Linalool and lilac aldehyde/alcohol in flower scents: Electrophysiological detection of lilac aldehyde stereoisomers by a moth

Stefan Dötterla, , , Dirk Burkhardtc, Bernhard Weißbeckerd, Andreas Jürgensb, Stefan Schützd and Armin Mosandlc

aDepartment of Plant Systematics, University of Bayreuth, Universitätsstrasse 30, D-95440 Bayreuth, Germany

bHortResearch, Canterbury Research Centre, Gerald Street, Lincoln 8152, PO Box 51, New Zealand

cUniversity of Frankfurt, Department of Food Chemistry, Marie-Curie-Strasse 9, D-60439 Frankfurt/Main, Germany

dInstitute for Forest Zoology and Conservation, University of Göttingen, Büsgenweg 3, D-37077 Göttingen, Germany

Received 5 October 2005;  revised 2 February 2006;  accepted 6 February 2006.  Available online 20 March 2006.

Abstract

The stereoisomers of linalool and lilac aldehyde/alcohol were determined in the flower scent of 15 plant species using enantioselective multidimensional gas chromatography/mass spectrometry (enantio-MDGC/MS). Both linalool and all 8 stereoisomers of lilac alcohol and lilac aldehyde were detected, and there was a species-specific pattern. Single stereoisomers were collected by micropreparative-enantio-MDGC and were electrophysiologically tested on antennae of the noctuid moth Hadena bicruris, a species known to rely on lilac aldehyde for finding its host plant. The moth responded to all 8 stereoisomers, though only four stereoisomers were found in the scent of its host plant. The moth was less sensitive to some isomers than to others.

Keywords: Linalool; Lilac aldehyde-alcohol; Stereoisomers; Micropreparative-enantio-MDGC; Electroantennographic detection; Hadena bicruris

Article Outline

1. Introduction2. Experimental

2.1. Plant material and volatile collection

2.2. Insects2.3. Enantio-MDGC/MS2.4. Micropreparative-enantio-MDGC2.5. Electrophysiological analysis2.6. Statistical analyses

3. Results

3.1. Detected stereoisomers3.2. Pattern of lilac aldehydes3.3. Collection of single stereoisomers and electrophysiological detection of single isomers in Hadena

4. Discussion

4.1. Linalool as a precursor for lilac compounds4.2. Pattern of lilac aldehydes4.3. Lilac aldehyde stereoisomers and insects

AcknowledgementsReferences

1. Introduction

Many flowers emit volatiles for attraction of pollinators. Volatiles are especially useful at night when visual cues become insufficient. Floral volatiles can synthesized by several pathways, wherein benzenoids, isoprenoids, and fatty acid derivatives are the most typical chemical classes [1]. Several thousand compounds have been described so far, and many of them have stereogenic centres [2]. However, in most studies the exact determination of stereoisomers was neglected, most probably due to the difficult separation of single enantiomers. It is also mostly unknown, whether pollinators can electrophysiologically and/or behaviourally differentiate between different enantiomers.

From pheromone studies it is known that chirality is an important factor in insect communication. Two species of scarab beetles use different enantiomers of (Z)-5-(1-decenyl)oxacyclopentan-2-one (japonilure) as sex pheromone. Anomala osakana produces and uses (S)-japonilure as a female sex pheromone, and Popillia japonica uses (R)-japonilure as a sex pheromone [3] and [4]. When offered additionally the “wrong” enantiomer to the species in a biotest, male attraction was strongly inhibited [3]. On the other hand, in the case of the Douglas-fir cone gall midge, Contarinia oregonensis, males detected and responded only to (Z,Z)-4,7-tridecadien-(S)-2-yl acetate, the female sex pheromone, whereas the corresponding (R)-enantiomer was electrophysiologically and behaviourally inactive [5].

In addition to pheromonal tests it is known from studies on plant-herbivore interactions that chirality is an important factor. For the white pine cone beetle, Conophthorus coniperda, it was shown that (−)-α-pinene, and not (+)-α-pinene significantly increased catches to traps baited with female sex pheromone [6]. Another beetle, the pine weevil Hylobius abietes responded better to (+)-α-pinene than to the (−)-enantiomer in an elecrophysiological study [7]. For the moth Helicoverpa armigera it was shown in a recent study that an electroantennogram response to α-

pinene was higher in the (−)-form than in the (+)-form. However, this electrophysiological result was contrary to behavioural results, where naïve moths showed an innate preference for (+)-α-pinene [8].

However, only few examples determining the absolute configuration of optically active compounds in floral scents, and additional testing the single isomers to the pollinating insects are known (e.g. [9]). Therefore, it is unknown in most pollination systems whether or not chirality plays a role in pollination systems. Among the most complicated chiral floral scent compounds are the oxygenated monoterpenoids lilac aldehydes and alcohols. Each of these compounds has three stereogenic centres, and therefore eight different aldehyde and eight alcohol stereoisomers are possible (Fig. 1). The lilac compounds occur in plant species of many families, such as Caryophyllaceae (Dianthus spp. [10], Silene spp. [11], [12] and [13]), Rosaceae (Prunus padus [14]), Lamiaceae (Origanum vulgare [15]), Onagraceae (Gaura longiflora [16]), Orchidaceae (Platanthera spp. [17]), Polemoniaceae (Phlox paniculata [15]), Salicaceae (Salix spp. [18]), Violaceae (Viola etrusca [19]), Actinidiaceae (Actinidia [20]), Rubiaceae (Cephalanthus occidentalis [15]), Vitaceae (Vitis vinifera cv. Moscato bianco [21]), Asteraceae (Artemisia pallens [22], Eupatorium cannabinum [15]), Oleaceae (Syringa vulgaris [23]), and are found in several honeys [24], [25], [26] and [27].

