Summary Beech (Fagus sylvatica L.) seedlings were grownin an ambient or elevated CO2 concentration ([CO2]) either insmall stands in microcosms for three to four seasons or individ-ually in pots fertilized at different nutrient supply rates. Leavesat different stages of development, as well as stems and roots atthe end of the growing season, were used for analysis of struc-tural biomass and lignin. In elevated [CO2], lignification ofleaves was slightly retarded compared with structural biomassproduction and showed a strong correlation with the activitiesof ionically, cell-wall-bound peroxidases but not with total sol-uble peroxidases or covalently wall-bound peroxidases. Theeffect of elevated [CO2] on lignin concentration of mature tis-sues was dependent on nutrient supply rate. In leaves and roots,elevated [CO2] increased the lignin concentration in dry massin N-limited plants. In seedlings grown with high nutrient sup-ply, the lignin concentration in dry mass was unaffected or di-minished by elevated [CO2]. Because elevated [CO2] enhancedseedling growth in the high nutrient supply treatments, the totalamount of lignin produced per seedling was higher in thesetreatments. We predict that long-term sequestration of carbonwill increase as long as biomass production is stimulated by el-evated [CO2] and that tissue quality will change depending ondevelopmental stage and nutrient availability.

Keywords: carbon sequestration, climate change, develop-ment, lignin, nitrogen, structural biomass.

IntroductionIn woody plants, structural components such as cellulose andlignins constitute major sinks for reduced carbon. Estimatessuggest that lignins represent about 25% of total terrestrialbiomass (Boudet et al. 1995). Lignins constitute a consider-able carbon fraction not only in stem wood of trees but also inother tissues such as leaves and roots. The incorporation of

lignins renders plant cell walls mechanically rigid, water re-pellent and chemically resistant, thereby determining tissuequality and biodegradability (Monties 1989, Lewis and Yama-moto 1990, Dean and Erikson 1994, Boudet et al. 1995, Polleet al. 1997a). Physiologically, lignification marks an impor-tant developmental stage that is completed by programmedcell death during the formation of structural elements such asfibers, tracheids and vessels. Thus, growth and development aswell as tissue maturation rely on the ability of trees to allocateconsiderable carbon resources to lignin production.

In recent years, our understanding of the enzymatic steps inthe biosynthesis of lignin precursors at the molecular level hasincreased considerably (Boudet 1998, Whetten et al. 1998).Much less is known about the processes that control polymer-ization of lignin in the cell wall, and the interactions betweenthese processes and environmental factors, which may modu-late lignin production at the tissue or whole-plant level.Among environmental factors, the balance between atmo-spheric CO2 concentration ([CO2]) and pedospheric nitrogen(N) supply appears to be critical. Higher lignin concentrationsand lignin/N ratios were found in leaf litter from yellow poplar(Liriodendron tulipifera L.), ash (Fraxinus sp.), birch (Betulasp.), sugar maple (Acer saccharum Marsh.) and Sitka spruce(Picea sitchensis (Bong.) Carrire) grown at elevated [CO2]than at ambient [CO2] (Cotrufo et al. 1994, Boerner andRebbeck 1995). In contrast, in a free-air CO2 enrichment ex-periment with wheat, elevated [CO2] had no apparent effect oneither lignin concentration or its chemical composition (Akinet al. 1995). Similarly, Poorter et al. (1997) found no effect ofelevated [CO2] on tissue lignin concentrations in a range ofplant species, including woody species. In Picea abies (L.)Karst. and Pinus palustris Mill., neither varied N nutrition norelevated [CO2] affected lignin concentrations in stem wood(Httenschwiler et al. 1996, Entry et al. 1998). However, inPinus palustris, significant decreases in lignins were found inlateral roots of trees grown with high N supply (Entry et al.

Tree Physiology 22, 469477 2002 Heron PublishingVictoria, Canada

Lignification in beech (Fagus sylvatica) grown at elevated CO2concentrations: interaction with nutrient availability and leafmaturation

L. BLASCHKE,1 M. FORSTREUTER,2 L. J. SHEPPARD,3 I. K. LEITH,3 M. B. MURRAY3 andA. POLLE4,51 Albert-Ludwigs-Universitt Freiburg, Institut fr Forstbotanik und Baumphysiologie, Am Flughafen 17, 79085 Freiburg, Germany2 Technische Universitt Berlin, Institut fr kologie, Fachgebiet kologie der Gehlze, Knigin-Luise Str. 22, 14195 Berlin, Germany3 Centre for Ecology and Hydrology, Edinburgh, Bush Estate, Penicuik, Midlothian EH26 0QB, Scotland4 Georg-August-Universitt Gttingen, Forstbotanisches Institut, Bsgenweg 2, 37077 Gttingen, Germany5 Author to whom correspondence should be addressed (apolle@gwdg.de)

Received February 26, 2001; accepted October 14, 2001; published online April 1, 2002

1998). Based on these and other findings, Coteaux et al.(1999) concluded that lignin concentration was modified byelevated [CO2], and that the effect was species-specific, butthat no clear positive or negative trend was apparent. At vari-ance with this opinion, ONeill and Norby (1996) suggestedthat lignin/N ratios were significantly affected by environ-mental factors such as light, nutrient availability and the grow-ing conditions of the roots (pots or field). A possible explana-tion of the contrasting observations reported in the literature isthat growth in elevated [CO2] affects ontogeny.

This study was undertaken to investigate this suggestion andto establish a physiological basis for estimating carbon se-questration into lignins in woody plants in future atmospheric[CO2] scenarios. To assess whether changing environmentalconditions affect lignification and cause changes in tissuelignin concentrations, we studied the effects of elevated [CO2],varied nutrient supply rates and growth conditions (pots orsmall stands) on the amounts of lignin and structural biomassin beech (Fagus sylvatica L.) seedlings. To determine the roleof nutrition in lignification, beech seedlings were grown indi-vidually in pots in open-top chambers at ambient or elevated[CO2] with low, medium or high nutrient availability. Previousstudies have shown that these differences in nutrient supply re-sult in high (2.65%), medium (2.05%) and low (1.52%) N con-centrations in foliage of beech seedlings grown at ambient[CO2] (Polle et al. 1997b, Linder and Murray 1998). Seedlingsgrown in elevated [CO2] had N concentrations about 15%lower than seedlings grown in ambient [CO2] (Polle et al.1997b). Elevated [CO2] caused increased growth and biomassproduction in trees with a medium or high nutrient supply, buthad no effect on growth of trees with a low nutrient supply rate(Linder and Murray 1998).

