Determination of de novo and pool emissions of terpenesfrom four common boreal/alpine trees by 13CO2 labellingand PTR-MS analysispce_2104 781..792

ANDREA GHIRARDO1, KRISTINE KOCH1, RISTO TAIPALE2, INA ZIMMER1, JÖRG-PETER SCHNITZLER1 &JANNE RINNE2

1Karlsruhe Institute of Technology, Institute of Meteorology and Climate Research (IMK-IFU), Kreuzeckbahnstrasse 19,82467 Garmisch-Partenkirchen, Germany and 2Department of Physics, PL-64, 00014 University of Helsinki, Finland

ABSTRACT

Boreal forests emit a large amount of monoterpenes intothe atmosphere. Traditionally these emissions are assumedto originate as evaporation from large storage pools. Thus,their diurnal cycle would depend mostly on temperature.However, there is indication that a significant part of themonoterpene emission would originate directly from denovo synthesis. By applying 13CO2 fumigation and analyzingthe isotope fractions with proton transfer reaction massspectrometry (PTR-MS) and classical GC-MS, we deter-mined the fractions of monoterpene emissions originatingfrom de novo biosynthesis in Pinus sylvestris (58%), Piceaabies (33.5%), Larix decidua (9.8%) and Betula pendula(100%). Application of the observed split between de novoand pool emissions from P. sylvestris in a hybrid emissionalgorithm resulted in a better description of ecosystem scalemonoterpene emissions from a boreal Scots pine foreststand.

Key-words: 13C; boreal forest; DMADP; GDP; isoprene;monoterpene; stable isotope labelling.

INTRODUCTION

The boreal region comprises the largest forest area on ourplanet. It covers 15 million km2 in Eurasia and NorthernAmerica (FAO 2001). In addition to being one of the mainCO2 sinks, it is an important source of reactive volatileorganic compounds (VOCs) emitted into the atmosphere.These biogenic VOCs (BVOCs) have an effect on the tro-pospheric ozone production, carbon monoxide, hydroxylradical levels, methane lifetime, and formation and growthof secondary aerosols (Trainer et al. 1987; Chameides et al.1988; Jacob & Wofsy 1988; Novakov & Penner 1993). Thus,BVOCs play an important role for air quality and climatedynamics. VOC emissions from boreal forests are oftencharacterized by the dominance of monoterpenes, a class ofC10H16 hydrocarbons (Tarvainen et al. 2007; Rinne, Bäck &

Hakola 2009), which take part in the aerosol growth pro-cesses (Tunved et al. 2006). Because of the large borealforest area and its impacts on the above-mentioned pro-cesses, reliable emission estimates are needed underpresent and predicted future climate.

Boreal forests have low tree species diversity (FAO2001). Dicotyledon trees, such as birches and willows, arecommon but rarely occur as dominant species. Coniferousspecies, which are the dominant tree species in theseregions, store large amounts of monoterpenes into specialinternal storage structures such as resin canals and ducts,whereas dicotyledons do not possess these structures.Thus, the monoterpene emission from dicotyledons can beassumed to be closely related to biosynthesis.

Traditionally, monoterpenes emitted by boreal conifer-ous trees have been assumed to originate as evaporationprocess from the storage structures (Guenther, Monson &Fall 1991; Grote & Niinemets 2008). The evaporationdepends mostly on saturation vapour pressure of monoter-penes, which is related to the needle temperature. As aresult, monoterpene emissions from coniferous trees arecommonly calculated by temperature-dependent algo-rithms (Tingey et al. 1980; Guenther et al. 1991; Guentheret al. 1993). Even though specific storage structures areabsent in most dicotyledon tree species, the temperature-dependent algorithm is often used also to describe themonoterpene emissions from these species as well (e.g.Tarvainen et al. 2007).

Conversely, isoprene and monoterpene emissions frommany dicotyledon trees originate from recently fixedphotosynthetic intermediates by de novo biosynthesis in alight- and temperature-dependent manner (Tingey et al.1979; Loreto et al. 1996, 2000) when photosynthesis is notimpaired by environmental stresses (Kreuzwieser et al.2002; Fortunati et al. 2008; Teuber et al. 2008).

Studies of light and temperature dependency of mono-terpene emission from some coniferous tree species suchas Scots pine (Steinbrecher et al. 1999; Shao et al. 2001;Tarvainen et al. 2005), Norway spruce (Steinbrecher 1989;Schürmann et al. 1993) and Stone pine (Staudt et al. 1997)have indicated a significant contribution of de novo mono-terpene emissions, although, a proper quantification was

Correspondence: J. Rinne. Fax: +358 9191 50860; e-mail: janne.rinne@helsinki.fi

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not achieved. The separation of monoterpene emissionsoriginating from de novo synthesis and evaporation fromstorage pools by measuring light and temperature depen-dencies is often inconclusive. In measurements conductedunder field conditions, light and temperature tend to corre-late with each other, making the separation of their effectsdifficult. Therefore, our knowledge on the partitioning ofmonoterpene emissions evaporating from storage poolsand the ones originating directly from de novo biosynthesisis insufficient for improving existing algorithms or develop-ing new process-based models.

