Macrolides and Alcohols as Scent Gland Constituents of the

Article pubs.acs.org/jnp

Macrolides and Alcohols as Scent Gland Constituents of the Madagascan Frog Mantidactylus femoralis and Their Intraspecific Diversity Dennis Poth,†,‡ Pardha Saradhi Peram,† Miguel Vences,*,§ and Stefan Schulz*,† †

Institut für Organische Chemie, Technische Universität Braunschweig, Hagenring 30, 38106 Braunschweig, Germany Zoologisches Institut, Technische Universität Braunschweig, Mendelssohnstraße 4, 38106 Braunschweig, Germany

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S Supporting Information *

ABSTRACT: Acoustic and, to a lesser degree, visual signals are the predominant means of signaling in frogs. Nevertheless, certain lineages such as the mantelline frogs from Madagascar use the chemical communication channel as well. Males possess femoral glands on the hind legs, which recently have been shown to contain volatile compounds used in communication as pheromones. Many mantelline species occur in sympatry, and so far species recognition is regarded to occur mainly by acoustic signals. The analysis of the gland constituents of Mantidactylus femoralis by GC/MS revealed the presence of volatile macrolides and secondary alcohols. The new natural products mantidactolides A (4) and B (6), as well as several methyl carbinols, were identified, and their structures were confirmed by synthesis. The analysis of individuals from different locations of Madagascar revealed the presence of two groups characterized by specific patterns of compounds. While one group contained the alcohols and mantidactolide B, the other showed specific presence of the macrolides phoracantholide I (1) and mantidactolide A (4). Genetic analysis of some individuals showed no congruence between genetic relatedness and gland constituents. Several other individuals from related species had different gland compositions. This suggests that a basic set of biosynthetic machinery might be available to a broader group of related species.

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Mantidactylus multiplicatus, M. betsileanus, and several mantellid species of the genus Gephyromantis, it was shown that these scent-emitting femoral glands of the males contain speciesspecific mixtures or compounds, many of them with unknown structure.5 These compounds likely are used as pheromones, as has been shown for M. multiplicatus. Although a high interspecific variability in gland composition is obvious, the intraspecific diversity of the composition is as yet unstudied. The present study focuses on analyzing the volatile compounds from the femoral glands of Mantidactylus femoralis as well as the variation of these substances within populations of M. femoralis and closely related species in the subgenus Ochthomantis of Madagascan river bank frogs. Individual genetic analysis of the frogs determined their genetic relatedness, and the characterization of new compounds from M. femoralis glands demonstrated a considerable geographic variation of femoral skin secretions in this species.

mphibians are an animal class in which the knowledge on chemical communication is relatively scarce. The predominant means of signaling in anurans (frogs) is acoustic and, to a lesser degree, visual.1 Variations in the advertisement calls within species of frogs are typically restricted to the effects of temperature, individual body size, or hormone-related sexual motivation.2 Qualitative differences in call structure are, on the contrary, indicative of species-level differentiation and usually correlate with strong genetic differences. Despite the undoubted importance of bioacoustic communication in frogs, a considerable proportion of these animals are characterized by sexually dimorphic macroglands at different parts of their bodies,3 suggesting that the secretions of these glands may be of importance in their mating or territorial behavior. Although it is well established that amphibians use pheromones, until recently only water-soluble compounds such as peptides or prostaglandins have been identified as pheromones.4 In a recent study we have demonstrated that volatile alcohols or macrolides are also used by frogs as pheromones.5 In Madagascar and the Comoros, one endemic group of more than 200 species of frogs, the Mantellidae, contains various genera that are characterized by distinct glands on the underside of male thighs. These so-called femoral glands6 are particularly distinct in the genus Mantidactylus, where rudiments are also visible in females. In a previous study of © 2013 American Chemical Society and American Society of Pharmacognosy

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RESULTS AND DISCUSSION Identification of Volatile Femoral Gland Constituents of Mantidactylus femoralis. During the analysis of CH2Cl2 extracts of femoral glands from various Madagascan mantelline Received: February 11, 2013 Published: September 4, 2013 1548

dx.doi.org/10.1021/np400131q | J. Nat. Prod. 2013, 76, 1548−1558

Journal of Natural Products

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Figure 1. Gas chromatogram of the femoral gland extract of a single male Mantidactylus femoralis (individual number ZCMV 11251) collected at the Fotsialanana River. The main constituents A and B occur in high concentrations and are accompanied by a mixture of fatty acid ethyl esters and glycerides. X: artifact.

frogs we encountered an individual of M. femoralis from the Fotsialanana river collection site (in the Makira Reserve, eastern Madagascar) that contained two volatile compounds in high concentrations (Figure 1). Compound A was identified as phoracantholide I (1) due to its characteristic mass spectrum by comparison with data from the literature.7 It was first described as a constituent of the defense secretion of the Australian beetle Phoracantha synonyma, where it occurs as the R-enantiomer. Synthetic reference material of both enantiomers was obtained by hydrogenation of its unsaturated analogue phoracantholide J (2), which was previously synthesized.5 Gas chromatography on a chiral phase revealed that natural phoracantholide I (1) found in M. femoralis has the S-configuration (Figure S1), while beetles produce the opposite enantiomer.7

as phoracantholides I and J are likely derived from fatty acid biosynthesis. A common motif for the formation of methyl branches in fatty acid biosynthesis is the incorporation of methylmalonate instead of malonate during biosynthesis.9 This will result in the location of methyl groups at even-numbered carbon atoms. The resulting target structures are shown in Scheme 1, and the building blocks are highlighted. Because the Scheme 1. Possible Structures of Compound B According to Fatty Acid Biosynthesis

mass spectra of cyclic compounds do not allow easy location of methyl groups in cyclic compounds, all four structures were synthesized, and their mass spectra were compared with that of the natural compound. An approach using ring-closing metathesis10 with C6F6 activation10c followed by hydrogenation was used in all cases. 2-Methyl-9-decanolide (3) was synthesized from available phoracantholide J (2) by α-methylation of 2 with NaHMDS/ MeI, followed by hydrogenation of lactone 7 (Scheme 2). A short synthesis was developed for 4-methyl-9-decanolide (4, Scheme 3). Ethyl 3-bromopropionate (8) was coupled with isopropenylmagnesium bromide under Li2CuCl4 catalysis to form ethyl 4-methylpent-4-enoate (9). Saponification with KOH yielded 4-methylpent-4-enoic acid (10). Acid 10 was then coupled with hept-6-en-2-ol (12), which was obtained by the addition of 3-butenylmagnesium bromide (11) to

