Dynamics of the enhanced emissions of monoterpenes and methyl salicylate, and decreased uptake of formaldehyde, by Quercus ilex leaves after application of jasmonic acid

Authors

  • Iolanda Filella,

    1. Unitat Ecofisiologia CSIC-CEAB-CREAF, Center for Ecological Research and Forestry Applications (CREAF), Universitat Autònoma de Barcelona, 08193 Bellaterra (Barcelona), Spain
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  • Josep Peñuelas,

    1. Unitat Ecofisiologia CSIC-CEAB-CREAF, Center for Ecological Research and Forestry Applications (CREAF), Universitat Autònoma de Barcelona, 08193 Bellaterra (Barcelona), Spain
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  • Joan Llusià

    1. Unitat Ecofisiologia CSIC-CEAB-CREAF, Center for Ecological Research and Forestry Applications (CREAF), Universitat Autònoma de Barcelona, 08193 Bellaterra (Barcelona), Spain
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Author for correspondence: Iolanda Filella Tel: +34 93 5814670 Fax: +34 93 5814151 Email: i.filella@creaf.uab.es

Summary

  • • Jasmonic acid (JA) is a signalling compound with a key role in both stress and development in plants, and is reported to elicit the emission of volatile organic compounds (VOCs). Here we studied the dynamics of such emissions and the linkage with photosynthetic rates and stomatal conductance.
  • • We sprayed JA on leaves of the Mediterranean tree species Quercus ilex and measured the photosynthetic rates, stomatal conductances, and emissions and uptake of VOCs using proton transfer reaction mass spectrometry and gas chromatography after a dark–light transition.
  • • Jasmonic acid treatment delayed the induction of photosynthesis and stomatal conductance by approx. 20 min, and decreased them 24 h after spraying. Indications were found of both stomatal and nonstomatal limitations of photosynthesis. Monoterpene emissions were enhanced (20–30%) after JA spraying. Jasmonic acid also increased methyl salicylate (MeSa) emissions (more than twofold) 1 h after treatment, although after 24 h this effect had disappeared. Formaldehyde foliar uptake decreased significantly 24 h after JA treatment.
  • • Both biotic and abiotic stresses can thus affect plant VOC emissions through their strong impact on JA levels. Jasmonic acid-mediated increases in monoterpene and MeSa emissions might have a protective role when confronting biotic and abiotic stresses.

Introduction

Jasmonic acid (JA) is a naturally occurring signalling compound found in higher plants that plays a key role in both stress and development (Creelman & Mullet, 1997; Gfeller & Farmer, 2004). Jasmonic acid and its methyl ester, methyl jasmonate (MeJA), collectively referred as jasmonates, constitute a major signalling mechanism in stress-induced gene expression (Wasternack & Parthier, 1997). Endogenous levels of JA increase in plants in response to a wide range of biotic and abiotic stresses such as water deficit (Creelman & Mullet, 1997; Gao et al., 2004); salinity (Mopper et al., 2004); low temperatures (Kondo et al., 2004); and ozone (Koch et al., 2000; Rao et al., 2000), and it has been suggested that jasmonates could mediate defensive responses to these environmental stresses (Wilen et al., 1994; Tsonev et al., 1998; Mackerness et al., 1999). Although natural seasonal cycles and/or environmental stress induce more significant changes in JA than herbivory (Mopper et al., 2004), JA is best known for its role in response to herbivory and mechanical wounding (Baldwin, 1998; Reymond et al., 2004).