Stable isotope 13CO2-labelling technique offers a possibil-ity to better define de novo monoterpene biosynthesis uti-lizing recently fixed carbon. This technique has previouslybeen used mainly to study isoprene and monoterpeneemissions from dicotyledon trees (Delwiche & Sharkey1993; Loreto et al. 1996). However, even when terpeneemissions originate completely from de novo synthesis, theincorporation of 13C is incomplete, reaching maximumvalues of approximately 80% under non-stressed condi-tions (e.g. Loreto et al. 1996; Kreuzwieser et al. 2002). Thisfact has hampered the correct determination of de novo andpool emissions in monoterpene storing species.

For these reasons, this 13CO2-labelling study aims to: (1)analyze comprehensively the metabolic origin of isopreneand monoterpenes in leaves and needles of some majorEurasian-boreal/alpine tree species; (2) determine theunderlying pool sizes of terpene precursors in subcellularcompartments; and (3) quantify the species-specific contri-bution from storage pools and de novo synthesis to theoverall emission of monoterpenes in order to improve thealgorithms used in emission models.

MATERIALS AND METHODS

Plant material

For all the experiments, 2-year-old saplings of Silver birch(Betula pendula L.), Scots pine (Pinus sylvestris L.),Norway spruce [Picea abies (L.) Karst.] and Europeanlarch (Larix decidua L.) were used. Silver birches werecollected locally at Garmisch-Partenkirchen, Germany,while Scots pine, European larch and Norway sprucesaplings were obtained from a pre-alpine plant nursery(Pflanzgarten, Laufen, Germany). Additionally, 5-year-oldHolm oak trees (Quercus ilex L.), originating from anItalian plant nursery (Balducci, Pistoia, Italy) were used asplant model (Grote et al. 2006) with high light-dependentmonoterpene emissions.

Plants were placed into 20 cm high pots with diameter of25 cm, containing 50% (v/v) of perlite (Agriperl Dämm-stoff GmbH, Dortmund, Germany), 25% (v/v) of silicasand with particle size of 1–3 mm, 25% (v/v) of potting soil(Fruhstorfer Einheitserde, Bayerische Gärtnereigenossen-schaft, Aschheim, Germany) and 10 g of Osmocote andTriabon fertilizer (Scotts International GmbH, Nordhorn,Germany). The potted plants were watered weekly. Theplants were grown outside, in the garden of IMK-IFU at

Garmisch-Partenkirchen throughout the growing season.The institute is located in the alpine area, at 780 AMSL,next to a forest dominated by Scots pine and Norwayspruce trees. Summer temperature ranges from 10 to 30 °Cand photosynthetic photon flux density (PPFD) from 100to 1500 mmol m-2 s-1. The experiments were carried out inthe first two weeks of September. A few needles at theintersection from the current to the old needle age classwere removed and the stem enwrapped with Teflon tape inorder to fit them to the enclosures. This procedure mini-mized the effect of mechanical stress, and respectivemonoterpene emission, when twigs were later on enclosedgas-tight in the cuvette.

Total needle area was calculated using the projected areaand the cross-sectional shape for each needle-species asdescribed by Chen et al. (1997). Because twigs were imme-diately frozen after the experiment, total needle area wascalculated by counting the number and measuring thelength of the needles before the experiment, while the pro-jected area was taken from similar needles of other plants.

Cuvette measurements of BVOC andphotosynthetic gas exchange

For the experiments, two custom-made dynamic cuvettesystems with cuvette volume of 450 cm3 (for details, seeLoivamäki et al. 2007) were coupled to a portable gasexchange system (GFS-3000, Heinz Walz, Germany) andrun in parallel. The on-line quantification of labelled andunlabelled BVOCs was achieved by proton-transfer-reaction mass-spectrometer (PTR-MS, Ionicon Analytik,Austria; Hansel et al. 1995).

The plants were transferred into the IMK-IFU phyto-chamber and twigs were enclosed into the cuvettes in theafternoon the day before feeding the plant with 13CO2.Light intensities were set to a photosynthetic photon fluxdensity (PPFD) of 1000 mmol m-2 s-1 during the light phaseand circa 0 mmol m-2 s-1 during night phase. Leaf/needletemperature was kept constant at 30 °C. Cuvettes wereflushed continuously with synthetic VOC-free air (21.0%v/v O2, 79.0% v/v N2, BASI Schöberl, Germany) with dewpoint of 4 °C, and containing 385 ppmv of CO2 with anatural abundance of 13CO2. The cuvette inlet flow wasadjusted to 600 mL min-1 leading to air residence time ofless than 47 s. From the outlet air, 70 mL min-1 was takeninto the PTR-MS for VOCs monitoring. In addition, 4 L ofoutlet air was trapped in a three-bed-adsorbent tube (90 mgCarbotrap C, 60 mg Carbotrap, 60 mg Carbopack X,Supelco Bellafonte, PA, USA) by sampling at 133 mL min-1

for 30 min. Tubes were analyzed by GC-MS as described bySchnitzler et al. (2004a). The outlet air (250 mL min-1) fromthe cuvette passed the GFS-3000 and the remaining air flowwas continuously monitored to ensure the flow system wasoperating correctly.