Full-size image (62K)

Fig. 1. Stucture of lilac aldehyde (A) and lilac alcohol (B) stereoisomers.

However, the exact composition of stereoisomers was determined only in one case. In Syringa vulgaris only four out of all eight lilac aldehydes as well as alcohols were found [23]. In this plant lilac compounds were synthesised via the deoxy-xylulose pathway, and linalool is an important precursor [23] and [28]. Linalool is also an optically active compound with two enantiomers, and only (S)-linalool was found in Syringa. The stereogenic centre of the precursor linalool was not affected during biosynthesis of lilac compounds. This is why only (5′S)-lilac aldehydes and alcohols occur in Sy. vulgaris.

Lilac aldehyde is of special interest for pollinators. It is emitted in high amounts, especially in nocturnal plant species, and it is known to be highly attractive to the nocturnal moth species Autographa gamma and Hadena bicruris [29] and [30]. In the case of A. gamma, highly specific and sensitive olfactory receptors were found in the antennae [31]. Lilac aldehyde is also known to elicit strong antennal signals in butterfly species [32] and [33]. However, electrophysiological and behavioural experiments were conducted in last-mentioned studies only with isomeric mixtures of these oxygenated monoterpenoids, and the role of single stereoisomers remains unknown.

Recently a method has been developed allowing the separation and identification of all eight stereoisomers of lilac aldehydes and 7 of all eight possible lilac alcohol isomers [34]. In the present paper this method is used to determine the exact isomeric composition of lilac compounds in several plant species of different plant families. Additionally, the enantiomeric distribution of linalool was determined. Finally, single isomers of lilac aldehydes were collected using micro preparative enantio-MDGC (enantioselective multidimensional gas chromatography), and the single isomers were subsequently tested on the moth H. bicruris in a GC-EAD (gas chromatography with electroantennographic detection) study.

2. Experimental

2.1. Plant material and volatile collection

Linalool, lilac aldehyde and lilac alcohol isomers of 15 different plant species belonging to the 9 plant families Actinidiaceae (Actinidia arguta (Sieb Et Zucc.)), Asclepidiaceae (Cynanchum auriculatum Royle ex Wight), Caprifoliaceae (Viburnum opulus L.), Caryophyllaceae (Silene otites (L.) Wibel, S. alba L., S. vulgaris (Moench) Garcke), Oleaceae (Syringa vulgaris L.), Polemoniaceae (Linanthus dichotomus Benth., Phlox divaricata L.), Ranunculaceae (Anemone nemorosa L.), Rosaceae (Prunus padus L., P. laurocerasus L., P. spinosa L.), and Thymelaeaceae (Daphne cneorum L., Struthiola dodecandra Wynberg) were selectively differentiated by enantio-MDGC analysis. The volatile samples of the species were collected using dynamic headspace [35]. Therefore, flowers were enclosed in an oven bag (Nalophan, 10 cm × 10 cm, respectively, 15 cm × 15 cm, depending on size of flowers or inflorescences) for 2 h and emitted volatiles were trapped in an adsorbent tube filled with 20 mg of a 1:1 mixture of Tenax-TA 60–80 and Carbotrap 20–40 (Supelco, Germany). The air was sucked from the bag over the adsorbent by a membrane pump (G12/01 EB, Rietschle Thomas, Puchheim, Germany). Volatiles were eluted with 70 μl of acetone (SupraSolv, Merck, Germany) to get the samples for the enantioselective multidimensional gas chromatography (see below). The samples of Phlox divaricata and Linanthus dichotomus were provided by Robert Raguso. Roman Kaiser provided a sample of Struthiolia dodecandra, and Adam Mattich provided the Actinidia arguta sample.

2.2. Insects

Three specimen of a Hadena bicruris Hufn. (Lepidoptera: Noctuidae) culture were used for the experiments. The larvae were reared on artificial diet as described in [36]. Adults were fed immediately after hatching with a sugar solution (30%). Two- to four-day-old naïve female and male moths were used for the experiments. Each moth was tested in the GC-EAD/MS study to all fractions (see below).

2.3. Enantio-MDGC/MS

The enantio-MDGC-MS was performed with a Siemens SiChromat 2–8, with two independent column oven programs and a live T-switching device, coupled to the transfer line of a Finnigan MAT GCQ, using an open split interface.