Jasmonates have also been associated with the increased production of volatile organic compounds (Hopke et al., 1994; Boland et al., 1995; Rodriguez-Saona et al., 2001; van Poecke & Dicke, 2004). Increases in the biosynthesis of isoprene from recently fixed carbon caused by exogenous JA have been also found (Ferrieri et al., 2005). Application of exogenous JA has been used to stimulate induced plant resistance without damaging the plant (Thaler et al., 1996), particularly in studies of herbivory: JA induces systemic accumulation of compounds with antiherbivore properties and elicits volatile emissions that attract the natural enemies of herbivores (Thaler et al., 2001). The application of JA has also been described as inducing the release of volatile sesquiterpenes in Zea mays (Schmelz et al., 2001) and increasing the emission of volatiles in Lima bean (Heil, 2004). The exogenous application of the methyl ester, MeJa, has also been described as triggering a twofold increase in monoterpene and sesquiterpene accumulation in needles and a fivefold increase in total terpene emissions in the foliage of Norway spruce (Martin et al., 2003), as well as dramatic increases in terpenoid emissions in Nicotiana attenuata (Halitschke et al., 2000).

Proton transfer reaction mass spectrometry (PTR-MS) has emerged as a useful tool that permits us to monitor online, almost simultaneously, a large number of different volatile organic compound (VOC) species with a fast time response (< 1 s) and with a low detection limit (parts per trillion by volume). By comparing PTR-MS and GC-PTR-MS measurements, Warneke et al. (2003) showed that PTR-MS accurately measured some VOCs (isoprene, monoterpenes, methanol, acetonitrile, acetaldehyde, acetone, benzene, toluene and higher aromatic VOCs). However, GC-MS is also needed to identify different compounds with the same mass (e.g. the various monoterpenes) that PTR-MS cannot separate.

The holm oak (Quercus ilex L.) is one of the dominant Mediterranean forest species. No specialized storage structures for monoterpenes have been found in its leaves or bark, and emissions appear to be mainly influenced by temperature and light, although water availability is also influential because of the dependence of terpene production on metabolites originating in the photosynthetic processes (Staudt & Seufert, 1995; Loreto et al., 1996; Bertin et al., 1997; Owen et al., 1997; Llusià & Peñuelas, 1999, 2000; Peñuelas & Llusià, 1999). The emissions of VOCs other than terpenoids have also been studied in Q. ilex (Kesselmeier et al., 1997; Holzinger et al., 2000; Peñuelas & Llusià, 2001), although no study that we are aware of has focused on trees’ VOC emissions as responses to JA.

We compared emissions from Q. ilex leaves sprayed with JA (JA-S) with those from control leaves by means of PTR-MS and GC-MS analyses. Our aim was to study the online dynamics of the effect of JA on emissions of terpenes and other VOCs in this common Mediterranean species, which is often subject to stressful conditions associated with high temperatures, high irradiance and low water availability in summer (Peñuelas et al., 1998), and high irradiance and relatively low temperatures in winter (Oliveira & Peñuelas, 2000, 2001). In order to study the role of photosynthesis and conductance in these emissions, we submitted the measured leaves to a dark–light transition during emission monitoring and conducted CO2 response curves of JA-S and control leaves.

Materials and Methods

Plant material and jasmonic acid treatment

We used 2-yr-old Q. ilex plants grown in a nursery (Forestal Catalana, S.A., Breda, Spain) under typical Mediterranean environmental conditions (mean annual average temperature 16°C, mean annual precipitation 600 mm). They were grown in 2-l pots with a substrate composed of peat and sand (2 : 1).

Three plants were sampled. In each of the three plants, two leaves with similar characteristics (age, position and illumination) were measured. One leaf was treated as a control while the other was sprayed with JA (JA-S). In the JA-S leaves the abaxial and adaxial leaf surfaces were sprayed until runoff with a solution of 0.5 mm JA (Sigma, St Louis, MO, USA) (Thaler et al., 2001). When spraying, the sprayed leaf was isolated from the rest with a protecting plastic cuvette that prevented the rest of the plant from being sprayed. The solution was prepared by dissolving JA in acetone and then diluting this mixture in water to 0.5 mm. This dose, well below the level toxic to the plant (Thaler et al., 1996), apparently lies within the range of biologically meaningful concentrations and simulates the level of resistance induced by one fourth-instar Helicoperva zea feeding for 24 h on glasshouse-grown tomato plants (Thaler et al., 1996; Thaler et al., 2001). In the control leaves, the abaxial and adaxial leaf surfaces were sprayed until runoff with water solution which contained the same very small concentration of acetone as the solution used for the JA treatment. We measured gas exchange in control and JA-S leaves subjected consecutively to a sequence of 20 min darkness and 1 h light. After 24 h both JA-S and control leaves were measured again in another dark–light transition until stabilization of photosynthetic and emission rates (usually 30–40 min). Each JA-S leaf was compared with the corresponding control leaf from the same plant to compensate as far as possible for the high interindividual variation in emission rates.