13CO2-labelling was performed in light until incorpora-tion of 13C into isoprene and monoterpenes reached asteady state (8 h for conifers and 5 h for dicotyledons)by replacing the CO2 with natural 13C abundance with

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99% 13CO2 at 385 � 7.7 ppmv (Air Liquide, Griesheim,Germany) in morning at 0900 h CET, 3 h after onset of lightin the cuvette, when net CO2 assimilation and BVOC emis-sions were stable. Measurements were performed byswitching automatically between the two parallel cuvettesevery 15 min. The first 3 min of data after each switch werediscarded and the remaining points were averaged. Prior toeach analysis, the background of each empty cuvette wasmeasured.

PTR-MS analysis of isoprene andmonoterpene emission

The PTR-MS operating parameters were E/N = 110 Td(E = electric field; N = number of density of the drift tubemolecules; Td = 1 Townsend = 10-17 cm2 V molecule-1),p-drift = 1.73 mbar, T = 43 °C, V = 400, O2

+ and NO+ < 2%of H3O+. The low E/N ratio was maintained in order tominimize fragmentation of monoterpenes. Although lowdrift pressure lowered the detection, the fragmentation rateof the most abundant monoterpenes (a-pinene, sabinene,limonene) decreased consequently (m81/m137 = 1.7),giving higher signal for the non-fragmented monoterpeneisotopes. In order to determine a possible influence of watervapour pressure (WVP) on monoterpene fragmentationpattern (Tani et al. 2004), a daily humidity-dependent cali-bration (1–50 ppbv gas standards with 5000–15000 ppmvH2O) was performed with 11 compound VOC standard inN2 (Apel-Riemer Environmental, Denver, CO, USA). Thiswas needed as the WVP in the cuvette changes betweenlight and dark measurement, that is, during open and closedstomata.

The detection efficiency of the PTR-MS changes accord-ing to the molecular mass of the compounds following atypical transmission factor (TF) curve (Steinbacher et al.2004). For the quantification of the monoterpene isotopes(i.e. m137–147), which lie in the descending part of the TFcurve (e.g. Taipale et al. 2008), each isotope was quantifiedusing a relative transmission factor (rTF) based on the frac-tionation rate of monoterpene from standard and the massdiscrimination of the instrument (i.e. TF). We checked thattotal monoterpene emission before labelling of Holm oakwas equal to the sum of all monoterpene isotope emissionscalculated with rTF after short 13C labelling period.

The limit of detection (LOD, reported in each graph) wascalculated separately at each measurement and set to twicethe standard deviation of the count per second (cps) signalof background measurement divided by the instrument’ssensitivity (cps ppv-1).

Measurement of subcellular DMADP and totalGDP pools

At the end of 13CO2-labelling, the twigs/leaves were imme-diately frozen in liquid N2 and stored at -80 °C. Dimethyl-allyl diphosphate (DMADP) and geranyl diphosphate(GDP) contents were measured according to Nogues, Brilli

& Loreto (2006) using a modified extraction protocol ofBrüggemann & Schnitzler (2002): 50 mg of frozen homog-enized leaf powder was solubilized with 1 mL buffer (50 mmTris/HCl pH8, 20 mm DTT, 20 mm MgCl2, 5% glycerin) con-taining 10 mm sodium molybdate as phosphatase inhibitorand centrifuged at 20 000 g for 15 min. Supernatant(100 mL) was placed into a 2 mL gas-tight white glass vial(Supelco) and incubated at 70 °C for 90 min with 100 mL ofH3PO4 (17% v/v). Afterwards, the sample was cooled downfor 3 min on ice and then the reaction stopped by adding100 mL of 9.2 m NaOH and the vial was left at room tem-perature. Finally, the protonated masses of isoprene and itsisotopes, 69–74 amu (Loivamäki et al. 2007) and for linaloolat masses 97–104 amu (Nogues et al. 2006) were measuredwith PTR-MS. Each sample was analyzed in triplicate.

