Research
Caterpillars of Euphydryas aurinia (Lepidoptera:
Nymphalidae) feeding on Succisa pratensis leaves induce
large foliar emissions of methanol
Blackwell Publishing, Ltd.
Josep Peñuelas1, Iolanda Filella1, Constantí Stefanescu2 and Joan Llusià1
Unitat Ecofisiologia CSIC-CREAF CREAF, Edifici C, Universitat Autònoma de Barcelona, E−08193 Bellaterra, Barcelona, Spain; 2Butterfly Monitoring
1
Scheme, Museu de Granollers de Ciències Naturals, Francesc Macià, 51, E−08400 Granollers, Spain
Summary
Author for correspondence:
Josep Peñuelas
Tel: +34 935812199/935811312
Fax: +34 935814151
Email: [email protected]
Received: 4 March 2005
Accepted: 14 March 2005
• A major new discovery made in the last decade is that plants commonly emit large
amounts and varieties of volatiles after damage inflicted by herbivores, and not
merely from the site of injury. However, analytical methods for measuring herbivoreinduced volatiles do not usually monitor the whole range of these compounds and
are complicated by the transient nature of their formation and by their chemical
instability.
• Here we present the results of using a fast and highly sensitive proton transfer
reaction–mass spectrometry (PTR-MS) technique that allows simultaneous on-line
monitoring of leaf volatiles in the pptv (pmol mol−1) range.
• The resulting on-line mass scans revealed that Euphydryas aurinia caterpillars
feeding on Succisa pratensis leaves induced emissions of huge amounts of methanol
– a biogeochemically active compound and a significant component of the volatile
organic carbon found in the atmosphere – and other immediate, late and systemic
volatile blends (including monoterpenes, sesquiterpenes and lipoxygenase-derived
volatile compounds).
• In addition to influencing neighboring plants, as well as herbivores and their
predators and parasitoids, these large emissions might affect atmospheric chemistry
and physics if they are found to be generalized in other plant species.
Key words: biogenic volatile organic compounds (VOCs), Euphydryas aurinia,
herbivory and leaf wounding, lipoxygenase-derived (LOX) volatiles, methanol,
monoterpenes, sesquiterpenes, Succisa pratensis.
New Phytologist (2005) 167: 851–857
© New Phytologist (2005) doi: 10.1111/j.1469-8137.2005.01459.x
Introduction
Numerous studies carried out over the last decade have shown
that plants release volatiles in response to herbivore attack
(Dicke et al., 1990; Turlings et al., 1990; Paré & Tumlinson,
1999; Llusià & Peñuelas, 2001), both from the exact area of
feeding damage and from undamaged plant parts (Karban
& Baldwin, 1997; Mumm et al., 2003). This induction of
volatiles results in a change in plant odor that may be quantitative (i.e. the same volatiles are released by the undamaged
and damaged plants, although in differing amounts) or
qualitative (i.e. the damaged plant produces components that
www.newphytologist.org
are not emitted by the undamaged plant) (Boland et al., 1999;
Dicke, 1999; Paré & Tumlinson, 1999). Interest in these
herbivore-induced volatile emissions has greatly increased in
recent years given that these emissions influence neighboring
plants, as well as herbivores and their predators and parasitoids
(Turlings et al., 1990; Peñuelas & Llusià, 2001; van Poecke &
Dicke, 2004). However, there is another reason why these
emissions are worth considering closely: they may also considerably affect atmospheric chemistry and physics (Peñuelas
& Llusià, 2003).
The analytical methods for measuring these herbivoreinduced volatile organic compounds (VOCs) do not usually
851
852 Research
monitor their whole range and are complicated by their transient nature after leaf herbivore grazing and by their chemical
instability. Sampling and analysis of plant VOCs by gas
chromatography can be very time-consuming and only recently
has it become generally feasible to follow the dynamics of VOC
emissions caused by most kinds of biotic stresses (Steeghs
et al., 2004). Here we used a relatively recent methodology
using proton transfer reaction–mass spectrometry (PTR-MS)
that allows for the almost simultaneous on-line monitoring of
these leaf volatile compounds in the pptv (pmol mol−1) range.
This technique has emerged as a useful tool that allows us to
study accurately large numbers of different VOCs with a
rapid time response (< 1 s) and with a low detection threshold
(10–100 pptv) (Lindinger et al., 1998; Warneke et al., 2003).