Gas-exchange measurements: CO2, H2O and VOCs

Intact leaves were clamped in a Parkinson leaf cuvette (Std Broad 2.5, PP Systems, Hitchin, UK) and the leaf temperature was maintained at 25°C. A calibrated Ciras-2 IRGA-porometer (PP Systems) was used for determining rates of CO2 and H2O exchange.

Part of the air exiting the leaf cuvette flowed through a T-system to the PTR inlet. The PTR-MS apparatus consists of three parts: the ion source, where ions are produced by a hollow cathode discharge using water vapour as the molecular source of ions; the drift tube, where VOCs with a higher proton affinity than that of water (166.5 kcal mol−1), which include most unsaturated and almost all oxygenated hydrocarbons, undergo a proton-transfer reaction with H3O+; and the ion detector, which provides sensitive detection of mass-selected ions characteristic of the molecules under study. PTR-MS and its use in VOC analysis has been described in detail elsewhere (Lindinger et al., 1998; Fall et al., 1999).

In our experiment 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. 3 × 106 counts per second. With the PTR-MS, we measured all protonated masses that had presented detectable emissions in pretreatment screening scans of the Q. ilex seedlings and all those previously reported in other Q. ilex studies in the literature (Holzinger et al., 2000): M28, M29, M31, M33, M45, M47, M59, M61, M67, M69, M70, M71, M72, M73, M75, M81, M83, M85, M87, M89, M91, M93, M95, M101, M105, M121, M135, M136, M137, M153, M155, M157, M179 and M205.

For each measurement we also sampled the VOCs with carbon-trap adsorption tubes. We also conducted GC-MS analyses to confirm VOC identities and to separate the different monoterpenes. Part of the air exiting the chamber flowed through a T-system to a glass tube (11.5 cm long, 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), all separated by plugs of quartz wool and treated as described by Peñuelas et al. (2005a). After VOC sampling, the adsorbent tubes were stored at −20°C until analysis (within 24–48 h). VOC analyses were conducted by GC-MS (Hewlett Packard HP59822B, Palo Alto, CA, USA) as described by Peñuelas et al. (2005a).

Before taking measurements, we measured the background concentrations of VOCs by sampling the Parkinson cuvette empty, and these data were used to calculate the emission/uptake of every compound.

We conducted an additional study to measure the response of net C assimilation by leaves to their intercellular CO2 concentration (A/Ci curves) (photosynthesis system LCpro+ with a broad head, ADC BioScientific Ltd, Hodesdon, UK) in order to examine possible changes in the maximum velocity of RuBP carboxylation by Rubisco (Vcmax) and the maximum electron transport rate contributing to RuBP regeneration (Jmax) caused by JA-S treatment. The A/Ci curves were conducted in the leaves of plants with the same age, position and orientation as those analysed with the PTR-MS system, and were submitted to the same JA treatment. Net photosynthetic rates were measured over a range of intracellular CO2 levels (50–800 nmol mol−1) at 1000 µmol m−2 s−1 PPFD and 25°C.

Data treatment and statistical analyses

We used t-tests, one-way ANOVA and post hoc tests (statistica, StatSoft Inc., Tulsa, OK, USA) to compare leaf emissions after treatments. Regression and correlation analyses were also conducted for the variables studied. Estimations of Vcmax and Jmax were made by fitting a maximum likelihood regression below and above the inflexion of the A/Ci response using the method described by McMurtrie & Wang (1993). We used the mean values of the measured variables once they reached stability to calculate the ratios between JA-sprayed leaves and control leaves (see Figs 1b, 3b, 4b, 5b).

