Identification of Components of Male-Produced Pheromone of Coffee White Stemborer, Xylotrechus quadripes
- First Online:
- Cite this article as:
- Hall, D.R., Cork, A., Phythian, S.J. et al. J Chem Ecol (2006) 32: 195. doi:10.1007/s10886-006-9360-0
- 291 Views
The coffee white stem borer, Xylotrechus quadripes Chevrolat (Coleoptera: Cerambycidae), is the foremost pest of arabica coffee in India, Sri Lanka, China, Vietnam, and Thailand. Previous work showed that female beetles were attracted to traps baited with male beetles. Analyses of volatiles from male X. quadripes of Indian origin by gas chromatography (GC) linked to electroantennographic (EAG) recording from a female beetle antenna showed three male-specific components comprising more than 90% of the volatiles, two of which elicited EAG responses. The major EAG-active component was produced at up to 2 μg hr−1 insect−1 and was identified as (S)-2-hydroxy-3-decanone (I) by comparison of GC data, and mass (MS), infrared, and nuclear magnetic resonance (NMR) spectra with those of synthetic standards. The second component was identified as 3-hydroxy-2-decanone (II) produced in part by isomerization of I under the conditions of the GC analysis, although the NMR spectrum suggested it is naturally produced at up to 7% of I. The minor component that elicited an EAG response, present at 7% of the amount of I, was identified as (S,S)-2,3-dihydroxyoctane (III) from GC and MS data. 2-Hydroxy-3-octanone (0.2–0.5% of I), 2,3-decanedione (2% of I), 2-phenylethanol (3% of I), and octanoic acid (4% of I) were also identified in volatiles from male beetles. A general, stereospecific synthetic route to the enantiomers of 2-hydroxy-3-alkanones from the enantiomers of ethyl lactate was developed. The enantiomers of III were synthesized from (E)-2-octene by Sharpless asymmetric dihydroxylation. (S)-(I) was attractive to male X. quadripes in laboratory bioassays, but addition of (S,RS)-(III) at 10% of I reduced attractiveness. In field trials carried out in India with sticky, cross-vane traps, (S)- and (RS)-(I) attracted male X. quadripes and addition of (S,S)-(III) at 10% of I reduced attractiveness. Significant numbers of female Demonax balyi Pascoe (Coleoptera: Cerambycidae) were sometimes caught in traps baited with (S)-(I) alone.
Key WordsXylotrechus quadripescoffee white stem borerColeopteraCerambycidaemale sex pheromone(S)-2-hydroxy-3-decanone3-hydroxy-2-decanone(S,S)-2,3-dihydroxyoctaneDemonax balyi
The coffee white stem borer, Xylotrechus quadripes Chevrolat (Coleoptera: Cerambycidae) is the foremost pest of arabica coffee in India, Sri Lanka, China, Vietnam, and Thailand (Le Pelley, 1968; Rhainds et al., 2002). In India, it is estimated that over 9 million trees have to be destroyed each year because of attack by X. quadripes, costing around $40 m annually for replacement and loss of production.
The adults of X. quadripes have distinct flight periods, and the females lay eggs in crevices in the bark. The larvae tunnel into the trunk, blocking the tunnel with frass behind them to deter predators. The tunneling in the trunk and roots rapidly kills young plants of up to 7–8 yr old, whereas older plants may survive for a few seasons but eventually succumb. Current control measures include maintenance of good shade because the adults are more active in sunlight, but this can reduce photosynthesis and yield. Removal of loose bark or scrubbing the bark at the base of the tree discourages oviposition, and collar pruning of infested trees may be effective if the pest has not reached the roots. Benzene hexachloride applied to the trunk at critical times during the flight periods is still the only effective insecticide against young larvae of X. quadripes, but this has now been banned for agricultural use in India and many other countries. Once larvae have penetrated into the stem, trees should be removed and destroyed to prevent spread of infestation (Le Pelley, 1968; Sreedharan and Kumar, 2001; Rhainds et al., 2002).
Male X. quadripes were shown to attract female beetles in the field in India by Venkateshu et al. (1986). Production of a sex pheromone by males of a related species, the grape borer X. pyrrhoderus, was demonstrated by Iwabuchi (1982), and the chemical structures of the components were identified by Sakai et al. (1984). The same components were shown to be produced by the mulberry borer, X. chinensis (Iwabuchi et al., 1987; Kuwahara et al., 1987). Identification and synthesis of the male sex pheromone of X. quadripes could provide the basis for development of a monitoring system for this pest in order to determine its distribution, and to time control measures directed at young larvae more effectively. Mass trapping with pheromone-baited traps could provide a method for control of this pest because it occurs at relatively low density and the pheromone attracts female beetles, thus directly affecting egg-laying.