Moreover, the lack of sample treatment in PTR-MS facilitates
the detection of species such as organic acids, peroxides, and
doubly oxygenated species that are otherwise difficult to measure.
This technique is thus a more general detection method than,
for example, gas chromatography–mass spectrometry (GCMS), in which different columns may be required to target
different classes of compounds. However, the identity of the
compound still needs to be confirmed by other methods such
as GC-MS (Steeghs et al., 2004).
We used both PTR-MS and GC-MS to asses the dynamics
and the quantitative significance of herbivore-induced
emissions. We studied the emissions induced by Euphydryas
aurinia larvae (Lepidoptera: Nymphalidae) feeding on Succisa
pratensis (Dipsacaceae), one of the two host plants commonly
used by the butterfly in north-east Spain and the only host
plant used in damp meadows throughout much of its
European range (Wahlberg et al., 2002). We analysed the
compounds emitted by E. aurinia caterpillars immediately
after leaf attack (first hour) to check for the presence of a
rapid-induced response and after 1 d of continuous herbivorous
attack to check for the presence of a more delayed induced
response, as well as those emitted by leaves not attacked on
attacked plants (systemic response). We also monitored the
compounds emitted by the caterpillars.
Materials and Methods
System studied: caterpillars, plants and experimental
design
In spring females of E. aurinia Rottemburg lay large batches
of about 200–300 eggs on the underside of the leaves of S.
pratensis Moench. Larvae hatch in 3 wk and spin a large silken
web around the leaves, within which they feed for c. 3 wk.
Immediately after their third molt, larvae enter diapause in a
winter web at the base of the plant and do not resume feeding
until early next spring.
The experiments were carried out with 60 larvae from a
single egg-cluster and six S. pratensis plants from a single population. The plants were chosen randomly and were transplanted
New Phytologist (2005) 167: 851– 857
Fig. 1 Time course dynamics of the lipoxygenase-derived (LOX)
volatiles (m83 + m85 + m97 + m101 + m115 + m143) and
monoterpene emission rates of Succisa pratensis leaves before and
after Euphydryas aurinia caterpillars started feeding on them.
with their roots intact in Mediterranean-like environmental
conditions in a glasshouse until the experiment started 3 d
later. As soon as they hatched, the larvae were fed with fresh
cut leaves of a wild S. pratensis plant and, after molting into
their second instar, they were divided into three groups of
20 and each group was placed on a leaf of a different potted
S. pratensis plant. The other three potted S. pratensis plants
(i.e. those with no larvae) were used as controls.
We measured the VOC emission rates and the net photosynthetic rates of leaves from unattacked S. pratensis plants, of
attacked leaves (20 larvae per leaf, Fig. 1) and of unattacked
leaves from plants with an attacked leaf (systemic response).
All these gas exchange measurements were conducted before
the attack, immediately after, and 1 d later. After these foliar
gas exchange rate measurements had been conducted, the
caterpillars were removed from the leaves and their own VOC
emissions were also measured.
In all the measurements leaves were clamped in a 90 cm3
PLC-2 ADC cuvette connected to an IRGA porometer (LCA4, ADC; Hoddeson, Hertfordshire, UK). The VOC-free zero
air was fluxed into the cuvette and the air coming out of the
cuvette flowed through a T-system to either a PTR-MS system
or to an absorption glass tube and was analysed by GC-MS.
Measurements of CO2 and H2O exchange and VOC sampling
www.newphytologist.org © New Phytologist (2005)
Research
and analysis were thus conducted simultaneously. Because the
flow through the cuvette was higher than the flow needed in
the PTR-MS, part of the flow was channeled out of the laboratory through an overflow outlet. The cuvette was lined with
Teflon and only Teflon tubing, connectors, and valves were
used in order to reduce the memory effects caused by surface
interactions in the system. The gas exchange measurements
were also conducted with the empty cuvette as an additional
control. A lamp suspended 20 cm above the cuvette provided
photosynthetic photon fluence rate (PPFR) of 700 µmol m−2
s−1. Leaf temperature was kept at 25°C by air conditioning.
All these leaves were afterwards harvested to measure their surface area in a Li-Cor 3100 Area Meter (Li-Cor Inc., Lincoln,
NE, USA), and to calculate their dry mass (by drying at 60°C
until constant weight was reached).