Subcellular DMADP pool sizes were calculated measur-ing the total DMADP pool (DMADPtot), the isotope ratiosof both isoprene and of DMADPtot during steady stateconditions after 13CO2 labelling. Assuming that there isinsignificant exchange of DMADP between cytoplasm andchloroplast (Wu et al. 2006), and that the biosynthesis ofisoprene is mediated by the MEP-pathway (Lichtenthaler1999), the pattern of 13C incorporation into isoprene mustreflect the pattern of chloroplastidic DMADP. The chloro-plastic DMADP pool (DMADPchl) can be calculated asfollowing:

DMADPf DMADP

fchl

DMADP tot

Isoprene

= (1)

where fDMADP is the fraction of fully labelled DMADP (i.e.only chloroplastic); the fIsoprene the fraction of fully labelledisoprene.The cytosolic DMADP pool (DMADPcyt) is there-fore calculated by following subtraction:

DMADP DMADP DMADPcyt tot chl= − (2)

Calculation of monoterpene emission fromde novo biosynthesis

De novo monoterpene emission was calculated by takinginto account the 13C-labelling pattern of emitted monoter-penes and isoprene. The MEP-pathway is responsible forthe production of both immediate isoprene and monoter-pene precursors; therefore, the 13C-labelling pattern ofplastidic DMADP and GDP must be identical. Althoughisoprene emission is always entirely de novo, the emittedisoprene does not become completely labelled after 13CO2

feeding because of alternative C-sources. We have takeninto account the other C-sources of isoprene and monoter-pene biosynthesis by calculating the de novo monoterpenebiosynthesis fraction (F, in percentage) as following,

FMI

= 100% (3)

where M and I are the fraction of monoterpenes and iso-prene labelled with 13C, respectively.

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Isoprene is biosynthesized in chloroplasts during lightmainly by enzymatic (ISPS) conversion of freshly syn-thesized chloroplastic DMADP (for review, see Grote &Niinemets 2008). In small amounts, isoprene also originatesfrom non-enzymatic conversion of DMADP (cytosolic andchloroplastic) under physiological pH (Lehning et al. 1999;Brüggemann & Schnitzler 2002).Therefore, before calculat-ing the fraction of monoterpenes originating from de novosynthesis, we subtracted isoprene emission measured duringdark, which is because of non-enzymatic conversion, fromall measured emissions to yield enzymatic emissions.

Quantification of endogenous monoterpenecontents and 13C isotopic signature

Endogenous monoterpene contents and pattern were mea-sured by gas chromatography flame ionization detection(GC-FID) (Fischbach et al. 2002), whereas the isotopic splitof the samples was determined by PTR-MS (Loivamäkiet al. 2007). Fifty milligrams of frozen homogenized leafmaterial was dissolved in a 2 mL gas tight vial with 1 mL ofpentane and shaken at 4 °C for 3 h, and then let to restovernight at 5 °C in darkness. Finally, 1 mL of this solutionwas transferred into a gas tight vial and incubated for 1 h atroom temperature before PTR-MS and GC-FID analyses.The PTR-MS signal was recorded at masses 137–147 amu.

Light- and temperature-dependent hybridmodel and flux measurements

The hypothesis of the two origins of the monoterpene emis-sion from conifers can be written formally as:

E E E= +synth pool (4)

where Esynth represents the monoterpene emission originat-ing directly from biosynthesis and Epool that originatingfrom specialized storage structures.

By using the formulation by Guenther (1997) for iso-prene emission for Esynth and a classical formulation formonoterpene emission (Guenther 1997) for Epool, we canwrite

E E f C C f= + −( )[ ]0 1denovo T L denovo γ (5)

where CT and CL are synthesis activity factors for tempera-ture and light, respectively; g is temperature activity factorfor vapourization from pool; fdenovo is the fraction of theemission originating directly from synthesis; and E0 is nor-malized emission potential.

In order to test the applicability of the hybrid model(Eqn 5), we used ecosystem scale monoterpene emissiondata measured by the disjunct eddy covariance (DEC)method with PTR-MS (for details, Rinne et al. 2007 andTaipale et al. 2008) at a Scots pine forest. We set parameterfdenovo to 0.58, that is, the fractions for pool and de novoemissions to 42 and 58%, as indicated for the Scots pine bythe labelling experiment.

A more detailed description of the emission algorithmand flux measurements can be found in Appendix S1.

Statistical analysis

All experiments were performed independently withthree replicates. Statistical analysis (t-test) was performedusing the Software package Origin (version 7.0; OriginLab,Northampton, MA, USA).

RESULTS

Incorporation of 13C into monoterpenes andisoprene upon labelling with 13CO2

The day before 13CO2-labelling, twigs (or leaves) wereenclosed and acclimated to their new environment. Imme-diately after insertion into the enclosure, European larch,Norway spruce and Scots pine twigs (Fig. 1a–c, respectively)showed very high monoterpene emission rates which expo-nentially declined during the rest of the light phase andthroughout the night.This monoterpene emission was likelybecause of mechanical stress during enclosure of the twigs,with the consequent release of monoterpenes from storagestructures. Contrary to conifers, mechanical stress didnot affect monoterpene emissions of Silver birch (Fig. 1d)and Holm oak (Fig. 1e), in which monoterpene emissionsadapted slowly to 30 °C and 1000 mmol m-2 s-1 PPFD. Thenext morning, monoterpene emission rates increased whenlight was switched on. As temperature in the enclosure waskept at 30 °C throughout the whole experiment, most ofthe increase in emission identified was because of light-dependent process.Monoterpene emission from Silver birchand Holm oak showed a completely light-dependent behav-iour with practically zero emission overnight (Fig. 1d,e) andsimilar emission before and after dark period.