Figure 1.

Quercus ilex net photosynthetic rates and stomatal conductances in a dark–light (1000 µmol m−2 s−1) transition. (a) Examples of the dynamics in the dark–light transition 1 and 24 h, after jasmonic acid (JA) spraying; (b) ratio of net photosynthetic rates and stomatal conductances after the dark–light transition in JA-sprayed (JA-S) leaves to control leaves of the same plant, 1 and 24 h after JA treatment or absence of treatment (n = 3) (+, P < 0.1; *, P < 0.05, ANOVA).

Figure 3.

Quercus ilex M153 (methyl salicylate) emissions in a dark–light (1000 µmol m−2 s−1) transition. (a) Examples of the dynamics of the emissions in the dark–light transition 1 and 24 h after jasmonic acid (JA) spraying; (b) M153 emissions ratio after the dark–light transition in JA-sprayed (JA-S) leaves to control leaves of the same plant, 1 and 24 h after JA treatment or absence of treatment (n = 3) (**, P < 0.01, ANOVA).

Figure 4.

Quercus ilex M31 (formaldehyde) emissions in a dark–light (1000 µmol m−2 s−1) transition. (a) Examples of the dynamics of the emissions in the dark–light transition 1 and 24 h after jasmonic acid (JA) spraying; (b) M31 emissions ratio after the dark–light transition in JA-sprayed (JA-S) leaves in relation to control leaves of the same plant, 1 and 24 h after JA treatment or absence of treatment (n = 3) (*, P < 0.05, ANOVA).

Results

Decreases in photosynthetic rate and stomatal conductance

A delay in the induction of photosynthesis and stomatal conductance as a response to dark–light transition (Pearcy, 1999) was observed in the JA-S leaves. After a light transition conducted following JA application, JA-S leaves took 20–25 min to start showing positive net photosynthetic rates, whereas this effect occurred almost immediately in control leaves (Fig. 1a). The JA-S leaves took 30 min to start increasing stomatal conductance, whereas control leaves took only 10 min (Fig. 1a). After light exposure, net photosynthetic rates and stomatal conductances decreased by approx. 40% in JA-S leaves with respect to control leaves when the dark–light transition was conducted 24 h after treatment (Fig. 1a,b).

The analyses of A/Ci curves showed a 20% decrease in Vcmax and Jmax 1 h after JA spraying (20.73 ± 6.47 and 20.44 ± 7.15%, respectively). At 24 h after JA spraying only Jmax showed a reduction trend, although differences were not significant (23 ± 14%).

Jasmonic acid-induced emission changes

Application of JA significantly changed the emission of only five of the measured protonated masses (probably corresponding to three VOC species): mass 137 and mass 81 (monoterpenes); mass 67 (possibly a monoterpene fragment ion); mass 153 (MeSa); and mass 31 (formaldehyde).

Enhanced monoterpene emissions  Monoterpene (mass 137 + mass 81) emissions increased from dark to light, as did net photosynthetic rates and stomatal conductances (Fig. 2a,b). As a result they were correlated with photosynthetic rate and stomatal conductance (r = 0.63, P < 0.0001 and r = 0.47, P < 0.0001, respectively). However, monoterpene emission rates from the JA-S leaves were approx. 20% higher than those of the control leaves (Fig. 2a,b) despite their lower photosynthetic rates (Fig. 1). Thus JA treatment had opposite effects on photosynthetic rates and on monoterpene emissions under light conditions, when photosynthesis was active. The monoterpene emissions after 1 h were higher than after 24 h, probably caused by an indirect effect of the treatment: JA spraying wetted the surface of the measured leaf, and wetting increases monoterpene emissions (Schade et al., 1999; personal observation in Q. ilex), perhaps through increased cuticular permeability.

Figure 2.

