Microbial response to exudates in the rhizosphere of young beech trees (Fagus sylvatica L.) after dormancy

Soil Biology & Biochemistry 41 (2009) 1976–1985

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Microbial response to exudates in the rhizosphere of young beech trees (Fagus sylvatica L.) after dormancy J. Esperschu¨tz a, b, F. Buegger b, J.B. Winkler c, J.C. Munch a, M. Schloter b, *,1, A. Gattinger a, d,1 a

Technical University Munich, Chair of Soil Ecology, Center of Life and Food Sciences Weihenstephan, Ingolstaedter Landstr. 1, D-85764 Neuherberg, Germany ¨ nchen, GmbH, German Research Center for Environmental Health, Institute of Soil Ecology, Department of Terrestrial Ecogenetics, Helmholtz Zentrum Mu Ingolstaedter Landstr. 1, D-85764 Neuherberg, Germany c ¨ nchen, GmbH, German Research Center for Environmental Health, Institute of Soil Ecology, Department of Environmental Engeneering, Helmholtz Zentrum Mu Ingolstaedter Landstr. 1, D-85764 Neuherberg, Germany d Geohumus International GmbH, Carl-Benz-Str. 1; D-60386 Frankfurt, Germany b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 17 February 2009 Received in revised form 28 June 2009 Accepted 5 July 2009 Available online 16 July 2009

Plants act as an important link between atmosphere and soil: CO2 is transformed into carbohydrates by photosynthesis. These assimilates are distributed within the plant and translocated via roots into the rhizosphere and soil microorganisms. In this study, 3 year old European beech trees (Fagus sylvatica L.) were exposed after the chilling period to an enriched 13C–CO2 atmosphere (d13C ¼ 60& – 80&) at the time point when leaves development started. Temporal dynamics of assimilated carbon distribution in different plant parts, as well as into dissolved organic carbon and microbial communities in the rhizosphere and bulk soil have been investigated for a 20 days period. Photosynthetically fixed carbon could be traced into plant tissue, dissolved organic carbon and total microbial biomass, where it was utilized by different microbial communities. Due to carbon allocation into the rhizosphere, nutrient stress decreased; exudates were preferentially used by Gram-negative bacteria and (mycorrhizal) fungi, resulting in an enhanced growth. Other microorganisms, like Gram-positive bacteria and mainly micro eucaryotes benefited from the exudates via food web development. Overall our results indicate a fast turnover of exudates and the development of initial food web structures. Additionally a transport of assimilated carbon into bulk soil by (mycrorhizal) fungi was observed. Ó 2009 Elsevier Ltd. All rights reserved.

Keywords: 13 C labelling Microbial biomass Cmic PLFA Rhizosphere Rhizodeposition Beech Dormancy Chilling C dynamics

1. Introduction With an amount of 80% of the terrestrial biomass (Saugier et al., 2001), forest ecosystems represent the major natural vegetation of European landscapes (Ellenberg, 1996). Woody plants, representing the majority of the above-ground part of a forest ecosystem, determine the quantity and quality of resources translocated (as plant litter and by exudation) into the soil of these ecosystems. This allocation of carbon (C) compounds into plants and soil has been frequently studied (e.g. Bauhus and Barthel, 1995; Wiemken et al., 2001; Dyckmans et al., 2002; Hackl et al., 2004), but still there is a lack of information about C dynamics between plant and soil. Despite its importance for soil organic matter and nutrient

* Corresponding author. German Research Center for Environmental Health, Institute of Soil Ecology, Department of Terrestrial Ecogenetics, Ingolstaedter Landstr. 1, D-85764 Neuherberg, Germany. Tel.: þ49 89 31872304; fax: þ49 89 31873376. E-mail address: [email protected] (M. Schloter). 1 Both the authors equally contributed to the study. 0038-0717/$ – see front matter Ó 2009 Elsevier Ltd. All rights reserved. doi:10.1016/j.soilbio.2009.07.002

transformation processes (Wiemken et al., 2001), the influence of plant derived C on microbial community structure and function in the rhizosphere is still poorly understood, although it is generally accepted that bacteria, fungi and archaea are strongly influenced by root C inputs (Brant et al., 2006). Recent studies suggest that half or more of the soil activity is driven by recent photosynthates (Ekblad and Ho¨gberg, 2001; Ho¨gberg et al., 2008). The rhizosphere has been defined as the volume of soil influenced by plant roots, which describes as a zone of high microbial activity (Hiltner 1904). C-compounds and nutrients are released by plant roots into the rhizosphere by ‘‘rhizodeposition’’, which describes the total C entering the soil in form of water-soluble exudates, secretions, lysates, gases and mucilage (Grayston et al., 1996). Within rhizodeposits mainly carbohydrates, carboxylic acids and amino acids are highly attractive for microorganisms and therefore most responsible for microbial growth (Lynch and Whipps, 1990). Since rhizosphere microbial communities are strongly influenced by root exudates (Brant et al., 2006), it has been hypothesised that plants select for beneficial microbial communities in their rhizosphere (Singh et al., 2007). In temperate forests,