In a second experiment, seedlings were grown directly insoil in microcosms in ambient or elevated [CO2], with otherenvironmental conditions maintained near ambient (Over-dieck and Forstreuter 1995). Seedlings formed small standswith a relatively dense canopy, with leaf area indices of 4.4and 6.2 at ambient and elevated [CO2], respectively (Forstreu-ter 1998). The height and biomass of beech seedlings grown inelevated [CO2] were 58 and 67% greater, respectively, thanthose of seedlings grown in ambient [CO2] (Overdieck andForstreuter 1995, Lee et al. 1998). We used material from thisexperiment to determine lignin concentrations in sun andshade leaves, stems and roots, and to assess the amount oflignin sequestered in above- and belowground biomass. Tocharacterize the influence of elevated [CO2] on leaf develop-ment, time courses of structural biomass accumulation andlignin formation were determined from bud break to leafmaturity in seedlings grown in microcosms. Developmentalchanges in activities of soluble and wall-bound peroxidaseswere also determined and related to lignin production rates.

Materials and methods

Growth of seedlings in microcosms and seedling harvestTwo-year-old beech seedlings (F. sylvatica, provenance Nord-

deutsches Tiefland) were planted in a loamy sand mixture insemi-closed acrylic microcosms at the Institute of Ecology,Berlin, Germany (described in detail by Forstreuter 1995).Climatic conditions in the microcosms tracked ambient condi-tions with the following deviations: air temperature 0.5 C;relative air humidity 15%; and photosynthetic photon fluxdensity 17%. The microcosms were located outside and wereexposed to natural sunlight. Each microcosm covered a groundarea of 0.8 0.8 m2 and had a height of 1.2 m. Two micro-cosms were supplied with ambient air (371 46 mol mol1CO2), whereas the other two microcosms were supplied withCO2-enriched air (701 10 mol mol1 CO2; Forstreuter1995). Seedlings were grown for three consecutive seasonsunder these conditions. The experiment began with 48 seed-lings per microcosm, and the numbers were reduced to 36 and25 in subsequent years (Forstreuter 1995, Overdieck and For-streuter 1995). In fall 1993, the seedlings were harvested. Indi-vidual seedlings were separated into different organs. Leaveswere collected according to canopy layer (Forstreuter 1995).Height growth, leaf area index and biomass of leaves, stemsand roots have been reported by Overdieck and Forstreuter(1995; see also Lee et al. 1998). Oven-dried tissues (85 C)were analyzed for lignin.

To determine the developmental pattern of lignification,leaves were harvested over two consecutive years from beechseedlings exposed for 3 and 4 years to ambient and elevated[CO2] in microcosms as described above. Samples were takenfrom Day 5 up to Day 32 after full expansion of the leaves ofthe first flush. The leaves were frozen at 70 C until assayedfor peroxidase activities. Aliquots of the leaves were oven-dried (40 C) for subsequent analysis.

Growth of potted seedlings in open-top chambersBeech (F. sylvatica, Accession No. 91(439)F, provenanceHungary) seeds were germinated in pots (0.02 m3) in ambientand elevated [CO2]. In the first year, pots were located in tun-nels, and subsequently they were placed in open-top chamberswith a floor area of 7 m2 and a height of 2.3 m (TerrestrialEcology, Edinburgh, U.K.; for a detailed description seeFowler et al. 1989). Four chambers received ambient air(about 355 mol mol1 CO2) and four received air with ele-vated [CO2] (700 80 mol mol1 CO2) by addition of pureCO2 to the air stream as decribed by Murray et al. (1994).Seedlings were repotted each March in compost consisting ofsphagnum peat, 5-mm quartz and sterilized loam (13:4:3, v/v).Seedlings were supplied with one of three Ingestad nutrientsolutions (high = 2 optimum, medium = 0.5 optimum andlow = 0.1 optimum) applied weekly (Ingestad and Lund1986) to provide three nutrient regimes (high N, medium Nand low N). Optimum nutrient solution was taken to be theamount of N solution required to produce 2.7% N in the fo-liage (Linder and Murray 1998). Each chamber containedeight seedlings per nutrient treatment. In August 1994, oneleaf per seedling was harvested from each of the eight cham-bers. Leaves collected from the eight plants per nutrient treat-ment were pooled within each chamber to give four independ-ent replicates per nutrient treatment. Stems were harvested in

470 BLASCHKE, FORSTREUTER, SHEPPARD, LEITH, MURRAY AND POLLE

TREE PHYSIOLOGY VOLUME 22, 2002

January 1995 from five seedlings chosen at random. The sam-ples were oven-dried at 80 C for 3 days and milled. Biomassdata have been reported by Lee et al. (1998).Extraction of cell walls for lignin analysesDry milled plant powder (200 mg) was suspended in 20 ml ofwashing buffer (100 mM K2HPO4/KH2PO4, pH 7.8, 0.5% Tri-ton X-100), stirred slowly for 30 min at room temperature andcentrifuged at 5,500 g for 20 min. The pellet was resuspendedin washing buffer and washed again. Subsequently, the pelletwas washed four times (30 min each time) in 100% MeOH.The resulting pellet consisted mainly of cell walls, i.e., struc-tural biomass (SBM). The SBM pellet was dried (12 h at80 C), weighed, and used to determine lignin content by thethioglycolate method (Bruce and West 1989).Lignin analysesAliquots of 12 mg of SBM pellet (three replicates per sam-ple) were weighed into Eppendorf tubes and mixed with1.5 ml of 2 N HCl and 0.3 ml of thioglycolic acid (adapted af-ter Bruce and West 1989). Samples were incubated at 95 Cfor 4 h with mixing, rapidly cooled on ice and centrifuged for10 min at 15,000 g. The supernatant was discarded and the pel-lets washed three times with 1 ml of distilled water. Pelletswere then incubated with 1 ml of 0.5 N NaOH for 18 h on ashaker at room temperature. The suspension was centrifugedfor 10 min at 15,000 g and transferred to a 2-ml Eppendorftube. The pellet was resuspended in 0.5 ml of 0.5 N NaOH,mixed vigorously and centrifuged. The resulting supernatantwas combined with the first, alkaline supernatant and mixedwith 0.3 ml of concentrated HCl. Samples were incubated for4 h at 4 C to precipitate lignothioglycolate derivates. Sampleswere centrifuged, the supernatant discarded, and the pelletsolubilized in 1 ml of 0.5 N NaOH. The absorbance of the re-sulting solution was measured at 280 nm. Standard curves,generated with increasing amounts (0.5 to 2.5 mg) of commer-cial alkaline lignin (Aldrich) subjected to the same procedureas above, were used for calculation of lignin concentration.