Quercus ilex monoterpene emissions in a dark–light (1000 µmol m−2 s−1) transition. (a) Examples of the dynamics of the emissions in the dark–light transition 1 and 24 h after jasmonic acid (JA) spraying; (b) monoterpene emissions ratio after the dark–light transition in JA-sprayed (JA-S) leaves in relation to control leaves of the same plant, 1 and 24 h after JA treatment or absence of treatment (n = 3) (**, P < 0.01, ANOVA).

The emission of mass 67 also increased after dark–light transition, and reached significantly higher levels in JA-treated leaves (data not shown). Mass 67 has been described as a fragment ion of monoterpenes (Hewitt et al., 2003; Tani et al., 2003). Holzinger et al. (2000) also observed emissions of mass 67 in Q. ilex and attributed it, with uncertainty, to cyclopentadiene. To check this possible monoterpene origin of M67, we examined the fragmentation pattern of an α-pinene standard under our experimental PTR-MS conditions, and found that M67 was one of the fragments measured. Moreover, there was a strong correlation between M67 and the masses for monoterpenes M137 + M81 (r2 = 0.71, P < 0.0001). Hence it is likely that mass 67 was a fragment of monoterpene. Mass 67 emission was also correlated with photosynthetic rates and stomatal conductances (r = 0.48, P < 0.0001 and r = 0.32, P < 0.0001, respectively), as it also occurred with the other monoterpene fragments.

By using GC-MS, we determined the relative composition of measured monoterpenes emitted from the Q. ilex control and JA-S leaves, 1 and 24 h after treatment. Jasmonic acid treatment altered neither the composition nor the relative abundance of the volatile monoterpenes emitted by Q. ilex plants (Table 1).

Table 1.  Relative composition of measured monoterpenes emitted from Quercus ilex control (C) and jasmonic acid-sprayed (JA-S) leaves, 1 and 24 h after treatment (no significant differences were found between treatments)
 Monoterpene emission composition (%)
1 h24 h
CJA-SCJA-S
  1. Values are means (n = 3) ± SE.

α-pinene25.63 ± 1.4323.49 ± 0.6018.63 ± 6.5723.30 ± 1.33
Camphene 0.00 0.09 ± 0.09 2.25 ± 1.87 0.16 ± 0.16
α-phellandrene 1.95 ± 0.78 7.07 ± 5.84 5.38 ± 5.40 4.12 ± 2.39
β-pinene17.18 ± 0.2316.76 ± 0.79 9.75 ± 1.5611.54 ± 3.44
β-myrcene 3.85 ± 2.86 4.47 ± 3.07 0.25 ± 0.25 3.54 ± 3.55
Limonene50.67 ± 0.3148.13 ± 1.4763.74 ± 11.955.51 ± 6.44
Rt 9.52 0.72 ± 0.720.000.00 1.84 ± 0.35

Fluxes of methyl salicylate  There was an increase in emission rates of MeSa in JA-S leaves from approx. 4 to 8 pmol m−2 s−1 after the dark–light transition (Fig. 3a), although only in the measurements taken 1 h after JA spraying; this increase was not found 24 h after the JA-S treatment (Fig. 3a,b). Control leaves also increased their emission of MeSa after the dark–light transition, although this increase was lower than for JA-S leaves (Fig. 3b) and, as in JA-S leaves, no emission increases were found 24 h after the JA-S treatment (Fig. 3a,b).

Decreases in formaldehyde uptake  The leaf cuvette concentrations of mass 31 (formaldehyde) decreased after the dark–light transition in both control and JA-S leaves, indicating a formaldehyde leaf uptake of approx. 500 pmol m−2 s−1 in light, when stomata were open. This leaf uptake was significantly lower (half the rates) when leaves were treated with JA 1 d after treatment (Fig. 4a,b). Net photosynthetic rates and stomatal conductances were directly correlated with formaldehyde uptake (r = 0.56, P < 0.0001 and r = 0.21, P < 0.01, respectively) during the dark–light transition.