Isoprene emission rates of all species displayed asimilar pattern; a rapid decline of emission in darkness andcomparable emission rates during the next day (Fig. 2).Overall, isoprene emission rates (ranging from 0.02 to0.7 nmol m-2 s-1) of all species were much lower comparedwith monoterpene emissions and the portion of isoprene inrespect to the overall terpene emission was only 12 � 3%,23 � 4% and 13.9 � 0.6% for European larch, Norwayspruce and Scots pine (Fig. 2a–c). The GC-MS analysis con-firmed that isoprene detected with PTR-MS was not mis-taken with a fragment of 2-methyl-3-buten-2-ol (MBO)(data not shown).

In the morning, the measurements were continuedfor 180 min after the light had been switched on inorder to ensure a steady state of BVOC emissions andphotosynthetic gas exchange (net CO2 assimilation:European larch 2.4 � 0.4, Norway spruce 4.5 � 0.9,Scots pine 2.2 � 0.2, Silver birch 1.7 � 0.3, Holm oak1.6 � 0.2 mmol m-2 s-1 � SD) before 12CO2 was replaced by13CO2 (99% 13C).

Isoprene became rapidly enriched in 13C in all specieswith a similar final labelled fraction of approximately 80%

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(Fig. 2). The incorporation of 13CO2 into monoterpenes dis-played different behaviour between conifers and dicotyle-dons (Fig. 1). European larch, Norway spruce and Scotspine incorporated 13C into monoterpenes incompletely(Fig. 1a–c). This can be assumed to be because of the sig-nificant non-labelled monoterpene emissions from storagestructures in needles and bark tissue (for endogenousmonoterpenes, see Fig. 6). However, all conifers emittedsignificant amounts of recently fixed C as monoterpenes

(European larch 6.6% � 1.6%, Norway spruce 23 � 5%,Scots pine 39 � 7%). Scots pine twigs increased theirlabelling for over 480 min before reaching a stable andhigh incorporation. The most labelled monoterpeneswere a-pinene, b-pinene, camphene and limonene (data notshown).

Silver birch and Holm oak incorporated 13C more com-pletely (Fig. 1d,e). Within 30 min, approximately 80% of Cin monoterpenes became 13C-labelled. These values weresimilar to the 13C incorporation into isoprene (Fig. 2d,e). In

Figure 1. Monoterpene emissions ( ) from conifer trees,(a) European larch, (b) Norway spruce and (c) Scots pine, fromdicotyledon trees (d) Silver birch and (e) Holm oak and 13Clabelling ( ). The labelling 13CO2 was applied at time 0 (verticaldash line). The horizontal dot line represents the LOD and thelight grey square the night phase (each point represents a 30 minaverage; n = 3 � SE; spruce n = 5).

Figure 2. Isoprene emission ( ) from conifer trees,(a) European larch, (b) Norway spruce and (c) Scots pine, fromdicotyledon trees (d) Silver birch and (e) Holm oak and 13Clabelling ( ). The labelling 13CO2 was applied at time 0 (verticaldash line). The horizontal dot line represents the LOD and thelight grey square the night phase (each point represents a 30 minaverage; n = 3 � SE; spruce n = 5).

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Silver birch, the variability of monoterpene emission ratesbetween the samples was very high, while the incorporationof 13C was similar (Fig. 1d).

Subcellular DMADP, total GDP pool sizes

We quantified the precursors of isoprene and monoterpe-nes, DMADP and GDP, of leaf/needle samples takenimmediately after gas exchange analysis. The analysisrevealed distinct differences in absolute concentrations:generally, total DMADP pools were higher than totalGDP pools (Fig. 3a,b, respectively) in all species. Withinspecies, Scots pine displayed significantly higher DMADPcontents compared with other conifer species, whereas theisoprene rate was the lowest (comparison Figs. 2 and 3).Incorporation of 13C into the isoprene precursor DMADPwas very low, in particular the number of fully labelledDMADP molecules (i.e. where all 5 12C atoms have beensubstituted by 13C, data not shown). The calculation ofplastidic to cytosolic DMADP ratio showed that inNorway spruce, Scots pine and Silver birch, approximately20–25% of the overall DMADP pool is localized in chlo-roplasts (Fig. 3a), whereas such calculation was not pos-sible in European larch and Holm oak because of unclearDMADP labelling pattern.

The 13C incorporation into the overall GDP pool (Fig. 3b)was approximately 7–18% with significant difference(P < 0.01) only between Scots pine (18.1 � 0.6%) andEuropean larch (9.2 � 1.5%).