The other major component was an unknown natural product with a mass spectrum similar to that of 1, pointing to a macrolide structure. High-resolution mass spectrometry revealed an m/z of 184.1483, corresponding to a molecular formula of C11H20O2 and indicating the presence of an additional CH2 compared to phoracantholide I. Phoracantholide I exhibits a gas chromatographic retention index (I) of 1268, while compound B showed I = 1336. The difference (ΔI = 68) ruled out the possibility of an extended chain, as, for example, in 9- or 10-undecanolide, because this would require a ΔI of about 100.8 Therefore, compound B was likely a branched 9-decanolide in which the additional CH2 is located as a methyl group somewhere along the chain. Macrolides such 1549



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Scheme 2. Synthesis of 2-Methyl-9-decanolide (3)a

by hydrogenation of the double bond of unsaturated macrolide 19. The synthesis of 8-methyl-9-decanolide (6) is shown in Scheme 5. 4-Pentenoic acid (20) was converted into the corresponding acid chloride, coupled with (S)-4-benzyloxazolidin-2-one and stereoselectively methylated to form (S)-4benzyl-3-((S)-2-methylpent-4-enoyl)oxazolidin-2-one (21). Reduction with LAH led to alcohol 22. The oxidation of 22 to the corresponding aldehyde was followed by a Grignard reaction with methylmagnesium bromide to furnish (S)-3-methylhex-5en-2-ol (23). Next 23 was coupled with 5-hexenoic acid to yield the unsaturated ester 24. Cyclization with Grubbs-II catalyst formed the unsaturated lactone 25, which was hydrogenated in the last step to yield 8-methyl-9-decanolide (6). After the completion of the syntheses, the mass spectra and gas chromatographic retention times of the synthesized macrolides were compared (Figure 2). All compounds showed different mass spectra, and only that of 4-methyl-9-decanolide matched that of the natural compound B. We then set out to establish the absolute configuration of the natural compound by synthesizing an enantiomerically enriched stereoisomer (Scheme 6). Commercially available (+)-(β)-citronellene (26) was selectively epoxidized with mCPBA at the trisubstituted double bond.11 The resulting epoxide 27 was converted into the aldehyde 28 by a Lemieux−Johnson oxidation12 and further oxidized by Jones oxidation to yield the chiral acid 29. (S)-Hex5-en-2-ol (33) was obtained via a copper-mediated coupling of allyl bromide (32) and (S)-propylene oxide.13 Esterification of acid 29 with alcohol 33 led to the unsaturated ester 30, which was converted into the unsaturated lactone 31 via ring-closing metathesis. A final hydrogenation of the double bond formed the desired (4R,9S)-4-methyl-9-decanolide (4R,9S-4). The other three enantiomers were synthesized via the same route using appropriately configured starting materials. The building block (R)-4-methylhex-5-enoic acid (R-29) was synthesized from (−)-(β)-citronellene and (R)-hex-5-en-2-ol (R-33) obtained by the addition of allyl bromide to (R)-propylene oxide. The combination of the different enantiomers of 29 and 33 gives access to all four enantiomers of 4 via the respective ester 30. Using gas chromatography on a chiral Lipodex-G phase it could then be demonstrated that the natural compound from M. femoralis has the shown (4R,9S)-configuration (Figure S2). For this new natural product we propose the name mantidactolide A. In conclusion, the femoral gland constituents of this frog consisted of the two major components phoracantholide I (1) and mantidactolide A (4). Chemical Diversity of Femoral Gland Contents of Individuals. In addition to the specimen from the Fotsialanana River containing the two macrolides described above, femoral gland extracts of 11 individuals of M. femoralis collected at five different locations throughout Madagascar were analyzed. These samples contained a varying mixture of volatile compounds, some of which have not been described before as natural products. Figure 3 shows a total ion chromatogram of the femoral gland extract of an individual collected in Bemanevika containing the volatile compounds C−F. Surprisingly, neither macrolide 2 nor 4 was present in this sample. Instead a new macrolide with I = 1342 (F) was found. Compound F’s mass spectrum (panel F; Figure 2) was identical to that of macrolide 6 prepared previously. The retention index matched that of the later eluting diastereomer. Comparison of the H−H coupling constant J8,9 of the minor, first-eluting

a Conditions: (a) NaHMDS, MeI, CH2Cl2, 2 h, 72%; (b) H2, 10% Pd/ C, MeOH, 5 h, 62%, CH2Cl2.


(a) Li2CuCl4, isopropenylmagnesium bromide, THF, 0 °C, 12 h, 77%; (b) KOH, EtOH, H2O, 4 h, 80%; (c) (i) CuCN, Et2O, (ii) propylene oxide, 0 °C, 12 h, 81%; (d) 10 + 12, EDC·HCl, DMAP, CH2Cl2, 0 °C, 3 h, 75%; (e) Hoveyda−Grubbs II, C6F6, toluene, 80 °C, 3 h, 80%; (f) H2, 10% Pd/C, MeOH, 5 h, 59%. a

propylene oxide. After formation of unsaturated ester 13 a ringclosing metathesis using the Hoveyda−Grubbs II catalyst formed the unsaturated lactone 14. The target compound 4 was obtained via hydrogenation. The synthesis of 6-methyl-9-decanolide (5) was carried out in six steps starting from 6-methyltetrahydro-2H-pyran-2-one (15) (Scheme 4). Introduction of a methyl group in the αScheme 4. Synthesis of 6-Methyl-9-decanolide (5)a

a

Conditions: (a) LDA, MeI, THF, 93%; (b) LAH, Et2O, 56%; (c) MePPh3Br, nBuLi, NaHMDS, THF, −78 °C to rt, 31%; (d) 4pentenoic acid, EDC·HCl, DMAP, CH2Cl2, 0 °C, 3 h, 82%; (e) Hoveyda−Grubbs II, C6F6, toluene, 80 °C, 6 h, 48%; (f) H2, 10% Pd/ C, MeOH, 5 h, 31%.

position with LDA/MeI and the subsequent partial reduction with LAH furnished 3,6-dimethyltetrahydro-2H-pyran-2-ol (16), which was transformed by a Wittig reaction into alcohol 17. Coupling with 4-pentenoic acid furnished the ester 18, the precursor for the following ring-closing metathesis using the Hoveyda−Grubbs II catalyst. Finally, macrolide 5 was obtained 1550



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a Conditions: (a) oxalyl chloride, Et2O, rt, 67%; (b) (S)-4-benzyloxazolidin-2-one, THF, rt, 84%; (c) NaHMDS/MeI, THF, rt, 12 h, 74%; (d) LAH, Et2O, rt, 3 h, 68%; (e) oxalyl chloride, DMSO, Et3N, CH2Cl2, 0 °C; CH3MgBr, rt, 1 h, 40%, 2 steps; (f) 5-hexenoic acid, EDC·HCl, DMAP, CH2Cl2, rt, 5 h, 73%; (g) Grubbs II, toluene, 80 °C, 3 h, 45%; (h) H2, 10% Pd/C, MeOH, 5 h, 76%.