Differential extraction and determination of peroxidaseactivities

Soluble peroxidase Frozen leaf material was powdered inliquid nitrogen and 300 mg of the frozen powder was trans-ferred to 10 ml of extraction buffer (50 mM 2-N-morpholino-ethansulfonic acid, pH 6.5, 1% Triton X-100, 1% soluble poly-vinylpyrolidone K-30) and mixed twice for 30 s each time witha blender at maximum speed. The mixture was stirred for 1 h onice and centrifuged at 5,000 g for 10 min at 4 C. The super-natant (S1) was kept on ice. The pellet was washed three timeswith 20 ml of extraction buffer (S2, S3, S4). The pellet was des-ignated CW pellet and kept for further extraction.

Analysis of peroxidase (POD) activities revealed that S1,S2, S3 and S4 contained 78, 12, 10 and 0% of soluble POD ac-tivities, respectively. For routine analysis, 2 ml of S1 and 4 mlof each of S2, S3 and S4 were combined, resulting in a volumeof 14 ml. This volume was combined with 20 ml of ice-coldacetone (20 C), mixed, incubated for 5 min and then centri-

fuged at 5,000 g for 10 min at 4 C. The supernatant was dis-carded and the pellet was briefly dried in air. The pellet wasdissolved in 2 ml of 50 mM 2-N-morpholino-ethansulfonicacid, pH 6.5 and used for determination of total soluble POD.

Ionically wall-bound POD The CW pellet was mixed with5 ml of saline buffer (50 mM 2-N-morpholino-ethansulfonicacid, pH 6.5, 1% Triton X-100, 1 M NaCl) and stirred for 16 hon ice. The mixture was centrifuged at 5,000 g for 10 min at4 C. The saline-treated CW pellet was kept for further analy-sis. The supernatant was passed over Sephadex G-25 (PD-10column, Pharmacia, Freiburg, Germany) and eluted with50 mM 2-N-morpholino-ethansulfonic acid, pH 6.5. The ex-tract was used for determination of ionically bound POD.

Covalently wall-bound POD The saline-treated CW pelletwas washed three times for 30 min with 20 ml of washingbuffer (100 mM Tris-HCl, pH 7.8). The final pellet, corre-sponding to about 60 mg of cell walls, was suspended in a finalvolume of 5 ml of washing buffer and used for determination ofcovalently wall-bound POD activities (McDougall 1993).Determination of POD activities Soluble and ionicallybound POD activities were determined according to Pedreno etal. (1989), employing up to 50 l of extract per assay andconiferyl alcohol as substrate (= 260 nm). The POD activitieswere calculated based on an extinction coefficient of 7500 lmol1 cm1 (Takahama 1993). Covalently wall-bound POD ac-tivities were determined with up to 70 l of cell wall suspensionand coniferyl alcohol as substrate (Otter and Polle 1994). Ab-sorption changes were corrected for unspecific changes result-ing from sedimentation. Each sample was analyzed with fivedifferent extract volumes. Individual POD activities were cal-culated by linear regression analysis.

Determination of contamination To determine if nonspecificbinding of symplastic components occurred during the extrac-tion procedure, malate dehydrogenase activity was assayed asa symplastic marker in the fractions used for determination oftotal soluble, ionically wall-bound, and covalently wall-boundPOD activites. Malate dehydrogenase activity was assayed asdescribed by Weimar and Rothe (1986).

Statistical analysisResults were evaluated by multivariate analysis of variancefollowed by a multiple range test using the STATGRAPHICssoftware package (Manugistics, Rockville, MD). Where ap-propriate, the data were analyzed by linear regression. The an-alytical error of individual samples was generally below 5%.Values presented in the tables and figures are means SD ofindependent chamber replicates (n = 4) or individual plants(n = 810).

Results

Structural biomass and lignin accumulation during foliarmaturation in relation to peroxidase activitiesFor two consecutive years we determined the relative increase

TREE PHYSIOLOGY ONLINE at http://heronpublishing.com

MODULATION OF LIGNIN PRODUCTION BY CO2 AND MINERAL NUTRIENTS 471

in dry mass, the accumulation of structural biomass and the in-crease in lignin concentrations during leaf maturation in beechseedlings grown in microcosms in ambient or elevated [CO2](Figure 1). The developmental pattern of dry mass accumula-tion was different between the 2 years and was affected by ele-vated [CO2] (Figures 1A and 1B). The relative increase in leafdry mass was significantly higher in elevated [CO2] than inambient [CO2] (Kendalls P = 0.0018 for the whole data set)(Figures 1A and 1B). The rate of dry mass production was de-layed by about 2 weeks in 1996 compared with 1995 (Figures1A and 1B).

In mature leaves, structural biomass, i.e., cell walls, repre-sented between 70 and 75% of the total dry mass (Figures 1Cand 1D). During foliar development, structural biomass pro-duction showed a pattern similar to that observed for dry massproduction (cf. Figures 1A and 1C with 1B and 1D). However,in contrast to leaf dry mass production, leaf structural dry mass

production was lower in seedlings grown in elevated [CO2]than in ambient [CO2] (Kendalls P = 0.0035, Figures 1C and1D).