GC conditions were as follows. Pre column: self prepared fused silica capillary (30 m × 0.23 mm I.D.), coated with a 0.23 μm film of SE-52; carrier gas helium, 1.85 bar; injector temperature 250 °C; flame ionization detection (FID), detector temperature 250 °C. Main column: self prepared fused silica capillary (30 m × 0.23 mm I.D.), coated with a 0.23 μm film of 0,4% heptakis(2,3-di-O-methyl-6-O-tert-butyldimethylsilyl) β-cyclodextrin (DIME-β-CD) (30%) in SE-52 (70%), carrier gas helium, 1.45 bar, MS-detector; transfer line temperature 250 °C, open split interface 250 °C; helium sweeping flow 1 ml/min; ion trap manifold 200 °C; EI 70 eV; oven temperature program: precolumn: 60 °C (5 min isothermal), raised at 3 °C/min to 250° (30 min isothermal); main column temperature program: 60 °C (25 min isothermal), raised at 1.5 °C/min to 200 °C. Cut times: lilac aldehyde 26.5–28.0 min. Data acquisition is done with the Xcalibur instrument software (Finnigan).

2.4. Micropreparative-enantio-MDGC

The micropreparative-enantio-MDGC was performed with a Siemens SiChromat 2–8, with two independent column oven programs. The two capillary columns of the MDGC were coupled with a press-fit connector. The flow of the chiral main column was splitted and the single stereoisomers were collected by stir bar sorptive extraction (SBSE) as individual eluent fractions (Fig. 2). During 1 h all the eight stereoisomers were reeluted from the stir bars with 1 ml of a mixture of pentane and diethylether 1:1 (v/v). After reducing the solvent to 100 μl the fractions were used for electrophysiological analysis.

Full-size image (28K)

Fig. 2. Collection of single stereoisomers using micro-preparative enantio-MDGC/SBSE.

GC conditions were as follows. Pre column: self prepared fused silica capillary (30 m × 0.23 mm I.D.), coated with a 0.23 μm film of SE-52; Main column: self prepared fused silica capillary (30 m × 0.23 mm I.D.), coated with a 0.23 μm film of 0,4% heptakis (2,3-di-O-methyl-6-O-tert-butyldimethylsilyl) β-cyclodextrin (DIME-β-CD) (30%) in SE-52 (70%), carrier gas helium, 2.0 bar; injector temperature 250 °C; flame ionization detection (FID), detector temperature 250 °C. Oven temperature program: pre column: 60 °C (5 min isothermal), raised at 3 °C/min to 150°C (30 min isothermal); main column temperature program: 60 °C (25 min isothermal), raised at 1.5 °C/min to 200 °C.

2.5. Electrophysiological analysis

Experiments were performed by means of a GC-EAD/MS system described in [37]. It consisted of a 6890N gas chromatograph and a 5973N quadrupole mass spectrometer (both Agilent, Palo Alto, USA). The GC was equipped with a type 7163 autosampler, a split/splitless injector, and a J&W Scientific HP-5MS column (Agilent, length 30 m, inner diameter 0.25 mm, film thickness 0.25 μm). The injector was operated in the pulsed splitless mode. Helium (purity 99.999%) was

used as carrier gas at a constant flow of 1 ml/min, gas vector 24 cm/s. The following temperature program was employed: start: 50 °C, hold for 1.5 min, ramp 6 °C/min to 200 °C, hold for 5 min.

A Graphpack 3D/2 flow splitter (Gerstel, Mülheim, Germany) was used in order to split the effluent from the column into two pieces of deactivated capillary leading to the mass spectrometer (length 1 m, I.D. 0.1 mm) and to the EAD set-up (length 1 m, I.D. 0.2 mm). The restriction of these capillaries resulted in an equal split of the gas flow into the two set-ups.

The mass spectrometer used electron ionization at 70 eV and was operated in the scan mode with a mass range from 35 to 300 mass units at a scan speed of 2.78 scans s−1.

The EAD set-up used a modified version of an “Olfactory Detector Port” (Gerstel, Mülheim, Germany). The ODP incorporated a flexible heating sleeve (230 °C), which guided the capillary out of the GC oven where the effluent of the capillary was mixed with humidified air. The airflow was directed to the insect antenna, which was housed in a detector cell made of PTFE modified according to [38] and [39]. Both ends of the antenna were contacted to Ag/AgCl electrodes via hemolymph Ringer solution [40]. EAG-potentials of the antenna were amplified by a factor of 100, and the amplified signal passed through a high pass filter with a cut-off frequency of 0.01 Hz to suppress the slow drift, which is often observed in the EAD signal. Afterwards, a constant voltage of 0.5 V was added to the signal. These steps were necessary in order to match the amplifier output to the input signal range (0–1 V) of a 35900E A/D converter (Agilent). The signal was recorded by the Agilent ChemStation software.

2.6. Statistical analyses

It is generally assumed that the lilac alcohols are generated by reduction of the lilac aldehydes. The substituents priority at the C2 position is changed during the reduction of the aldehyde function to the primary alcohol. Therefore, it is a clear correlation between lilac aldehyde and lilac alcohol stereoisomers. Since only the lilac aldehydes are reported to be behaviourally active, we only included lilac aldehydes in following analyses. We further excluded from these analyses the samples of Phlox divaricata and Silene vulgaris, because they strongly differed from all the other samples (see below).