Figure 2. Mass spectra of natural compounds B and F and of the synthesized macrolides 3−6.

The other two unknown volatile compounds C and E had mass spectra (Figure 4) similar to that of 8-methylnonan-2-ol (D), suggesting these to be secondary alcohols as well. The retention indices indicated these alcohols to contain a methyl branch either at the ω-1, as in D, or at the ω-2 position. The variation of the retention index depending on the position of the methyl branch along the chain can be estimated using an empirical gas chromatographic retention index system, developed by us.15 Both compounds 6-methyl- and 7methyloctan-2-ol were then synthesized. 3-Methylpentanol (34) was converted into the corresponding bromide 35, which was then coupled with propylene oxide under copper catalysis, forming 6-methyloctan-2-ol (36) (Scheme 7). 7-Methyloctan-2-ol was synthesized by a similar route starting from 1-bromo-4-methylpentane (see SI). A comparison of the mass spectra revealed slight differences (Figure 4), and the retention index I = 1070 for 7-methyloctan2-ol was higher than for 6-methyloctan-2-ol (36), I = 1065. The

diastereomer (J8,9 = 3.4 Hz) and the major later eluting diastereomer (J8,9 = 10.4 Hz) with those predicted by software simulation14 (2.8 and 8.2 Hz) led to the tentative assignment of relative configurations. The first-eluting diastereomer (I = 1329) shows an (8R*,9S*)-configuration, while the later eluting natural diastereomer (I = 1346) shows an (8R*,9R*)configuration. Therefore, compound F was identified as the new natural compound 8-methyl-9-decanolide (6), which we propose calling mantidactolide B, most likely exhibiting an (8R*,9R*)-configuration. In only one of the samples we found also trace amounts of the (8R*,9S*)-diastereomer besides the major (8R*,9R*)-diastereomer. Compound D was readily identified as 8-methylnonan-2-ol, which is a known femoral gland constituent from M. multiplicatus.5 Gas chromatography on a chiral phase revealed that the natural 8-methylnonan-2-ol from M. femoralis also has the R-configuration (Figure S3), identical to the configuration occurring in M. multiplicatus. 1551



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Scheme 6. Synthesis of (4R,9S)-4-Methyl-9-decanolide (4R,9S-4)a

a

Conditions: (a) mCPBA, NaOAc, CH2Cl2, 2 h, 90%; (b) NaIO4, dioxane, H2O, 24 h, 94%; (c) CrO3, H2SO4, acetone, 10 min, 49%; (d) (i) Mg, Et2O, (ii) CuCN, (iii) (S)-propylene oxide, 0 °C, 12 h, 51%; (e) 29 + 33, EDC·HCl, DMAP, CH2Cl2, 0 °C, 3 h, 92%; (f) Grubbs II, C6F6, toluene, 80 °C, 3 h, 69%; (g) H2, 10% Pd/C, MeOH, 5 h, 74%.

Figure 3. Gas chromatogram of a femoral gland extract of an individual male Mantidactylus femoralis collected at Bemanevika. In addition to the already known compounds D (8-methylnonan-2-ol), G (squalene), and H (cholesterol), several other volatile alcohols, C and E, as well as the macrolide compounds F were present. X: artifact.

Figure 4. Mass spectra of (a) 7-methyloctan-2-ol, (b) 6-methyloctan-2-ol (C, 36), and (c) 8-methyldecan-2-ol (E, 39) and characteristic mass spectrometric fragmentation.

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Alcohol 37 was converted into the corresponding tosylate, which was then coupled with hex-5-enylmagnesium bromide under copper catalysis to form alkene 38. The double bond was oxidized via a Wacker oxidation, and the resulting ketone was subsequently reduced using LAH to yield 8-methyldecan-2-ol (39). The mass spectrum as well as the retention index of the synthetic compound matched those of the natural compound (C). The minor component 7-methyl-2-nonanol (40), occurring in some individuals, was tentatively identified by its mass spectrum and I value. M. femoralis is found along the eastern portion of Madagascar.16 Although some geographic variation can be expected among populations of such a widespread species, the diversity of compounds encountered in their femoral gland extracts was unsuspected. All the data refer to the glands of adult males, as samples from the rudimentary glands of female frogs and from belly skin did not contain any volatile compounds and are not shown. Only eight of the 12 individuals analyzed contained volatile compounds in their femoral glands. The absolute concentration of the samples varied widely. Some individuals contained up to 500 μg of volatile material, while others contained nothing or very small amounts, only. This divergence can be explained be either recent usage of the gland or lack of femoral gland constituent accumulation due to physiological conditions. Among individuals, varying mixtures of the identified alcohols and the three macrolides were present, which was dependent on their collection site (Figure 5).

Scheme 7. Synthesis of Natural Alcohols 6-Methyloctan-2-ol (36) and 8-Methyldecan-2-ol (39)a

a Conditions: (a) PBr3, pyridine, Et2O, 2 d, 80%; (b) (i) Mg, Et2O; (ii) CuCN, THF; (iii) propylene oxide, 12 h, 0 °C, 64%; (c) pyridine, TsCl, CH2Cl2, 1 h, 95%; (d) hex-5-enylmagnesium bromide, Li2CuCl4, 0 °C, 24 h, 53%; (e) PdCl2, Cu(OAc)2, O2, DMF, H2O, 72 h, 85%; (f) LAH, Et2O, rt, 2 h, 88%.

latter matched the value found for natural compound C, I = 1066. In the mass spectra of both C and synthetic 36 a fragment with m/z = 97 was generated, which is absent in the mass spectrum of 7-methyloctan-2-ol (Figure 4). This fragment is formed by the loss of water and a C2H5 fragment, as indicated in Figure 4. Compound C was identified to be 6-methyloctan-2ol (36). The typical fragmentation of a ω-2-branched methylcarbinol was also present in the mass spectrum of compound E, exhibiting the characteristic ion at m/z 125. Thus 8-methyldecan-2-ol (39) was proposed for alcohol E. A short synthesis starting from 2-methylbutan-1-ol (37) verified the proposed structure (Scheme 7).

