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First published online November 30, 2007
Journal of Experimental Biology 210, 4335-4344 (2007)
Published by The Company of Biologists 2007
doi: 10.1242/jeb.009944
Structure, ratios and patterns of release in the sex pheromone of an aphid, Dysaphis plantaginea
1 School of Biological Sciences, University of Southampton, Bassett Crescent
East, Southampton, Hampshire, SO16 7PX, UK
2 BCH Division, Rothamsted Research, Harpenden, Hertfordshire, AL5 2JQ,
UK
3 East Malling Research, New Road, East Malling, Kent, ME19 6BJ,
UK
4 Department of Biological Sciences, Imperial College at Silwood Park,
Ascot, Berkshire, SL5 7PY, UK
* Author for correspondence (e-mail: asj{at}soton.ac.uk)
Accepted 18 September 2007
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Summary |
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Key words: Dysaphis plantaginea, nepetalactol, nepetalactone, oviparae, periodicity, pheromone ratios, rosy apple aphid, sex pheromone
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Introduction |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
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;
Dawson et al.,1989). Since
then, nepetalactol (1 or 2) and nepetalactone (5 or 6) have been found either
singly or as a mixture in the sex pheromone of numerous other species of aphid
(Table 1). However, because
analyses used achiral methods, most identifications did not confirm which
enantiomers were present nor whether the component was a mixture of
enantiomers, hence 1 or 2 and 5 or 6. In one species, Phorodon
humuli, the sex pheromone comprises two different nepetalactol
diastereoisomers (3 and 4) (Campbell et
al., 1990).
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When sampled by air entrainment, aphid oviparae (sexual females) have been
shown to release the two iridoid pheromone components in a ratio that is
relatively species-specific (Table
1); however, there are some studies that indicate that ratios
change with aphid age (Hardie et al.,
1990; Jeon et al.,
2003; Goldansaz et al.,
2004). Males generally show strongest behavioural responses
towards their typical conspecific ratio but some laboratory studies have
reported response to a broad range of synthetic ratios and even to a pheromone
ratio released by a different species of aphid
(Petterson, 1971;
Marsh, 1975;
Dawson et al., 1990;
Hardie et al., 1990;
Steffan, 1990;
Lilley and Hardie, 1996). A
similar lack of pheromone specificity has also been reported from field
trapping, as a given ratio sometimes catches many species of male aphid
(Boo et al., 2000;
Goldansaz et al., 2004). If
nepetalactol and nepetalactone are ubiquitous aphid sex pheromone components,
as has been suggested (Dawson et al.,
1990), ratio combinations are limited and additional mechanisms
for species isolation are likely to exist. Numerous factors have been proposed
and these include the use of different blends of diastereoisomers or
enantiomers of nepetalactol and nepetalactone
(Campbell et al., 1990;
Hardie et al., 1997), the
presence of unidentified additional constituents
(Guldemond et al., 1993;
Lilley and Hardie, 1996),
interactions with host-plant volatiles (Petterson et al., 1970a;
Campbell et al., 1990;
Hardie et al., 1994b;
Losël et al., 1996),
differences in oviparae colour and species-specific movements
(Steffan, 1990), genitalia
incompatibility, spatial and seasonal separation of populations
(Hardie et al., 1990), and
circadian or diel separation of pheromone release
(Guldemond and Dixon, 1994;
Thieme and Dixon, 1996).
It is known for Lepidoptera (butterflies and moths) that the timing of sex
pheromone release within the daily light:dark cycle can be an important
component for effective communication that also parallels appropriate
behavioural responsiveness in the conspecific receiver. There are typically
species-specific peaks in pheromone release during the light:dark cycle;
however, exact onset and duration is modulated by age and abiotic factors
(Dreisig, 1986;
Delisle and McNiel, 1987;
McNeil, 1991;
Rosén, 2002;
Silvegren et al., 2005;
Mazor and Dunkelblum, 2005).
Some have reported that aphids release sex pheromone only during the
photophase (Marsh, 1972;
Eisenbach and Mittler, 1980;
Eisenbach and Mittler, 1987)
and in sympatric situations there is some evidence that pheromone release
during the photophase is divided according to the aphid sub-species
(Guldemond and Dixon, 1994;
Thieme and Dixon, 1996).
However, in aphid studies so far, inferences about sex pheromone release
throughout the day have been indirect and categorical and are drawn from
observations of female calling behaviour and/or male responses (e.g.
Marsh, 1972;
Eisenbach and Mittler, 1980;
Eisenbach and Mittler, 1987;
Guldemond and Dixon, 1994;
Thieme and Dixon, 1996).
Although technically challenging, direct measurements of sex pheromone
chemicals are continuous and quantitative. Such a direct approach has recently
been attempted by Jeon et al. (Jeon et
al., 2003) but measurements were limited to a single day and
resolution was at 2 h intervals. With a greater understanding of sex pheromone
release, physiological and evolutionary aspects of aphid biology can be
addressed, implications for natural enemies who exploit this pheromone can be
evaluated, and improvements to aphid management approaches facilitated
(Hardie et al., 1991;
Hardie et al., 1994a;
Powell et al., 1993;
Powell et al., 1998;
Gabrys et al., 1997;
Boo et al., 1998;
Zhu et al., 1999).
The rosy apple aphid [Dysaphis plantaginea (Passerini); Homoptera,
Aphididae] is the most serious pest of apple in Europe
(Blommers, 1994). It is a
host-alternating species that, over the summer, reproduces parthenogenetically
on herbaceous Plantago lanceolata L. (Plantaginaceae). In early
autumn, a sexual generation is produced. First to develop are the gynoparae
(winged females), who return to the apple tree, where they lay the oviparous
stage. Once mature, the oviparae release a sex pheromone that approximately
corresponds with the development of winged males on P. lanceolata.