We have reported the identification of (S)-2-hydroxy-3-decanone as the major component of the male sex pheromone of X. quadripes from India (Hall et al., 1998). This was confirmed by Rhainds et al. (2001a), using insects from China, and those authors also reported the presence of 2,3-decanedione in volatiles from the male beetles. Here, we report in full the identification of the major component of the sex pheromone of male X. quadripes from India, identification of an alternative minor component, and laboratory and field bioassays of these components.
Methods and Materials
X. quadripes adults were collected as they emerged from infested logs held in a netted room at the Central Coffee Research Institute (CCRI) under ambient temperature and humidity. Insects were collected several times during the morning emergence hours (Venkatesha et al., 1995) to ensure that they did not mate, and were sexed (Visitpanich, 1994) and maintained in separate tubes. These were hand-carried from India and separately maintained in an insectary at 70% RH with 12:12 h L/D light and 27:23°C temperature cycles.
Volatiles were collected from 15 virgin male beetles held in a silanized, 2-liter round-bottomed flask containing a filter paper (15 cm diam; Whatman No. 52). Air was drawn into the flask through a filter of activated charcoal (10 × 2 cm; 10–18 mesh) at 2 l/min−1, and volatiles were trapped on a filter of Porapak Q (50–80 mesh; 200 mg; Waters Corp., USA) packed between silanized glass wool plugs in a Pasteur pipette (i.d. 4 mm). The Porapak Q was purified by Soxhlet extraction with chloroform for 8 hr, and filters were rinsed well with dichloromethane before use. Volatiles from 15 virgin female beetles were collected similarly. Trapped volatiles were eluted from the filter after 24–72 hr with dichloromethane (3 × 0.5 ml) or alternatively with CDCl3 (1 ml) or ether (1 ml) for nuclear magnetic resonance (NMR) analysis or microreduction, respectively. For the latter, a few grains of lithium aluminum hydride were added to the ether solution, and the mixture was shaken at intervals for 1 hr. Water (0.1 ml) was then carefully added and the mixture was shaken. The upper ether layer was removed, dried with magnesium sulfate and analyzed by GC.
A container wash was obtained by maintaining nine male beetles in a conical flask (250 ml) with a filter paper (7 cm diam; Whatman No. 52) for 48 hr and then removing the beetles and washing flask and filter paper with hexane (5 + 5 ml). A female beetle container wash was obtained similarly.
Whole-body washes were obtained by placing nine male or nine female beetles in hexane (2.5 ml) 3 hr after lights on. After 10 min, the solvent was removed, and the residue was rinsed with a further aliquot of hexane (1 ml).
Gas Chromatographic Analyses
Gas chromatographic (GC) analyses were carried out on Carlo Erba Mega instruments with fused silica columns (25 m × 0.32 mm i.d.) coated with polar CP Wax 52CB (Carbowax 20M equivalent; Chrompack, London, UK) or nonpolar CP Sil 5CB (methyl silicone; Chrompack). Carrier gas was helium at 0.5 kg cm−2 and oven temperature was programmed at 50°C for 2 min, then at 6°C min−1 to 230°C. Injection was splitless or split (50:1) (200°C), and detection was by flame ionization detection (FID; 220°C). GC retention times are reported as retention indices (RI) relative to those of normal hydrocarbons.
Enantiomeric purity was determined by GC analysis on cyclodextrin GC columns (CP-Chirasil-DexCB, 25 m × 0.32 mm i.d., Chrompack; or β-Cyclodextrin, 50 m × 0.22 mm i.d., SGE, Milton Keynes, UK.). These were operated isothermally at 130 or 135°C, respectively, with helium carrier gas at 0.5 or 1.5 kg cm−2, respectively, and injector and detector temperatures as described above.
Coupled Gas Chromatography-Mass Spectrometry
Gas chromatography-mass spectrometry (GC-MS) analyses were carried out on a fused silica column (25 m × 0.2 mm i.d.; CP Wax 52CB) directly linked to a Finnigan ITD 700 ion trap detector (Thermoquest, Hemel Hempstead, Herts., UK) operated in EI or CI (iso-butane) mode. GC conditions were as described above.