¨ tz et al. / Soil Biology & Biochemistry 41 (2009) 1976–1985 J. Esperschu

ectomycorrhizal fungi are the most important group of beneficial microbes. They live in close symbiosis with fine roots of deciduous and evergreen trees. Ectomycorrhizal fungi utilize high amounts of organic substances obtained from the plant and in turn provide water, nutrients and mobilized minerals to their host, especially when nutrient conditions in the rhizosphere are limited (Smith et al., 1997; Ho¨gberg, 2006). Probably one of the most common methods to characterize microbial communities in soil is the use of phospholipid fatty acids (PLFA) and phospholipid ether lipids (PLEL). Phospholipids are essential membrane components of all living cells (Zelles, 1999a, b; Gattinger et al., 2003). Since they are not synthesized as storage compounds and are rapidly degraded after cell death, phospholipids serve as good indicators for living organisms (White et al., 1979). Combing this method with the fumigation of the plants with 13C labelled CO2 it is possible to characterize those microbes that can utilize the rhizodeposits. Consequently using this approach, it is possible to follow carbon fluxes from the plant into the rhizosphere and to identify microbes that metabolize the exudates (Kuzyakov, 2001). In the present study, C fluxes between plants (Fagus sylvatica L.) and soil were investigated after chilling for a period of three weeks, using a continuous labelling setup. We postulated that due to low concentrations of easily degradable carbon during dormancy of the trees, growth conditions for microbes in the rhizosphere are limited; therefore after chilling, when exudates are released into the rhizosphere, an increase in activity of microbial communities utilizing plant C is expected. It was the aim of this paper to verify that (i) limited growth conditions in the rhizosphere during dormant periods influence abundances and physiological adaption of individual rhizosphere microbial communities. When leaves have been developed, (ii) photosynthetically fixed C is rapidly transported through the plant and released into the rhizosphere, where it is incorporated into the microbial biomass. Due to plant available C (iii) a shift takes place in the microbial community structure, favouring plant C utilizing microbes. 2. Material & methods 2.1. Experimental setup The soil used in the following experiments (34% sand, 46% silt and 20% clay) was taken from Ho¨glwald (0–40 cm), a mixed forest stand in Bavaria, Germany (48 17’21’’N, 114’7’’E). Soil pH (H2O) was 3.9 ( 0.1) and C/N analyses result in 6.4% ( 0.64) of total carbon, 0.3% ( 0.02) of total nitrogen and a C/N ratio of 19.9 ( 0.38). The d13C signature of the soil was 26.7  0.25& d13C Vienna-Pee Dee Belemnite (V-PDB), analysed by EA-IRMS as described below. Three-year-old nursery-grown beech trees (Fagus sylvatica L.; Staatliche Samenklenge Laufen, Germany) were grown in 10 l pots of 20 cm in diameter (one plant per pot) in autumn 2004. During the winter months the plants were kept at ambient air temperatures to fulfill dormancy. In spring 2005, 27 planted and 6 unplanted pots were placed into a greenhouse at a photoperiod of 14 hours daylight (additional light by sodium vapour discharge lamps, SON-T Agro 400, Philips, NL) and temperatures of 25  C (day) and 18  C (night). Relative humidity was maintained between 75% and 85% by a sprinkler system. Fertilization was performed based on a double strength Hoagland nutrient solution (Hoagland and Arnon, 1950; Kozovits et al., 2005) with 250 ml one week after placement into the greenhouse. Irrigation of the plants was performed once every 72 hours (300 ml) via irrigation tubes to adjust 40–50% of the maximum water holding capacity throughout the experiment. A total of 3 planted (plants, rhizosphere, bulk soil) and 3 unplanted

1977

pots (bulk soil) were harvested serving as unlabelled controls (time point 0), 4 weeks after the pots were placed into the greenhouse, immediately after the uppermost leaves were fully developed. The remaining 24 planted and 3 unplanted pots were placed at the same day into a tent (volume about 7000 l) built of transparent plastic foil (ethylene-tetrafluorethylene ETFE, film thickness 80 mm, Koch Membranen GmbH, 83253 Germany), to separate the plants from the outer greenhouse atmosphere. CO2-concentration in the tent’s atmosphere was reduced by plant photosynthesis during the daytime. When the CO2 concentration in the tent dropped below a minimum level of 350 mmol*mol1, enriched 13CO2 (d13C ¼ þ170&), Air Liquide, Du¨sseldorf, Germany) was added to the tent atmosphere, until CO2 concentration reached 400 mmol*mol1. Using this experimental setup, an enriched 13C-atmosphere of þ60& to þ80& V-PDB was established in the tent. At night time the tent’s atmosphere was pumped through vials containing soda lime (contains sodium and calcium hydroxide) using a membrane pump (N 0135.3 AN.18, KNF Neuberger, Freiburg, Germany) with a flow of about 200 l*min1 to reduce the CO2 produced by the plant due respiration. The CO2 depleted air was pumped again into the closed tent. This system’s capacity ensured a stable CO2-concentration of 350 mmol*mol1 over night. The CO2 concentration of the inner and outer tent atmosphere (day/night) was measured continuously with a photo acoustic CO2-controller (7MB1300, Siemens, Germany, calibration at 400–600 mmol*mol1  2%). During harvests the tent was left open for at most 5 minutes to minimize dilution with ambient CO2 from the outer tent atmosphere. Plants, rhizosphere and bulk soil were harvested in triplicates (randomly chosen) at 8 time points (0.5, 1, 1.5, 2.5, 3.5, 5.5, 10.5 and 20.5) days after fumigation started. A higher harvesting frequency at the beginning of the experiment (within the first 5 days of labelling) was chosen according to a fast carbon transport through the plants into rhizosphere organisms, suggested in other studies (Butler et al., 2004; Jones et al., 2004; Leake et al., 2006; Ho¨gberg et al., 2008). The adhering soil to the roots after shaking was defined as rhizosphere soil; bulk soil was taken at >5 mm distance from root. Rhizosphere soil and bulk soil was sieved <2 mm and stored at þ4  C for microbial biomass analyses and at 20  C for PLFA analyses. All d13C values, relating to the international V-PDB standard, were calculated as follows (Werner and Brand, 2001):

d13 C ð&Þ ¼

h

RSample  RV

. i RV *1000

(1)

RSample and RV represent the 13C to 12C ratios of sample and international standard V-PDB (0.0111802), respectively. To estimate the amount of soil autotrophic CO2 fixation, d13C in PLFA, Cmic and DOC was analysed in soil of three unplanted pots at the beginning and the end of the experiment. 2.2. Carbon analyses in bulk material After harvest, aliquots of leaves, twigs (annual growth), stems (perennial growth), fine roots (<2 mm) and coarse roots (>2 mm) were separated and subsequently dried at 65  C for 48 h, ballmilled (Retsch MM2, Retsch GmbH, Haan, Germany) and weighed into tin capsules (HEKAtech GmbH, Wegberg, Germany). Analyses for C content and d13C were performed by an Elemental Analyzer coupled with an Isotopic Ratio Mass Spectrometer (EA-IRMS; Eurovector, Milan, Italy coupled with a MAT 253, Thermo Electron, Bremen, Germany). 2.3. Microbial biomass and dissolved organic carbon Within 3 days after harvest, microbial biomass (Cmic) was estimated by chloroform-fumigation extraction according to