Figure 5. Occurrence of volatile compounds in femoral gland extracts of Mantidactylus femoralis from different localities in Madagascar. The different marks in the table indicate the amount of the compounds compared to the largest peak. ×××: 30−100%, ××: 10−30%, ×: 1−10%, ○: below 1%. The samples varied in absolute amount of the volatile compounds. The samples from Analabe contained only low amounts, while the samples from Angozongahy and Vohiparara and one sample from Bemanevika showed higher concentrations of gland constituents. The largest amounts were present in the sample from Fotsialanana-Makira and one sample from Bemanevika. The color code is used to show collection sites on the map: (1) Angozongahy (yellow); (2) Analabe (red, 3 individuals), including individual ZCMV 12228; (3) Bemanevika (green, 2 individuals); (4) Vohiparara (blue), including individual ZCMV 8029; (5) Fotsialanana-Makira (gray), including individual ZCMV 11251. 1553



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Figure 6. Phylogeny of the subgenus Ochthomantis in the genus Mantidactylus based on DNA sequences (3071 bp) of 5 mitochondrial genes. The phylogram is a Bayesian 50% majority-rule consensus tree with other compatible groupings also shown. Bayesian posterior probabilities (BPP) >0.95 are shown by circles at nodes. The specimens colored in green contained volatile compounds in their femoral glands. No such compounds were found in the single analyzed gland of M. mocquardi (red); no data are available for the other species. Undescribed (candidate) species are numbered according to previous work.16

differences in the femoral gland compounds, this might suggest that M. femoralis as currently understood contains cryptic or incipient species. Individuals from Bemanevika were genetically similar to those from Analabe and also showed the greatest qualitative similarity in femoral gland compounds to samples from Analabe. While evolutionary diversification is one possible explanation for the differing femoral gland composition, also a food dependence might exist, although direct uptake of the compounds by feeding and storage in the femoral glands is unlikely to be the only mechanism involved. In laboratory raised M. betsileanus macrolide 2 is produced even when fruit flies were used as sole food source, which did not contain the macrolide (D. Poth, M. Vences, S. Schulz, unpublished results). This differs from poison frogs, which sequester dietary alkaloids in their skin and lack these toxins when captive-raised on alkaloid-free food.17 In individual samples of high concentration several trace components were detected that are prominent constituents of other individuals. The Fotsialanana sample contains traces of compound 6 and methyl 2-octenoate. One sample from Bemanevika contained the respective methyl ketones of the secondary alcohols as trace components. These findings might suggest that a relatively broad biosynthetic flexibility is present in the frogs, allowing an easy evolutionary alteration of major gland constituents, depending on the species or even the individual. In order to understand the geographic and species-specific nature of the differences in the femoral gland compositions, a molecular phylogenetic analysis based on the DNA sequences of mitochondrial genes was performed on samples from different locations, which also confirmed the taxonomy of the collected samples (Figure 6). This analysis shows that the analyzed M. femoralis indeed group together despite the genetic

The results showed that the individuals can be classified into two groups with similar chemistry. While the samples from Angozongahy, Analabe, and Bemanevika (group 1) contained secondary alcohols, these were lacking in extracts from Vohiparara and Fotsialanana (group 2). In contrast, phoracantholide I (1) and mantidactolide A (4) occurred only in individuals of group 2. Furthermore, mantidactolide B (6) is the major component of most individuals of group 1, but can be found only in traces in group 2. Nevertheless, these groups are not uniform in composition and differences can be seen between individuals. The methyl-branched alcohols D, 36, and 39 were present in all femoral gland extracts of group 1. (R)-8-Methylnonan-2-ol (D) was always the major secondary alcohol except in one sample from Bemanevika, in which 6-methyl-2-octanol (36) dominated. Alcohols 39 and 40 were mostly minor metabolites. Mantidactolide A (4) was the major component in the extract from Fotsialanana, but only a minor component in the Vohiparara individual, while both contained 1 in major and 6 in trace amounts. In one sample from Bemanevika macrolide 6 was only a minor component besides the major secondary alcohol D. Most surprising are the strong differences among the samples from Fotsialanana and Angozongahy, as these two sites are very close to each other; both are located in the Makira Reserve. In fact, various specimens from the Makira Reserve were genetically heterogeneous, as exemplified by those from Hevirina (which yielded only empty femoral gland extracts) and Fotsialanana (Figure 6). According to 16S rRNA sequence data (not shown; all sequences deposited in GenBank), the Angozongahy specimen was also somewhat genetically divergent despite the small geographic distances between these three sites (less than 5 km). Together with the strong 1554



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very vocal.16 When occurring in sympatry, they appear to differ in the pheromone cocktail of their gland secretions. On the other hand, at least the most widespread species M. femoralis presents a remarkable variation in these compounds across its distribution, even in neighboring populations such as those of Makira, and the possibility of incipient speciation processes involving chemical communication as a premating isolation mechanism should be researched further.

differences found among some of them (see above) and that the lineage of all M. femoralis samples is genetically distinct from all related species. Diversity between Species. To better understand patterns of intra- and interspecific variability of the femoral gland chemistry in mantellid frogs, extracts of various additional species related to M. femoralis, all belonging to the subgenus Ochthomantis (Figure 6), were analyzed. No volatile compounds were identified in the single sample available for M. mocquardi, but a variety of such compounds, which remain to be studied in detail, were found in M. majori and the undescribed candidate species Mantidactylus sp. 47 and Mantidactylus sp. 63. As revealed by the phylogenetic analysis, these species with M. femoralis represent all the major clades within Ochthomantis (Figure 6), suggesting that volatile compounds in femoral glands are probably universal within this subgenus. Although the amount of data is rather limited, the chemical composition of the gland secretions appears to be particularly different where species occur in sympatry. For instance, in the Ranomafana region, where M. majori and Mantidactylus sp. 47 occur together with M. femoralis (sample from Vohiparara), these three species do not share major or minor components of their glands. Phoracantholide I (1) was a major compound in M. femoralis from the Ranomafana area (Vohiparara) but at the same locality was not detected in M. majori or Mantidactylus sp. 47. The major compounds in three of four of the individuals of M. majori were identified as 8methylnonan-2-one and 10-methylundecan-2-one, while all four specimens of Mantidactylus sp. 47 contained primarily methyl 2-octenoate and, in lower concentrations, 9-methyldecan-2-one. Nevertheless, the close biosynthetic similarity to the alcohols from M. femoralis is evident for the ketones, and traces of methyl 2-octenoate can be found in some of the M. femoralis individuals. Of the two samples of Mantidactylus sp. 63 studied, both from the Analabe forest, one contained no volatile compounds, but the other shared compounds with various specimens of M. femoralis. For instance, it contained phoracantholide I (1), but remarkably this compound was not observed in the syntopic specimens of M. femoralis from Analabe. However, macrolide 1 was observed in other, geographically distant populations of M. femoralis from Fotsialanana and Vohiparara. In addition, the sample of Mantidactylus sp. 63 also contained phoracantholide J (2), which was previously identified from other species of Mantidactylus.5 Furthermore, several unidentified compounds occurred in these species.