Coupled Gas Chromatography-Electroantennography
Linked gas chromatography-electroantennography (GC-EAG) analyses were carried out as described by Cork et al. (1990) with effluent from the GC column split equally between the FID and a small glass reservoir in the column oven, the contents of which were expelled at 15-sec intervals with nitrogen (500 ml min−1 for 3 sec) over an EAG preparation. In the GC analysis, we used a fused silica column (25 m × 0.32 mm i.d.; CP Wax 52CB) programmed at 50°C for 2 min, then 20°C min−1 to 100°C, and then 4°C min−1 to 260°C. For EAG preparations, the head of the beetle was excised and impaled on the indifferent glass capillary electrode. The tip of one antenna was then inserted into the recording glass electrode and the preparation positioned so that the stimulus was directed at the terminal segments of the antenna. The electrodes contained 0.1 M aqueous KCl solution with 10% polyvinylpyrrolidone to reduce evaporation, and were inserted into stainless steel electrode holders (Syntech, Hilversum, The Netherlands) mounted on micromanipulators (Leica Microsystems, Wetzlar, Germany). Electrical contact was made with chloridized silver wires to the amplifier (UN-06, Syntech), and both EAG and GC signals were collected and processed with Turbochrom 4 software (Perkin Elmer, Beaconsfield, Bucks., UK).
NMR spectra were recorded in CDCl3 on a JEOL EX270 machine at 270 MHz for 1H and 67.8 MHz for 13C. Enantiomeric purity was determined after addition of the chiral cosolvent, (R)-(−)-2,2,2-trifluoro-1-(9-anthryl)ethanol (10 mg; Sigma-Aldrich, Gillingham, Dorset, UK). Infrared (IR) spectra were recorded as thin films with a Perkin Elmer 298 grating spectrophotometer (Perkin Elmer, Beaconsfield, Bucks., UK).
Use of 2-tetrahydropyranyl or tert-butyl dimethylsilyl protecting groups gave 2-hydroxy-3-decanone without detectable racemization as shown in Figure 1. The former protecting group was preferred for large-scale synthesis, as described below.
Freshly distilled 3,4-dihydro-2H-pyran (50.4 g, 0.6 mol) was added dropwise to a solution of (S)-ethyl lactate (59 g, 0.5 mol; Sigma-Aldrich, Gillingham, Dorset, UK) and a few crystals of p-toluenesulfonic acid (PTSA) in dry diethyl ether (250 ml) cooled to 0°C under nitrogen. The reaction mixture was stirred for 4 hr at room temperature. Solid K2CO3 was added to the solution, and the mixture was extracted twice with saturated NaHCO3 solution. The organic phase was dried over MgSO4, filtered, and the solvent was removed on a rotary evaporator. The residue was distilled to give the product in 92% yield (92 g; b.p. 76–81°C/0.5 mmHg). 1H NMR: δ 1.27 (t, J = 7.2 Hz, 3H, CH2CH3), 1.45 (d, J = 7.2 Hz, 3H, CHCH3), 1.55–1.80 (m, 6H, CH2), 3.68 (m, 2H, O–CH2), 4.16 (q, J = 7.2 Hz, 2H, CH2CH3), 4.20 (q, J = 3.3 Hz, 1H, CH3CH–OH), 4.40 (d, J = 6.9 Hz, 1H, O–CH–O); MS: m/z (%) 200 (2), 184 (1), 157 (3), 144 (2), 130 (3), 119 (trace), 101 (10), 85 (100), 67 (5), 57 (5), 43 (10); GC analysis on the polar column showed the two diastereoisomers in a 68:32 ratio.
Lithium hydroxide monohydrate (4.6 g, 0.1 mol) was added to a stirred solution of ethyl 2-(2-tetrahydropyranyloxy)propanoate (20.2 g, 0.1 mol) in ethanol (50 ml) under nitrogen. After 4 hr of stirring, GC analysis showed no trace of the starting material. Ethanol was removed under reduced pressure and the residue was washed twice with petroleum ether (100 ml) to give lithium 2-(2-tetrahydropyranyloxy)propanoate as a white solid in 100% yield (18.0 g). 1H NMR: δ 1.40 (d, J = 7.3 Hz, 3H, CHCH3), 1.65–1.80 (m, 6H, CH2), 3.60 (m, 2H, O–CH2), 4.20 (q, J = 7.3 Hz, 1H, O–CHCH3), 4.40 (d, J = 6.9 Hz, 1H, O–CH–O).
Heptyl lithium was prepared from heptyl bromide (17.9 g, 0.1 mol) and lithium (1.75 g, 0.25 mol) in dry ether (100 ml) under argon. The heptyl lithium (95 ml, 66 mmol) was added dropwise to a mechanically stirred suspension of lithium 2-(2-tetrahydropyranyloxy)propanoate (10.8 g, 60 mmol) in dry ether (50 ml) under nitrogen and cooled in an ice bath. After stirring at room temperature overnight, the contents were poured onto ice and saturated NH4Cl solution. The aqueous layer was extracted twice with ether and the extracts were washed with saturated NH4Cl, dried with MgSO4, and filtered through a pad of Florisil. The solvent was evaporated under reduced pressure to give the crude 2-(2-tetrahydropyranyloxy)-3-decanone (14.7 g; 95%). Kugelrohr distillation of an analytical sample gave the pure product as a colorless oil (90%), b.p. 98°C/0.06 mmHg; MS: m/z (%) 211 (2, M−C2H4O)+, 182 (2), 173 (5), 155 (3), 137 (2), 127 (5), 114 (3), 95 (trace), 85 (100), 67 (3), 57 (5), 41 (10); GC analysis on the polar column showed the two diastereoisomers in a 68:32 ratio.
