prod. 2004, 67, 1953−1955. (42) Riatto, V. B.; Pilli, R. A.; Victor, M. M. Tetrahedron 2008, 64, 2279−2300. (43) Baraban, E. G.; Morin, J. B.; Phillips, G. M.; Phillips, A. J.; Strobel, S. A.; Handelsman, J. Tetrahedron Lett. 2013, 54, 4058−4060.

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