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EXPERIMENTAL SECTION

General Experimental Procedures. Specific rotations were obtained with a Propol digital automatic polarimeter (Dr. Kernchen) with a 1 cm cuvette at a wavelength of 578 nm using the solvents reported. NMR spectra were obtained with the following instruments: Bruker DPX-200 (1H 200 MHz, 13C 50.5 MHz), DRX-400 (1H 400 MHz, 13C 101 MHz), or AV II-600 (1H 600 MHz, 13C 151 MHz). Chemical shifts are reported in ppm relative to tetramethylsilane as an internal standard (δ = 0). High-resolution MS data were obtained with a gas chromatograph (GC 6890, Agilent Technologies) equipped with a Phenomenex ZB5-MS column (30 m × 0.25 mm i.d. × 0.25 μm) coupled to a time-of-flight mass spectrometer (JMS-T100GC, GCAccuTOF, JEOL, Japan) in EI mode (70 eV). JEOL MassCenter workstation software was used. The system was tuned with PFK to achieve a resolution of 5000 (fwhm) at m/z 292.9824. GC-MS was performed on a HP 6890 gas chromatograph coupled to an MSD 5973 (EI 70 eV) (Hewlett-Packard) and on a GC 7890A coupled to an MSD 5975C (Agilent Technologies). Separation was performed on a fused-silica capillary column BPX-5 (SGE Inc., 25 m × 0.22 mm i.d. × 0.25 μm) and an HP5-MS (Agilent Technologies, 30 m × 0.25 mm i.d. × 0.25 μm). Chiral phase gas chromatography was performed using a Hydrodex-6-TBDMS phase (Macherey-Nagel, 25 m × 0.25 mm i.d.) or a Lipodex-G phase (Macherey-Nagel, 50 m × 0.25 mm i.d.). Commercially available starting material and solvents were purchased from Sigma-Aldrich and used without further purification. Technical solvents were distilled before use. All reactions involving watersensitive chemicals were performed in heat gun-dried glass equipment with magnetic stirring under a nitrogen atmosphere. TLC was performed on Polygram SIL G/UV254 plates (Macherey-Nagel) with detection by UV (254 nm) or by immersion in a 10% ethanolic solution of phosphomolybdic acid, followed by heating. Flash chromatography was performed on silica gel M60 (0.04−0.063 mm, 230−400 mesh ASTM) (Macherey-Nagel) under pressure or on a flash chromatograph (Combi Flash Companion, Teledyne Isco) with the eluent mentioned. Sample Preparation and Analysis. A total of 25 samples of different species were collected from sites within north and northeast Madagascar during the rainy periods of the years 2008−2012 and analyzed to investigate the gland constituents of individuals of the subgenus Ochthomantis. The femoral glands were excised and stored in vials containing dichloromethane. The samples were filtered, and the solution was analyzed using GC/MS. The detected volatile compounds were identified by comparison of their mass spectra, fragmentation patterns, and gas chromatographic retention indices with those of reference compounds, synthesized as described above. Gas chromatographic co-injection experiments verified their identity. Control samples of the belly skin and the femoral skin of females were also analyzed, but they did not contain any volatile compounds. It was not possible to obtain NMR data of the natural compounds due to the diverse sample compositions and low concentrations in the biological material. Molecular Genetics. In order to verify the species identity of sampled specimens, and due to the high morphological similarity of most species of Ochthomantis, a fragment of the mitochondrial 16S rRNA gene of all the relevant specimens was sequenced, following established protocols.18 These sequences were compared with a comprehensive sequence data set from a previous study.16 To comprehend the phylogeny of these frogs, sequences of additional mitochondrial genes were determined using standard methods and primers commonly used in Madagascan frogs.18,19,20 The final

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CONCLUSION In this study the structures of all the volatile compounds identified in the femoral gland extracts of M. femoralis collected at different Madagascar localities were confirmed by synthesis. Furthermore, the absolute configuration of the new macrolide natural product mantidactolide A (4), as well as the configuration of the already described compounds (R)-8methylnonan-2-ol (D) and (S)-phoracantholide I (1), was determined using gas chromatographic methods with chiral phases. In addition, mantidactolide B (6) was described as a new natural compound. The data herein demonstrate that, although the femoral gland constituents are mostly different among related species and especially in sympatry, macrolides are in general frequently occurring in the femoral glands of mantellid frogs. Species of the subgenus Ochthomantis are morphologically cryptic and not 1555