After bud break, the rate of lignin production in leaves waslower in elevated [CO2] than in ambient [CO2] (Figures 1E and1F), and the decrease was more pronounced in 1995 than in1996. When all data were analyzed together, leaves in the ele-vated [CO2] treatment contained less lignin than leaves in theambient [CO2] treatment (Kendalls P = 0.0018, Figures 1Eand 1F). Because lignin concentrations were expressed on thebasis of structural biomass, the observed decrease in lignifica-tion was not caused by a decrease in structural biomass pro-duction.

To determine whether the decrease in lignin production inresponse to elevated [CO2] was associated with CO2-inducedchanges in the activities of PODs, soluble, ionically wall-bound and covalently wall-bound POD activities were assayedin beech leaves during different stages of development. Thedifferential fractionation of wall-bound PODs compared withtotal soluble PODs was highly specific because malate dehy-drogenase activity, which was used as a contamination markerfor symplastic components, was more than 66-fold depleted inthe fraction containing ionically bound components comparedwith POD, and was not detected in the cell wall fraction (Ta-ble 1). The POD activities in the different fractions showedsignificant seasonal variation with higher activities in youngleaves than in mature leaves (not shown). Rank correlationanalysis revealed no effects of elevated [CO2] on the fractionof total soluble PODs (P = 0.8348); however, wall-bound PODactivities were generally slightly lower in leaves from the ele-vated [CO2] treatment than in leaves from the ambient [CO2]treatment (P = 0.0606 for the ionically wall-bound PODs, P =0.0478 for the covalently wall-bound PODs). Correlations be-tween lignification rate calculated from the data in Figures 1Eand 1F and the various POD activities indicated that solublePOD activities were unrelated to lignification rates (Fig-ure 2A). The highest POD activities and the closest correlationwith lignification rates were observed for the ionically wall-bound PODs (Figure 2B). Covalently wall-bound PODsshowed intermediate behavior (Figure 2C).Structural biomass and lignin at the whole-plant levelTo determine if elevated [CO2] affected tissue lignin concen-

472 BLASCHKE, FORSTREUTER, SHEPPARD, LEITH, MURRAY AND POLLE

TREE PHYSIOLOGY VOLUME 22, 2002

Figure 1. Accumulation of relative dry mass (SBM) (A, B), structuraldry mass (C, D) and lignins (E, F) and their calculated productionrates during maturation of beech (Fagus sylvatica) leaves in two con-secutive years. Seedlings were grown in soil in microcosms in ambi-ent (, ) or elevated [CO2] (, - - -). Symbols refer to left y-axis,lines to right y-axis. In each year, sampling started 5 days after full fo-liar expansion of the first flush. Top-layer leaves from a canopy ofbeech seedlings were analyzed (n = 4, SD).

Table 1. Relative peroxidase (POD) and malate dehydrogenase(MDH) activities in different subcellular fractions of beech (Fagussylvatica) leaves. Mature leaves obtained from microcosms were ex-tracted as described in Materials and methods. Peroxidase and MDHactivities in total soluble extracts were set as 100% (n = 3, SD). Ab-breviation: ND = not detected.

Fraction POD (%) MDH (%) MDH/POD

Total soluble extract 100 5 100 7 1Ionically soluble extract 231 38 3.5 0.4 0.015Cell wall 28.3 7.2 ND 0

trations and the overall amount of lignin produced per seed-ling, beech seedlings grown in microcosms in elevated orambient [CO2] for 3 years were harvested and analyzed fortotal biomass, structural biomass and lignin (Table 2) in sunleaves, shade leaves, stems and roots (Overdieck and For-streuter 1995).

The relative amounts of structural biomass isolated from thedifferent organs were slightly increased by elevated [CO2] inleaves (+10%) and roots (+7%) but not in stems (Table 2).Structural biomass of leaves contained lignin concentrationssimilar to those in roots (Table 2). The highest lignin concen-trations were found in stem cell walls (Table 2). Growth in ele-vated [CO2] caused a significant decrease in lignin concentra-tions in stems, irrespective of whether the data were expressedon a dry matter or a structural biomass basis (Table 2). Thelignin concentration of roots was unaffected by elevated[CO2]. Elevated [CO2] tended to decrease lignin concentrationof sun leaves and increase lignin concentration of shade leaves(Table 2).

Although the effect of elevated [CO2] on lignin concentra-tions per unit dry mass or structural biomass was modest, theoverall lignin content per seedling increased considerably inresponse to CO2 enrichment (Figure 3) as a result of a strongstimulation of photosynthesis and growth that was maintainedthrough three consecutive growth periods (Forstreuter 1995,Overdieck and Forstreuter 1995). Compared with seedlingsgrown in ambient [CO2], elevated [CO2] increased whole-plant lignin contents by factors of 1.64 (roots), 1.69 (stems)and 1.74 (leaves).

Lignin in potted beech seedlings grown with differentnutrient supply ratesLignin concentrations and structural biomass were investi-gated in stems and leaves of beech seedlings grown in pots for3 years at elevated or ambient [CO2] and with different nutri-ent supply rates. Stem structural biomass was unaffected bygrowth [CO2] or N nutrition and amounted to 0.721 0.019 gg1 of dry mass (Table 3). By contrast, leaves of seedlingsgrown with high N supply contained 10% more structural bio-mass than leaves of seedlings grown with medium or low Nsupply (0.719 0.032 versus 0.653 0.010 g g1 of dry mass,Table 3). Elevated [CO2] had no effect on the structural bio-mass of leaves (Table 3).

Both nutrient supply and elevated [CO2] significantly af-fected lignin concentrations per unit of structural biomass inleaves (Figure 4A, Table 3). The lowest lignin concentrationswere found in leaves of seedlings grown with a high nutrientsupply rate in elevated [CO2] (Figure 4A). However, differentnutrient supply rates did not affect lignin concentration perunit of structural biomass in stems, whereas elevated [CO2] re-sulted in slightly elevated lignin concentrations comparedwith ambient [CO2] (Figure 4B). Lignin concentrations instem cell walls were about 1.55 times higher than in leaf cellwalls.