We used the CNESS (chord-normalized expected species shared, see [41]) distance index to determine the differences between the single samples. Therefore, relative quantities (percentage) of single lilac aldehyde isomers were used. We calculated CNESSm8 indices using the updated version of the COMPAH program [42], provided by E. Gallagher at UMASS/Boston (http://www.es.umb.edu/edgwebp.htm). We used nonlinear multidimensional scaling (NMDS) to detect meaningful underlying dimensions and to visualise similarities between samples (see [43]). To evaluate how well the particular configuration produces the observed distance matrix (in our case the matrix of CNESS) the stress value is given. The smaller the stress value, the better is the fit of the reproduced ordination to the observed distance matrix [44].

3. Results

3.1. Detected stereoisomers

The stereoisomers of linalool, lilac aldehyde and lilac alcohol found in the 15 studied species are shown in Table 1. Both isomers of linalool as well as all conceiveable lilac aldehyde and lilac alcohol isomers were found. Linalool was found in 10 species, lilac aldehyde in all studied species and lilac alcohol in 11 species. In five of the linalool containing plants only the (S)-isomer was found. In these species only 5′S configured lilac aldehyde and/or lilac alcohol isomers were detected. In five species also (R)-linalool was detected with the highest amount in Phlox divariacata. In this species only (5′R)-configured lilac aldehyde and lilac alcohol stereoisomers were detected. In the other (R)- and (S)-linalool containing species only (5′S)-lilac isomers were found.

Table 1.

Relative amount of linalool, lilac aldehyde, and lilac alcohol stereoisomers found in 15 plant species

A. a.

A. n.

C. a.

D. c.

L. d.a

P. d.a

P. I.

P. p.

P. s.

S. a.a

S. d.

S. o.a

S. v.

Sy. v.

V. o.

Linalool S41

–98

100

100

545

100

– – –100

–100

81

R59

– 2 – – 9555

– – – – – – –19

Lilac aldehydes

(2S, 2′S, 5′S)58

48

39

91 79 –42

3543

4283

5711

3833

(2R, 2′S, 5′S)10

–13

7 8 – 5 12 8 30 9 3410

3016

(2R, 2′R, 5′R)

– – – – – 8 – – – – – – 6 – –

(2S, 2′R, 5′R) – – – – – 5 – – – – – – 1 – –

(2S, 2′S, 5′R) – – – – – 5 – – – – – –26

– –

(2R, 2′S, 5′R) – – – – – 83 – – – – – –44

– –

(2S, 2′R, 5′S)13

52

20

1 4 –44

4341

28 3 8 2 1938

(2R, 2′R, 5′S)19

–28

1 9 – 9 10 8 1 5 2 1 1313

Lilac (2R, 2′S, 5′S) 8 7 7 91 – – – – – 44 8 77 5 40 3

A. a.

A. n.

C. a.

D. c.

L. d.a

P. d.a

P. I.

P. p.

P. s.

S. a.a

S. d.

S. o.a

S. v.

Sy. v.

V. o.

alcohols 0 0 2 2

(2S, 2′S, 5′S) 9 – 4 3 – – – – – 26 6 1421

3229

(2S, 2′R, 5′R) – – – – – – – – – – – – 4 – –

(2R, 2′R, 5′R)

– – 10 0 – –

(2R, 2′S, 5′R) – – – – – 82 – – – – – –67

– –

(2S, 2′S, 5′R) – – – – – 8 – – – – – – – – –

(2R, 2′R, 5′S) 630

1 6 – – – – – 6 7 8 – 1622

(2S, 2′R, 5′S) 5 –88

– – – – – – 25 5 – 3 1218

A. a. (Actinidia arguta), A. n. (Anemone nemorosa), C. a. (Cynanchum auriculatum), D. c. (Daphne cneorum), L. d. (Linanthus dichotomus), P. d. (Phlox divaricata), P. l. (Prunus laurocerasus), P. p. (Prunus padus), P. s. (Prunus spinosa), S. a. (Silene alba), S. d. (Struthiola dodecandra), S. o. (Silene otites), S. v. (Silene vulgaris), Sy. v. (Syringa vulgaris), V. o. (Viburnum opulus).a Mean of 2 samples was built.

In five species no linalool was detected, though lilac compounds were found. In all of these species, and with the exception of Silene vulgaris, only (5′S)-configured lilac compounds were found. Silene vulgaris was the only plant where all eight possible four (5′S)- and four (5′R)-configured lilac aldehyde and lilac alcohol stereoisomers were detected.

Therefore, 13 of the 15 studied species contained only the (5′S)-configured lilac stereoisomers, whereas in Phlox divaricata only (5′R)-lilac compounds were found, and in Silene vulgaris all eight lilac aldehyde and lilac alcohol isomers were identified.