¨ tz et al. / Soil Biology & Biochemistry 41 (2009) 1976–1985 J. Esperschu

1978

Vance et al (1987). The K2SO4 extracts were stored at 20  C until measurement. Total organic C contents in the extracts were measured as CO2 in a Total Carbon Analyzer (TOC 5050, Shimadzu Corporation, Tokyo, Japan). Microbial biomass was calculated using a kEC-factor of 0.45 (Wu et al., 1990). Dissolved organic C (DOC) was extracted by shaking soil samples (equivalent to 5 g oven-dried soil) in 0.01 M CaCl2 solution (1:5; w/v) on a rotary shaker for 30 minutes. Subsequently, the soil suspension was centrifuged, the supernatant filtered through polycarbonate filters of 0.4 mm pore-size (Whatman Nucleopore Track-Etch Membrane filters). The filtered extracts were stored at 20  C until measurement. Total organic C contents in the extracts were determined on the Shimadzu TOC 5050. Measurement of d13C in K2SO4 and CaCl2 extracts was done by on-line coupling of liquid chromatography and stable isotope ratio mass spectrometry (LCIRMS, Thermo Electron, Bremen, Germany) according to (Krummen et al., 2004). The d13C in microbial biomass (d13CBio) was calculated as follows (Marx et al., 2007a):



d13 CBio ð&Þ ¼ d13 Cfum *Cfum  d13 Cn-fum *Cn-fum

. CBio

(2)

d13Cfum and d13Cn-fum are d13C values in fumigated (fum) and nonfumigated (n-fum) extracts, respectively; Cfum and Cn-fum are C concentrations (in mg C*l1) of fumigated and non-fumigated extracts, and CBio represents the microbial C concentration [mg C*l1]. 2.4. PLFA analyses For PLFA analyses based on Zelles et al. (1995), 10 g soil (dry weight) was extracted with 125 ml methanol, 63 ml chloroform and 50 ml phosphate buffer (0.05 M, pH 7). After 2 hours of horizontal shaking, 63 ml water and 63 ml chloroform were added to promote phase separation. After 24 h the water phase was removed and discarded. The total lipid extract was separated into neutral lipids, glycolipids and phospholipids on a silica-bonded phase column (SPE-SI 2 g/12 ml; Bond Elut, Analytical Chem International, CA, USA). The phospholipid fraction was further separated into saturated (SATFA), monounsaturated (MUFA) and polyunsaturated (PUFA) fatty acids (see Zelles et al., 1995 for details) to facilitate the identification of fatty acids as well as to obtain a good baseline separation of peaks for isotopic calculations. PLFA were analysed as fatty acid methyl esters (FAME) on a gas chromatograph/mass spectrometry system (5973MSD GC/MS Agilent Technologies, Palo Alto, USA) linked via a combustion unit to an isotope ratio mass spectrometer (GC/MS-C-IRMS, DeltaPlusAdvantage, Thermo Electron Cooperation, Bremen, Germany). Separation and detection of FAME was performed via GC/MS, while isotopic composition of fatty acids was detected after combustion (GC Combustion III, Thermo Electron Cooperation, Bremen, Germany) in the IRMS. FAME were separated on a polar column (BPX-70, SGE GmbH, Griesheim, Germany), 60 m  0.25 mm  0.25 mm, coated with 70% of cyanopropyl polysilphenylene-siloxane. To separate SATFA and MUFA (underivatized), an initial GC-temperature of 50  C was kept for 2 minutes, then increased at 55  C*min1 to 136  C, and subsequently at 2  C*min1 to 250  C. PUFA fractions were separated by an initial temperature of 150  C, raised at 1.5  C*min1 to 210  C. To identify the position of the double bonds, derivatised MUFA (see Zelles et al., 1995 for details) were again measured at 60  C, kept for 2 minutes and followed by several ramps (120  C*min1 to 200  C; 20  C*min1 to 203  C; 0.13  C*min1 to 210  C; 5  C*min1 to 250  C). The final temperature of all GC-programs was held for 10 minutes. The mass spectra of the individual FAME were identified by comparison with established fatty acid libraries (Solvit, CH 6500 – Luzern, Switzerland) using MSD Chemstation (Version D.02.00.237).

Fatty acids are designated as the total number of C-atoms followed by the number of double bonds and their location (u) after the colon. The prefixes ‘‘cy’’, ‘‘i’’ and ‘‘a’’ indicate cyclopropyl-groups, and iso- and anteiso- branching, respectively. Saturated straightchained fatty acids were indicated by ‘‘nor’’. ‘‘10Me’’ indicates a methyl-branching at the 10th C-atom, whereas ‘‘br’’ indicates methyl-branching at an unknown position. According to Zelles (1999b), iso- and anteiso- branched PLFA have been used as indicators for Gram-positive bacteria, whereas Gram-negative bacteria have been represented by monounsaturated and cyclopropyl PLFA. Additionally PLFA 16:1u5 was used as an indicator for Gram-negative bacteria, since arbuscular mycorrhiza (which is also represented by PLFA 16:1u5) is not likely to be present in high numbers in combination with beech trees. PLFA 18:2u6.9 was taken as an indicator for fungal biomass. For eukaryotic biomass, PLFA n22:0, n24:0, 18:3 and 20:4 have been chosen. The ratio of u7 MUFA to its cyclopropyl derivates was calculated on the basis of 16:1u7c and 18:1u7 to cyclopropylic cy17:0 and cy19:0 (Reichhardt et al.,1997; Ratledge and Wilkinson, 1988) indicative for stressed microbial communities. Samples were measured in duplicates and measurements were repeated when a fluctuation of >0.5& d13CV-PDB in the internal standard occurred. According to Werner and Brand (2001), differences in the d13CV-PDB value of the internal standard resulting from measurements by EA-IRMS and GC/MS-C-IRMS (nonadecanoic acid methyl ester, d13CV-PDB ¼ 30.5& V-PDB or myristic acid methyl ester in PUFA fractions, d13CV-PDB ¼ 28.7& V-PDB) were included into the d13CV-PDB ratio of individual FAME obtained from the measurement (using mass spectrometer software Isodat 2.5). The actual PLFA ratio (d13CPLFA) was obtained by calculating the one C-atom in the methyl group that has been added during derivatisation into the C isotope ratios of the FAME (d13CFAME). Since no isotopic fragmentation in this step is known (Abrajano et al., 1994), the calculation was done as follows:

h

d13 CPLFA ¼ ðn þ 1Þ*d13 CFAME  1*d13 CMethanol

i.

n

(3)

where n is the number of C atoms in the PLFA and d13CMethanol the d13CV-PDB ratio of methanol used for derivatization (38.5& V-PDB, determined by LC-IRMS). Related to the individual d13CV-PDB content in the labelling atmosphere, the percentage of newly incorporated carbon into individual PLFA-biomarker was calculated using a fractionation factor acomp of each individual PLFA relative to the atmosphere (Farquhar et al., 1989):

acomp ¼

h

d13 CCO2 =1000 þ 1

.

d13 Ccomp =1000 þ 1

i

(4)

for d13CCO2 a mean value of d13CV-PDB of 11& was measured and the calculated d value (after equation 3) was applied for d13Ccomp. The newly incorporated carbon (Cnew) proportion was calculated based on equations 5 and 6

d13 Cmax ¼

h



i

d13 Caltered =1000 þ 1 *acomp  1 *1000

.   13 13 d13 Cmax  d13 Ccomp *100 Cnew ¼ d Cnew  d Ccomp

(5) (6)

d13Cmax is calculated as the maximum label incorporation possible with respect to the individual acomp and the d value of the altered atmosphere (d13Caltered), and d13Cnew the d value of the individual component measured under d13Calt, respectively. 2.5. Statistical analysis Statistical analyses were performed with SPSS 15.0. Significant differences (p < 0.05) were analysed comparing means of three

¨ tz et al. / Soil Biology & Biochemistry 41 (2009) 1976–1985 J. Esperschu

coarse roots also showed significant 13C incorporation (p < 0.05) after 20.5 days but to a lower extent (6d13C ¼ 15&).

independent, randomly chosen replicates. Results obtained at any harvesting time point have been tested against the data obtained from the unlabelled control pots (time point 0), using a univariate analysis of variance (ANOVA) followed by Duncan’s post hoc tests. Differences between rhizosphere soil and bulk soil samples have been tested at p < 0.05 using student’s t-test for paired samples (rhizosphere soil and bulk soil in the same planted pot). All results were illustrated as means  standard deviations (S-Plus 6.2).

3.2. Distribution of labelled carbon in microbial biomass and dissolved organic carbon in the rhizosphere and in bulk soil samples Both Cmic and DOC showed higher abundance levels in the rhizosphere soil compared to the bulk soil (210 to 330 mg C kg1 DW and 200–250 mg C kg1 DW, respectively) in the course of the experiment (data not shown). Fig. 2 illustrates the carbon translocation into the below-ground part of the plant soil system: The d13C values in the rhizosphere Cmic (Fig. 2 a) were significantly increased after 10.5 days of labelling (p < 0.05) compared to the beginning of the experiment. Also DOC was significantly enriched in 13C after 10.5 days of labelling, however to a lower amount compared to Cmic (Fig. 2 b). In contrast no 13C increase was observed in DOC and Cmic of bulk soil samples. Analysis of 13C enrichment in microbial biomass of unplanted soil samples did not show 13C enrichment (data not shown) indicating that microbial autotrophic CO2 fixation did not play a role in this experiment.

3. Results 3.1. Distribution of

13

1979

C labelled carbon in different plant parts

Fig. 1 illustrates the incorporation of 13C into different plant parts of 13C-CO2 fumigated beech trees. Significant incorporation (p < 0.05) was detected in leaves after 3.5 day of labelling. In twigs and stems, 13C incorporation was detected at p < 0.05 after 10.5 days. The highest increase of d13C in plant parts was detected in fine roots at the end of the experiment (6 d13C ¼ 47&). Like fine roots,

30

∗

leaves

15 0

∗

∗

∗

-15

twigs

0

∗

∗

∗

∗

-15

stem

15 0

-15

fine roots

15 0

∗

-15

coarse roots

13

C signature in plant tissue [δ13C ‰ V-PDB]

15

15 0

∗

-15

-30 0

2

4

6

8

10

12 18

22

labelling time [d] 13

13

Fig. 1. C incorporation in total C of beech plant parts (d C&V-PDB) at different harvesting time points (means  standard deviation (n ¼ 3). Differences p < 0.05 compared to the beginning of the experiment (day 0) were indicated by asterisks (*).