Analysis of VOCs by PTR-MS and GC–PTR-MS
Part of the air exiting the leaf cuvette flowed through a Tsystem to the PTR-MS inlet. The PTR-MS apparatus was a
highly sensitive device (PTR-MS-FTD hs; Ionicon Analytik,
Innsbruck, Austria) consisting of three parts: the ion source,
where ions are produced by a hollow cathode discharge using
water vapor as the molecular source of ions; the drift tube,
where proton transfer reactions to the trace constituents in
the air occur (VOCs with a higher proton affinity than that
of water (166.5 kcal mol−1), including most unsaturated and
almost all oxygenated hydrocarbons, undergo a proton-transfer
reaction with H3O+); and the ion detector, which provides
sensitive detection of the mass selected ions that are
characteristic of the molecules of interest. Both PTR-MS and
its use in VOC analysis has been described in detail elsewhere
(Lindinger et al., 1998; Fall et al., 1999). Here, the PTR-MS
drift tube was operated at 2.1 mbar and 40°C, with a drift
field of 600 V cm−1. The parent ion signal was maintained
at c. 2 × 106 counts s−1 during the measurements. We conducted
scans of all masses between 21 and 205 to determine which
masses changed emissions during the different periods after
different herbivore attack conditions.
For each of the measurement periods in unattacked and
attacked leaves of attacked plants, as well as in the control
plants and caterpillar emissions, we also sampled VOCs in
carbon trap adsorption tubes and conducted GC-MS analyses
in order to confirm VOC identities by an alternative method.
Part of the air exiting the chamber flowed through a T-system
into a glass tube (11.5 cm long and 0.4 cm internal diameter)
manually filled with terpene adsorbents Carbotrap C (300 mg),
Carbotrap B (200 mg), and Carbosieve S-III (125 mg) (Supelco,
Bellefonte, PA, USA) separated by plugs of quartz wool and
treated as described by Peñuelas et al. (2005). After VOC
sampling, the adsorbent tubes were stored at −30°C until
analysis (within 24–48 h). The VOC analyses were conducted
in a GC-MS (HP59822B; Hewlett Packard, Palo Alto, CA,
USA), as described by Peñuelas et al. (2005).
© New Phytologist (2005) www.newphytologist.org
Statistical analyses
We used t-tests and one-way  and post hoc tests (Statistica;
StatSoft Inc., Tulsa, OK, USA) to compare the emissions from
leaves before the attack with (1) emissions from the same leaves
immediately after the attack, (2) emissions from the same
leaves 24 h after the attack, (3) emissions from unattacked
leaves on attacked plants, and (4) emissions from unattacked
leaves on control plants. There were three plants per group.
A t-test was also used to compare VOC emissions from the
caterpillars alone with the empty chamber.
Results and Discussion
Volatiles emitted de novo or in greater amounts
Time-course The PTR-MS technique and the verification
with GC revealed that a blend of 13 compounds was
consistently emitted de novo or in greater amounts by leaves
immediately after attack by E. aurinia caterpillars: acetaldehyde,
ethanol, hexanals, hexenals, hexenols, mass 88, hexadienal,
mass 116, monoterpenes, hexenyl acetates, mass 163, mass
172 and the sesquiterpene α-caryophyllene (Table 1). The
time-course dynamics showed that the emission of these
volatiles started to increase after the caterpillar attack (see
examples in Fig. 1).
Another blend of 15 compounds, was observed 1 d after
the attack started: methanol, mass 58, butanone, hexenols,
hexanals, hexanols, hexyl acetates, mass 102, heptanals, mass
124, mass 127, monoterpenes, mass 139, mass 175, and the
sesquiterpene α-caryophyllene. Finally, a blend of four of
these compounds (mass 58, butanone, mass 127, monoterpenes, and hexenyl acetates), was also found to be emitted
systemically (i.e. emitted by unattacked leaves of attacked
plants; Table 1). Increased emissions were not found either
when measuring caterpillars alone or with no leaves in the
gas exchange chamber, or when measuring the unattacked
control plants (data not shown).
Emission rates When studied 24 h after the start of the
caterpillar feeding (Fig. 2), attacked leaves emitted up to
20 nmol m−2 s−1 of methanol more than unattacked leaves.