The crude 2-(2-tetrahydropyranyloxy)-3-decanone (14.7 g) was dissolved in methanol (120 ml), a few crystals of PTSA were added, and the solution was stirred at room temperature. GC analysis on the polar column indicated reaction was complete after 6 hr. A small amount of solid K2CO3 was added and most of the methanol was removed on a rotary evaporator under reduced pressure. The residue was dissolved in 1:1 diethyl ether/hexane, washed with saturated NaHCO3 and dried over MgSO4. After removal of solvents under reduced pressure, the residue was distilled to give a main fraction in 68% yield (b.p. 57–60°C/0.04 mmHg; 7.0 g). IR (film): ν 3420 cm−1 (O–H), 1710 cm−1 (C=O); 1H NMR: δ 0.88 (t, J = 6.9 Hz, 3H, CH2CH3), 1.27 (bm, 8H, CH), 1.38 (d, J = 6.9 Hz, 3H, CH3CHOH), 1.61 (m, 2H, CH2–CH2–C=O), 2.41 (dt, J = 16.8, 7.6 Hz, 1H, CH2–CHH–C=O), 2.52 (dt, J = 16.8, 7.6 Hz, 1H, CH2–CHH–C=O), 3.55 (d, J = 4.5 Hz, 1H, CH–OH), 4.24 (dq, J = 7.1, 4.5 Hz, 1H, CH3CHOH); 13C NMR: δ 14.05, 19.86, 22.59, 23.63, 29.00, 29.18, 31.63, 37.54, 72.56, 212.72; MS: m/z (%) 173 (M + 1)+ (15), 155 (3), 137 (5), 127 (40), 57 (100), 45 (60), 29 (30).
By GC analysis on the polar GC column, all samples of 2-hydroxy-3-decanone contained 5–10% of the isomeric 3-hydroxy-2-decanone. GC analysis of the enantiomers on the 25-m cyclodextrin column at 130°C gave baseline separation (retention times R 23.7 min, S 24.0 min). Both synthetic enantiomers showed 98% enantiomeric excess (ee), similar to those of the starting ethyl lactates. A high level of enantiomeric purity was also indicated by the 1H NMR spectrum of the S enantiomer in the presence of a chiral cosolvent using either the signals due to the CH3–CH–OH doublet or the CH3–CH–OH multiplet.
(S)-2-Hydroxy-3-octanone was synthesized similarly from (S)-ethyl lactate and 1-bromopentane in 62.5% yield (b.p. 110°C/15 mm). Spectroscopic data were identical with those of Mori and Otsuka (1985).
2,3-Dihydroxyoctane was first synthesized as a 40:60 mixture of the S,S and S,R diastereoisomers by reduction of (S)-2-hydroxy-3-octanone with lithium aluminum hydride in ether. The erythroS,R diastereoisomer was predicted to be the major product of this reduction, assuming coordination of the aluminum by the 2-hydroxy group, and this was confirmed by synthesis of the separate enantiomers (below). This diastereoisomeric mixture was used without further purification in laboratory bioassays.
(S,S)-2,3-Dihydroxyoctane was synthesized by reaction of (E)-2-octene (Sigma-Aldrich) with the Sharpless reagent “AD-mix-α” (Sigma-Aldrich) in tert-butyl alcohol in the presence of methanesulfonamide (Sharpless et al., 1992). The product was obtained in 91% yield after chromatography and Kugelrohr distillation (b.p. 135°C/4 mmHg). Similar reaction of (E)-2-octene with the “AD-mix-β” (Sigma-Aldrich) gave (R,R)-2,3-dihydroxyoctane in similar yield. Both compounds were completely free (≤0.2%) of erythro diastereoisomers by GC analysis. 1H NMR: δ 0.89 (t, J = 6.6 Hz, 3H, CH3–CH2–), 1.18 (d, J = 6.3 Hz, 3H, CH3–CH(OH)–), 1.50–1.22 (m, 8H, 4 × CH2), 2.47–2.41 (br m, 2H, 2 × OH), 3.48–3.46 (br m, 1H, CHOH), 3.58 (quint, J = 6.3, 1H, CH3–CHOH); 13C NMR: δ 14.0, 19.4, 22.6, 25.2, 31.8, 33.3, 70.8, 76.2. These data were identical with literature values (e.g., Mori and Otsuka, 1985).