Males are attracted and mate with the oviparae, who then lay cold-hardy
overwintering eggs on the apple tree. Further details of aphid life cycles can
be found in Dixon (Dixon,
1998). In the present study, we determine a typical ratio of
nepetalactol and nepetalactone for D. plantaginea, confirm the
enantiomeric form of the iridoids and present uninterrupted hourly sampling
data on the pheromone released from a group of same-aged oviparae over a
20-day period. A simple and inexpensive sequential sampling device was
developed to examine release periodicity through consecutive diel cycles and
also to investigate the stability of ratios throughout the day and over
increasing aphid age. This is the first report on the full identification and
ratio of the two major components of D. plantaginea sex pheromone and
is also the most detailed temporal study on sex pheromone release from any
species of aphid.
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Materials and methods |
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In 2003, an excised branch and detached leaves heavily infested with mixed-aged oviparae of D. plantaginea and uninfested plant material were collected from apple, cv. Braeburn overgrafted onto Katy on a MM106 rootstock. In addition, a same-aged cohort of oviparae (N=35) were reared outdoors from gynoparae of D. plantaginea that were enclosed overnight by mesh on a branch of potted apple, cv. Cox's Orange Pippin on MM106 rootstock. Samples collected in 2003 from the oviparae were used to determine the ratio of the two major components of D. plantaginea sex pheromone. Samples from the detached leaves were used for full structural identification.
In 2004, gynoparae (N=125) were collected in the apple orchard cv. Cox's Orange Pippin on MM106 and were placed onto the leaves of a fresh branch of apple (cv. Cox's Orange Pippin, 45 cm long) standing in a pot of distilled water. This branch and gynoparae were then enclosed in mesh and placed at the onset of scotophase in a controlled environment (CE) room that mimicked outside conditions [L:D 12 h:12 h; photophase 09.00–21.00 h BST (British summer time), 16±0.5°C; scotophase 21.00–09.00 h BST, 12±0.5°C]. After 24 h, the gynoparae were removed, leaving a cohort of same-aged oviparae (N=95). The mesh bag was replaced and the branch with oviparae returned to the CE room. When adult, these oviparae were used in the temporal study.
Collection of pheromone for identification and ratios (aphids collected in 2003)
All volatiles were trapped onto Porapak Q 50/80 (50 mg; Supelco,
Bellefonte, PA, USA) held in glass tubing (5 mm o.d.) by two plugs of
silanised glass wool. The Porapak Q was conditioned by washing with
redistilled diethyl ether (5 ml) and heating at 132°C for 2 h under a
stream of nitrogen. After the air entrainment, the volatiles were eluted from
the Porapak with redistilled diethyl ether (750 µl) and samples were stored
in a freezer (–22°C).
The infested apple branch and detached leaves were placed in two glass vessels with volumes of 2 litres and 500 ml, respectively. Air that had been purified by passage through an activated charcoal filter (BDH, Poole, Dorset, UK; 10–14 mesh, 50 g) was pushed into (800 ml min–1) and pulled out of (700 ml min–1) the vessels. The difference in air volume passed outwards at seals that were not airtight. Volatiles were entrained onto Porapak Q for 4 days. The same procedure was performed on an uninfested branch and leaves.
The same-aged adult oviparae (N=35) reared on the potted Cox's plant were moved onto a single leaf that was enclosed by a cleaned cylindrical glass vessel that had a slot for the petiole (length 8 cm, depth 5 cm). This slot was plugged with a ball of silanised glass wool, which prevented aphids from escaping but allowed excess air volume to pass outwards. For comparison, a second vessel enclosed an uninfested leaf. Air that had been purified by passage through an activated charcoal filter (BDH, 10–14 mesh, 50 g) was pushed into (400 ml min–1) and pulled out of (350 ml min–1) both vessels. Volatiles were entrained onto Porapak Q for 24 h.
The sequential sampling machine
The mains power source (240 V AC) was split, and half was transformed to
power the 24 V DC circuit. Twelve 24 h timers (ETU2000; Timeguard Ltd, London,
UK) and 240 V AC DPDT power relays (Maplin Electronics, Wombwell, Yorkshire,
UK) were wired as pairs, in parallel, into the 240 V circuit. Twelve 24 V
two-way solenoid valves (Valeader Pneumatics Ltd, Cambridge, UK) were wired in
parallel into the 24 V DC circuit. Since relays interrupted power to
individual solenoid valves, the valves were normally in a closed position.
However, when activated by its respective timer, the relay would complete the
circuit for an individual valve and thus cause it to open. When in an open
state, each valve allowed air to be pulled through one of the 12 sampling
lines that it regulated; by using flow-meters, flow through each arm was
confirmed to be equal. The electronic components were arranged inside a wooden
box and timers surrounded a central piece of glassware (the distribution
chamber) (Fig. 2). The
distribution chamber was a tube of glass (12 mm i.d.) sealed at one end with
six paired side arms, each with a PTFE tap (4 mm hole). The open top end of
the chamber was the inlet through which volatiles would be drawn and,
depending on which valve was activated, the volatiles would be pulled out
through one of the side arms.
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). The open end at the bottom of the vessel was closed using
two semicircular aluminium plates that, when put together, left a small hole
in the middle for the apple stem to pass through, and any gap was sealed with
balls of PTFE tape. Air that had been purified by passage through activated
charcoal (BDH, 10–14 mesh, 50 g) was then pushed in at the inlet (1.2 l
min–1) and the aphids were allowed to settle on the new
branch in the chamber for 24 h. Excess air volume passed outwards through the
gap loosely sealed by PTFE tape.