3.2. Pattern of lilac aldehydes

When analysing the pattern of lilac aldehyde isomers in the 13 species that contained only the (5′S)-lilac in more detail, we found specific patterns for different plant species (Fig. 3). In most of the samples we found all four (5′S)-isomers, while in Anemone nemorosa and in one sample of Silene otites (S.o. 1) only two isomers were found. In A. nemorosa the (2S, 2′S, 5′S)- and (2S, 2′R, 5′S)-isomers were detected, while in the S. otites sample the (2R, 2′S, 5′S)-isomer was found instead of the (2S, 2′R, 5′S)-one. The other S. otites (S.o. 2) samples was similar to the two

Silene alba samples with high amounts of the (2S, 2′S, 5′S)-, (2R, 2′S, 5′S)-, and (2S, 2′R, 5′S)-isomers. The three Prunus species and Viburnum opulus were dominated by (2S, 2′S, 5′S)- and the (2S, 2′R, 5′S)-isomer, whereas in Syringa vulgaris the (2R, 2′S, 5′S)- reached higher amounts than the (2S, 2′R, 5′S)-isomer. Daphne cneorum was strongly dominated by the (2S, 2′S, 5′S)-isomer (91%), while this isomer reached about 80% in Struthiolia dodecandra and Linanthus dichotomus. (2S, 2′S, 5′S) was also the most abundant isomer in the Actinidia and Cynanchum samples, where also the (2R, 2′R, 5′S)-isomer was found in relatively high amounts.

Full-size image (24K)

Fig. 3. Nonmetric multidimensional scaling of lilac aldehyde pattern in studied plant species containing solely (5′S)-configured isomers (Silene vulgaris, containing all 8 isomers and Phlox divaricata, containing only (5′R)-configured isomers were excluded from analysis) based on the CNESS index (stress = 0.06). A. a. (Actinidia arguta), A. n. (Anemone nemorosa), C. a. (Cynanchum auriculatum), D. c. (Daphne cneorum), L. d. (Linanthus dichotomus), P. d. (Phlox divaricata), P. l. (Prunus laurocerasus), P. p. (Prunus padus), P. s. (Prunus spinosa), S. a. (Silene alba), S. d. (Struthiola dodecandra), S. o. (Silene otites), S. v. (Silene vulgaris), Sy. v. (Syringa vulgaris), V. o. (Viburnum opulus).

3.3. Collection of single stereoisomers and electrophysiological detection of single isomers in Hadena

An isomeric mixture of lilac aldehyde isomers was synthesised according to [34]. All eight lilac aldehyde stereoisomers were separated using enantioselective MDGC-MS. The flow of the chiral main column from the enantioselective MDGC was split and the single stereoisomers were collected by stir bar sorptive extraction (SBSE) as individual eluent fractions. All eight stereoisomers were re-eluted from the stir bars. The fractions were controlled by enantio-MDGC-MS. Fig. 4 shows the main column chromatogram of fraction 8. The enrichment of stereoisomer 8 is clearly visible.

Full-size image (11K)

Fig. 4. Enantio-MDGC analysis of lilac aldehyde stereoisomers (A) and of fraction 8 (B) containing isomer 7 and isomer 8.

The three specimen of Hadena bicruris electrophysiologically tested responded to all 8 isomers of lilac aldehyde in the GC-EAD study (Table 2). However, the moths were more sensitive to some isomers than to other isomers. All three tested individuals responded to the (2R, 2′R, 5′R)- and (2R, 2′R, 5′S)-isomers only in the higher doses, while the other isomers were detected at least by two individuals also in the lower doses.

Table 2.

Antennal responses of three different Hadena bicruris moths to two different amounts of the eight lilac aldehyd stereoisomers

Delivered amounts(ng) Hadena 1 Hadena 2 Hadena 3

(2S, 2′S, 5′S) 0.4; 8.3 ++ ++ ++

(2R, 2′S, 5′S) 0.4; 12.5 + ++ ++

(2R, 2′R, 5′R) 0.6; 5.8 + + +

(2S, 2′R, 5′R) 0.3; 19.0 ++ ++ ++

(2S, 2′S, 5′R) 0.8; 18.0 + ++ ++

(2R, 2′S, 5′R) 0.7; 9.8 ++ ++ ++

(2S, 2′R, 5′S) 1.0; 11.5 ++ ++ ++

(2R, 2′R, 5′S) 0.5; 24.0 + + +

+: moths detected stereoisomers in amounts >5 ng; ++: moths detected isomers also in amounts ≤1 ng.

4. Discussion

This study revealed a specific pattern of linalool and especially lilac aldehyde and lilac alcohol isomers in the floral scent of 15 different plant species. Furthermore, all 8 possible lilac aldehyde as well as alcohol isomers occur in nature, and it was demonstrated that a moth responded electrophysiologically to all eight lilac aldehyde isomers.

Though the lilac compounds have been known for more than 30 years from flower volatiles of Syringa vulgaris [45], the structural elucidation and chromatographic behaviour of the individual stereoisomers has been known for only since 2003 [34]. In Syringa vulgaris as well as in 12 other species in this study, only the four (5′S)-configured lilac compounds occur, whereas only (5′R)-lilac compounds were found in Phlox divariacata, and both (5′S)- and (5′R)-configured compounds were found in Silene vulgaris. Therefore, (5′S)-configured lilac compounds are more widespread than the (5′R)-stereoisomers.

4.1. Linalool as a precursor for lilac compounds

A main precursor of lilac compounds, at least in Syringa vulgaris, is linalool. This oxygenated monoterpene is a widespread floral scent compound and it is described as a behaviourally attractive compound to many insects [46]. It is further known from cephalic secretions of different Colletes species, such as C. cunicularius, where it acts as female sex pheromone. This bee species produces only S-linalool, and male bees respond more strongly to this enantiomer compared to the (R)-one. Nevertheless, the bees did not discriminate between both isomers in a GC-EAD study [47].