Analyses of the synthetic 2,3-dihydroxyoctanes on the 50 m cyclodextrin GC column at 135°C showed good separation of the two enantiomers. The S,S enantiomer (retention time, 28.2 min) had an ee of only 83%, whereas that of the R,R enantiomer (28.7 min) was 90%. The lower ee of the former was thought to be due to the prolonged reaction time (24 hr at 0°C and 17 hr at room temperature) compared with that used in the second reaction (15 hr at RT).
2,3-Decanedione was prepared in 50% yield by oxidation of (S)-2-hydroxy-3-decanone with pyridinium chlorochromate and sodium acetate in dichloromethane (Corey and Suggs, 1975) followed by flash chromatography and Kugelrohr distillation (b.p. 80°C/0.06 mm). 1H NMR: δ 0.86 (t, J = 7 Hz, 3H, CH3–CH2), 1.27 (m, 8H, 4 × CH2), 1.56 (quint, J = 7 Hz, 2H, CH2–CH2–CH2–C=O), 2.31 (s, 3H, CH3–C=O), 2.71 (t, J = 7 Hz, 2H, CH2–C=O); 13C NMR: δ 14.0, 22.6, 23.1, 23.7, 29.0, 29.1, 31.6, 35.7, 197.7, 199.6; MS m/z (%) 127 (22), 109 (8), 83 (1), 81 (1), 67 (9), 58 (4), 57 (100), 55 (11), 43 (79), 42 (12), 41 (61) (c.f. slightly different data in Rhainds et al., 2001a).
Controlled release dispensers for the synthetic pheromone were heat-sealed polyethylene vials (22 × 8 × 1.5 mm thick; Just Plastics Ltd., London, UK) or polyethylene sachets (2.5 cm × 2.5 cm × 120 μ thick) made from heat-sealed layflat tubing (Transatlantic Plastics, Southampton, UK) and contained 100 μl (approximately 80 mg) or 200 μl pheromone. Release rates from dispensers were measured by periodic weighing of duplicate dispensers maintained in a wind tunnel at 27°C and 8 km hr−1 wind speed.
Bioassays were carried out in an insectary maintained at 27°C with 5000 lux lighting, between 10:00 and 12:00 hours coinciding with the period of maximum activity of the beetles in the field (Venkatesha et al., 1995). Two types of bioassay apparatus were used, both constructed of Plexiglas.
The “swastika” bioassay consisted of a central circular chamber (19 × 5 cm high) with four equally spaced entry/exit openings in the side wall. The openings led to tubes (3.8 cm diam, total length 55 cm) with a right angle bend in the middle to give a swastika appearance. Plexiglas sample bottles (6 × 4 cm) with fine mesh tops and bottoms were fixed to the distal ends of the tubes. Air was drawn into the apparatus through the arms and out through a tube attached to the center of the top of the central chamber at 20 l/min−1. Initial studies were carried out at NRI with 25 male and 25 female insects so that insects were used as lures and/or test individuals more than once, but never on the same day. A single insect was placed at the center of the central chamber with appropriate treatments in the sidearms. Insect behavior was observed for 10 min: if an insect entered one of the sidearms this was scored, otherwise it was scored as remaining in the central chamber. The apparatus was rotated through 90° between each observation to avoid any possible effects due to uneven lighting or visual cues. Ten insects were tested in each experiment and data were analyzed by a χ2 test assuming equal distribution of responding beetles between the arms as null hypothesis.
The tube bioassay utilized a Plexiglas tube (1 m × 3.8 cm i.d.) with a Plexiglas sample bottle (6 cm × 4 cm) with fine gauze top and bottom attached at one end. The test source was put in the sample bottle and a single test beetle placed at the opposite end of the tube. Air was drawn in through the source bottle and out through the other end of the tube at 10 l min−1. The tube was marked at 10-cm intervals, and the nearest approach to the source by the test insect during 10 min of observation was scored according to the interval mark crossed (i.e., 1.0 = reached source, 0.5 halfway, 0.0 = no upwind movement). Each lure was tested with 10 insects, the data were subjected to arcsin transformation before analysis of variance, and differences between means were tested for significance by Duncan's multiple range test (DMRT).
Field trials were carried out in estates round the Central Coffee Research Institute, Chikmagulur, Karnataka, the Coffee Research Sub-Station, Chettalli, Karnataka, and the Regional Coffee Research Station, Thandigudi, Tamil Nadu, India, during the two flight seasons, April–June and October–December, in 1997 and 1998. Traps were sticky delta traps (20 × 20 floor, 15-cm walls; Agrisense, Treforest, UK) or sticky white cross-vane traps (60 × 30 cm; Agrisense) fastened to wooden poles at a height of approximately 1.5 m. Treatments were tested in randomized complete blocks with 20 m between traps, and catches were recorded and discarded every 2–4 d. Total catches were ranked and analyzed by the nonparametric Kruskal–Wallis or Mann–Whitney U tests. Critical differences were calculated to test differences in catches for significance.