The sequential hourly sampling of headspace started at the beginning of the second day of the adult stadium at the onset of scotophase (21.00 h BST). Timers were synchronised with the CE room, and each was programmed to open their respective solenoid valve for 1 h at two different times 12 h apart (e.g. solenoid 1 09.00–10.0 h BST and 21.00–22.00 h BST). Sampling tubes therefore had to be changed once every 12 h before the sequence started again. PTFE taps were closed for changeover of tubes so that the sample being collected was not disrupted.
The sampling machine was connected to the outlet of the glass vessel by a short length of PTFE tubing (6 mm i.d.), and the air was pulled out of the vessel at a rate of 1 l min–1 through one of the 12 sampling tubes containing Porapak Q positioned on the distribution chamber. This system of greater inlet flow than outlet sampling, although causing a loss of 1/6th of the headspace (corrected for in quantitation), ensured that the glass chamber remained uncontaminated by outside sources. Samples were captured onto Porapak Q, eluted with redistilled diethyl ether (750 µl), and an internal standard of dodecane (400 ng) was added to each. Samples were reduced to 40 µl under a stream of purified nitrogen before analysis.
Laboratory calling behaviour
Pheromone-releasing plaques, or pseudorhinaria, are located on the hind
tibia of aphids, and calling behaviour is characterised by the raising of hind
legs and abdomen (Petterson,
1970b). In the laboratory, the number of oviparae displaying
calling behaviour was recorded every hour on three different days. When the
oviparae were 14, 18 and 21 days into the adult stadium, starting at 09.00 h
(12 h of scotophase or 0 h of photophase), a count of how many oviparae were
calling was made. Every hour, further counts were made until the onset of
scotophase (21.00 h), and the last count was made 5–10 min into the
scotophase using a red light.
Analysis of pheromone samples
Samples entrained from the infested branch and leaves in 2003 were analyzed
(1 µl) by gas chromatography (GC) on non-polar (HP-1, 50 mx0.32 mm
i.d.x0.52 µm film thickness; J%W Scientific, Falcon, CA, USA) and
polar (HP-wax, 30 mx0.23 mm i.d.x0.5 µm film thickness)
capillary columns, using a 6890 GC machine (Agilent Technologies UK Ltd,
Stockport, Cheshire, UK) fitted with a cool on-column injector and a flame
ionisation detector (FID). The oven was kept at 30°C for 1 min, heated at
5°C min–1 to 150°C and then 10°C
min–1 to 250°C (220°C for the wax column), where it
was maintained for 20 min. The carrier gas was hydrogen.
Samples entrained in 2003 from oviparae reared from gynoparae and a selection of samples collected in 2004 for the temporal study were initially analysed on a Hewlett-Packard 5890 Series II GC (Hewlett-Packard Co., Palo Alto, CA, USA) linked to a Hewlett-Packard 5971 Quadrupole Mass Selective Detector ionising by electron impact at 70 eV. The column used was a non-polar fused capillary column (HP-1MS, 30 mx0.25 mm i.d.x0.25 µm film thickness). The carrier gas was helium (constant 62 034 Pa) and oven temperature was held at 40°C for 2 min then programmed at 5°C min–1 to 150°C then 10°C s–1 to 250°C and held for 16 min. Samples (1 µl) were injected into a split/splitless injector (220°C, splitless 1 min) and data were captured and analysed by Enhanced ChemStation software (v. A.03.00; Hewlett-Packard Co.). After tentative identification of nepetalactol and nepetalactone, samples were also run on a Hewlett-Packard 5890 Series II GC fitted with split/splitless injector (220°C, splitless 30 s) and FID. This instrument was used for analysis of samples from the temporal study. The principal capillary column was non-polar (SOLGEL-1, 30 mx0.32 mm i.d.x0.25 µm film thickness, HP-1 equivalent; SGE, Melbourne, Victoria, Australia) and for confirmation a polar column was used (CARBOWAX, 30 mx0.32 mm i.d.x0.25 µm film; Alltech, Stamford, Lincolnshire, UK). The oven was kept at 40°C for 1 min, heated at 10°C min–1 to 250°C (220°C for Wax) and held for 2 min. The carrier gas was helium (constant 35 cm s–1). Samples were injected (2 µl) in splitless mode and data were captured using a 35900 HPIB interface (Hewlett-Packard Co.) and analysed using HP 3365 Series 2 ChemStation (v. A.03.01).
Tentative identifications were confirmed to the enantiomeric pair by peak enhancement with authenticated samples of (1R,4aS,7S,7aR)-nepetalactol and (4aS,7S,7aR)-nepetalactone on both non-polar (HP-1 or SOLGEL-1) and polar (HP-wax or CARBOWAX) columns. Quantities were initially determined using a multiple point external standard method but for the temporal study an internal standard was used. For the temporal study, when peaks were very small or not detected, samples were further concentrated using nitrogen before reinjection.
Confirmation of nepetalactol enantiomer by derivatisation and nuclear magnetic resonance (NMR)
Full structural identification of the enantiomer of nepetalactol was
achieved by microscale NMR spectroscopy after derivatisation. Air entrainment
samples of D. plantaginea oviparae, containing 84 µg of
nepetalactol (1) (GC approximation) were concentrated under a stream of
purified nitrogen and dissolved in dichloromethane (0.5 ml) under nitrogen. A
solution of
(S)-(+)-
-methoxy--(trifluoro-methyl)phenylacetyl
chloride (40 mg, 0.16 mmol) and pyridine (25 µl) in dichloromethane (0.5
ml), prepared under nitrogen, was added, together with a few crystals of
dimethylaminopyridine, and the reaction stirred overnight. The solvent was
then removed under a stream of nitrogen and the residue partially redissolved
in 10% diethyl ether in petroleum ether (40–60°C boiling fraction).