Analysis of in vitro enzyme activity demonstrated thatNorway spruce, Scots pine and Holm oak needles/leavescontained mono-TPS activities, which were not detectablein European larch and Silver birch (Fig. S1). The a-pinene,b-pinene and sabinene were the predominant monoterpe-nes synthesized in Holm oak, while Scots pine and Norwayspruce formed predominantly a-pinene and b-pinene,respectively (Fig. S1). Qualitative monoterpene analysisshowed similar pattern of mono-TPS activities and overallemission for Holm oaks, whereas the pattern largely dif-fered in conifer species (Fig. 4) reflecting the influence ofthe two origins of the emission.

The proportion of monoterpenes originating from denovo biosynthesis differed in European larch, Norwayspruce and Scots pine (Fig. 5). In European larch, newlysynthesized monoterpenes contributed marginally (rang-ing 9.8 � 1.0%) to the overall monoterpene emission ofthese species. This is in contrast to Norway spruce andScots pine. In these species, the light-dependent biosynthe-sis of monoterpenes amounted to 34 � 8% and 58 � 4%of the total emission, respectively. In birch and oak, themonoterpene emissions were practically entirely de novo(Fig. 5).

Distribution of monoterpenes among specificand unspecific storage structures inneedles/leaves and bark

The endogenous pools of monoterpenes differed stronglybetween conifers (Fig. 6a–f) and dicotyledon species(Fig. 6g–j), as well as within the group of conifers. All coni-fers stored monoterpenes in needles as well as in the bark.In Norway spruce (Fig. 6e,f), the concentration of monoter-penes were almost equal in both tissues, whereas inEuropean larch (Fig. 6a,b) and Scots pine (Fig. 6e,f), theconcentration of monoterpenes (84.5 and 76.4%, respec-tively) was much higher in the bark tissue of the twigs.

In European larch, the pattern of stored monoterpenes inneedles was similar to that in bark, with a-pinene andb-pinene being predominant compounds. In Norway spruceneedles, the endogenous pool was dominated by camphene,limonene and a-pinene, whereas in the bark b-pinene anda-pinene were the most abundant monoterpenes. In Scotspine needles, a-pinene was the overall dominating com-pound followed by camphene. Interestingly, the bark ofScots pine twigs contained high amounts of b-myrcene, acompound which was less abundant in the needles (Fig. 6).

Incorporation of 13C into storage pools was significantlydetectable (P < 0.05) only in Scots pine needles(1.2% � 0.4) but not in its bark, nor in any other coniferousspecies. This indicates that the incorporation of 13C into theemitted monoterpenes was not because of the incorpora-tion into monoterpene storages, and that the emissionsoriginating from de novo synthesis by-passes the storage.The two tree species without specific storage pools (birchand oak) stored monoterpenes only marginally within theirtissue (Fig. 6g,i).

Figure 3. (a) Total cellular ( ) and subcellular chloroplastic ( )DMADP pools and percentage of chloroplastic DMADP ontotal ( ); (b) Total cellular GDP pools ( ) and 13Cincorporations ( ) (n = 3 � SE, spruce n = 5). Statisticallysignificant differences are indicated by different letters(minuscule and majuscule compare pools and percentage,respectively) without (P < 0.05) or with asterisk (P < 0.01). n.d.,not detectable.

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Influence of the de novo synthesis onmodelling ecosystem scalemonoterpene emission

The diurnal cycle calculated by the traditional pool emis-sion model (Eqn 3 in Appendix S1) resembled the mea-sured emission but underestimated the daytime emissionsand overestimated the night-time emission (Fig. 7).The dayto night-time variation in the ecosystem scale monoterpeneemission were better described by the hybrid model(Eqn 5) taking the fractionation between de novo andpool emission from the laboratory results. The correlationcoefficient (r2) between the measured and modelledfluxes was 0.78 and 0.86 for the pool-only and hybridmodel, respectively. The standard emission potentials were0.66 mg m-2 h-1 and 0.83 mg m-2 h-1 (1.3 mmol m-2 s-1 and1.7 mmol m-2 s-1), respectively.

DISCUSSION

The present analysis shows that the fraction of monoterpeneemission originating from de novo biosynthesis differedstrongly between fully mature needles of the three coniferspecies (Scots pine 58%; Norway spruce 33.5%; Europeanlarch 9.8%). On the contrary, in both Silver birch and Holmoak leaves, this fraction was similar (98–100%), indicatingthat the emission originated totally from de novo synthesis.

A major step forward for correct estimations of de novomonoterpene emissions in conifers consisted in consideringthe isoprene labelling pattern. Isoprene is immediatelyemitted after synthesis because of the absence of storageand its high volatility. Both isoprene and monoterpenes aresynthesized via the MEP-pathway, sharing the same inter-mediates [DMADP and its isomer isopentenyl diphosphate(IDP) form GDP] with similar 13C labelling pattern (Lich-tenthaler 1999). Even under optimal photosynthetic condi-tions, the maximal incorporation of 13C into total isoprenecarbon, when the plants are fed by 13CO2, is approximately

Figure 4. Isoprene and monoterpene emissions from (a)European larch, (b) Norway spruce and (c) Scots pine, fromdicotyledon trees (d) Silver birch and (e) Holm oak twigs.a-thujene ( ), a-pinene ( ), camphene ( ), sabinene andb-pinene together ( ), myrcene ( ), D-3-carene ( ), cymene ( ),b-phellandrene ( ), limonene ( ), ocimenes ( ), 1,8-cineole ( )and isoprene ( ) (n = 3 � SE, for spruce n = 5). Emission is givenper leaf/needle area.