Seedling N concentration was significantly affected bygrowth [CO2] and by nutrient supply rate (Polle et al. 1997b,Linder and Murray 1998). To investigate the relationship be-tween lignin concentration and N concentration, we plottedlignin concentration against tissue N concentration based ondata from both experiments (soil-grown plants in microcosmsand potted plants in open-top chambers). Lignin concentrationwas highly correlated with N concentration in leaves and rootsbut not in stems (Figure 5).

DiscussionIn a range of plant species, elevated [CO2] accelerates ontog-eny (e.g., rice (Oryza sativa L.): Jitla et al. 1997; tobacco(Nicotiana tabacum L.): Miller et al. 1997; oak (Quercusrubra L.): Anderson and Tomlinson 1998, Tomlinson and An-derson 1998; cherry (Prunus avium L.): Centritto et al. 1999).Therefore, in response to atmospheric CO2 enrichmment, wepredicted that foliar maturation, measured as an accumulationof structural components including lignins, would be acceler-ated, especially when there was an increased supply of re-duced carbon. However, in beech, foliar growth phenology is

TREE PHYSIOLOGY ONLINE at http://heronpublishing.com

MODULATION OF LIGNIN PRODUCTION BY CO2 AND MINERAL NUTRIENTS 473

Figure 2. Activities of total soluble (A), ionically wall-bound (B) andcovalently wall-bound peroxidases (PODs) (C) in beech (Fagus syl-vatica) leaves in relation to lignin accumulation rates. Beech seed-lings were grown in soil in microcosms. The POD activities wereassayed in subcellular fractions of the leaves analyzed for lignin (Fig-ure 1). Symbols: POD activities in leaves from trees grown at ambient() or elevated [CO2] (). Lignin production rates were calculatedfrom the data shown in Figure 1.

unaffected by elevated [CO2] (Murray and Ceulemans 1998),and we found that developmental lignification was slightly de-layed (Figure 1). It has been shown that repression of ligninbiosynthesis promotes growth in transgenic trees (Hu et al.1998). Murray and Ceulemans (1998) observed that the beechtrees studied here form larger leaves when grown in elevated[CO2] than in ambient [CO2], a response that may be associ-ated with our finding that elevated [CO2] delayed developmen-tal lignification in beech leaves (Figure 1).

Elevated [CO2] decreased lignin concentrations of matureleaves if the seedlings were not nutrient-limited (Figure 1, Ta-ble 2, Figure 4). In contrast to lignins, soluble phenolic con-centrations have frequently been observed to increase inresponse to elevated [CO2] (Penuelas et al. 1996, Poorter et al.

1997, Entry et al. 1998, Gebauer et al. 1998), but not always(Heyworth et al. 1998). In agreement with our findings, theCO2 response of secondary carbon-based components is tis-sue-specific and depends on soil fertility (Entry et al. 1998,Gebauer et al. 1998). We have previously observed that, inbeech seedlings grown at ambient [CO2], the carbon incorpo-rated into lignins during leaf development is predominatelyderived from internal storage compounds (65%) with a smallerfraction (35%) derived from newly assimilated carbon (Dyck-mans et al. 2000). In elevated [CO2], this balance is shifted to-ward an increased use of new assimilates for lignin production(Dyckmans et al. 2000). To explain our finding of lower ligninsynthesis (35% in 1995 and 15% in 1996, derived from Fig-ure 1) in young expanding leaves (5 days after bud break) ofbeech seedlings grown in elevated [CO2] compared with ambi-ent [CO2], we postulate that increased growth in response toelevated [CO2] resulted in a transient shortage of carbon forlignin production.

An assessment of the budget of above- and belowgroundlignin contents of beech plants grown in ambient and elevated[CO2] is presented in Figure 3. We note that these values areonly rough estimates of the absolute amounts of lignin presentbecause of difficulties inherent in the methods for lignin deter-mination. The thioglycolic acid method probably underesti-

474 BLASCHKE, FORSTREUTER, SHEPPARD, LEITH, MURRAY AND POLLE

TREE PHYSIOLOGY VOLUME 22, 2002

Table 2. Effect of elevated CO2 concentration [CO2] on structural biomass and lignin in different tissues of beech (Fagus sylvatica). Seedlingswere grown for 3 years in soil in microcosms in ambient or elevated [CO2]. Abbreviations: DM = dry mass; and SBM = structural biomass. Valuesare means of 8 to 10 individual plants ( SD). Values in columns followed by different letters are significantly different at P 0.05.

Tissue [CO2] (mol mol1) Structural biomass (mg g1 DM) Lignin (mg g1 SBM) Lignin (mg g1 DM)

Sun leaves 371 583 12 a 56.2 8.3 ab 32.9 6.2 ab701 612 16 ab 50.2 2.5 a 30.7 1.7 a

Shade leaves 371 601 44 a 54.3 6.1 ab 32.5 3.5 ab701 643 88 b 58.4 10.4 b 36.7 2.8 a

Stems 371 835 16 e 78.4 10.5 d 65.3 7.6 e701 832 17 e 67.3 6.6 c 55.6 5.9 d

Roots 371 738 8 c 53.0 7.6 ab 39.2 5.8 c701 788 21 d 52.9 7.9 ab 41.7 6.5 c

Figure 3. Estimated lignin content of roots, stems and leaves of beech(Fagus sylvatica) seedlings grown for 3 years in ambient (371 molmol1) or elevated [CO2] (701 mol mol1) in soil in microcosms.Plant biomass data for the same experiment were taken from Over-dieck and Forstreuter (1995) and lignin data are from Table 2.

Table 3. Calculated P-values for the effects of elevated CO2 concen-tration ([CO2]), nitrogen supply (N) and their interaction ([CO2] N)on structural biomass (SBM) and lignin concentrations in stems andleaves of beech (Fagus sylvatica) seedlings. The seedlings weregrown in pots in open-top chambers. Data refer to the experimentshown in Figure 4.