Discussion

Decreases in photosynthetic rate and stomatal conductance

These results show that lower photosynthetic rates and stomatal conductances can be induced by JA treatment. Several photosynthetic parameters appear to be involved: plants treated with exogenous JA have been reported to have lower CO2 fixation and RUBPase activity, but also significant increases in rates of dark respiration and photorespiration; in CO2 compensation point value; in stomatal resistance (Popova & Vaklinova, 1988; Horton, 1991); and in transpiration rates (Herde et al., 1997). MeJA has also been described as a promoter of stomatal closure (Sanz et al., 1993; Creelman & Mullet, 1997; Beltrano et al., 1998; Evans, 2003).

The mechanisms involved in these decreases in net photosynthetic rate and stomatal conductance are not fully known. However, a breakdown in the biosynthesis of Rubisco (Weidhase et al., 1987; Popova & Vaklinova, 1988), inhibition of the Hill reaction activity, and a number of changes in the kinetic characteristics of flash-induced O2 evolution and in the organization of the photosynthetic membranes (Maslenkova et al., 1990; Maslenkova et al., 1995), have all been reported to occur as a result of JA and MeJA treatments. Methyl jasmonate concentration-dependent effects on guard cell K+ channels and K+ fluxes that promote stomatal closure have been demonstrated (Evans, 2003). It has been suggested that the modifications in photosynthetic membrane structure and function caused by mediators of plant stress response such as JA allow the photosynthetic apparatus to adapt to changing environmental conditions (Maslenkova et al., 1995).

In this study, the parallel response of photosynthetic rates and conductance, both immediately after JA treatment and 24 h later, indicates that decreases in rates of photosynthesis appear to be linked mainly to reductions in stomatal conductance, although the A/Ci curve analysis also showed a nonstomatal limitation of photosynthesis caused by JA. The decrease in Vcmax could indicate a loss and/or inactivation of Rubisco, or could be caused by an underestimation of Vmax caused by CO2 transfer conductance (gi) limitations (Ethier & Livingston, 2004). The decrease in Jmax indicates that the decrease in photosynthetic capacity was accompanied by a reduction in the ability to regenerate RuBP. Regeneration of RuBP could be limited either by an inability to supply reductants and ATP from electron transport, or by inactivation and loss of Calvin cycle enzymes (Nogués & Baker, 2000). Regarding the response of stomatal conductance to JA, although little is known about the mechanisms by which jasmonates affect stomatal behaviour, the fairly quick loss of potassium from the cytoplasm of guard cells, as described by Evans (2003) in Vicia faba guard cells exposed to JA, could explain the observed first response of lower stomatal conductances in JA-S leaves. A decrease in the synthesis or activity of Rubisco caused by JA (Maslenkova et al., 1995) would agree with the decrease in Vcmax found in JA-S plants, and could be related to the delay in the response of photosynthetic rates to the dark–light transition in JA-S leaves, as Rubisco regeneration and activity limitations could be the key to this process when the dark period lasts for some minutes (Sassenrath-Cole et al., 1994; Pearcy, 1999). The modifications in photosynthetic membrane structure and function caused by JA (Maslenkova et al., 1995) could explain the day-after decrease in photosynthetic rates.

Whatever the mechanism(s), the results show that JA acts as a stress signal that elicits a plant response, the aim of which is to reduce activity and therefore CO2 and H2O exchange.

Enhanced monoterpene emissions

The relationship between monoterpene emission and photosynthetic rates found during the dark–light transitions agrees with the reported relationship between monoterpene emission and CO2 assimilation in Q. ilex (Staudt & Seufert, 1995; Loreto et al., 1996; Bertin et al., 1997; Kesselmeier et al., 1997; Peñuelas & Llusià, 1999; Llusià & Peñuelas, 2000), as its monoterpene emission is driven mainly by its current biosynthesis. However, under light conditions, when photosynthesis was active, JA treatment caused the opposite effects on photosynthetic rates and monoterpene emissions, and the enhancement of monoterpene emissions by JA was not avoided by the decrease in net photosynthetic rates that also occurred as a result of JA.