The absolute configuration of linalool (S or R) defines the configuration of lilac compounds at position 5′ in Sy. vulgaris. Since in this plant only (S)-linalool is generated via the 1-deoxy-xylulose 5-phosphate (DOX) pathway, only (5′S)-configured lilac aldehydes and lilac alcohols are found [23].

In present study, (S)-linalool was identified in five plant species (including Syringa), and in all these species the corresponding (5′S)-configured lilac compounds were detected. In five other species (R)- and (S)-linalool were detected, but in four of them only (5′S)-configured lilac compounds were detected, and only in Phlox divaricata (5′R)-configured lilac isomers were found. There are two possible explanations, (i) enantioselective enzymes are producing (5′S)-configured lilac compounds, and (R)-linalool is not an appropriate substrate, or (ii) (5′R)-lilac stereoisomers were produced, but the amount was too low for detection. The latter explanation may be the case in Cynanchum auriculatum and Viburnum opulus, where the (S)-configured isomer of linalool was strongly dominant. However, in the case of Actinidia arguta and Prunus laurocerasus both linalool isomers were found in high amounts. In these cases there could be an enantioselective enzyme producing exclusively (5′S)-configured lilac stereoisomers. It is also conceivable that the enzymes important for lilac compounds are discriminating between (R)- and (S)-precursors and preferably use the (S)-configurated precursors resulting in high amounts of (5′S)-lilac isomers, and amounts below the limit of detection of (5′R)-configured lilac compounds.

Enantiodiscrimination was observed in Syringa, where the lilac producing enzymes showed a discrimination in favor of the nongenuine (R)-linalool [23] during incorporation experiments. Enantioselective enzymes are also known from the literature. The best known is (S)-linalool synthase, which stereoselectively converts geranyl pyrophosphate to (S)-linalool [48].

More data from incorporation experiments are needed to determine whether the enzymes converting linalool to lilac compounds are working stereoselectively.

In five plant species linalool could not be detect though lilac aldehyde stereoisomers were found. Either linalool is glycosidically bound and therefore not detectable in headspace samples (compare with [49]), or the enzymes converting linalool to lilac compounds are very sensitive and linalool is converted into other compounds before it can be released. It seems unlikely that lilac compounds are not synthesised via linalool.

4.2. Pattern of lilac aldehydes

A multivariate analysis using relative amounts of single lilac aldehyde isomers in the solely (5′S)-containing taxa revealed specific patterns in different plant species. In most samples all four isomers were found, while in Anemone nemorosa and in a single Silene otites sample only two isomers were detected. The small absolute quantity of lilac compounds was probably responsible for this observation, because the total amount of lilac compounds was very low in these two samples in comparison to all other samples, which contained high amounts of lilac compounds. In the second S. otites sample the isomers (2S, 2′S, 5′S) and (2R, 2′S, 5′S) were dominant, and these two isomers were also found in S. otites 1. Therefore it is assumed that also in sample S. otites 1 all four isomers were present, but the two minor isomers were below the detection limit. The same is hypothesised for Anemone nemorosa.

Two specimens were also analysed of Silene alba and Linanthus dichotomus, and we found low variability between samples within species. Furthermore, we analysed three species of the genus Prunus, and all three species were characterised by very similar lilac aldehyde profiles. Similarly, the two Thymelaeaceae, Daphne cneorum and Struthiola dodecandra, were very similar and both emitted high relative amounts of the (2S, 2′S, 5′S)-isomer. Therefore it seems that phylogenetic relationships strongly influence the pattern in lilac aldehydes in these cases. However, this study also shows that closely related species can emit quite different lilac aldehyde isomers, as in the case of the different Silene species, where two specis emit only the (5′S)-isomers, and one species emits additionally the 5′R isomers. On the other hand, species of quite different plant families can emit the lilac aldehydes in the same relative proportions (e.g. Linanthus dichotomus and Struthiolia dodecandra).

Also pollinators could influence the pattern of lilac aldehydes, because the plant species are pollinated by different insects. In the Silene species, for example, nocturnal pollinators such as moths are important for pollination [50] and [51], whereas in Prunus spinosa or Viburnum opulus bees are the most important pollinators [52] and [53]. Perhaps different groups of pollinators are attracted by different isomers of lilac aldehydes.

4.3. Lilac aldehyde stereoisomers and insects

The lilac aldehydes are important compounds for the attraction of pollinators. They are known to be the key compounds for attraction of the specialized nursery pollinator (i.e. insects that pollinate and use the developing seeds as a resource for their progeny) Hadena bicruris to Silene alba [29]. These moths strongly prefer S. alba for egg deposition (Dötterl, unpublished data) and olfactorily discriminate between S. alba and S. vulgaris [54], though these two Silene species emit almost the same compounds with lilac aldehydes dominating the scent [12]. As both plants emit different lilac aldehyde isomers, the lilac compounds could be used by the moth to discriminate between the two plant species. However, this is only possible, if Hadena bicruris can detect the different isomers (5′S, 5′R). In the present study we show that the moth responds electrophysiologically to all eight lilac aldehyde isomers, but that some isomers are more sensitively detected than others. However, we do not know whether the moths have different olfactory receptors for detection of the single lilac aldehyde isomers or whether the moth posses enantiospecific odorant binding proteins. This would be an essential prerequisite for the moths to be able to differentiate between different patterns of lilac aldehydes, and use these patterns for host plant finding.