Material Effect SBM Lignin Lignin(mg g1 DM) (mg g1 SBM) (mg g1 DM)

Leaves [CO2] 0.2154 0.0372 0.0718N 0.0017 0.0000 0.0013[CO2] N 0.0905 0.0659 0.2596

Stems [CO2] 0.1570 0.0371 0.1143N 0.5092 0.6801 0.2682[CO2] N 0.3627 0.4948 0.7770

mates lignin concentrations because it relies on the specificcleavage of -O-4-side bonds (Boudet et al. 1995). A furtherdifficulty is associated with the calibration of the spectropho-tometric method. The use of different standards, e.g., sprucelignin or coniferyl alcohol, results in differences in apparentlignin concentrations (Blaschke 1998). Gravimetric methodsyielded higher lignin concentrations in beech than those re-ported here (beech petioles about 10% and stem wood about22%; Brauns and Brauns 1960, Miksche and Yasuda 1977);however, gravimetric methods are unsuitable for small sam-ples such as those used in our study, and tend to overestimate

lignin because of contamination with residual protein and sug-ars (Monties 1989, Reeves 1993). We found that ligninsamples from beech leaf and stem tissue samples, which wereprepared for gravimetric determinations according to themethod of Van Soest (1963), contained 5 and 2% N, respec-tively (Dyckmans et al. 2002). This finding indicates that grav-imetric methods, though frequently used, are unreliable forcomparative studies of lignin concentrations in different planttissues. Because of these difficulties, estimates of carbon se-questered in lignins should be used with caution. Baumgartenet al. (2000), who also used the thioglycolic acid method,found lignin concentrations in beech leaves similar to those re-ported in Figures 1 and 3 and Table 2 (cf. Dyckmans et al.2000). Despite uncertainty about absolute lignin concentra-tions in plant tissues, our data showed that elevated [CO2] re-sulted in an overall increase by a factor of about 1.6 in theamount of lignin per seedling, which corresponds with thestimulation of growth (Figure 3).

ONeill and Norby (1996) reported that the lignin/N ratioincreases in plants grown in elevated [CO2]. As in other stud-ies (Poorter et al. 1997, Medlyn et al. 1999), elevated [CO2] inour study caused substantial decreases in tissue N concentra-tions in beech seedlings, especially at low nutrient supply rates(8 to 16%; Polle et al. 1997b), but increases in lignin con-centrations were not consistently observed (+10 to 10%; Ta-ble 2, Figure 4). Under our experimental conditions, therewere no increases in the lignin/N ratio in leaves of seedlingsgrown with a high nutrient supply, but the ratio increased inseedlings grown with a low nutrient supply. Because thedegree of lignification is considered an important factor in tis-sue degradability, further studies are necessary to determinewhether fluctuations in the lignin/N ratio, as a result ofchanges in N rather than lignin, affect litter decompositionrates.

We found a significant correlation between lignin and Nconcentrations in leaves and roots (Figure 5). Because phenyl-propanoid subunits comprising lignins do not contain N, onemight assume that an increasing availability of reduced carbonin plants grown in elevated [CO2] would enhance lignification.However, because lignin is a metabolic endproduct and thus isnot reused, plants might preferentially invest surplus carbon ingrowth and recyclable resources such as sugars and starch, es-pecially in tissues with a relatively high turnover comparedwith stems. In beech seedlings, the enhanced supply of re-duced carbon was used for growth (Overdieck and Forstreuter1995, Lee et al. 1998) and was not preferentially sequesteredin metabolically inert compounds when N was not severelylimiting (Figures 3 and 4). The lignin concentration increasedonly when tissue N concentration was low (Figure 5). Theseobservations suggest that lignins constitute a pool for seques-tration of additional carbon per unit of plant material when nu-trients are severely limited. Nevertheless, carbon sequestrationinto lignins will likely increase as long as elevated [CO2]drives increased biomass production.

We speculate that, under conditions of nutrient-restrictedgrowth and, presumably, overflowing pools of carbohydrates,metabolism shifts to the production of lignin as an additional

TREE PHYSIOLOGY ONLINE at http://heronpublishing.com

MODULATION OF LIGNIN PRODUCTION BY CO2 AND MINERAL NUTRIENTS 475

Figure 4. Lignin concentrations in the structural biomass of leaves (A)and stems (B) of beech (Fagus sylvatica) seedlings grown in ambient(black bars) and elevated [CO2] (white bars) at high, medium or lownutrient supply rates for 3 years. The seedlings were grown in pots inopen-top chambers. Leaves were collected in the third week of Au-gust 1994 and stems in the second week of January 1995. Data aremeans SD (n = 4).

Figure 5. Dependency of lignin (% of DM) on nitrogen concentration(% of DM) in stems (S), roots (R) and leaves (L) of beech seedlingsgrown with different nutrient supply rates in elevated or ambient[CO2] (Figure 4, Table 2). Symbols: = plants grown individually inpots; = plants grown in microcosms.

carbon sink. However, this scenario is difficult to reconcilewith lignin biosynthesis, which is linked to amino acid metab-olism through phenylalanine as the precursor for phenylpro-panoid synthesis (Whetten and Sederoff 1995). The physio-logical factors regulating lignification are poorly understood(Polle et al. 1997a). It has been proposed that cell-wall-associ-ated POD activities limit lignification (Lagrimini et al. 1993).However, our data show that, despite a close correlation be-tween lignification rates and cell-wall-associated PODs, the invitro activities of these enzymes were five orders of magnitudehigher than those required for lignin formation in situ (Fig-ure 2). This suggests that other factors, e.g., substrate forma-tion, limited lignification. To understand and predict potentialchanges in the degree of lignification in response to elevated[CO2], it will be necessary to elucidate the steps controllingfluxes between amino acid synthesis for protein productionand those for lignin production.

Overall, our results suggest that CO2 availability does notdirectly affect lignin concentrations, but affects them indi-rectly through effects on or an interaction with N supply andgrowth. In seedlings, elevated [CO2] reduced lignin concentra-tion on a dry mass basis, indicating diminished wood qualityin a CO2-enriched atmosphere (Table 2). Foliar lignin concen-trations increased significantly with decreasing nutrient avail-ability (Figure 4, Table 3), but it is not known if these changesin the degree of lignification were caused by changes in ana-tomical structures, e.g., a shift in the proportion of cell typeswith different lignin concentrations such as parenchyma andscelerenchyma cells. The observation that elevated [CO2] de-layed the accumulation of structural compounds, includinglignins, in leaves suggests that the juvenile phase was slightlyprolonged.