The insoluble material was discarded by decanting off the soluble portion. The
solvent was then removed under a stream of nitrogen and the residue
redissolved in deuteriochloroform for NMR analysis. 1H,
13C and 19F NMR spectroscopy was performed using a
Bruker 500 Avance NMR spectrometer (Billerica, MA, USA) with 1H
referenced to CDCl3 (7.25 p.p.m.), 13C to
CDCl3 (77.0 p.p.m.) and 19F to CFCl3 (0
p.p.m.). Quantitative 1H NMR spectroscopy was performed using a
pulse angle of 30°, an acquisition time of 5T1 (with T1 measured to be 2.5
s) and a delay of 5 s. The NMR analysis was compared to diastereoisomeric
derivatives from the two enantiomers of synthetic nepetalactol (1 and 2), and
full details are described elsewhere
(Goldansaz et al., 2004).
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transformed. All
data were checked for normality and homogeneity of variance using the
Anderson-Darling and Levene's tests. Paired Student's t-tests were
used to determine whether pheromone release was significantly increased during
photophase relative to the preceding scotophase and whether there were
significant decreases during scotophase relative to the preceding photophase.
In order to test for peaks in sex pheromone release within the photophase,
time periods of 0–3, 3–6, 6–9 and 9–12 h into
photophase were ascribed, and mean release rate for each was calculated and
subjected to analysis of covariance (ANCOVA) in which maximum pheromone
release for that photoperiod was factored as the covariate to account for age
influences. The same test was used to investigate scotophase changes in
pheromone quantity. The effect of aphid age on percentage nepetalactol in the
mixture was investigated by one-way analysis of variance (ANOVA). A two-way
ANOVA tested for scotophase and photophase differences in ratios.
Post-hoc Tukey's tests were used to identify where significant
differences lay within the above. All data were analysed on Minitab 13.1
(Minitab Inc., State College, PA, USA), and differences were considered
significant if P
0.05. |
Results |
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The mean ratio of the two identified components, based on entrainment samples taken from four different batches of D. plantaginea from two different apple cultivars over two years, was calculated as 3.7:1 (1R,4aS,7S,7aR)-nepetalactol: (4aS,7S,7aR)-nepetalactone (Table 2).
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Diel patterns in pheromone release
The data show a distinct cycling in the levels of pheromone collected (as
measured by summed quantities of nepetalactol and nepetalactone), with
significant differences between the photophase and scotophase
(Fig. 4) (both analyses
P<0.001, N=20). With the onset of photophase, pheromone
production increased gradually for the first three hours of photophase and
thereafter remained at high levels until the end of the photophase sampling
(P<0.001, N=20; 0–3 h lower than 3–6,
6–9 or 9–12 h). With the onset of scotophase, pheromone production
decreased rapidly, although low levels were detected throughout the scotophase
(P<0.001, N=20; 0–3 h higher than 3–6,
6–9 or 9–12 h).
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Changes in the ratio of nepetalactol:nepetalactone
Over adult stadium ages 2–21 days, the cohort of 95 oviparae produced
a total of 76.6 µg of
(1R,4aS,7S,7aR)-nepetalactol (1) and 22.9
µg of (4aS,7S,7aR)-nepetalactone (5), thus
giving a blend of 3.34:1 (77.0% nepetalactol). However, analysis found a
significant age effect (P<0.001)
(Fig. 6A), with ratios
remaining steady until the fourteenth day of the adult stadium, and thereafter
the relative amount of nepetalactol in the mixture declined. Total quantity of
pheromone based on ages 2–14 days of the adult stadium were 58.4 µg
and 15.8 µg for 1 and 5, respectively, which gives a ratio of 3.69:1 (78.7%
nepetalactol).
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Observations of calling behaviour
On all three days that behaviour was recorded, a similar pattern of
characteristic behaviours attributed to sex-pheromone release was observed
(Fig. 7). In the laboratory, no
calling behaviour was exhibited at the onset of photophase (12 h of
scotophase, also 0 h of photophase). During the first 3 h of photophase
(0–3 h), an increasing number of oviparae were observed to display
calling behaviours; for the remainder of the photophase (4–11 h),
>90% of the oviparae were exhibiting calling behaviour. Whilst >90% of
oviparae were calling, oviparae were never observed all calling at the same
time. The last observation of the day was made 5–10 min into scotophase,
and no oviparae were observed to be calling at this time. It was assumed that
throughout the rest of the scotophase there was no calling behaviour; this is
what others have reported in Aphis spiraecola (e.g.
Jeon et al., 2003).
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Discussion |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
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; Pickett et al.,
1992). Since 1 and the nepetalactone are believed to be on the
same biosynthetic pathway, and given that D. plantaginea produces 1,
the enantiomer of nepetalactone is most likely to be
(4aS,7S,7aR)-nepetalactone (5) because inversion of
three chiral centres is unlikely to occur easily. A typical ratio of
components for D. plantaginea would be 3.7:1 (1:5) and, in an
olfactometer, male D. plantaginea are attracted to a synthetic
mixture of these components (J.D.F., unpublished data).
Diel patterns in pheromone release
The sequential sampling device has enabled us to gather the most in-depth
dataset on sex pheromone release for any species of aphid. The data presented
here substantiates, in part, previous conclusions made by other workers who
recorded calling behaviour only during the photophase
(Marsh, 1972;
Eisenbach and Mittler, 1987;
Guldemond and Dixon, 1994;
Thieme and Dixon, 1996;
Jeon et al., 2003). We
observed no calling behaviour at the onset of photophase or shortly after the
onset of scotophase but did record intense calling behaviour during most of
the photophase. Following hourly analysis of pheromone release, it can be seen
that there is a clear cycling in the release of sex pheromone from D.
plantaginea oviparae. High levels of pheromone were always associated
with the photoperiods, there was then a rapid drop, and low levels were
detected throughout the scotophase. Other workers have reported similar
findings; Dawson et al. (Dawson et al.,
1990) and Jeon et al. (Jeon et
al., 2003) both collected headspace samples from oviparae
(Schizaphis graminum and A. spiraecola, respectively) during
the scotophase when calling behaviour was not exhibited, and both detected
pheromone in low quantity relative to photophase measurements. These
scotophase measurements could have been contamination left over from the
previous photophase; for example, on the glass parts of the apparatus.