Emissions from unattacked leaves (3.6 nmol m−2 s−1) were
within the range of methanol foliar emissions reported in the
available literature in data sets for other species (Galbally &
Kirstine, 2002); emissions of attacked leaves were within the
maximum upper range of fluxes described after grass-cutting
for hay (Karl et al., 2001).
This large increase in methanol emissions accounted for
almost 0.5% of the net photosynthetic rates in S. pratensis
leaves, which were, on average, 4.8 µmol m−2 s−1 before the
caterpillar attack (Table 2). This 0.5% is higher than the range
found to date for the ratio of methanol emission to net photosynthetic rates for various types of higher plants (between 0.01%
New Phytologist (2005) 167: 851–857
853
854 Research
Table 1 Blend of volatile compounds emitted by Succisa pratensis leaves at higher rates after feeding by Euphydryas aurinia larvae
m33 (methanol)
m45 (acetaldehyde)
m51 (methanol-water cluster)
m58 (isotope of hexenal)
m65 (ethanol-water cluster)
m73 (butanone)
m83 (hexenol + hexanal)
m85 (hexanol, hexyl acetate)
m88
m97 (2E, 4E hexadienal)
m101 (hexenal)
m102 (isotope of hexenal)
m115 (heptanal)
m116
m124
m127
m137 (monoterpenes)
m139
m143 (hexenyl acetates)
m163
m172
m175
M205 (sesquiterpenes)
Leaf before
any feeding
(pmol m−2 s−1)
Attacked leaf
(at first hour)
(pmol m−2 s−1)
Attacked leaf
(at 24 h)
(pmol m−2 s−1)
Unattacked leaf
(at 24 h)
(pmol m−2 s−1)
3636.49 (1507.75)
34.45 (2.79)
30.08 (12.50)
−7.83 (4.94)
−1.42 (1.65)
−27.98 (16.18)
6.68 (6.52)
1.15 (1.75)
−2.35 (2.19)
−8.48 (6.68)
−4.90 (6.80)
−0.65 (3.37)
−4.78 (4.08)
−0.24 (2.95)
6.84 (0.56)
−2.77 (5.28)
1.36 (2.84)
−2.51 (3.11)
−1.51 (4.13)
2.62 (3.37)
0.0
1.34 (1.69)
− 0.22 (1.34)
7405.15 (2507.80)
62.98 (2.95)
76.23 (20.37)
5.13 (2.03)
3.53 (0.36)
−13.82 (47.98)
72.31+ (21.08)
9.82 (13.44)
4.23 (1.35)
8.84 (2.43)
9.18 (4.14)
0.20 (2.56)
4.93 (3.69)
5.41 (1.54)
6.34 (2.92)
−0.61 (3.25)
14.07+ (3.13)
−1.97 (7.64)
7.01 (1.54)
6.38 (1.00)
1.33 (0.42)
0.89 (1.83)
2.94+ (0.71)
24097.93 (1738.69)
51.31 (16.96)
208.32 (40.99)
18.65 (5.04)
1.64 (2.77)
34.91 (7.92)
98.30+ (32.61)
12.53 (1.71)
3.47 (5.01)
−8.80 (6.75)
15.14 (16.96)
9.93 (2.18)
7.60+ (3.91)
5.12 (6.23)
3.09 (0.62)
2.47 (0.24)
14.86+ (2.40)
1.39 (0.55)
7.81 (4.22)
3.72 (6.92)
1.29 (0.89)
2.96 (0.35)
5.94+ (2.06)
3765.14 (2795.81)
50.71 (34.09)
32.86 (35.95)
13.77 (4.66)
− 0.12 (0.34)
27.64 (12.80)
24.42 (27.34)
12.83 (5.72)
0.68 (2.88)
−1.29 (4.84)
7.78 (9.79)
0.97 (2.73)
4.53 (6.45)
6.55 (6.01)
7.41 (3.95)
7.03 (3.11)
11.19 (4.93)
−1.59 (3.49)
11.32 (1.23)
2.32 (8.85)
0.76 (0.95)
0.06 (1.85)
0.12 (1.08)
Data are means with SEM in parenthesis; n = 3. Emission rates significantly different from control (before any feeding) (P < 0.05, +P < 0.1) are
in bold type. Only those masses with emission rates higher than 1 pmol m−2 s−1 are given.