GC Retention Indices (RI) of Components in Volatiles from Male X. quadripes and Synthetic Compounds
Further evidence for these assignments was obtained from the IR spectrum of a volatile collection assayed at approx. 1.5 mg and evaporated onto a sodium chloride plate. The IR spectrum showed a broad hydroxyl absorption at 3400 cm−1 and a sharp carbonyl absorption at 1710 cm−1. NMR spectra were obtained from a volatile collection eluted with CDCl3 and assayed by GC at 0.5 mg of (A), the ratio of (A) to (B) being 1:0.14. The 1H and 13C spectra were consistent with those for 2-hydroxy-3-decanone (I) by comparison with corresponding data for 2-hydroxy-3-octanone reported by Bel-Rhlid et al. (1989) and Mori and Otsuka (1985), respectively. In the 1H NMR spectrum, a singlet at δ 2.20 corresponding to the CH3–C=O in isomer (II) (cf. Bel-Rhlid et al., 1989) integrated approximately 7% relative to the CH3–CH–OH doublet at δ 1.38 for the major component (I). This suggested that II was indeed produced by the insect, although some isomerization of I to II also occurred during GC analysis.
A collection of volatiles from male beetles was eluted from the Porapak with ether and reduced with LiAlH4. GC analysis showed components (A) and (B) had reacted to give two new components in 40:60 ratio at RI 2105 and 2144 on the polar GC column. The two components had identical mass spectra, which were analogous to that reported for 2,3-dihydroxyoctane by Kuwahara et al. (1987), with ions at m/z 129 and 111 compared with m/z 101 and 83 in the latter compound. These two products were proposed to be two diastereoisomers of 2,3-dihydroxydecane (Table 1), and identical results were obtained on similar reduction of synthetic (S)-2-hydroxy-3-decanone (I).
Synthesis and Configuration of Proposed Pheromone Components
The enantiomers of threo 2,3-dihydroxyoctane were each synthesized by Sharpless asymmetric dihydroxylation of (E)-2-octene. These threo enantiomers had identical mass spectra and retention times on polar and nonpolar GC columns to those of the minor component (C) in volatiles from male X. quadripes, and none (<0.5%) of the erythro diastereoisomer could be detected. Mass spectra and GC retention times of the corresponding acetates were also identical to those of a minor component in a collection of volatiles from male X. quadripes after acetylation with acetic anhydride and pyridine (Table 1).
During this work, the corresponding eight-carbon analogue of the major pheromone component (I), i.e., (S)-2-hydroxy-3-octanone was found to be present at 0.2–0.5% of I in the volatiles collected from male X. quadripes by comparison of GC retention times and mass spectra with those of the synthetic material. Also, following the report of Rhainds et al. (2001a), 2,3-decanedione was synthesized, and reexamination of GC-MS results showed that this compound was present at 2% of the major component (I). The only other significant components detected were 2-phenylethanol (3% of I), octanoic acid (4% of I), and two unidentified components at RI 1894 and 2384, respectively, on the polar GC column (each approximately 1% of I) having mass spectra similar tothat of 2,3-decanedione. No significant EAG responses from female X. quadripes were observed to these components in GC-EAG analyses of male volatile collections.
Release of synthetic (S)-2-hydroxy-3-decanone (I)from both polyethylene vials and sachets was zero order under constant laboratory conditions. The release rate from the sachets was 0.32 mg hr−1 at 27°C, essentially commencing immediately after the sachets were made up and continuing at a constant level until the contents were exhausted. Release from the vials took 3 d to start, while the material permeated through the wall of the vial, but then remained constant at 0.018 mg hr−1 for the period of measurement (60 d).
Total Catchesa of X. Quadripes in Cross-Vane Traps Baited with Enantiomers of 2-Hydroxy-3-Decanone (I) during October 1997 (Four Replicates at each of Three Sites; 30 d)
Total Catchesa of Cerambycid Beetles in Cross-Vane Traps Baited with (S)-2-Hydroxy-3-Decanone (I) (April–May 1998; Three Replicates at Three Sites)
Catchesa of X.quadripes Beetles in Cross-Vane Traps Baited with (S)-2-Hydroxy-3-Decanone (I) or a Blend of I with 10% (S,S)-2,3-Dihydroxyoctane (III) at six Sites in India (November 1998–January 1999)
No. trap days
Mean/trap ± SE
Mean/trap ± SE
1.50 ± 0.50
0.21 ± 0.16
I + III
0.50 ± 0.26
0.08 ± 0.08
0.08 ± 0.08
0.08 ± 0.08
As previously reported (Hall et al., 1998), the major component in volatiles from male X. quadripes beetles from India eliciting an EAG response from antennae of female beetles was (S)-2-hydroxy-3-decanone (I), and this was confirmed by Rhainds et al. (2001a) using insects from China. However, in collections from Indian insects this compound represented up to 90% of the volatile materials and was produced at up to 2 μg hr−1 beetle−1, whereas the GC traces shown in Rhainds et al. (2001a) contained at least six other peaks of similar size to the major component.