However, in our experiment, adsorption onto the glass with continual slow
release would be unlikely over such extended periods with continuous aeration
(Stewart-Jones and Poppy,
2006). Another possibility is that pheromone was absorbed into the
waxy cuticles of the apple leaves or the aphids themselves. Again, this
explanation is unlikely because pheromone levels would be expected to tail and
gradually decrease with time into the scotophase. Rather than decreasing
during scotophase, in the first five days of the experiment pheromone levels
appeared to be increasing during scotophase
(Fig. 4) and this might reflect
a gradual development of biosynthetic pathway capacity for pheromone
production as the aphids matured. An important factor to remember relates to
the morphology of oviparae, who produce the pheromone in their hind tibia from
glandular cells just below a highly thinned area of cuticle that is traversed
by many large secretory canals (Petterson,
1970b; Zeng et al.,
1992). In contrast to most female Lepidoptera, who physically
expose the sex pheromone gland from within their bodies, aphid oviparae have
no such behavioural control over release. Essentially what is biosynthesised
by the glandular cells of oviparae probably diffuses out with little other
control. Thus, whilst calling behaviour in Lepidoptera can regulate release at
a secondary level to biosynthesis, for example in Trichoplusia ni
(Noctuidae) (Tang et al.,
1989), calling behaviour in aphid oviparae probably serves simply
to aid dispersal. Although unknown for aphids, changes in biosynthesis are
likely to be mediated by neurohormones in the PBAN (pheromone biosynthesis
activating peptide) family, as has been found to be the case in five other
insect orders (Rafaeli and Jurenka,
2003). Therefore, the changes we recorded are most likely the
result of pathway upregulation and downregulation. While we know that
pheromone is biosynthesised de novo
(Pickett et al., 1992), our
results and those of others (Dawson et al.,
1990; Jeon et al.,
2003) suggest that, although downregulated during scotophase,
biosynthesis does not cease completely. Future study into pathway regulation
might focus on gene expression analysis of pheromone biosynthesis genes using
genomic techniques such as aphid microarrays or real time-polymerase chain
reaction (rt-PCR).
From the current study, what is not known is whether pheromone production
is truly under circadian control, i.e. is it driven by an endogenous clock or
is it driven by external stimuli only? Others using behavioural measures have
concluded that pheromone release is driven by an endogenous circadian rhythm
(Marsh, 1972;
Eisenbach and Mittler, 1980).
By analysing pheromone collected in headspace from three different-aged aphids
under constant darkness, Jeon et al. also concluded that pheromone release was
under circadian control (Jeon et al.,
2003).
Diel peaks in pheromone release
Other than a photophase–scotophase pattern in pheromone release,
there is no laboratory evidence to suggest that D. plantaginea
oviparae have narrow time periods within the photophase during which
particularly high levels of sex pheromone are released. In the autumns of
2003, 2004 and 2005, mating in the field was observed throughout the
photoperiod and such observations would suggest that distinct periods of high
pheromone release do not occur in the field either. D. plantaginea
were observed mating in the field at 09.30, 10.45, 11.15, 11.20, 13.00, 13.20,
14.50 and 16.30 h BST (A.S.-J., unpublished data).
During photophase, laboratory-recorded patterns of pheromone release were
in agreement with laboratory observations of calling behaviour. Nearly all
oviparae were calling 3 h into the photophase and continued to do so until the
onset of scotophase. Similarly, Marsh concluded that maximum pheromone release
is reached 2 h into the photophase (Marsh,
1972). Although Marsh found that calling in M. viciae
waned before the onset of scotophase in old and young oviparae
(Marsh, 1972), in the current
study this was not found to be the case. In the laboratory under constant
conditions, calling might indicate release of pheromone; however, there are
significant age influences on quantity (see below). In the field, where rain
or wind can curtail calling behaviour
(Goldansaz and McNeil, 2003),
we stress that, due to the morphology of aphid oviparae (discussed above), an
observed absence of calling behaviour may not necessarily indicate no release
of pheromone. In rain or high wind conditions, which might occur regularly in
the field, release of pheromone without calling could still serve to attract
males in the immediate vicinity. Aphids are relatively weak fliers compared
with Lepidoptera and it has been suggested that when trying to locate oviparae
at close range, upwind walking by the males is particularly important and this
is only punctuated by brief flights when weather conditions are favourable
(Goldansaz and McNeil, 2006).
If aphids such as D. plantaginea were to restrict sex pheromone
release to a narrow time period within the photophase, the probability of a
successful mating would be dramatically reduced given weather variability. We
therefore think that, in contrast to Lepidoptera, most aphids are unlikely to
have temporally narrow periods of very high sex pheromone release, and for
D. plantaginea this is supported by our data. In a diverging species
complex or races that coexist sympatrically on the same plant, there are
reports of temporally narrow periods of calling behaviour (e.g.
Guldemond and Dixon, 1994;
Thieme and Dixon, 1996). In
these particular situations, temporal partitioning might exist, but this would
need to be verified by actual measurements of pheromone release.