¨ tz et al. / Soil Biology & Biochemistry 41 (2009) 1976–1985 J. Esperschu

In total 35 individual PLFA have been identified; cy19:0 and a16:0 were the most common fatty acids detected at all harvesting dates (Table 1). Relative abundance of most MUFA (16:1u5, 16:1u9, 16:1u7c/t, 17:1u8c/t, 18:1u9, 18:1u7) have been significantly increased (p < 0.05) after 10.5 days. Significant higher relative abundance values for MUFA 16:1u7c/t and 18:1u9 have been observed even earlier after 5.5 days. Fastest response time was measured for MUFA 17:1u8c/t, where a significant relative increase was already observed 0.5 days after the start of the experiment. Also PUFA 18:2u6.9 has been relatively increased already within the first days of the experiment (5.5 days). However the relative abundance decreased from that time point on until the end of the labelling period. In contrast SATFA a16:0 decreased continuously relative to total PLFA over the whole experimental period. Also other saturated PLFA (br11.17, i/a17:0, n17:0) have been relatively reduced mainly during the end of the labelling period, but a coherent response pattern was not observed. The ratio of MUFA precursors to cyclopropyl PLFA was calculated on the basis of the sum of u7 MUFA (16:1u7c and 18:1u7) to its cyclopropyl derivates, cy17:0 and cy19:0 (Fig. 3). During the experimental period an increase in this ratio was observed indicating a decrease in stress conditions in the rhizosphere. 3.4. Incorporation of labelled C into rhizosphere microbial communities Over the experimental period about half of the detected PLFA showed significant enrichment of 13C. The fastest significant incorporation of the 13C label was observed in the PLFA n20:0, n22:0 and n:24:0 half a day after the labelling has been started. In addition a relative fast significant incorporation of 13C labelled carbon was observed in SATFA i15:0. However most monounsaturated (16:1u9, 16:1u7c, 18:1u7, 18:1u9), cyclopropyl fatty acids (cy17:0 and cy19:0) and PLFA a16:0 showed a significant enrichment of 13C after 10.5 days of labelling (Table 2). Branched PLFA a14:0 and a15:0 as well as PLFA i16:0 showed 13C enrichment mainly at the end of the experiment. The maximum 13C incorporation compared to day 0 ranged between 2& (cy19:0) and 30& (18:2u6.9) at the end of the experiment. Based on equations (4), (5) and (6), PLFA 18:2u6.9, 18:3 and 20:4 showed the highest proportions of newly assimilated C derived from labelled plant assimilates in the rhizosphere ranging from 20% to 55% after 20.5 days. To reduce sample size for PLFA analysis, bulk soil samples have been analysed only at the beginning, after 10.5 days and after 20.5 days of experimental duration. Unsaturated PLFA 18:1u7 and 18:2u6.9 were significantly increased in d13 at around (6 d13C ¼ 5.45& respectively 9.85&) after 20.5 days of labelling. Furthermore a slight increase was also observed for PLFA a14:0 (6 d13C ¼ 2.25&) and n22:0 (6d13C ¼ 1.05&, respectively 1.25&)at the end of the experiment; however the data did not reach significance level (p < 0.05). 4. Discussion 4.1. Distribution of

13

to other parts of the plant. Higher d13C values in twigs compared to stems may be a result of carbon utilization in the still growing, annual twigs. Probably due to the length of the translocation pathway and anabolism of carbohydrates of woody plants (Kozlowski et al., 1991), labelled carbon was detected in the belowground plant parts not until then 20.5 days. Coarse roots basically serve as a storage site of carbon, resulting in a lower amount of incorporated freshly assimilated carbon compared to fine roots. Fine roots, known as a highly active plant tissue, require high amounts of carbon for growth, resulting in high extents of recently assimilated carbon. Significant incorporation of labelled 13C into the microbial biomass was observed 10.5 days after the experiment has been started. However individual microbes responded much faster to the root exudates as indicated by the occurrence of the 13C label in some PLFA (see below) shortly after the experiment has been started. It can be speculated if the slow occurrence of the label in most of the biomass is related to high metabolic activity and respiration of the microbes after dormancy of the trees, slow generation times of most rhizosphere microbes (as most lipids are formed during cell membrane synthesis) or by a highly structured food web, with only some microbes responding first to the exudates. 4.2. Distribution of labelled carbon in microbial biomass and dissolved organic carbon in the rhizosphere and in bulk soil samples From roots, large amounts of C are released into the soil in form of root exudates, containing 5% to 21% of photosynthetically fixed C

a δ13C in total DOC [‰ V-PDB]

3.3. Influence of rhizodeposition on microbial community structure in the rhizosphere

rhizosphere soil bulk soil

-15

-20

*

-25

-30

b δ13C in totalCmic[‰ V-PDB]

1980

*

-15

*

-20

-25

C labelled carbon in different plant parts -30

Results from 13C analyses of different plant parts reflect the allocation and distribution of recently fixed C within young beech trees. 3.5 days after the start of labelling, 13C has been detected in leaves, where the assimilated carbon is used for leaf formation in the initial phase of growth (Dyckmans et al., 2002) after dormancy. In leaves, assimilates are transformed into sugars and amino acids, which are transported via the woody plant tissue (twigs and stems)

0

2

4

6

8

10

12 18

20

22

labelling time [d] Fig. 2. 13C incorporation (d13C&V-PDB) into dissolved organic carbon (DOC; a) and microbial biomass (Cmic; b) in the course of the experiment (mean values  standard deviations of n ¼ 3). Differences p < 0.05 compared to time point 0 are indicated by asterisks (*).

¨ tz et al. / Soil Biology & Biochemistry 41 (2009) 1976–1985 J. Esperschu

1981

Table 1 Relative abundance of PLFA (mol%) in rhizosphere soil (means of n ¼ 3  standard deviations) at different harvesting time points during the experiment. Significant differences compared to the beginning (t0) are indicated by an asterisks (p < 0.05). PLFA [mol%]

br12:0  i14:0  a14:0  i15:0  a15:0  n15:0  15:1u8  i16:0  a16:0  16:1u5  16:1u9  16:1u7c  16:1u7t  i17:0  br11.17:0  a17:0  n17:0  17:1u8c  17:1u8t  cy17:0  br10.18:0  i18:0  n18:0  18:1u9  18:1u7  18:2u6.9  cy18:0  18:3  br10.19:0  cy19:0  n20:0  20:4  n22:0  n24:0  dic22:0 

labelling time [d] 0

0.5

1.0

1.5

2.5

3.5

5.5

10.5

20.5

0.21 0.10 0.35 0.01 2.03 0.40 8.63 0.34 4.03 0.13 0.72 0.09 0.03 0.03 5.99 0.36 20.18 0.66 0.25 0.28 0.06 0.07 0.27 0.24 0.05 0.05 8.30 0.27 1.42 0.03 1.74 0.14 0.68 0.04 0.02 0.02 0.05 0.05 3.20 0.19 1.72 0.08 0.85 0.01 4.04 0.16 0.71 0.72 0.44 0.33 2.02 1.41 0.42 0.01 0.30 0.22 1.11 0.17 17.26 1.04 3.40 0.26 0.43 0.17 5.51 0.30 3.03 0.23 0.87 0.42