GC analysis with splitless or split injection of natural or synthetic 2-hydroxy-3-decanone produced variable amounts of the isomeric 3-hydroxy-2-decanone (II), although this did not elicit a response from female beetles in GC-EAG analyses. Rhainds et al. (2001a) reported that this isomerization did not occur with cool, on-column injection and that this isomer is not produced by the male beetle. However, the 1H NMR spectrum of crude material from the Indian insects showed a signal corresponding to this isomer integrating at approximately 7% of the major isomer, suggesting that 3-hydroxy-2-decanone was naturally produced or isomerized during the collection process.
Neither 2-hydroxy-3-decanone (I) nor 3-hydroxy-2-decanone (II) has previously been reported from insects although analogues are produced by other cerambycid species. The eight-carbon analogue of I, (S)-2-hydroxy-3-octanone, is produced by males of the grape borer, X. pyrrhoderus Bates (Iwabuchi, 1982; Sakai et al., 1984), and the mulberry borer, X. chinensis Chevrolat (Kuwahara et al., 1987; Iwabuchi et al., 1987). A six-carbon analogue of II, (R)-3-hydroxy-2-hexanone, is produced by the old house borer, Hylotrupes bajulus L. (Fettköther et al., 1995), Pyrrhidium sanguineum L. (Schröder et al., 1994), and Anaglyptus subfasciatus Pic. (Leal et al., 1995). The latter species also produces the eight-carbon analog, (R)-3-hydroxy-2-octanone (Leal et al., 1995).
The Indian insects were shown to produce a third component that caused at least as large an EAG response from females as the major component in GC-EAG analyses, although it was only present at approximately 7% of the major component. This was identified as (S,S)-2,3-dihydroxyoctane (III), found previously as the major component in volatiles from male X. pyrrhoderus (Iwabuchi, 1982; Sakai et al., 1984). This was not reported by Rhainds et al. (2001a), although it would have eluted before the major component in their analyses. These authors did find a second component present in relatively small amounts but highly EAG-active, and this was identified as 2,3-decanedione. This compound was present at approximately 2% of the major component (I) in volatiles from Indian insects. Small amounts [0.2–0.5% of the major component (I)] of 2-hydroxy-3-octanone were also detected, but neither of these two compounds elicited an EAG response from female X. quadripes in GC-EAG analyses.
At least three syntheses of (S)-2-hydroxy-3-octanone have been reported, but these were not suitable for large-scale production of 2-hydroxy-3-decanone (I), because of the unavailability of starting materials (Mori and Otsuka, 1985) or low yields (Bel-Rhlid et al., 1989). Sakai et al. (1984) reported the synthesis of 2-hydroxy-3-octanone from lactic acid without any details, but it was thought that this could provide a good route to I from readily available starting materials. Numerous attempts to react heptyl lithium with free (S)-lactic acid under a variety of conditions in a range of solvents failed to give any significant quantity of the desired product. However, protection of the hydroxyl group and hydrolysis of the ester followed by reaction with heptyl lithium in diethyl ether and deprotection gave the desired product (I).
Initially, it was considered that the deprotection step should be as mild as possible to avoid isomerization of I to II, and the benzyl protecting group was selected (R1 = benzyl in Figure 1). The reaction scheme proceeded well, the benzyl group being put on with sodium hydride and benzyl bromide in THF at room temperature and removed smoothly by hydrogenation over palladium on charcoal in ethanol. However, the product (I) was essentially racemic, and examination of intermediates by chiral GC showed that the starting (S)-ethyl lactate was optically pure but that conversion to the benzyl ether had caused racemization. However, use of the tetrahydropyranyl or tert-butyldimethylsilyl protecting groups gave (I) without any significant racemization, the former being favored because the reagents were cheaper and yields higher. A further useful modification of the procedure involved hydrolysis of the protected ester with one equivalent of lithium hydroxide in aqueous ethanol and isolation of the lithium salt. In the case of the tetrahydropyranyl-protected derivative, the lithium salt was a white amorphous solid, and this reacted with one equivalent of heptyl lithium rather than the two equivalents that would be required for the free acid.