In conclusion, our results show that all 8 possible lilac aldehyde as well as alcohol isomers occur in nature, and that chirality may play a role in moth-pollination systems. The pattern of lilac isomers was species specific and may be used by pollinators for host plant finding. The specialised nursery pollinator Hadena bicruris detected electrophysiologically all 8 lilac aldehyde isomers, though the sensitivity to different isomers varied.

Acknowledgements

We are grateful to Adam Mattich, Roman Kaiser und Robert Raguso for providing scent samples. Angelika Täuber helped by rearing the moths, and Josef Woodring gave valuable comments on the manuscript.

References

[1] J.T. Knudsen, L. Tollsten and L.G. Bergström, Phytochemistry 33 (1993), p. 253. Abstract |

PDF (3063 K) | View Record in Scopus | Cited By in Scopus (399)

[2] G. Ohloff, Scent and Fragrances, Springer-Verlag, Berlin, Heidelberg (1994).

[3] W.S. Leal, Proc. Natl. Acad. Sci. U.S.A. 93 (1996), p. 12112. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (43)

[4] J.H. Tumlinson, M.G. Klein, R.E. Doolittle, T.L. Ladd and A.T. Proveaux, Science 197 (1977), p. 789. View Record in Scopus | Cited By in Scopus (85)

[5] R. Gries, G. Khaskin, G. Gries, R.G. Bennett, G.G.S. King, P. Morewood, K.N. Slessor and W.D. Morewood, J. Chem. Ecol. 28 (2002), p. 2283. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (29)

[6] D.R. Miller, C.M. Crowe, C. Asaro and G.L. Debarr, J. Chem. Ecol. 29 (2003), p. 437. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (9)

[7] A. Wibe, A.K. Borg-Karlson, M. Persson, T. Norin and H. Mustaparta, J. Chem. Ecol. 24 (1998), p. 273. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (39)

[8] C.D. Hull, J.P. Cunningham, C.J. Moore, M.P. Zalucki and B.W. Cribb, J. Chem. Ecol. 30 (2004), p. 2071. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (9)

[9] M. Ayasse, F.P. Schiestl, H.F. Paulus, F. Ibarra and W. Francke, Proc. R. Soc. Lond. B 270 (2003), p. 517. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (42)

[10] A. Jürgens, T. Witt and G. Gottsberger, Biochem. Sys. Ecol. 31 (2003), p. 345. Article | PDF (109 K) | View Record in Scopus | Cited By in Scopus (18)

[11] S. Dötterl, L.M. Wolfe and A. Jürgens, Phytochemistry 66 (2005), p. 203. View Record in Scopus | Cited By in Scopus (46)

[12] A. Jürgens, T. Witt and G. Gottsberger, Biochem. Syst. Ecol. 30 (2002), p. 383. Article | PDF (90 K) | View Record in Scopus | Cited By in Scopus (33)

[13] J.T. Knudsen and L. Tollsten, Bot. J. Linn. Soc. 113 (1993), p. 263. Abstract | Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (138)

[14] H. Surburg, M. Güntert and B. Schwarze, J. Ess. Oil Res. 2 (1990), p. 307.

[15] S. Andersson, L.A. Nilsson, I. Groth and G. Bergström, Bot. J. Linnean Soc. 140 (2002), p. 129. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (44)

[16] S. Kint, R. Teranishi, P.D. Lingren, T.N. Shaver and J.R. Raulston, J. Essent. Oil Res. 5 (1993), p. 201. View Record in Scopus | Cited By in Scopus (9)

[17] L. Tollsten and G. Bergström, Nord. J. Bot. 13 (1993), p. 607. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (28)

[18] L. Tollsten and J.T. Knudsen, Pl. Syst. Evol. 182 (1992), p. 229. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (25)

[19] G. Flamini, P.L. Cioni and I. Morelli, Flavour Fragr. J. 17 (2002), p. 147. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (12)

[20] A.J. Matich, H. Young, J.M. Allen, M.Y. Wang, S. Fielder, M.A. McNeilage and E.A.

MacRae, Phytochemistry 63 (2003), p. 285. Article | PDF (600 K) | View Record in Scopus | Cited By in Scopus (23)

[21] G. Mazza, P. Cascio and E. Barbieri, Rivista Vitic. Enol. 56 (2003), p. 57.

[22] L.N. Misra, A. Chandra and R.S. Thakur, Phytochemistry 30 (1991), p. 549. Abstract | PDF (343 K) | View Record in Scopus | Cited By in Scopus (19)

[23] M. Kreck, S. Puschel, M. Wüst and A. Mosandl, J. Agric. Food Chem. 51 (2003), p. 463. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (26)

[24] E. Alissandrakis, D. Daferera, P.A. Tarantilis, M. Polissiou and P.C. Harizanis, Food Chem.