Table 2 The CO2 and H2O exchange induced by leaves in the measuring chamber
CO2 (µmol m−2 s−1)
H2O (mmol m−2 s−1)
Leaf before
any feeding
Attacked leaf +
caterpillars
(At first hour)
Attacked leaf +
caterpillars
(At 24 h)
Unattacked leaf of
attacked plant (at 24 h)
4.81 ± 0.29a
1.46 ± 0.08
4.99 ± 0.49a
1.44 ± 0.07
3.54 ± 0.25b
1.06 ± 0.23
3.39 ± 0.16b
1.54 ± 0.19
The CO2 and H2O exchange explained by caterpillars alone has been taken into account in the calculations for attacked leaves. Different letters
after values indicate significant differences (P < 0.05).
and 0.24%, Galbally & Kirstine, 2002). This percentage
was highest in attacked leaves and in the unattacked leaves of
attacked plants, given that a systemic decrease in net photosynthetic rates of unattacked leaves on attacked plants was
detected (Table 2), in agreement with previous studies showing
that caterpillar feeding has adverse effects on photosynthesis
of leaf areas beyond the leaf areas in which caterpillars removed
tissue (Zangerl et al., 2002). However, other studies have found
compensatory increases of photosynthetic rates (Thomson
et al., 2003) or no effects (Aldea et al., 2005). Therefore, more
studies are warranted to elucidate the indirect effects of insect
herbivory on leaf CO2 and water exchange.
All the other immediate, late, and systemic increases in
absolute volatile emission rates 24 h after the attack were
between two and four orders of magnitude lower than those
New Phytologist (2005) 167: 851– 857
of methanol, although most of them increased several-fold
relative to unattacked plants. Lipoxygenase-derived (LOX)
volatile compounds such as hexenols, hexanals and hexenyl
acetate, which are primarily responsible for the odor of cut
grass, were also very responsive to E. aurinia feeding. They
increased their emission rates by c. 130 pmol m−2 s−1 (Fig. 2),
which is in the range described for other species exposed to
high ozone concentrations, pathogen attacks, or wounding
(Heiden et al., 2003). Grazed leaves also increased by c. 15-fold
their monoterpene emission fluxes (up to 75 pmol m−2 s−1)
compared with leaves with no herbivory (Fig. 2), that is, a
similar increase to that which has been described for the
needles of Douglas firs and ponderosa pines subjected to
mechanical wounding simulating herbivore-damage (Litvak
et al., 1999).
www.newphytologist.org © New Phytologist (2005)
Research
Fig. 2 Herbivore-induced emissions of foliar volatiles. The figures
show huge Succisa pratensis foliar emissions of methanol (a),
significant increases of lipoxygenase-derived (LOX) volatile
compounds such as hexenols, hexanals, and hexenyl acetate (b) and
monoterpenes (c) immediately after attack by Euphydryas aurinia
caterpillars, after 1 d of continuous attack, and systemically (i.e.
emitted by unattacked leaves of attacked plants). +, P < 0.10; *,
P < 0.05; **, P < 0.01 (Student t-test comparison with control leaves
from plants before the attack).
Biological functions and possible atmospheric effects
Most of these compounds emitted de novo or in greater
amounts after caterpillar attack are commonly emitted by
many plants as a response to both biotic (herbivory, pathogen
attack) and abiotic (ozone, wounding) stressors (Turlings
© New Phytologist (2005) www.newphytologist.org
et al., 1990; Fall et al., 1999; Peñuelas & Llusià, 2001;
van Poecke & Dicke, 2004), a fact that implies that these
emissions occur as a general response in plants to stress
and have connected metabolic pathways. In addition to an
immediate and direct defensive function as ways of repelling
or intoxicating herbivores and pathogens, they are also
involved in plant-to-plant communication and act as easily
detectable and distinctive chemical cues for the predators and
parasitoids of the herbivores feeding on the plants (i.e. they
serve as indirect defenses) (Turlings et al., 1990; Peñuelas &
Llusià, 2001; van Poecke & Dicke, 2004).