It is conceivable that JA triggers not only local, but also systemic responses on monoterpene emissions, as for instance observed in Succisa pratensis upon caterpillar attack (Peñuelas et al., 2005b), making even more significant the relative increase found in JA-S vs control leaves 24 h after spraying (Fig. 2). This observed pattern of increasing monoterpene emissions after JA application agrees with the response to jasmonates reported in other species (Halitschke et al., 2000; Schmelz et al., 2001; Martin et al., 2003; reviewed by van Poecke & Dicke, 2004). Although the precise mechanism by which JA affects volatile production is as yet unknown, there is evidence that this hormone can induce expression of terpene synthase-like genes. Fäldt et al. (2003a) found in Arabidopsis thaliana an increase in the transcript levels of AtTPS03, a gene encoding a (E)-β-ocimene synthase, and a parallel increase in (E)-β-ocimene emissions from JA-sprayed leaves. Treatment with MeJA has also been reported to induce an increase in enzyme activities and elevated levels of transcripts of monoterpene synthases and diterpene synthases in Norway spruce (Martin et al., 2002; Fäld et al., 2003b). In Q. ilex different monoterpene synthases appear to produce different monoterpenes (Staudt et al., 2004), and JA treatment altered neither the composition nor the relative abundance of the emitted monoterpenes. Thus it seems that JA would affect the activity of all these enzymes or their gene transcription similarly. Recently, however, there has been some evidence suggesting that the effect of JA on terpenoid emission is related to its impact on the substrate supply feeding its biosynthesis pathway, instead of a JA elicitation of the kinetics of these pathway enzymes (Arimura et al., 2004; Ferrieri et al., 2005).

The role of these JA-mediated increases in monoterpene emissions may be related to their properties in helping to protect against both abiotic and biotic stresses, given that these isoprenoids may be of particular relevance in the adaptation of plant species to adverse environmental conditions (Peñuelas & Llusià, 2003, 2004; Peñuelas & Munné-Bosch, 2005). Monoterpenes can act as antioxidants protecting plant membranes against peroxidation and reactive oxygen species such as singlet oxygen, as has been described for Q. ilex (Loreto et al., 2004), because of their double bonds. Furthermore, given that they are small lipophilic molecules, they may also assist hydrophobic interactions in membranes that result in their stabilization (Havaux, 1998; Munné-Bosch & Alegre, 2002, 2003; Velikova et al., 2004).

Emissions of methyl salicylate

The observed pattern of increased MeSa emissions was also found by Martin et al. (2003) in MeJA-treated Norway spruce leaves. The results of Ament et al. (2004) with JA-synthesis mutant tomato plants suggest that induced emissions of MeSA depend on JA at the transcriptional level. Methyl salicylate has also been found to be released after herbivore damage in other species, and to attract the enemies of herbivores (Dicke et al., 1999; Kessler & Baldwin, 2001). However, Heil (2004) reported a decrease in MeSa emission in Lima bean leaves sprayed with a dose of JA (double the dose used in this study), and attributed this decrease to downregulation between these two antagonist phytohormones.

Both the JA-induced increase and decrease in MeSA emissions are possible outcomes of the complex interactions between these two hormones and their metabolic pathways. The response of MeSa emissions to JA appears to depend on the JA dose and on plant conditions. Jasmonic acid and salicylic acid (SA) appear to regulate separate biochemical pathways and have different functions (Karban & Baldwin, 1997), although their signalling pathways can function independently or can interact through crosstalk in additive or negative ways (Chao et al., 1999; Knight & Knight, 2001; Stotz et al., 2002). Significant induction of MeSa has been found to require an optimal concentration of JA in a JA-synthesis mutant tomato plant (Ament et al., 2004). In the same study, Ament et al. (2004) also stated that the optimal dose of exogenous JA required to induce MeSa emission may depend on the amounts of other plant defence intermediates (such as SA) present at the moment of application, as SA may antagonize downstream JA responses.