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= 6.3 Hz), 0.81 (3H, d, J = 6.8 Hz); 13C NMR (100 MHz, CDCl3) δ 174.3, 72.8, 34.6, 32.5, 31.5, 30.2, 28.1, 23.5, 21.0, 20.7, 19.3; EIMS (70 eV) m/z (%) 184 (1) [M]+, 169 (1), 124 (8), 112 (50), 96 (20), 83 (61), 69 (44), 55 (100); HREIMS m/z 184.1458 (calcd for C11H20O2, 184.1463); I = 1336. (3S)-3-Methylhex-5-en-2-yl hex-5-enoate (24). (3S)-3-Methylhex5-en-2-ol (23) (0.19 g, 1.7 mmol), 5-hexenoic acid (0.18 g, 1.57 mmol), and DMAP (21 mg, 0.17 mmol) were added to absolute CH2Cl2 (10 mL) at 0 °C. Then EDC·HCl (0.33 g, 1.75 mmol) was added to the above solution.24 The reaction mixture was stirred at 0 °C for 1 h and at rt for 5 h until the complete consumption of the alcohol (23) was observed by TLC. The reaction mixture was diluted with Et2O (40 mL) and washed with saturated NaHCO3 (2 × 30 mL). The organic phase was dried using MgSO4, and the crude product was purified by column chromatography to yield (3S)-3-methylhex-5-en-2yl hex-5-enoate (24) (0.26 g, 1.23 mmol, 73%). Rf = 0.4 (pentane/Et2O, 30:1); 1H NMR (300 MHz, CDCl3) δ 5.76 (2H, m), 5.01 (4H, m), 4.85 (1H, m), 2.29 (2H, t, J = 7.6 Hz), 2.19 (1H, m), 2.09 (2H, m), 1.89 (1H, m), 1.73 (3H, m), 1.16 (3H, d, J = 7.0 Hz), 0.91 (3H, d, J = 8.1 Hz); 13C NMR (75 MHz, CDCl3) δ 173.1, 137.7, 136.7, 116.2, 115.3, 73.2, 37.6, 37.2, 34.0, 33.1, 24.2, 17.0, 16.3; EIMS (70 eV) m/z (%) 166 (1), 128 (1), 114 (4), 99 (2), 97 (42), 81 (27), 72 (2), 69 (47), 68 (8), 55 (87), 43 (22), 41 (100), 39 (39). (5Z,8S)-8-Methyl-5-decen-9-olide (25). (3S)-3-Methylhex-5-en-2-yl hex-5-enoate (24) (77 mg, 0.36 mmol) was dissolved in dry toluene (250 mL), and hexafluorobenzene (4.4 g, 23 mmol) was added to the above solution. Then Grubbs II catalyst (1,3-bis(2,4,6-trimethylphenyl)-2-imidazolidinylidene)dichloro(phenylmethylene) (tricyclohexylphosphine)ruthenium (40 mg, 0.076 mmol) was added, and the reaction mixture was heated to 80 °C for 3 h. The reaction was cooled to rt and washed with saturated NaHCO3. The organic phase was dried using MgSO4. The organic solvents were evaporated, and the Grubbs catalyst was filtered off on a silica gel filled microcolumn. The column was washed (four column volumes) with pentane/Et2O (40:1). The column fractions were evaporated, and the crude prodcut was purified by column chromatography to yield (5Z,8S)-8-methyl-5decen-9-olide (25) (30 mg, 0.16 mmol, 45%). Rf = 0.52 (pentane/Et2O, 10:1); 1H NMR (300 MHz, CDCl3) δ 5.36 (2H, m), 4.71 (1H, qd, J = 6.5, 4.1 Hz), 2.27 (3H, m), 2.17 (2H, m), 1.91 (2H, m), 1.73 (2H, m), 1.20 (3H, d, J = 6.4 Hz), 0.98 (3H, d, J = 6.6 Hz); 13C NMR (75 MHz, CDCl3) δ 175.3, 133.9, 131.0, 75.0, 38.3, 33.7, 32.2, 26.5, 26.0, 17.3, 14.1; EIMS (70 eV) m/z (%) 182 (12) [M]+, 164 (4), 149 (5), 138 (14), 126 (12), 110 (25), 99 (89), 95 (32), 93 (17), 84 (37), 81 (71), 79 (40), 68 (26), 67 (81), 55 (76), 53 (41), 41 (100), 39 (80). (8S,9RS)-8-Methyl-9-decanolide (6). (5Z,8S)-8-Methyl-5-decen-9olide (25) (30 mg, 0.16 mmol) was dissolved in MeOH (30 mL, HPLC grade), and 10 mg of 10% palladium on activated carbon was added. Then hydrogen gas was passed into the reaction solution at a pressure of 2 bar for 5 h. Next the catalyst was filtered off on a Celitefilled microcolumn, and the column was washed with MeOH. The MeOH fractions were collected and evaporated to yield (8S,9RS)-8methyl-9-decanolide (6) (23 mg, 0.12 mmol, 76%). The ratio (8S,9S): (8S,9R) was 1:0.7. 1 [α]21.5 D −8.3 (c 0.7, CHCl3); H NMR (600 MHz, CDCl3) δ 4.8 (1H, qd, J = 6.8, 3.4 Hz), 4.4 (1H, dq, J = 10.4, 6.2 Hz) 2.4 (2H, m) 2.2 (1H, ddt, J = 10.8, 7.2, 3.6 Hz), 2.1 (1H, m), 2.0 (2H, m), 1.8 (1H, m), 1.7 (1H, m), 1.6 (2H, m), 1.4 (8H, m), 1.2 (4H, m), 1.1 (3H, d, J = 6.2 Hz), 1.1 (3H, d, J = 6.8 Hz), 0.8 (3H, d, J = 7.0 Hz), 0.7 (3H, d, J = 7.0 Hz); 13C NMR (151 MHz, CDCl3) (8S,9S)-6 δ 173.3, 75.7, 35.7, 35.1, 30.6, 27.7, 24.2, 22.5, 20.4, 19.8, 12.8; (8S,9R)-6 δ 174.0, 77.7, 40.0, 34.7, 29.7, 27.0, 25.9, 22.6, 20.6, 18.9, 17.8; EIMS (70 eV) m/z (%) 184 (1) [M]+, 166 (2), 148 (2), 140 (11), 123 (2), 112 (17), 109 (3), 98 (62), 94 (6), 83 (19), 81 (12), 69 (24), 67 (14), 56 (26), 55 (71), 53 (13), 45 (14), 42 (46), 41 (100), 39 (62); HREIMS m/z 184.1466 (calcd for C11H20O2 184.1463); I(8S,9S)‑6 = 1346, I(8S,9R)‑6 = 1329. 6-Methyloctan-2-ol (36). 3-Methylmagnesium bromide was prepared by dropwise addition of 1-bromo-3-methylpentane (35,