The most significant result of this study was that the
attacked leaves emitted far greater amounts of methanol
(20 nmol m−2 s−1) than unattacked leaves (Fig. 2). There is
ample evidence to suggest that methanol is produced from
pectin demethylation in the cell walls (Gaffe et al., 1994; Fall
& Benson, 1996; Galbally & Kirstine, 2002) and since this
process occurs in the apoplast, methanol proves to be a common constituent of the transpiration stream in plants (Fall &
Benson, 1996), which is consistent with an expected release
from cut leaves. Our results show that methanol emission
started to slightly but not significantly increase after the
caterpillars started to feed, and that the greatest increase in
emissions occurred most intensively 1 d after the attack started
(Fig. 2). Unfortunately, because we did not analyse each leaf
continuously for a whole day, we cannot know whether a peak
of methanol or any other volatile emission occurred between
the first hour and 24 h after feeding started.
The lifespan of methanol is several days in the boundary
layer and a few weeks in the upper troposphere (i.e. much
longer than the chemical lifetime of other common biogenic
VOCs such as isoprene or monoterpenes, which are far more
reactive). The up to eight-times higher foliar methanol
emissions described here, together with their relatively long
lifespan, might have considerable impact on tropospheric
oxidants if these increased emissions were found to be case in
other plant species. For example, despite the poor reactivity
and large solubility of methanol in water (properties that
make deposition more likely to occur in this compound than
in most reactive VOCs), if increased methanol emissions were
generalized in response to herbivores, they might significantly
increase the 1–2% of O3 tropospheric concentrations, on
average, accounted for by methanol (Tie et al., 2003).
The increased emissions of the other volatiles reported
here, such as those of the lipoxygenase-derived volatiles and
the monoterpenes, might also significantly affect local oxidative tropospheric chemistry and play an important role in
perturbing local ozone dynamics (Litvak et al., 1999; Heiden
et al., 2003), if they were generalized and occurred in other
more abundant plants. Furthermore, LOX volatiles might
actually be emitted at higher rates a few hours after the stress
(Heiden et al., 2003; Wildt et al., 2003). This possibility was
hinted at by the continuously increasing emission rates after
the caterpillar attack (Fig. 1), but could not be monitored by
New Phytologist (2005) 167: 851–857
855
856 Research
our experimental set-up, which only monitored the first hour
and 24 h after the caterpillars started feeding.
In any case, to actually have a significant effect on the atmosphere, the responses found in S. pratensis should be generalizable
to many other species. Further studies are thus warranted by the
results reported here. Moreover, the actual amounts of volatiles
emitted by this (and other possible) species, and therefore
their overall atmospheric effects, depend on the biological
performance of the herbivore populations, which vary from
season to season. For example, E. aurinia populations experience
dramatic yearly changes of abundance, mainly as a result of
variations in the rates of parasitic attacks by specialist braconid
parasitoids Cotesia melitaerum and Cotesia bignellii (Ford &
Ford, 1930; Kankare et al., 2005). Thus, following a year of
low parasitism, larval densities can attain very high levels and
inflict serious damage to S. pratensis populations. However,
the impact of VOC emissions by S. pratensis as a result of
E. aurinia attack seems to be of limited scope. Thus, although
noticeable defoliation might occur in years of high larval
density, this happens at a reduced scale, for which reason E.
aurinia cannot be considered as an outbreaking insect species
(Berryman, 1987; Hunter, 1991). Euphydryas aurinia populations depending on this host are typically structured in small
patches of habitat, usually of less than 2 ha (Warren, 1994;
Wahlberg et al., 2002; own pers. obs.). Although in these
patches S. pratensis densities can be quite high (normally
about 100 plants ha−1, with maxima recorded at about 700
plants ha−1; Wahlberg et al., 2002), insect population sizes
remain rather small. For example, Warren (1994) estimated
the peak of the largest English population in a very good
year at about 30 000 adult butterflies, but almost invariably
populations are far smaller (in the order of 100 to 1000 adult
butterflies).
Despite the necessary caution implicit in a study of such
limited scope dealing with only one plant species and one
insect herbivore species, the results reported here show that it
is worth considering herbivore-induced emissions in our continuing efforts to quantify methanol (Galbally & Kirstine,
2002) and other VOC emissions (Guenther, 2002) in the Earth’s
major ecosystems. Thus, these studies should be repeated for
other more widely spread plant species and emissions generated
by plagues must also be fully studied.
Acknowledgements
This research was partly supported by the European Commission contract MC-RTN-CT-2003–504720 (ISONET) and
by Spanish MCYT grants REN2003-04871 and CGL200401402/BOS.