Effect of age on pheromone quantity and calling behaviour
Oviparae of potato aphids (Macrosiphum euphorbiae) do not display
calling behaviour until 2–3 days into the adult stadium
(Goldansaz and McNeil, 2003),
and the pheromone components were not detected until the second day into the
adult stadium for the vetch aphid (M. viciae)
(Hardie et al., 1990). Our
first day of sampling was on the second day of the adult stadium, and low
levels of pheromone were detected. Data show that sex-pheromone release in
D. plantaginea was highest on the eighth day of the adult stadium
before gradually decreasing again. This is consistent with other behavioural
studies that found male M. viciae responsiveness to calling oviparae
to be strongest when oviparae were 6 days into the adult stadium
(Marsh, 1972). Also, Aphis
fabae calling behaviour was observed to intensify until day 8 of the
adult stadium (Thieme and Dixon,
1996). Similarly, chemical studies that sampled for 24 h periods
from M. viciae found pheromone release to be highest on day 6
(Hardie et al., 1990). These
slight variations in age-related maximal pheromone output/attractiveness might
be species characteristics or may be due to different temperatures used in
rearing.
The maximum pheromone output for the vetch (M. viciae) and spiraea
(A. spiraecola) aphids has been reported to be over 200 ng per
ovipara per day. However, for both species, more typical levels were in the
region of 100–150 ng per ovipara per day
(Hardie et al., 1990;
Jeon et al., 2003). For D.
plantaginea, the maximum we calculated was 99.6 ng per ovipara per day,
with typical levels in the region of 60–90 ng per aphid per day. Even
after 3 weeks, D. plantaginea were still calling and able to release
pheromone. However, the quantity of pheromone being released was in decline
and there may have been only limited release beyond the 3-week period. This
long period over which adult D. plantaginea oviparae can release sex
pheromone might be particularly relevant to mating success in the field
because, in autumn, extended periods of poor weather, which delays mate
location by the males, may be quite common.
Comparison of our recorded calling behaviour and the quantified levels of
pheromone released showed a poor relationship. Whereas observations of aphids
aged 14, 18 and 21 days into the adult stadium showed no significant
difference in the number of oviparae calling
(Fig. 7), direct measurement of
pheromone during these photophases found significant differences (means of
45.9, 26.7, 12.0 ng per aphid during respective photophases). Similarly, Jeon
et al. noted intense calling behaviour in 3-week-old oviparae but detected no
pheromone (Jeon et al., 2003).
Again, these examples reinforce the idea that calling behaviour is an
unreliable indicator of quantity of pheromone being released.
Effect of age on (1R,4aS,7S,7aR) nepetalactol:(4aS,7S,7aR)-nepetalactone ratios
Unlike another study, which reported a significant fluctuation in ratios
with age of M. viciae (Hardie et
al., 1990), we did not find significant changes in iridoid ratios
for D. plantaginea, and only a slight decrease in the relative amount
of nepetalactol was recorded when the aphids were more than 14 days into the
adult stadium. By sampling 24 times per day, we have certainty in our
measurements, and the low variability in our dataset does support the idea
that species-specific ratios are an important component of intraspecies
recognition. Interestingly, the two other studies in which ratios have been
quantified over age also report a slight decrease in the relative amount of
nepetalactol with age (Jeon et al.,
2003; Goldansaz et al.,
2004). There may be a biological function for this change –
for example, to optimise attractant or aphrodisiac properties of the pheromone
– as suggested by Hardie et al.
(Hardie et al., 1990).
Alternatively, this decrease might suggest that feedback pathways between
biosynthesis of nepetalactol and selective oxidation to nepetalactone may be
less synchronous in older aphids, or it may be a symptom of senescence in the
glandular pheromone-producing cells.
Conclusions
Dysaphis plantaginea oviparae predominantly release sex pheromone
during the photophase; however, low levels are released throughout the
scotophase. Pheromone release is greatest on the eighth day of the adult
stadium and this species releases a typical blend of 3.7:1
(1R,4aS,7S,7aR)-nepetalactol:(4aS,7S,7aR)-nepetalactone.
Calling behaviour is poorly correlated with quantities of pheromone released
across different ages; conversely, due to aphid morphology, lack of calling
may not necessarily indicate no release of pheromone. We therefore suggest
caution when extrapolating calling behaviour to make inferences on pheromone
release. Lack of a temporally narrow and distinct period of high sex-pheromone
release during photophase suggests that alternative mechanisms or factors for
species recognition and isolation exist. Further study should aim to
understand how aphids in the field avoid cross-communicating, given that many
share the same iridoid components in their sex pheromone. Questions relating
to the influence of graded environmental variables, such as wind, light or
temperature, could be explored with regard to measured release of pheromone
and the ratio of components. Of particular interest would be a better
understanding of the mechanisms regulating biosynthesis of sex pheromone in
aphids. Aphids represent a fascinating superfamily of insects with unusual
life cycles. Many species are of great agricultural importance, and continued
progress in our understanding is vital if we are to successfully exploit our
biological and chemical knowledge of these insects for human benefit.