alignment contained fragments of the genes cytochrome b, cytochrome oxidase subunit I, 12S rRNA, 16S rRNA, and ND1. The sequences were resolved on an ABI 3130XL automated sequencer (Applied Biosystems). All newly determined sequences were submitted to GenBank (accession numbers KF426665-KF426724). The sequences were checked, and reading errors were corrected manually in CodonCode Aligner (CodonCode Corp.). The alignments were done in MEGA 521 and were unambiguous (only a few gaps required in the rRNA genes, mostly to accommodate the outgroup sequences). MrModeltest version 2.322 was used to select the best fitting nucleotide model of evolution under the Akaike information criterion (a GTR+I+G model). Phylogenetic analysis based on Bayesian inference was computed with MrBayes v3.0b423 using Markov chain Monte Carlo (MCMC) sets for 20 × 106 generations and sampled every 1000 generations. The trees corresponding to the first 10 × 106 generations were conservatively discarded as burn-in after empirically assessing the log-likelihood values of the sampled trees. Synthesis. (S)-Hex-5-en-2-yl (S)-4-methylhex-5-enoate (30). (S)4-Methylhex-5-enoic acid (29, 40 mg, 0.31 mmol), (S)-hex-5-en-2-ol (33, 36 mg, 0.36 mmol), and DMAP (4 mg, 0.03 mmol) were dissolved in 10 mL of absolute CH2Cl2 and cooled to 0 °C. N-EthylN′-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC, 76 mg, 0.4 mmol) was added in one portion, and the reaction was stirred for 1 h at 0 °C and for 2 h at room temperature (rt), similar to the procedure described by Patel et al.24 Then the reaction mixture was diluted with tert-butyl methyl ether, washed with saturated NaHCO3, and dried with MgSO4. After solvent removal under reduced pressure the crude product was purified by column chromatography on silica gel, yielding pure (S)-hex-5-en-2-yl (S)-4methylhex-5-enoate (30, 34 mg, 0.29 mmol, 92%). Rf = 0.53 (pentane/tert-butyl methyl ether, 40:1); 1H NMR (200 MHz, CDCl3) δ 5.52−5.96 (2H, m), 4.81−5.12 (5H, m), 2.20−2.33 (2H, m), 1.96−2.19 (3H, m), 1.45−1.80 (4H, m), 1.15−1.25 (3H, d, J = 6.1 Hz), 0.97−1.04 (3H, d, J = 6.8 Hz); 13C NMR (50 MHz, CDCl3) δ 173.4, 143.5, 137.8, 114.9, 113.5, 70.2, 37.5, 35.1, 32.5, 31.5, 29.7, 20.1, 19.9; EIMS (70 eV) m/z (%) 153 (1), 128 (40), 111 (36), 82 (57), 67 (72), 55 (100), 41 (49). (4S,5Z,9S)-4-Methyl-5-decen-9-olide (31). A solution of (S)-hex-5en-2-yl (S)-4-methylhex-5-enoate (30, 20 mg, 0.095 mmol) and hexafluorobenzene (1.8 mL, 6 mmol) was prepared in 150 mL of dry toluene according to the procedure of Rost et al.25 The Hoveyda− Grubbs-II catalyst dichloro[1,3-bis(2,4,6-trimethylphenyl)-2imidazolidinylidene](2-isopropoxyphenylmethylene)ruthenium(II) (12 mg, 0.019 mmol) was added to this solution, and the reaction was stirred for 3 h at 80 °C. The reaction was quenched by the addition of saturated NaHCO3 solution after cooling to rt, and the phases were separated. The organic layer was dried with MgSO4, and the solvent was removed under reduced pressure. After column chromatographic purification on silica gel (4S,5Z,9S)-4-methyl-5-decen-9-olide (31, 12 mg, 0.066 mmol, 69%) was obtained. Rf = 0.55 (pentane/tert-butyl methyl ether, 19:1); 1H NMR (400 MHz, CDCl3) δ 5.28−5.46 (1H, m), 4.98−5.15 (1H, m), 4.61 (1H, m), 2.16−2.25 (2H, m), 2.10−2.16 (1H, m), 1.94−2.05 (2H, m), 1.46−1.58 (4 H, m), 1.09 (3H, d, J = 6.5 Hz), 1.01 (3H, d, J = 7.0 Hz); 13 C NMR (100 MHz, CDCl3) δ 175.8, 132.6, 129.0, 71.1, 34.5, 33.1, 32.5, 32.0, 29.8, 21.7, 15.2; EIMS (70 eV) m/z (%) 182 (5) [M]+, 140 (5), 125 (7), 109 (15), 85 (100), 67 (51), 55 (31). (4R,9S)-4-Methyldecan-9-olide (4R,9S-4). A solution of (4S,5Z,9S)-4-methyl-5-decen-9-olide (31) (12 mg, 0.066 mmol) in 1 mL of absolute MeOH was prepared in a 1 mL vial, and 2 mg of 10% palladium on activated charcoal was added to hydrogenate the double bond according to the procedure of Kitahara et al.7 Hydrogen was bubbled through this solution with a pressure of 1 bar for 5 h. Then the catalyst was filtered off, and methanol was carefully evaporated in a gentle stream of nitrogen. Pure (4R,9S)-4-methyldecan-9-olide (4, 9 mg, 0.049 mmol, 74%) was obtained. +10.3 (c 0.74, CH2Cl2); 1H NMR (400 MHz, CDCl3) δ [α]26.7 D 4.88−5.00 (1H, m), 2.07−2.21 (2H, m), 1.77−1.96 (2H, m), 1.63− 1.75 (2H, m), 1.51−1.62 (3H, m), 1.25−1.43 (4H, m), 1.20 (3H, d, J 1556



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500 mg, 3.05 mmol) to a solution of activated magnesium turnings (0.9 g, 4 mmol) in 5 mL of absolute Et2O following the procedure of Yu et al.26 The solution was stirred for 3 h and then transferred into a cooled solution (−40 °C) of copper cyanide (0.3 g, 0.37 mmol) in 3 mL of absolute THF. After 20 min propylene oxide (177 mg, 3.05 mmol) was added slowly. The reaction mixture was stirred for 12 h at 0 °C and then quenched with saturated NH4Cl solution. The aqueous layer was extracted three times with ethyl acetate, and the combined organic layers were washed with brine and dried with MgSO4. The crude product was purified by column chromatography on silica gel (pentane/tert-butyl methyl ether, 9:1) after evaporation of the solvent to yield 6-methyloctan-2-ol (36, 280 mg, 1.94 mmol, 64%). Rf = 0.6 (pentane/tert-butyl methyl ether, 2:1); 1H NMR (400 MHz, CDCl3) δ 3.79 (1H, sxt, J = 6.0 Hz), 1.8 (1H, brs) 1.28−1.51 (7H, m), 1.17−1.21 (3H, d, J = 6.0 Hz), 1.06−1.16 (2H, m), 0.86 (3H, t, J = 7.6 Hz), 0.85 (3H, d, J = 6.8 Hz); 13C NMR (100 MHz, CDCl3) δ 68.0, 39.6, 36.5, 34.3, 29.4, 23.4, 23.2, 19.1, 11.3.; EIMS (70 eV) m/z (%) 129 (3), 111 (5), 97 (24), 83 (9), 69 (42), 57 (24), 55 (44), 45 (100), 41 (34), 39 (11); HREIMS m/z 129.1283 (calcd for C8H17O1 (M-15), 129.1279); I = 1065 8-Methyldecan-2-ol (39). 8-Methyldecan-2-one (162 mg, 0.96 mmol) was dissolved in dry Et2O (30 mL) under an inert atmosphere, and the flask was cooled to 0 °C. Following the procedure described by Hansen et al.,27 LiAlH4 (36.5 mg, 0.96 mmol) was added to the solution and the reaction was stirred at rt until the complete consumption of starting material was observed by TLC. The reaction was quenched using saturated NH4Cl, and the aqueous phase was separated after dissolution of the formed aluminum with concentrated HCl. After extraction with Et2O (3 × 30 mL) the combined organic phases were dried with MgSO4, the solvent was removed, and the crude product was purified by column chromatography to yield pure 8methyldecan-2-ol (39, 145 mg, 0.84 mmol, 88%). Rf = 0.5 (pentane/tert-butyl methyl ether, 5:1); 1H NMR (400 MHz, CDCl3) δ 3.79 (1H, sext, J = 6.0 Hz), 1.34−1.51 (3H, m), 1.24−1.34 (9H, m), 1.19 (3H, d, J = 6.1 Hz), 1.06−1.16 (1H, m), 0.85 (3H, t, J = 7.1 Hz), 0.84 (3H, t, J = 6.3 Hz); 13C NMR (100 MHz, CDCl3) δ 68.2, 39.4, 36.6, 34.4, 30.0, 29.5, 27.1, 25.8, 23.5, 19.2, 11.4; EIMS (70 eV) m/z (%) 157 (1), 125 (10), 97 (9), 83 (22), 70 (27), 55 (38), 45 (100), 41 (53); HREIMS m/z 157.1592 (calcd for C10H21O1 (M-15), 157.1589); I = 1268.

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framework of collaborations with the Département de Biologie Animale, Université d’Antananarivo. We are grateful to the Madagascar Institute for the Conservation of Tropical Environments MICET and the Valbio biological station for logistic support, and to the Madagascan authorities for granting research and export permits. We thank the Deutsche Forschungsgemeinschaft for research grant SCHU 984/10-1.