Figure 5. De novo biosynthesis during light condition forisoprene ( ) and monoterpenes ( ) (n = 3 � SE; for sprucen = 5).

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80%. Therefore, additional C-sources provide the remain-ing 12C to the MEP-pathway (Kreuzwieser et al. 2002;Schnitzler et al. 2004b). Under stress conditions, and limitednet CO2 assimilation (e.g. drought), isoprene is no longerclosely related to photosynthesis, instead other ‘old’ carbonmaintain the isoprene biosynthesis (Brilli et al. 2007). In thisstudy, we have taken into account the alternative C-sourcesof de novo monoterpene biosynthesis. Conceptually, thisapproach would allow estimating de novo emissions ofmonoterpenes from conifers even under stress conditions,when the contribution of ‘fresh’ photosynthetic carbon islimited.

Compared with earlier studies, our data show a highercontribution of the light-dependent de novo emission to

the overall monoterpene emission in Scots pine andNorway spruce. Using 13CO2, Shao et al. (2001) reportedfor Scots pine needles a contribution of 20–30% from denovo biosynthesis. From correlation analysis of night andday time emission rates of Scots pine, Steinbrecher et al.(1999) attributed 25–37% of the emission to a light-dependent process. In a comparable study with Stone pine(Pinus pinea L.), Staudt et al. (1997) found a-pinene,b-myrcene and linalool emission rates were reduced about3, 4 and 20 times, respectively, when plants were placed indarkness. In the same study, trans-b-ocimene and 1,8-cineole emissions were two orders of magnitude higherduring light condition than in darkness. For Norwayspruce, Steinbrecher (1989) and Schürmann et al. (1993)

Figure 6. Endogenous monoterpenepatterns in needles/leaves and bark tissueof (a,b) European larch, (c,d) Norwayspruce and (e,f) Scots pine, fromdicotyledon trees (g, h) Silver birch and(i,j) Holm oak. a-thujene ( ), a-pinene( ), camphene ( ), sabinene ( ),b-pinene ( ), myrcene ( ), limonene( ) and 1,8-cineole ( ). (n = 3 � SE, forspruce n = 5). n.d., not detectable.

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demonstrated a rapid incorporation of 13C in monoterpenefragments of the GC-MS analysis when needles whereexposed to 13CO2. However, they reported no quantitativelabelling rates. Using their data, we estimated a 20–30%contribution of de novo biosynthesis to the overall mono-terpene emission in Norway spruce needles. Even whenthe reported data records are very few, the present data arein the same range. Nevertheless, we assume that the presenthigher values are more realistic because of the considerationof non-labelled de novo synthesized monoterpenes.

By means of the labelling pattern of isoprene and itsprecursor DMADP, we were able to calculate the subcellu-lar distribution of DMADP. Because the cross-talk betweenchloroplast and cytosol of DMADP and IDP is minimal(Lichtenthaler 1999; Loreto et al. 2004; Wu et al. 2006), fullylabelled DMADP is formed in the chloroplast shortly after13CO2 fumigation (Loreto et al. 2004). Because chloroplasticDMADP molecules must have the 13C pattern of isoprene,the chloroplastic DMADP pool can be therefore calcu-lated. Our values are in the range of other studies (Loretoet al. 2004; Magel et al. 2006; Rasulov et al. 2009), showingthat in the light, approximatively 20–30% of total DMADPis localized in the chloroplasts.

In general, GDP pools were higher in Holm oak than inNorway spruce and higher in Scots pine than in Europeanlarch. The high 13C incorporation into GDP in Scots pine,compared with European larch, agrees very well with the,respectively, higher de novo monoterpene biosynthesis. Ingeneral, total concentrations of GDP were lower thanDMADP concentrations. This observation differs with datafrom Nogues et al. (2006), reporting higher values of GDPthan DMADP in Holm oak leaves. The reason for thisdifference might lie in our protocol: in contrast to the earlierwork, we performed a hydrophilic extraction of solublemetabolites to avoid terpene contamination from storagestructures and for the same reason we removed the cell

debris prior to hydrolysis in order to minimize any conver-sion of other unsaturated hydrocarbons into linalool duringthe acid catalyzed reaction.

We also analyzed terpene (isoprene and monoterpene)synthase activities in all species. Enzyme activities weredetected in Holm oak, Scots pine and Norway spruceextracts, reflecting the de novo biosynthesis and 13C-labelling pattern (Fig. S1). The pattern of in vivo monoter-pene formation is also reflected in the pattern ofmonoterpenes becoming labelled during 13CO2 fumigation.We found that a-pinene, b-pinene, camphene and limonenewere the most labelled monoterpenes in all species, whereasb-myrcene was not found labelled nor present in in vitroanalysis. When light-dependent monoterpene emission isnot impaired by emission from storage structures, thepattern of mono-TPS products and monoterpene emissionsfit fairly well (Fischbach et al. 2002; Schnitzler et al. 2004a).