Acknowledgments

Technical assistance was provided by C. Fliegauf. Financial supportfrom the German National Science foundation (DFG-Schwerpunkt:Ov25/1-1, Po362/3-1), the European Union (ECOCRAFT EV5V-CT92-0127; EG-EV 50 CT92-0127) and a travel grant to A.P. underCOST 614 are gratefully acknowledged.

References

Akin, D.E., L.L. Rigsby, G.R. Gamble, W.H. Morrison, III, B.A.Kimball, P.J. Pinter, Jr., G.W. Wall, R.L. Garcia and R.L. LaMorte.1995. Biodegradation of plant cell walls, wall carbohydrates, andwall aromatics in wheat grown in ambient or enriched CO2 concen-trations. J. Sci. Food Agric. 67:399406.

Anderson, P. and P. Tomlinson. 1998. Ontogeny affects response ofnorthern red oak seedlings to elevated CO2 and water stress. I. Car-bon assimilation and biomass production. New Phytol. 140:477491.

Baumgarten, M., H. Werner, K.H. Hberle, L.D. Emberson, P. Fabianand R. Matyssek. 2000. Seasonal ozone response of mature beechtrees (Fagus sylvatica) at high altitude in the Bavarian forest (Ger-many) in comparison with young beech grown in the field and inphytotrons. Environ. Pollut. 109:431442.

Blaschke, L. 1998. Untersuchungen zu Photosynthese, Lignifizierungund Wachstum unter erhhtem CO2. Ph.D. Diss., Univ. Freiburg,143 p.

Boerner, R.E.J. and J. Rebbeck. 1995. Decomposition and nitrogenrelease from leaves of three hardwood species grown under ele-vated O3 and/or CO2. Plant Soil 170:149157.

Boudet, A. 1998. A new view of lignification. Trends Plant Sci. 3:6771.

Boudet, A.M., C. Lapierre and J. Grima-Pettenati. 1995. Biochemis-try and molecular biology of lignification. New Phytol. 129:203236.

Brauns, F.E. and D.A. Brauns. 1960. The chemistry of lignin. Aca-demic Press, New York, 138 p.

Bruce, R.J. and C.A. West. 1989. Elicitation of lignin biosynthesisand isoperoxidase activity by pectic fragments in suspension cul-tures of castor bean. Plant Physiol. 91:889897.

Centritto, M., H.S.J. Lee and P.G. Jarvis. 1999. Interactive effects ofelevated CO2 and drought on cherry (Prunus avium) seedlings. I.Growth, whole-plant water use efficiency and water loss. NewPhytol. 141:129140.

Cotrufo, M.F., P. Ineson and A.P. Rowland. 1994. Decomposition oftree leaf litters grown under elevated CO2: effect of litter quality.Plant Soil 163:121130.

Coteaux, M.M., C. Kurz, P. Bottner and A. Raschi. 1999. Influenceof increased atmospheric CO2 concentrations on quality of plantmaterial and litter decomposition. Tree Physiol. 19:301311.

Dean, J.F.D. and K.E. Erikson. 1994. Laccase and the deposition oflignin in vascular plants. Holzforschung 48:2133.

Dyckmans, J., H. Flessa, A. Polle and F. Beese. 2000. The effect of el-evated CO2 on uptake and allocation of 13C and 15N in beech(Fagus sylvatica L.) during leafing. Plant Biol. 2:113120.

Dyckmans, J., H. Flessa, K. Brinkmann, C. Mai and A. Polle. 2002.Carbon and nitrogen dynamics in acid detergent fibre lignins ofbeech (Fagus sylvatica L.) during the growth phase. Plant Cell En-viron. In press.

Entry, J.A., G.B. Runion, S.A. Prior, R. Mitchell and H. Rogers. 1998.Influence of CO2 enrichment and nitrogen fertilization on tissuechemistry and carbon allocation in longleaf pine seedlings. PlantSoil 200:311.

Forstreuter, M. 1995. Bestandesstruktur und Netto-Photosynthesevon jungen Buchen (Fagus sylvatica L.) unter erhhter CO2 Kon-zentration. Verh. Ges. Oekol. 24:283292.

Forstreuter, M. 1998. What can we learn from microcosms? In Euro-pean Forests and Global Change: the Likely Impacts of Rising CO2and Temperature. Ed. P.G. Jarvis. Cambridge University Press,Cambridge, pp 274292.

Fowler, D., J.N. Cape, J.D. Deans, I.D. Leith, M.B. Murray, R.I.Smith, L.J. Sheppard and M.H. Unsworth. 1989. Effects of acidmist on the frost hardiness of red spruce seedlings. New Phytol.113:321335.

Gebauer, R., B. Strain and J. Reynolds. 1998. The effect of elevatedCO2 and N availability on tissue concentrations and whole plantpools of carbon-based secondary compounds in loblolly pine(Pinus taeda). Oecologia 113:2936.

Httenschwiler, S., F.H. Schweingruber and C. Krner. 1996. Treering responses to elevated CO2 and increased N deposition in Piceaabies. Plant Cell Environ. 19:13691378.

Heyworth, C.J., G.R. Iason, V. Temperton, P.G. Jarvis and A.J.Duncan. 1998. The effect of elevated CO2 concentration and nutri-ent supply on carbon based secondary metabolites in Pinus syl-vestris L. Oecologia 115:344350.

Hu, W.-J., S.A. Harding, J. Lung, J.L. Popko, J. Ralph, D.D. Stokke,C.-J. Tsai and V.L. Chiang. 1999. Repression of lignin biosynthesispromotes cellulose accumulation and growth in transgenic trees.Nature Biotechnol. 17:808812.

476 BLASCHKE, FORSTREUTER, SHEPPARD, LEITH, MURRAY AND POLLE

TREE PHYSIOLOGY VOLUME 22, 2002

Ingestad, T. and A.B. Lund. 1986. Theory and techniques for steadystate mineral nutrition and growth of plants. Scand. J. For. Res.1:439453.

Jitla, D., G. Rogers, S. Seneweera, A. Basra, R. Oldfield and J. Con-roy. 1997. Accelerated early growth of rice at elevated CO2. Is it re-lated to developmental changes in the shoot apex? Plant Physiol.115:1522.