References
Aldea M, Hamilton JG, Resti JP, Zangerl AR, Berenbaum MR,
DeLucia EH. 2005. Indirect effects of insect herbivory on leaf gas
exchange in soybean. Plant, Cell & Environment 28: 402–411.
New Phytologist (2005) 167: 851– 857
Berryman AA. 1987. The theory and classification of outbreaks. In:
Barbosa P, Schultz JC, eds. Insect outbreaks. San Diego, CA, USA:
Academic Press, 3–30.
Boland W, Koch T, Krumm T, Piel J, Jux A. 1999. Induced biosynthesis
of insect semiochemicals in plants. In: Chadwick DJ, Goode JA, eds.
Insect–plant interactions and induced plant defence. Chichester, UK /New
York, NY, USA: Wiley, 110–131.
Dicke M. 1999. Evolution of induced indirect defence of plants. In: Tollrian R,
Harvell CD, eds. The ecology and evolution of inducible defenses. Princeton
NJ, USA: Princeton University Press, 62–88.
Dicke M, van Beek TA, Posthumus MA, Ben Dom N, van Bokhoven H,
de Groot Æ. 1990. Isolation and identification of volatile kairomone that
affects acarine predator–prey interactions: involvement of host plant in its
production. Journal of Chemical Ecology 16: 381–396.
Fall R, Benson AA. 1996. Leaf methanol – the simplest natural product from
plants. Trends in Plant Science 1: 296–301.
Fall R, Karl T, Hansel A, Jordan A, Lindinger W. 1999. Volatile organic
compounds emitted after leaf wounding: on-line analysis by protontransfer reaction mass spectrometry. Journal of Geophysical Research 104:
15963–15974.
Ford HD, Ford EB. 1930. Fluctuation in numbers and its influence on
variation in Melithaea aurinia, Rott. (Lepidoptera). Transactions of the
Royal Entomological Society of London 78: 345–351.
Gaffe J, Tieman DM, Handa AK. 1994. Pectin methylesterase isoforms
in tomato Lycopersicum esculentum tissue. Plant Physiology 105: 199–
203.
Galbally IE, Kirstine WV. 2002. The production of methanol by flowering
plants and the global cycle of methanol. Journal of Atmospheric Chemistry
43: 195–229.
Guenther A. 2002. The contribution of reactive carbon emissions from
vegetation to the carbon balance of terrestrial ecosystems. Chemosphere 49:
837–844.
Heiden AC, Kobel K, Langebartels C, Schuh-Thomas G, Wildt J. 2003.
Emissions of oxygenated volatile organic compounds from plants. Part I:
Emissions from lipoxygenase activity. Journal of Atmospheric Chemistry 45:
143–172.
Hunter AF. 1991. Traits that distinguish outbreaking and nonoutbreaking
Macrolepidoptera feeding on northern hardwood trees. Oikos 60: 275–
282.
Kankare M, Stefanescu C, van Nouhuys S, Shaw MR. 2005. Host
specialization by Cotesia wasps. (Hymenoptera: Braconidae) parasitising
species-rich Melitaeini (Lepidoptera: Nymphalidae) communities in
north-eastern Spain. Biological Journal of the Linnean Society. (In press.)
Karban R, Baldwin IT. 1997. Induced responses to herbivory. Chicago, IL,
USA: University of Chicago Press.
Karl T, Guenther A, Jordan A, Fall R, Lindinger W. 2001. Eddy covariance
measurement of biogenic oxygenated VOC emissions from hay harvesting.
Atmospheric Environment 35: 491–495.
Lindinger W, Hansel A, Jordan A. 1998. On-line monitoring of volatile
organic compounds at pptv levels by means of proton-transfer-reaction
mass spectrometry (PTR-MS). Medical applications, food control and
environmental research. International Journal of Mass Spectrometry. Ion
Processes 173: 191–241.
Litvak ME, Madronich S, Monson RK. 1999. The potential impact of
herbivore-induced monoterpene emissions from coniferous forests on
local tropospheric chemistry dynamics. Ecological Applications 9: 1147–
1159.
Llusià J, Peñuelas J. 2001. Emission of volatile organic compounds by apple
trees in response to spider mite attack and attraction of predatory mites.
Experimental and Applied Acarology 25: 65–77.