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Acknowledgments |
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References |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
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Blommers, L. H. M. (1994). Intergrated pest management in European apple orchards. Annu. Rev. Entomol. 39,213 -241.[CrossRef]
Boo, K. S., Chung, I. B., Han, K. S., Pickett, J. A. and Wadhams, L. J. (1998). Response of the lacewing Chrysopa cognata to pheromones of its aphid prey. J. Chem. Ecol. 24,631 -643.[CrossRef]
Boo, K. S., Choi, M. Y., Chung, I. B., Eastop, V. F., Pickett, J. A., Wadhams, L. J. and Woodcock, C. M. (2000). Sex pheromone of the peach aphid, Tuberocephalus momonis and optimal blends for trapping males and females in the field. J. Chem. Ecol. 26,601 -609.[CrossRef]
Campbell, C. A. M., Dawson, G. W., Griffiths, D. C., Petterson, J., Pickett, J. A., Wadhams, L. J. and Woodcock, C. M. (1990). Sex attractant pheromone of the damson-hop aphid Phorodon humuli (Homoptera: Aphididae). J. Chem. Ecol. 16,3455 -3465.[CrossRef]
Dawson, G. W., Griffiths, D. C., Janes, N. F., Mudd, A., Pickett, J. A., Wadhams, L. J. and Woodcock, C. M. (1987). Identification of an aphid sex pheromone. Nature 325,614 -616.[CrossRef]
Dawson, G. W., Griffiths, D. C., Merritt, L. A., Mudd, A., Pickett, J. A., Wadhams, L. J. and Woodcock, C. M. (1988). The sex pheromone of the greenbug, Schizaphis graminum. Entomol. Exp. Appl. 48,91 -93.[CrossRef]
Dawson, G. W., Janes, N. F., Mudd, A., Pickett, J. A., Slawin, A. M. Z., Wadhams, L. J. and Williams, D. J. (1989). The aphid sex-pheromone. Pure Appl. Chem. 61,555 -558.[CrossRef]
Dawson, G. W., Griffiths, D. C., Merritt, L. A., Mudd, A., Pickett, J. A., Wadhams, L. J. and Woodcock, C. M. (1990). Aphid semiochemicals – a review, and recent advances on the sex pheromone. J. Chem. Ecol. 16,3019 -3030.[CrossRef]
Delisle, J. and McNeil, J. N. (1987). Calling behaviour and pheromone titre of the true armyworm Pseudaletia unipuncta (Haw.) (Lepidoptera: Noctuidae) under different temperature and photoperiodic conditions. J. Insect Physiol. 33,315 -324.[CrossRef]
Dixon, A. E. F. (1998). Aphid Ecology (2nd edn). The Netherlands: Kluwer.
Dreisig, H. (1986). Timing of daily activities in adult Lepidoptera. Entomol. Gen. 12, 25-43.
Eisenbach, J. and Mittler, T. E. (1980). An aphid circadian rhythm – factors affecting the release of sex pheromone by oviparae of the greenbug, Schizaphis graminum. J. Insect Physiol. 26,511 -515.[CrossRef]
Eisenbach, J. and Mittler, T. E. (1987). Sex pheromone discrimination by male aphids of a biotype of Schizaphis graminum. Entomol. Exp. Appl. 43,181 -182.[CrossRef]
Gabrys, B. J., Gadomski, H. J., Klukowski, Z., Pickett, J. A., Sobota, G. T., Wadhams, L. J. and Woodcock, C. M. (1997). Sex pheromone of cabbage aphid Brevicoryne brassicae: identification and field trapping of male aphids and parasitoids. J. Chem. Ecol. 23,1881 -1890.[CrossRef]
Goldansaz, S. H. and McNeil, J. N. (2003). Calling behaviour of the potato aphid Macrosiphum euphorbiae oviparae under laboratory and field conditions. Ecol. Entomol. 28,291 -298.[CrossRef]
Goldansaz, S. H. and McNeil, J. N. (2006). Effect of wind speed on the pheromone-mediated behavior of sexual morphs of the potato aphid, Macrosiphum euphorbiae (Thomas) under laboratory and field conditions. J. Chem. Ecol. 32,1719 -1729.[CrossRef][Medline]
Goldansaz, S. H., Dewhirst, S., Birkett, M. A., Hooper, A. M., Smiley, D. W. M., Pickett, J. A., Wadhams, L. and McNeil, J. N. (2004). Identification of two sex pheromone components of the potato aphid, Macrosiphum euphorbiae (Thomas). J. Chem. Ecol. 30,819 -834.[CrossRef][Medline]
Guldemond, J. A. and Dixon, A. F. G. (1994). Specificity and daily cycle of release of sex pheromones in aphids: a case of reinforcement. Biol. J. Linn. Soc. Lond. 52,287 -303.[CrossRef]
Guldemond, J. A., Dixon, A. F. G., Pickett, J. A., Wadhams, L. J. and Woodcock, C. M. (1993). Specificity of sex pheromones, the role of host plant odour on the olfactory attraction of males, and mate recognition in the aphid Cryptomyzus. Physiol. Entomol. 18,137 -143.
Hardie, J., Holyoak, M., Nicholas, J., Nottingham, S. F., Pickett, J. A., Wadhams, L. J. and Woodcock, C. M. (1990). Aphid sex pheromone components: age-dependent release by females and species specific male response. Chemoecology 1, 63-68.[CrossRef]
Hardie, J., Nottingham, S. F., Powell, W. and Wadhams, L. J. (1991). Synthetic aphid sex pheromone lures female parasitoids. Entomol. Exp. Appl. 61,97 -99.[CrossRef]
Hardie, J., Nottingham, S. F., Dawson, G. W., Harrington, R., Pickett, J. A. and Wadhams, L. J. (1992). Attraction of field-flying aphid males to synthetic sex pheromone. Chemoecology 3,113 -117.[CrossRef]
Hardie, J., Hick, A. J., Holler, C., Mann, J., Merritt, L., Nottingham, S. E., Powell, W., Wadhams, L. J., Witthinrich, J. and Wright, A. F. (1994a). The responses of Praon spp. Parasitoids to aphid sex-pheromone components in the field. Entomol. Exp. Appl. 71,95 -99.[CrossRef]
Hardie, J., Storer, J. R., Nottingham, S. F., Peace, L., Harrington, R., Merritt, L. A., Wadhams, L. J. and Wood, D. K. (1994b). The interaction of sex pheromone and plant volatiles for field attraction of male bird-cherry aphid, Rhopalosiphum padi.Proc. Brighton Crop Prot. Conf. Pests Diseases 3,1223 -1230.