In any case, in our study, after the application of exogenous JA in plants that apparently were not stressed, we have found a JA-mediated volatilization of MeSa that provides evidence of crosstalk between JA and SA.

Decreases in formaldehyde uptake

The measurement of formaldehyde by PTR-MS is affected by humidity because of the similar proton affinity between formaldehyde and water (Kato et al., 2004). In typical environmental air conditions such as ours, measured formaldehyde levels can reach 60% of their actual value and will decrease as humidity increases (S. Kato, personal communication). Taking into account the relative formaldehyde signal change in response to humidity (S. Kato, personal communication), we estimated the influence of water vapour change, caused by changes in stomatal conductance, in the formaldehyde signal in our experiment and concluded that, although part of the measured formaldehyde change (approx. 10%) could be attributed to water vapour change, the influence of humidity alone cannot explain the observed change in formaldehyde concentration (50%).

The observed pattern of formaldehyde uptake agrees with previous reports of plants behaving as a general sink for formaldehyde (Wolverton et al., 1984). Several subsequent studies have demonstrated that exogenous formaldehyde can be incorporated into the metabolism of photosynthetic cells to be used as a C source (Giese et al., 1994) and be removed by different in vivo pathways (Hanson & Roje, 2001), although the main enzyme responsible for the metabolism of formaldehyde seems to be the glutathione-dependent formaldehyde dehydrogenase (FALDH) (Achkor et al., 2003). It has been proposed that the main function of this enzyme in biological organisms is to detoxify endogenous and exogenous formaldehyde. Diaz et al. (2003) reported that the gene coding for FALDH in Arabidopsis is downregulated by wounding and JA, and that in tobacco FADHL levels and enzymatic activity decreased 24 h after jasmonate treatment; this agrees with our results which show a decrease in formaldehyde uptake 1 d after JA application.

Our results contrast with those of Schaefer et al. (1995) and Kesselmeier et al. (1997), who reported evidence for the direct emission of formaldehyde in Q. ilex. However, Bode et al. (1996) and Kesselmeier et al. (1997) concluded that a bidirectional exchange of formaldehyde, emission as well as deposition, depends on the actual environmental concentration and on the leaf concentration resulting from the balance between production and consumption (Kesselmeier et al., 1993). In JA-S leaves, intracellular formaldehyde levels might be higher than in control leaves because of the effect of JA downregulating FADHL levels and activity. The increase in internal formaldehyde concentrations as a consequence of a decreased biogenic sink could partly explain the lower level of formaldehyde uptake in JA leaves than in control leaves. The lower conductance in JA-S leaves might have also partly influenced this lower uptake of formaldehyde, as the low Henry's Law constant of formaldehyde makes it strongly sensitive to stomatal conductance (Niinemets et al., 2004).

Conclusions

In summary, we show in this work that (i) exogenous JA elicited a plant response that lowered CO2 and H2O exchanges; (ii) JA appears to cause both stomatal and nonstomatal limitations; (iii) JA increased monoterpene emissions by 20–30% in Q. ilex; (iv) the effect of JA on monoterpene emissions increased with time, being stronger 24 h than 1 h after treatment; (v) the composition and relative abundance of the monoterpenes emitted were not altered by JA treatment; (vi) JA spraying induced a twofold increase in MeSa emissions; and (vii) after 24 h of treatment JA spraying decreased formaldehyde uptake by 50%. All these results suggest that, regardless of the nature of the stress factor that increases JA concentrations, the signalling molecule JA significantly affects plant VOC emissions.

Acknowledgements

We thank Dr S. Nogués for his technical assistance. This research was partially supported by Spanish MCYT grants REN2003-04871 and CGL2004-01402/BOS. We also gratefully acknowledge partial funding from the ISONET European Commission contract MC-RTN-CT-2003-504720, and a Fundación BBVA 2004 grant.

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