82 (2003), p. 575. Article | PDF (273 K) | View Record in Scopus | Cited By in Scopus (49)

[25] A.C. Soria, M. González, C. de Lorenzo, I. Martínez-Castro and J. Sanz, Food Chem. 85

(2004), p. 121. Article | PDF (297 K) | View Record in Scopus | Cited By in Scopus (32)

[26] A.C. Soria, I. Martinez-Castro and J. Sanz, J. Sep. Sci. 26 (2003), p. 793. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (30)

[27] A.L. Wilkins, Y.R. Lu and S.T. Tan, J. Agric. Food Chem. 41 (1993), p. 873. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (43)

[28] D. Burkhardt and A. Mosandl, J. Agric. Food Chem. 51 (2003), p. 7391. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (10)

[29] S. Dötterl, A. Jürgens, K. Seifert, T. Laube, B. Weißbecker and S. Schütz, New Phytol. 169 (2006), p. 707. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (28)

[30] D. Plepys, F. Ibarra and C. Löfstedt, Oikos 99 (2002), p. 69. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (27)

[31] D. Plepys, Odour-mediated nectar foraging in the silver Y moth, Autographa gamma, PhD Thesis in Department of Ecology, Lund University, Lund, 2001.

[32] S. Andersson, Chemoecology 13 (2003), p. 13. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (19)

[33] S. Andersson and H.E.M. Dobson, J. Chem. Ecol. 29 (2003), p. 2319. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (14)

[34] M. Kreck and A. Mosandl, J. Agric. Food Chem. 51 (2003), p. 2722. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (10)

[35] R.A. Raguso and O. Pellmyr, Oikos 81 (1998), p. 238. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (56)

[36] J.A. Elzinga, A. Biere and J.A. Harvey, Proc. Exper. Appl. Entomol. NEV Amsterdam 13 (2002), p. 109.

[37] B. Weissbecker, G. Holighaus and S. Schütz, J. Chromatogr. A 1056 (2004), p. 209. Article

| PDF (205 K) | View Record in Scopus | Cited By in Scopus (11)

[38] P. Färbert, U.T. Koch, A. Färbert, R.T. Staten and R.T. Cardé, Environ. Entomol. 26 (1997), p. 1105. View Record in Scopus | Cited By in Scopus (17)

[39] S. Schütz, B. Weißbecker, U.T. Koch and H.E. Hummel, Biosens. Bioelectron. 14 (1999), p.

221. Article | PDF (265 K) | View Record in Scopus | Cited By in Scopus (31)

[40] K.E. Kaissling and J. Thorson In: D.B. Satelle, L.M. Hall and J.G. Hildebrand, Editors, Receptors for Neurotransmitters, Hormones and Pheromones in Insects, Elsevier/North-Holland Biomedical Press, Amsterdam, Netherlands (1980), p. 261.

[41] D.D. Trueblood, E.D. Gallagher and D.M. Gould, Limnol. Oceanogr. 39 (1994), p. 1440. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (69)

[42] D.F. Boesch, Application of Numerical Classification in Ecological Investigations of Water Pollution, Special Scientific Report, Institute of Marine Science, Virginia (1977).

[43] I. Borg and J. Lingoes, Multidimensional Similarity Structure Analysis, Springer Verlag, Berlin (1987).

[44] K.R. Clarke, Aust. J. Ecol. 18 (1993), p. 117. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (2673)

[45] S. Wakayama, S. Namba, K. Hosoi and M. Ohno, Bull. Chem. Soc. Jpn. 46 (1973), p. 3183. Full Text via CrossRef

[46] R.A. Raguso and E. Pichersky, Pl. Spec. Biol. 14 (1999), p. 95. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (54)

[47] A.K. Borg-Karlson, J. Tengoe, I. Valterova, C.R. Unelius, T. Taghizadeh, T. Tolasch and W. Francke, J. Chem. Ecol. 29 (2003), p. 1. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (26)

[48] E. Pichersky, E. Lewinsohn and R. Croteau, Arch. Biochem. Biophys. 316 (1995), p. 803.

Abstract | PDF (429 K) | View Record in Scopus | Cited By in Scopus (53)

[49] J. Lücker, H.J. Bouwmeester, W. Schwab, J. Blaas, L.H.W. van der Plas and H.A. Verhoeven, Plant J. 27 (2001), p. 315. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (58)

[50] N.B.M. Brantjes and J.A.A.M. Leemans, Acta Bot. Neerl. 25 (1976), p. 281.

[51] A. Jürgens, T. Witt and G. Gottsberger, Bot. Acta 109 (1996), p. 316. View Record in Scopus | Cited By in Scopus (48)

[52] J. Guitián, P. Guitian and J.M. Sánchez, Pl. Syst. Evol. 185 (1993), p. 153. Full Text via CrossRef | View Record in Scopus | Cited By in Scopus (17)

[53] P.G. Krannitz and M.A. Maun, Can. Field Nat. 105 (1991), p. 13.

[54] S. Dötterl, Importance of floral scent compounds for the interaction between Silene latifolia (Caryophyllaceae) and the nursery pollinator Hadena bicruris (Lepidoptera: Noctuidae), Ph.D. Thesis in Department of Plant Systematics, University of Bayreuth, Bayreuth, 2004.

Corresponding author. Tel.: +49 921 552466; fax: +49 921 552786.

Journal of Chromatography AVolume 1113, Issues 1-2, 28 April 2006, Pages 231-238