Mumm R, Schrank K, Wegener R, Schulz S, Hilker M. 2003. Chemical
analysis of volatiles emitted by Pinus sylvestris after induction by insect
oviposition. Journal of Chemical Ecology 29: 1235–1252.
Paré PW, Tumlinson JH. 1999. Plant volatiles as a defense against insect
herbivores. Plant Physiology 121: 325–331.
www.newphytologist.org © New Phytologist (2005)
Research
Peñuelas J, Llusià J. 2001. The complexity of factors driving volatile organic
compound emissions by plants. Biologia Plantarum 44: 481– 487.
Peñuelas J, Llusià J. 2003. BVOCs: plant defense against climate warming?
Trends in Plant Science 8: 105–109.
Peñuelas J, Llusià J, Asensio D, Munné-Bosch S. 2005. Linking isoprene
with plant thermotolerance, antioxidants and monoterpene emissions.
Plant, Cell & Environment 28: 278 –286.
van Poecke RMP, Dicke M. 2004. Indirect defence of plants against
herbivores: using Arabidopsis thaliana as a model plant. Plant Biology 6:
387–401.
Steeghs M, Bais HP, De Gouw J, Goldan P, Kuster W, Northway M,
Fall R, Vivanco JM. 2004. Proton-transfer-reaction mass spectrometry
as a new tool for real time analysis of root-secreted volatile organic
compounds in Arabidopsis. Plant Physiology 135: 47–58.
Thomson VP, Cunningham SA, Ball MC, Nicotra AB. 2003.
Compensation for herbivory by Cucumis sativus through
increased photosynthetic capacity and efficiency. Oecologia 134:
167–175.
Tie X, Guenther A, Holland E. 2003. Biogenic methanol and its impacts
on tropospheric oxidants. Geophysical Research Letters 17: 1881.
Turlings TCJ, Tumlinson JH, Lewis WJ. 1990. Exploitation of
herbivore-induced plant odors by host-seeking parasitic wasps. Science
250: 1251–1253.
Wahlberg N, Klemetti T, Hanski I. 2002. Dynamic populations in a
dynamic landscape: the metapopulation structure of the marsh fritillary
butterfly. Ecography 25: 224–232.
Warneke C, de Gouw J, Kuster WC, Goldan GD, Fall R. 2003. Validation
of atmospheric VOC measurements by proton-transfer-reaction mass
spectrometry using a gas-chromatographic preseparation method.
Environmental Science and Technology 37: 2494–2501.
Warren MS. 1994. The UK status and suspected metapopulation structure
of a threatened European butterfly, the marsh fritillary Eurodryas aurinia.
Biological Conservation 67: 239–249.
Wildt J, Kobel K, Schuh-Thomas G, Heiden AC. 2003. Emissions of
oxygenated volatile organic compounds from plants – part II: emissions
of saturated aldehydes. Journal of Atmospheric Chemistry 45: 173–196.
Zangerl AR, Hamilton JG, Miller TJ, Crofts AR, Oxborough K,
Berenbaum MR, DeLucia EH. 2002. Impact of folivory on
photosynthesis is greater than the sum of its holes. Proceedings of the
National Academy of Sciences, USA 99: 1088–1091.
About New Phytologist
• New Phytologist is owned by a non-profit-making charitable trust dedicated to the promotion of plant science, facilitating projects
from symposia to open access for our Tansley reviews. Complete information is available at www.newphytologist.org.
• Regular papers, Letters, Research reviews, Rapid reports and both Modelling/Theory and Methods papers are encouraged.
We are committed to rapid processing, from online submission through to publication ‘as-ready’ via OnlineEarly – the 2004 average
submission to decision time was just 30 days. Online-only colour is free, and essential print colour costs will be met if necessary.
We also provide 25 offprints as well as a PDF for each article.
• For online summaries and ToC alerts, go to the website and click on ‘Journal online’. You can take out a personal subscription to
the journal for a fraction of the institutional price. Rates start at £109 in Europe/$202 in the USA & Canada for the online edition
(click on ‘Subscribe’ at the website).
• If you have any questions, do get in touch with Central Office ([email protected]; tel +44 1524 594691) or, for a local
contact in North America, the US Office ([email protected]; tel +1 865 576 5261).
© New Phytologist (2005) www.newphytologist.org
New Phytologist (2005) 167: 851–857
857