The present work also shows distinct differences in thepartitioning (pattern and amount) of monoterpenes intostorages in needles and in bark among the three coniferspecies confirming an earlier report (Sallas et al. 2003)where 23.6% (Scots pine, our values 24 � 3%) and 63.5%(Norway spruce, our values 51 � 4 %) of the endogenousmonoterpenes were found in needles. From the analysis ofthe tissue specific composition of monoterpenes in the threeconifer species, we conclude that different mono-TPSbecame differentially activated during needle development.This is because we found different endogenous monoter-pene patterns in needles compared with the bark. Similardifferences were reported by Sallas et al. (2003), witha-pinene being predominant in Scots pine needles, whereasa larger fraction of b-myrcene and limonene was observedin the monoterpene pool of the stem bark. In Norwayspruce, camphene was present largely in needles whereasb-pinene dominated in the bark stem monoterpene pool(Sallas et al. 2003).

Figure 7. Ecosystem scale monoterpeneemission measured by the disjunct eddycovariance (DEC) method (black dots) ata Scots pine forest, emissions calculatedby the pool model [blue line; (Eqn 3)correlation coefficient = 0.78], andemissions calculated by the hybridmodel [red line; (Eqn 5) correlationcoefficient = 0.85]. Emission is given perground area.

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Because of the large storage of monoterpenes in conifers,emissions have been assumed to be mainly a consequenceof thermal volatilization and thus traditionally modelled bytemperature-dependent algorithm (Guenther et al. 1991,1993). Light-dependent mechanisms were believed to bemarginal and therefore neglected in the models. However,ignoring the contribution of monoterpene emissions fromde novo biosynthesis resulted in discrepancies betweenmodelled and measured emission rates (Arneth et al. 2008).This effect can be seen for Scots pine during spring, whenthe vegetation is recovering and the model data do notmatch measured emission rates (Tarvainen et al. 2005).Running a hybrid model with the 58% contribution of light-dependent de novo emission for Scots pine (Fig. 7), we wereable to describe better the night- and daytime fluxes for thisspecific boreal forest stand. Even though the difference issignificant only at 90% confidence level, the result indicatesthat an introduction of an algorithm for the de novo bio-synthesis better describes the diurnal variation in monoter-pene emission from coniferous forests. Nevertheless, we canassume that the fraction of de novo terpene emission fromconifers undergoes seasonal variations because of the phe-nology of the trees (Fischbach 2001).Therefore, more infor-mation on the emission dynamics (e.g. seasonality andinfluence of biotic/abiotic stress) is needed to obtain a real-istic representation of de novo monoterpene emission in theemission models. Inclusion of: (1) boreal specific character-istics of de novo monoterpene biosynthesis and (2) a termdescribing the de novo emission of monoterpenes in semi-empirical models (e.g. Guenther et al. 1993); and (3) thedescription of evaporation from storages and storagedynamics in process-based emission models (Niinemetset al. 2002; Grote et al. 2009) will be the next logical step toachieve a more realistic description of terpene emissionsfrom boreal and alpine areas.

ACKNOWLEDGMENTS

The authors would like to thank R. Steinbrecher and B.Wolpert (IMK-IFU) for helping during GC-MS analysisand useful discussion, and M. Possell (Lancaster University,UK) for critical reading of the manuscript. Financialsupport was given to J.-P.S. by the Human Frontier ScienceProgram and to J.R. and R.T. by the Academy of Finland(1118615, 120434, 125238) and the Kone Foundation.

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Received 13 September 2009; received in revised form 8 December2009; accepted for publication 8 December 2009

SUPPORTING INFORMATION

Additional Supporting Information may be found in theonline version of this article:

Figure S1. Isoprene and monoterpene (mono-TPS) syn-thase activities in (A) Norway spruce, (B) Scots pine and(C) Holm oak. Enzyme (ISPS and mono-TPS) activities inconifer needles were analyzed with the respective buffersystem optimized in Fischbach et al. (2002). Analysis ofHolm oak enzyme activities was performed according to

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Schnitzler et al. (2004a). In European larch and Silver birchextracts, no enzyme activities could be detected. 1)a-thujene ( ), 2) a-pinene ( ), 3) camphene ( ), 4) sab-inene ( ), 5) b-pinene ( ), and 6) isoprene ( ) (n = 3 � SE,for Norway spruce n = 5; n.d. = not detectable).Appendix S1. Emission algorithm and flux measurements.

Please note: Wiley-Blackwell are not responsible for thecontent or functionality of any supporting materials sup-plied by the authors. Any queries (other than missing mate-rial) should be directed to the corresponding author for thearticle.

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