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(1) (a) Ryan, M. J. In The Evolution of the Amphibian Auditory System; Fritzsch, B., Ed.; New York, 1988; pp 637−677. (b) Hödl, W.; Amezquita, A. In Anuran Communication; Ryan, M., Ed.; Washington DC, 2001; pp 121−141. (2) (a) Schneider, H.; Sinsch, U. In Amphibian Biology, Vol. 7. Systematics; Heatwole, H. H., Tyler, M., Eds.; Chipping Norton, UK, 2006; pp 2892−2932. (b) Vences, M.; Wake, D. B. in Amphibian Biology, Vol. 6, Systematics; Heatwole, H. H., Tyler, M., Eds.; Surrey Beatty: Chipping Norton, UK, 2007; pp 2613−2669. (3) Thomas, E. O.; Tsang, L.; Licht, P. Copeia 1993, 1993, 133−143. (4) (a) Belanger, R. M.; Corkum, L. D. J. Herpetol. 2009, 43, 184− 191. (b) Houck, L. D. Annu. Rev. Physiol. 2009, 71, 161−176. (c) Woodley, S. K. J. Comp. Physiol. A 2010, 196, 713−727. (5) Poth, D.; Wollenberg, K. C.; Vences, M.; Schulz, S. Angew. Chem., Int. Ed. 2012, 51, 2187−2190. (6) Vences, M.; Wahl-Boos, G.; Hoegg, S.; Glaw, F.; Spinelli Oliveira, E.; Meyer, A.; Perry, S. Biol. J. Linn. Soc. 2007, 92, 529−539. (7) (a) Kitahara, T.; Koseki, K.; Mori, K. Agric. Biol. Chem. 1983, 47, 389−393. (b) Moore, B. P.; Brown, W. V. Aust. J. Chem. 1976, 29, 1365−1374. (8) van den Dool, H.; Kratz, P. J. Chromatogr. 1963, 11, 463−471. (9) Draeger, G.; Kirschning, A.; Thiericke, R.; Zerlin, M. Nat. Prod. Rep. 1996, 13, 365−375. (10) (a) Kingsbury, J. S.; Harrity, J. P. A.; Bonitatebus, P. J., Jr.; Hoveyda, A. H. J. Am. Chem. Soc. 1999, 121, 791−799. (b) Grubbs, R. H.; Chang, S. Tetrahedron 1998, 54, 4413−4450. (c) Rost, D.; Porta, M.; Gessler, S.; Blechert, S. Tetrahedron Lett. 2008, 49, 5968−5971. (11) Cernigliaro, G. J.; Kocienski, P. J. J. Org. Chem. 1977, 42, 3622− 3624. (12) Mori, K. Tetrahedron 2008, 64, 4060−4071. (13) Pungaliya, C.; Srinivasan, J.; Fox, B. W.; Malik, R. U.; Ludewig, A. H.; Sternberg, P. W.; Schroeder, F. C. Proc. Natl. Sci. Acad. U.S.A. 2009, 106, 7708−7713. (14) ACD/Structure Elucidator, version 12.01; Advanced Chemistry Development, Inc.: Toronto, ON, Canada, www.acdlabs.com, 2009. (15) (a) Schulz, S. Lipids 2001, 36, 637−647. (b) Dickschat, J. S.; Martens, T.; Brinkhoff, T.; Simon, M.; Schulz, S. Chem. Biodiversity 2005, 2, 837−865. (c) Nawrath, T.; Gerth, K.; Müller, R.; Schulz, S. Chem. Biodiversity 2010, 7, 2228−2253. (16) Randrianiaina, R. D.; Strauß, A.; Glos, J.; Glaw, F.; Vences, M. Contrib. Zool. 2011, 80, 17−65. (17) Daly, J. W.; Spande, T. F.; Garraffo, H. M. J. Nat. Prod. 2005, 68, 1556−1575. (18) Vences, M.; Kosuch, J.; Glaw, F.; Böhme, W.; Veith, M. J. Zool. Syst. Evol. Res. 2003, 41, 205−215. (19) Vences, M.; Vieites, D. R.; Glaw, F.; Brinkmann, H.; Kosuch, J.; Veith, M.; Meyer, A. Proc. R. Soc. B 2003, 270, 2435−2442. (20) Vences, M.; Nagy, Z. T.; Sonet, G.; Verheyen, E. Springer Prot. Meth. Mol. Biol. 2012, 858, 79−107. (21) Tamura, K.; Peterson, D.; Peterson, N.; Stecher, G.; Nei, M.; Kumar, S. Mol. Biol. Evol. 2011, 28, 2731−2739. (22) Nylander, J. A. A. MrModeltest version 2.3; Evolutionary Biology Centre, Uppsala University: Uppsala, Sweden, 2004. (23) Ronquist, F.; Huelsenbeck, J. P. Bioinformatics 2003, 19, 1572− 1574. (24) Patel, J.; Hoyt, J. C.; Parry, R. J. Tetrahedron 1998, 54, 15927− 15936. (25) Rost, D.; Porta, M.; Gessler, S.; Blechert, S. Tetrahedron Lett. 2008, 49, 5968−5971.

ASSOCIATED CONTENT

* Supporting Information S

Synthetic procedures, chromatography on chiral phases, and NMR spectra are available free of charge via the Internet at http://pubs.acs.org.

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REFERENCES

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Tel: (+49)-531-3915271 Fax: (+49)-531-3915272. Home page: http://www.oc.tu-bs.de/ schulz/index.html. Present Address ‡

Center for Marine Biotechnology and Biomedicine, Scripps Institution of Oceanography, UCSD, 8655 Kennel Way, La Jolla, CA 92037, USA. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We are grateful to F. Glaw, James and C. Patton, E. Rajeriarison, T. Rajoafiarison, R. D. Randrianiaina, F. Ratsoavina, and D. R. Vieites for their help in the field, and to J. Glos, S. Ndriantsoa, A. Rakotoarison, J. Riemann, and M.O. Rödel for providing samples. The fieldwork was supported by the Volkswagen Foundation and carried out in the 1557



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(26) Yu, M.; Alonso-Galicia, M.; Sun, C. W.; Roman, R. J.; Ono, N.; Hirano, H.; Ishimoto, T.; Reddy, Y. K.; Katipally, K. R.; Reddy, K. M.; Gopal, V. R.; Yu, J.; Takhi, M.; Falck, J. R. Bioorg. Med. Chem. 2003, 11, 2803−2821. (27) Hansen, D. B.; Starr, M. L.; Tolstoy, N.; Joullie, M. M. Tetrahedron: Asymmetry 2005, 16, 3623−3627.

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Macrolides and Alcohols as Scent Gland Constituents of the

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