– – 0.18* 0.05 1.64 0.27 7.04 0.41 3.26 0.22 0.56 0.03 0.05 0.01 5.31 0.26 19.64 0.12 0.49 0.05 0.12 0.02 0.66* 0.05 0.13 0.02 8.56 0.37 1.46 0.08 1.41 0.11 0.53 0.06 0.08* 0.01 0.19* 0.02 2.94 0.24 1.66 0.14 0.86 0.03 3.68 0.32 2.10 0.25 1.19 0.19 3.06 1.94 0.40 0.01 0.43 0.36 0.89 0.13 19.17 0.99 3.33 0.23 0.26 0.10 5.26 0.57 2.72 0.49 0.73 0.40

0.16 0.09 0.23 0.09 1.81 0.49 7.44 1.38 3.62 0.57 0.54* 0.04 0.05 0.01 4.64 0.40 18.72 1.21 0.38 0.15 0.08 0.03 0.55 0.14 0.07 0.03 7.58 0.28 1.34 0.04 1.60 0.12 0.56 0.07 0.06 0.00 0.14* 0.03 2.93 0.27 1.47 0.16 0.80 0.00 3.77 0.33 1.58 0.54 1.01 0.24 4.92* 1.83 0.38 0.09 0.83 0.28 0.90 0.16 17.41 0.68 3.67 0.21 0.32 0.07 6.14 0.19 3.09 0.27 1.27 0.67

0.14 0.07 0.28 0.06 1.89 0.18 7.35 0.41 3.46 0.19 0.64 0.06 0.04 0.02 5.20 0.34 17.99 1.03 0.41 0.14 0.10 0.04 0.63 0.17 0.12 0.04 8.21 0.20 1.33 0.06 1.50 0.09 0.61 0.04 0.07* 0.03 0.16* 0.03 3.10 0.07 1.87 0.11 0.82 0.01 3.82 0.14 1.98 0.56 1.19 0.30 3.75 1.91 0.43 0.07 0.70 0.47 1.14 0.07 16.67 1.14 3.54 0.15 0.36 0.16 5.92 0.17 3.65 0.11 0.93 0.77

0.17 0.08 0.27 0.10 1.77 0.42 7.17 1.31 3.72 0.27 0.70 0.09 0.06 0.01 5.87 0.45 17.27* 0.36 0.47 0.14 0.11 0.01 0.63 0.10 0.14* 0.03 8.65 0.37 1.31 0.09 1.54 0.12 0.64 0.09 0.08* 0.02 0.17* 0.03 2.99 0.06 2.05 0.11 0.82 0.02 3.63 0.13 2.12 0.18 1.15 0.05 4.60* 1.33 0.47 0.02 0.59 0.27 1.23 0.08 16.29 1.00 3.32 0.39 0.49 0.06 5.63 0.96 3.48 0.71 0.48 0.43

0.39 0.38 0.32 0.04 2.02 0.16 7.10 1.59 3.72 0.27 0.69 0.06 0.06 – 5.12 0.69 16.38* 2.26 0.46 0.18 0.10 0.03 0.58 0.20 0.10 0.05 7.29 1.56 1.08* 0.25 1.27* 0.37 0.45* 0.06 0.07* 0.02 0.16* 0.05 2.53 0.68 1.56 0.38 0.73 0.14 3.83 0.69 1.80 0.54 1.16 0.42 4.49* 2.54 0.28 0.07 0.72 0.50 0.92 0.30 14.33 3.63 3.03 0.32 0.49 0.22 4.89 0.39 2.87 0.30 9.99 16.35

0.38 0.09 0.37 0.02 2.22 0.07 7.75 0.11 4.08 0.91 0.67 0.13 0.07 0.00 4.68 1.41 17.90 1.10 0.58 0.12 0.14 0.03 0.74* 0.16 0.15* 0.06 7.77 0.77 1.27 0.05 1.28* 0.05 0.53 0.08 0.08* 0.02 0.20* 0.02 2.81 0.24 1.55 0.38 0.76 0.04 3.42 0.51 2.34* 0.30 1.43 0.20 7.79* 2.98 0.30 0.04 0.93 0.51 0.85 0.19 16.49 1.98 2.82 0.60 0.50 0.11 4.61 1.00 2.34 0.44 0.21 0.03

0.35 0.02 0.37 0.01 2.22 0.02 7.78 0.25 3.57 0.09 0.71 0.04 0.08 0.02 5.13 0.43 17.18* 0.20 0.75* 0.04 0.17* 0.01 1.09* 0.14 0.19* 0.01 7.61 0.43 1.23 0.04 1.55 0.08 0.54 0.09 0.12* 0.00 0.28* 0.02 2.72 0.15 1.66 0.30 0.75 0.04 3.58 0.49 3.13* 0.41 2.25* 0.74 6.22* 3.12 0.27 0.08 0.83 0.76 1.02 0.24 15.68 0.67 2.89 0.37 1.05 0.51 4.69 0.83 2.22 0.69 1.10 0.12

0.40 0.19 0.41 0.05 2.34 0.27 8.90 0.42 3.85 0.03 0.70 0.05 0.09 0.02 4.75 0.18 16.78* 0.33 1.03* 0.05 0.21* 0.03 1.35* 0.19 0.19* 0.01 6.71* 0.12 1.20 0.06 1.62 0.12 0.56 0.12 0.13* 0.01 0.38* 0.03 2.71 0.09 1.22 0.04 0.82 0.02 3.67 0.40 3.59* – 3.54* 1.18 3.93* 1.54 0.37 0.09 0.44 0.04 0.72 0.04 17.03 0.92 3.06 0.14 1.14 0.39 4.77 0.31 2.17 0.12 1.74 1.23