Several multistep syntheses of (S,S)-2,3-dihydroxyoctane (III) have been reported (Mori and Otsuka, 1985; Bel-Rhlid et al., 1989; Chattopadhyay et al., 1990; Kang et al., 1990; Bonini and Righi, 1992; Takahata et al., 1994; Bonini et al., 1995; Paolucci et al., 1995), but a single-step synthesis applicable to both enantiomers was used in this work. Reaction of (E)-2-octene with the Sharpless reagents “AD-mix-α” or “AD-mix-β” in tert-butyl alcohol in the presence of methanesulfonamide (Sharpless et al., 1992) gave (S,S)- or (R,R)-2,3-dihydroxyoctane (III), respectively, in 91% yield. Both compounds were completely free (≤0.2%) of erythro diastereoisomers by GC analysis. Conducting the reaction at room temperature for 15 hr gave product with 90% ee, although this could probably be improved further. An analogous synthesis of the enantiomers of 2,3-dihydroxyhexane was recently described by Lacey et al. (2004).
Attraction of female X. quadripes to male beetles or synthetic lures was demonstrated with crawling insects in swastika or tube bioassays in the laboratory. Attempts to carry out bioassays with a large wind tunnel, a similar swastika bioassay, or in cages under field conditions in India were less successful, although mating was observed as described by Venkatesha et al. (1995) and Rhainds et al. (2001a). These observations were consistent with those of Rhainds et al. (2001a), who were able to demonstrate weak attraction of female X. quadripes to natural or synthetic pheromone in a room or field cage but only by introducing large numbers of insects and after patient and detailed observations. These authors reported that mating on the ground was common, providing a possible explanation why bioassays with crawling beetles were more successful than those with flying beetles.
We also demonstrated the attraction of female X. quadripes beetles to synthetic lures in the field. The (S)- or racemic (RS)-2-hydroxy-3-decanone was attractive, but addition of (S,RS)- or (S,S)-2,3-dihydroxyoctane (III) at 10% of I reduced attractiveness in laboratory bioassays and field trapping trials, respectively. However, numbers trapped were extremely low. Rhainds et al. (2001a) essentially failed to trap any beetles in field tests using multifunnel or delta traps, although beetles were trapped in the funnel traps during the laboratory tests, presumably with much higher population densities. Rhainds et al. (2001a) suggested that our field trapping results (Hall et al., 1998) were attributable to higher population densities or favorable positioning of traps. Unfortunately, no independent estimations of population are available that can be compared with those of Rhainds et al. (2001a), but these were not thought to be particularly high, and traps were positioned at random. The most likely explanation for the difference in results is that we used much larger, sticky cross-vane traps, consistent with the mating behavior described by Venkatesha et al. (1995) and Rhainds et al. (2001a), in which females are attracted to the vicinity of males after which short range chemical or visual signals probably take over. Similar conclusions were reached by Morewood et al. (2002), who reported that cross-vane traps with a large silhouette were more effective than multifunnel traps for trapping cerambycid beetles.
Although small numbers of male insects were also trapped in these trials, numbers were not significantly different than those in unbaited traps. Thus, the pheromone of X. quadripes must be considered as a sex pheromone, similar to those in other Xylotrechus species (Iwabuchi, 1982; Iwabuchi et al., 1987). Lacey et al. (2004) recently reported production of an aggregation pheromone by males of Neoclytus acuminatus acuminatus, a Cerambycid belonging to the same subfamily and tribe as X. quadripes—Cerambycinae and Clytini, respectively. Lacey et al. (2004) did not test the natural pheromone, but the synthetic pheromone attracted significant numbers of both females and males, generally more of the former than the latter.
Trials are in progress in India to determine whether pheromone traps can cause any reduction in damage by X. quadripes, but, as suggested by Rhainds et al. (2001a), the chances of success at a cost-effective density would not seem to be high unless the effectiveness of the traps can be improved. Because adult X. quadripes do not feed (Visitpanich, 1994; Venkatesha et al., 1995 and observations of the authors), addition of feeding attractants as used with A. subfasciatus (Nakamuta et al., 1997) is probably not an option. Addition of oviposition attractants described by Rhainds et al. (2001b) would not be expected to have much effect in coffee plantations where these odors must be in abundance. However, the field studies reported here were limited in that only a single blend of the two EAG-active components was tested, and the effects of other potential pheromone components such as the 2-hydroxy-3-octanone and 2,3-decanedione were not evaluated. Thus, possibilities exist for optimizing the pheromone blend, the pheromone release rate, and the trap design in relation to mating behavior.
This publication is an output from research projects (R6928 and 7246) funded by the United Kingdom's Department for International Development (DFID). However, DFID accepts no responsibility for any information provided or views expressed. Demonax balyi beetles were identified by the CABI Identification Service.