Lagrimini, L.M., J. Vaughn, W.A. Erb and S.A. Miller. 1993.Peroxidase overproduction in tomato: wound-induced polyphenoldeposition and disease resistance. HortScience 28:218221.

Lee, H.S.J., D. Overdieck and P.G. Jarvis. 1998. Biomass, growth andcarbon allocation. In European Forests and Global Change: TheLikely Impacts of Rising CO2 and Temperature. Ed. P.G. Jarvis.Cambridge University Press, Cambridge, pp 126191.

Lewis, N.G. and E. Yamamoto. 1990. Lignin: occurence, biogenesisand biodegradation. Annu. Rev. Plant Physiol. Plant Mol. Biol.41:455496.

Linder, S. and M. Murray. 1998. Do elevated CO2 concentrations andnutrients interact? In European Forests and Global Change: TheLikely Impacts of Rising CO2 and Temperature. Ed. P.G. Jarvis.Cambridge University Press, Cambridge, pp 215235.

McDougall, G.J. 1993. Solubilization of wall-bound peroxidases bylimited proteolysis. Phytochemistry 33:765767.

Medlyn, B.E., F.W. Badeck, D.G.G. De Pury, et al. 1999. Effects of el-evated [CO2] on photosynthesis in European forest species: ameta-analysis of model parameters. Plant Cell Environ. 22:14751495.

Miksche, G.E. and S. Yasuda. 1977. ber die Lignine der Nadeln undBltter einiger Angiospermen und Gymnospermen. Holzforschung31:5759.

Miller, A., C.-H. Tsai, D. Hemphill, M. Endres, S. Rodermel andM. Spalding. 1997. Elevated CO2 effects during leaf ontogeny. Anew perspective on acclimation. Plant Physiol. 115:11951200.

Monties, B. 1989. Lignins. In Methods in Plant Biochemistry. Eds.P.M. Dey and J.B. Harborne. Academic Press, New York, pp113157.

Murray, M.B. and R. Ceulemans. 1998. Will tree foliage be large andlive longer? In European Forests and Global Change: The LikelyImpacts of Rising CO2 and Temperature. Ed. P.G. Jarvis. Cam-bridge University Press, Cambridge, pp 94125.

Murray, M.B., I.R. Smith, I.D. Leith, D. Fowler, H.S.J Lee, A.D.Friend and P.G. Jarvis. 1994. Effects of elevated CO2, nutrition andclimatic warming on bud phenology in Sitka spruce (Picea sitch-ensis) and their impact on the risk of frost damage. Tree Physiol.14:691706.

ONeill, E. and R.J. Norby. 1996. Litter quality and decompositionrates of foliar litter produced under CO2 enrichment. In TerrestrialEcosystem Response to Elevated Carbon Dioxide. Eds. G. Kochand H. Mooney. Academic Press, New York, pp 87103.

Otter, T. and A. Polle. 1994. The influence of apoplastic ascorbate onthe activities of cell wall-associated peroxidase and NADH oxidaseactivities in needles of Norway spruce (Picea abies L.) needles.Plant Cell Physiol. 35:12311238.

Overdieck, D. and M. Forstreuter. 1995. Stoffproduktion jungerBuchen (Fagus sylvatica L.) bei erhhtem CO2 Angebot. Verh.Ges. Oekol. 24:323330.

Pedreno, M.A., A. Ros Barcelo, F. Sabater and R. Munoz. 1989. Con-trol by pH of cell wall peroxidase activity involved in lignification.Plant Cell Physiol. 30:237241.

Penuelas, J., M. Estiarte, B. Kimball, et al. 1996. Variety of responsesof plant phenolic concentration to CO2 enrichment. J. Exp. Bot. 47:14531467.

Polle, A., T. Otter and H. Sandermann. 1997a. Biochemistry andphysiology of lignin synthesis. In The Physiology of Trees. Eds. H.Rennenberg, W. Eschrich and H. Ziegler. Backhuys Publishers,Leiden, pp 455475.

Polle, A., M. Eiblmeier, L.J. Sheppard and M.B. Murray. 1997b. Re-sponses of antioxidative enzymes to elevated CO2 in leaves ofbeech (Fagus sylvatica L.) seedlings grown under a range of nutri-ent regimes. Plant Cell Environ. 20:13171321.

Poorter, H., Y. Van Berkel, R. Baxter, et al. 1997. The effect of ele-vated CO2 on the chemical composition and construction costs ofleaves of 27 C3 species. Plant Cell Environ. 20:472482.

Reeves, J.B. 1993. Chemical studies on the composition of fiber frac-tions and lignin determination residues. J. Dairy Sci. 76:120128.

Takahama, U. 1993. Regulation of peroxidase-dependent oxidation ofphenolic by ascorbatedifferent effects of ascorbic acid on the ox-idation of coniferyl alcohol by apoplastic soluble and cell wall-bound peroxidases from epicotyls of Vigna angularis. Plant CellPhysiol. 34:809817.

Tomlinson, P. and P. Anderson. 1998. Ontogeny affects response ofnorthern red oak seedlings to elevated CO2 and water stress. II: Re-cent photosynthate distribution and growth. New Phytol. 140:493504.

Van Soest, P.J. 1963. Use of detergents in the analysis of fibrous feeds.II. A rapid method for the determination of fibre and lignin. J.Assoc. Off. Anal. Chem. 46:829835.

Weimar, M. and G.M. Rothe. 1986. Preparation of extracts from ma-ture spruce needles for enzymatic analysis. Physiol. Plant. 69:692698.

Whetten, R. and R. Sederoff. 1995. Lignin biosynthesis. Plant Cell 7:10011013.

Whetten, R.W., J.J. MacKay and R.R. Sederoff. 1998. Recent ad-vances in understanding lignin biosynthesis. Annu. Rev. PlantPhysiol. Plant Mol. Biol. 49:585609.

TREE PHYSIOLOGY ONLINE at http://heronpublishing.com

MODULATION OF LIGNIN PRODUCTION BY CO2 AND MINERAL NUTRIENTS 477