¨ tz et al. / Soil Biology & Biochemistry 41 (2009) 1976–1985 J. Esperschu

1982

13

C values determined in DOC of rhizosphere soil proves the release of photosynthetically derived assimilates into the rhizosphere. DOC is postulated as a good carbon source for microbial growth and productivity (Meyer et al., 1987; Paterson, 2003). The 13C labelled assimilates were incorporated into the microbial biomass fraction (Cmic), indicated by a d13C increase in Cmic in rhizosphere soil. The higher d13C values observed in microbial biomass compared to d13C values of DOC are consistent with previously reported data (Potthoff et al., 2003; Yevdokimov et al., 2006; Marx et al., 2007b) and reflect a high utilization of rhizodeposits. It has been suggested that rhizodeposits are rapidly metabolized by microorganisms, and thus the comparably low values of d13CV-PDB in DOC can be explained by a relatively small fraction of labelled rhizodeposits remaining in the pool of water-soluble organic C (Yevdokimov et al., 2006; Marx et al., 2007a).

0.4

ratio ∑-ω7/cy

0.3

0.2

* *

0.1

0.0 0

2

4

6

8

10

12 18

20

22

labelling time [d] Fig. 3. Ratios of –u7 PLFA (16:1u7 and 18:1u7) to its cyclopropyl fatty acids (cy17:0 and cy19:0) at individual harvesting time points during the experimental period (mean of triplicates  standard deviations). Asterisks *) indicate significant differences at p < 0.05 compared to the beginning of the experiment (day 0).

(Marschner, 1995). However depending on the degree of mycorrhization, this amount might be significantly reduced (Smith and Read, 2008). C-compounds released into soil contribute mainly to the pool of dissolved organic carbon (DOC). The latter was confirmed by results in this study, where higher total amounts of DOC were detected in rhizosphere soil than in bulk soil. Increasing

4.3. Influence of rhizodeposition on microbial community structure in the rhizosphere 40% of total PLFA were identified as cy19:0 and a16:0 indicating high proportions of Gram-negative and Gram-positive bacteria (Zelles, 1999b) in the rhizosphere of young beech trees. Increases in MUFA (16:1u5, 16:1u9, 16:1u7c/t, 17:1u8c/t, 18:1u7) during the experimental period showed a growth of Gram-negative organisms, whereas Gram-positive bacteria (mainly a16:0, but also br11.17:0 and a17:0) indicated a decline over time. MUFA have been detected in high amounts in Gram-negatives like e.g. Pseudomonas, Acetobacter, Azospirillum, Thiobacillus, Rhizobium, Agrobacterium,

Table 2 Relative increase of the 13C label in individual PLFAs in rhizosphere soil compared to t0 (means of n ¼ 3  standard deviations) at different harvesting time points during the experimental period. Only those PLFA are shown that show at least one significant difference compared to t0. Significant values are indicated by asterisks (p < 0.05). increase [%]

labeling time [d] 0.0

0.5

1.0

1.5

2.5

a14:0

0.00

i15:0  a15:0  i16:0  a16:0  16:1u5  16:1u9  16:1u7c  cy17:0  n18:0  18:1u7  18:2u6.9  18:3  cy19:0  n:20:0  20:4  n22:0  n24:0

0.00

0.13 0.18 0.12 0.03 0.08 0.15 0.04 0.17 0.10 0.07 0.17 0.16 0.07 0.12 0.13 0.20 0.10 0.12 0.16 0.05 0.27 0.58 0.14 0.05 0.02 0.59 0.05 0.05 0.16* 0.05 0.21 0.27 0.22* 0.05 0.30* 0.08

0.05 0.18 0.08 0.05 0.02 0.16 0.04 0.14 0.10 0.09 0.17 0.17 0.07 0.15 0.16 0.21 0.03 0.11 0.08 0.11 0.09 0.57 0.13 0.13 0.21 0.36 0.03 0.04 0.05 0.14 0.08 0.27 0.06 0.18 0.14 0.18

0.08 0.12 0.10 0.02 0.04 0.05 0.02 0.05 0.17 0.02 0.02 0.19 0.11 0.18 0.12 0.22 0.03 0.08 0.06 0.03 0.10 0.39 0.45 0.23 0.22 0.33 0.04 0.04 0.04 0.04 0.16 0.22 0.03 0.04 0.04 0.08

0.06 0.19 0.07 0.02 0.01 0.05 0.03 0.12 0.18 0.09 0.04 0.19 0.17 0.17 0.09 0.22 0.04 0.07 0.06 0.07 0.15 0.29 0.61 0.19 0.36 0.52 0.04 0.03 0.05 0.07 0.18 0.12 0.05 0.04 0.03 0.08

0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00

3.5 0.10 0.13 0.16* 0.07 0.12 0.08 0.10 0.09 0.32 0.14 0.19 0.18 0.13 0.15 0.18 0.26 0.02 0.11 0.08 0.10 0.02 0.71 0.85 0.40 0.66 0.87 0.04 0.05 0.01 0.06 0.23 0.18 0.02 0.04 0.04 0.11

5.5 0.19 0.15 0.18* 0.05 0.06 0.11 0.02 0.15 0.25 0.05 0.17 0.14 0.13 0.32 0.26 0.25 0.04 0.12 0.17 0.02 0.22 0.47 0.73 0.24 0.59 0.63 0.04 0.04 0.08 0.09 0.16 0.35 0.12 0.07 0.17 0.08

10.5 0.21 0.25 0.18* 0.06 0.09 0.14 0.18 0.12 0.80* 0.40 0.08* 0.14 0.17* 0.23 0.46* 0.42 0.17* 0.10 0.39* 0.17 0.65* 0.60 2.60* 1.07 3.62* 0.88 0.09* 0.08 0.01 0.07 0.45 0.40 0.03 0.09 0.04 0.20