Hardie, J., Peace, L., Pickett, J. A., Smiley, D. W. M., Storer, J. R. and Wadhams, L. J. (1997). Sex pheromone stereochemistry and purity affect field catches of male aphids. J. Chem. Ecol. 23,2547 -2554.[CrossRef]
Jeon, H., Han, K. S. and Boo, K. S. (2003). Sex pheromone of Aphis spiraecola (Homoptera: Aphididae): composition and circadian rhythm in release. J. Asia Pac. Entomol. 6, 159-165.
Lilley, R. and Hardie, J. (1996). Cereal aphid responses to sex pheromones and host-plant odours in the laboratory. Physiol. Entomol. 21,304 -308.
Lilley, R., Hardie, J., Merritt, L. A., Pickett, J. A., Wadhams, L. J. and Woodcock, C. M. (1995). The sex pheromone of the grain aphid, Sitobion avenae (Fab.) (Homoptera, Aphididae). Chemoecology 6,43 -46.[CrossRef]
Lösel, P. M., Lindemann, M., Scherkenbeck, J., Maier, J., Engelhard, B., Campbell, C. A. M., Hardie, J., Pickett, J. A., Wadhams, L. J., Elbert, A. et al. (1996). The potential of semiochemicals for control of Phorodon humuli (Homoptera: Aphididae). Pestic. Sci. 48,293 -303.[CrossRef]
Marsh, D. (1972). Sex pheromone in an aphid, Megoura viciae. Nat. New Biol. 238, 31-32.[CrossRef]
Marsh, D. (1975). Responses of male aphids to the female sex pheromone in Megoura viciae Buckton. J. Entomol. A 50,43 -64.
Mazor, M. and Dunkelblum, E. (2005). Circadian rhythms of sexual behavior and pheromone titers of two closely related moth species Autographa gamma and Cornutiplusia circumflexa. J. Chem. Ecol. 31,2153 -2168.[CrossRef][Medline]
McNeil, J. N. (1991). Behavioural ecology of pheromone mediated communication in moths and its importance in the use of pheromone traps. Annu. Rev. Entomol. 36,407 -430.[CrossRef]
Petterson, J. (1970a). Studies on Rhopalosiphum padi (L.). I. Laboratory studies on olfactometric responses to the winter host Prunus padus L. Lantbrukshogsk Ann. 36,381 -399.
Petterson, J. (1970b). An aphid sex attractant. 1. Biological studies. Entomol. Scand. 1, 63-73.
Petterson, J. (1971). An aphid sex attractant. II. Histological, ethological and comparative studies. Entomol. Scand. 2,81 -93.
Pickett, J. A., Wadhams, L. J. and Woodcock, C. M. (1992). The chemical ecology of aphids. Annu. Rev. Entomol. 37,67 -90.[CrossRef]
Powell, W., Hardie, J., Hick, A. J., Holler, C., Mann, J., Merritt, L., Nottingham, S. F., Wadhams, L. J., Witthinrich, J. and Wright, A. F. (1993). Responses of the parasitoid Praon volucre (Hymenoptera, Braconidae) to aphid sex-pheromone lures in cereal fields in Autumn – implications for parasitoid manipulation. Eur. J. Entomol. 90,435 -438.
Powell, W., Pennacchio, L. F., Poppy, G. M. and Tremblay, E. (1998). Strategies involved in the location of hosts by the parasitoid Aphidius ervi Haliday (Hymenoptera: Braconidae: Aphidiinae). Biol. Control 11,104 -112.[CrossRef]
Rafaeli, A. and Jurenka, R. A. (2003). PBAN regulation of pheromone biosynthesis in female moths. In Insect Pheromone Biochemistry and Molecular Biology (ed. G. J. Blomquist and R. G. Vogt), pp. 107-136. London: Elsevier Academic Press.
Rosén, W. Q. (2002). Endogenous control of circadian rhythms of pheromone production in the turnip moth, Agrotis segetum. Arch. Insect Biochem. Physiol. 50, 21-30.[CrossRef][Medline]
Silvegren, G., Löfstedt, C. and Rosén, W. Q. (2005). Circadian mating activity and effect of pheromone pre-exposure on pheromone response rhythms in the moth Spodoptera littoralis. J. Insect Physiol. 51,277 -286.[CrossRef][Medline]
Steffan, A. W. (1990). Courtship behaviour and possible pheromone spread by hindleg raising in sexual females of aphids (Homoptera, Aphidinea). Entomol. Gen. 15, 33-49.
Stewart-Jones, A. and Poppy, G. M. (2006). Comparison of glass vessels and plastic bags for enclosing living plant parts for headspace analysis. J. Chem. Ecol. 32,845 -864.[CrossRef][Medline]
Tang, J. D., Charlton, R. E., Jurenka, R. A., Wolf, W. A.,
Phelan, P. L., Sreng, L. and Roelofs, W. L. (1989).
Regulation of pheromone biosynthesis by a brain hormone in two moth species
Proc. Natl. Acad. Sci. USA
86,1806
-1810.
Thieme, T. and Dixon, A. F. G. (1996). Mate recognition in the Aphis fabae complex: daily rhythm of release and specificity of sex pheromones. Entomol. Exp. Appl. 79, 85-89.[CrossRef]
Zeng, R. G., Liu, S. K. and Shaefers, G. A. (1992). Morphology and histology of the sex pheromone gland of the strawberry aphid, Chaetosiphon fragaefolii (Cockerell). Acta Entomol. Sin. 35,290 -293.
Zhu, J., Cossé, A. A., Obrycki, J. J., Boo, K. S. and Baker, T. C. (1999). Olfactory reactions of the twelve-spotted lady beetle, Coleomegilla maculata and the green lacewing, Chrysoperla carnea to semiochemicals released from their prey and host plant: electroantennogram and behavioral responses. J. Chem. Ecol. 25,1163 -1177.[CrossRef]
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