20.5 0.39* 0.11 0.30* 0.11 4.62* 0.12 0.20* 0.10 0.78* 0.09 0.10* 0.13 0.31* 0.10 0.56* 0.31 0.22* 0.14 0.56* 0.15 0.85* 0.43 3.10* 0.38 4.63* 0.80 0.17* 0.05 0.13* 0.12 1.65* 0.33 0.12 0.13 0.31* 0.18

¨ tz et al. / Soil Biology & Biochemistry 41 (2009) 1976–1985 J. Esperschu

Escherichia or Flavobacterium (Kerger et al., 1986, The´berge et al., 1996, Jarvis et al., 1996, Zelles, 1997), commonly found in the rhizosphere of annual plants as well as trees. The composition of microbial communities in the rhizosphere can be altered by root exudates (Hodge and Millard, 1998), and the exudates’ composition varies depending on environmental and biological parameters, like for example ontogenetic stage (Baudoin et al., 2003). The experiment started when the uppermost leaves have been fully developed. During the dormant period carbon fluxes to the rhizosphere are reduced, which in turn causes limited growth conditions for rhizosphere organisms. During the labelling period, which started right after bud-break, the plants’ need for carbohydrates decreased and consequently higher amounts of carbon were allocated through the roots into the rhizosphere. In accordance with Kramer and Gleixner (2008), our results indicate Gram-negative bacteria benefit from plant exudates. However cyclopropyl fatty acids did not increase during the experimental period, like other biomarker for Gram-negatives. It has been reported earlier, that these derivates of u7 monounsaturated fatty acids operate as membrane stabilizing fatty acids and hence are formed under given environmental stress such as starvation (Findlay and Dobbs, 1993; Frostegard et al., 1993). Lower ratios of this stress indicator are correlated with a higher formation of cyclopropylic fatty acids, and therefore an increase during the experimental period illustrates better growth conditions for rhizosphere bacteria. Obviously, the rhizosphere was nutrient limited during bud-break, because of high needs of assimilates for the plant itself. The time when the experiment started and leaves were fully developed, nutrient limitation and hence starvation conditions for rhizosphere bacteria declined. PUFA 18:2u6.9 is frequently used as a biomarker for fungi (Frostegard and Bååth, 1996), or probably ectomycorrhizal fungi, as suggested by Priha et al. (1999) in combination with forest trees. MUFA 18:1u9 was recently detected to be positively correlated with 18:2u6.9 in forest soil (Ho¨gberg, 2006). Not surprisingly also in the present study a relative increase in 18:1u9 and in 18:2u6.9 was detected, however mainly at the end of the labelling period. As known from parallel experiments carried out in the same soil using plant material from the same nursery, the degree of mycorrhization of the plant roots was 80–90% at the start of the experiment (Pritsch et al., 2005). A high abundance of mycorrhizal fungi that is already present in the rhizosphere would promote soil nutrient and water uptake (Smith and Read, 2008) when demands of the aboveground plant parts are high, as it is given during bud-break. However, since PLFA 18:2u6.9 is also known to be widespread among the eukaryotic kingdom (Zelles, 1997), one cannot completely exclude that it originates from plant roots or root tips. Although harvest and sample preparation prior to extraction was accomplished with care, high fluctuation in 18:2u6.9 obtained from rhizosphere may be a result from root material. As plants have been transplanted only 8 month before the start of the experiment a total development of an ectomycorrhizal net in the pots has obviously not be started; therefore it was possible to clearly separate bulk soil from rhizosphere in one pot and plant effects for both compartments could be studied. This is also confirmed by the 13C incorporation data into PLFA 18:2u6.9 in bulk soil, which was very low and only increased at the end of the labelling period, indicating a transfer of the label more to saprophytic fungi, than a direct translocation into an ectomycorrhizal net. 4.4. Incorporation of labelled C into rhizosphere microbial communities Besides a clear incorporation of the 13C label into PLFA 18:2u6.9 in the rhizosphere, indicating the close interaction between plant and (mycorrhizal fungus), which has been also observed by other

1983

authors (Arao, 1999; Butler et al., 2003), mainly Gram-negative bacteria responded to an increased carbon allocation into the rhizosphere. Enhanced growth has been suggested as a result of higher amounts of plant carbon transported through the roots into the rhizosphere. Typical indicator lipids for Gram-negative bacteria like MUFA 16:1u9, 16:1u7c, 18:1u7, 18:1u9 as well as SATFA cy17:0 and cy19:0 showed 13C enrichment after 10.5 days of labelling, other Gram-negative PLFA like 16:1u5 and 17:1u8 t have been enriched in d13C even earlier. The interpretation of the fast occurrence of the 13C label in long chained PLFA (n20:0, 20:4, n24:0) at the beginning of the labelling period and at the end of the experiment might be related to the response of two different groups of microbes, as those PLFA occur in Gram-negative bacteria as well as in micro eucaryotes. Therefore the fast label incorporation after 0.5 days might be related to Gram-negative bacteria more than to micro eucaryotes and confirms the results discussed above. The high incorporation of the label at the end of the experiment could be interpreted as an indictor for food web development and the involvement of micro eucaryotes at that time point. Microeukaryotes and protists graze on bacteria (Bonkowski et al., 2000) and as secondary consumers they might have incorporated the labelled plant carbon. Consequently they would not have been directly influenced by changes within the carbon availability caused by rhizodeposition after bud-break. According to the high amount of labelled carbon (about 20% at the end of the experiment) detected in PUFA 20:4, the authors suggest a high activity of microeukaryotes and protists in the rhizosphere of beech trees at the end of the labelling period. Furthermore this data indicates strongly food web development at that time point. Coherently with increasing abundances of these PLFA, these results indicate a fast growth of Gram-negative bacteria due to plant exudates. A preferential use of plant C by Gram-negative bacteria was already postulated by Kramer and Gleixner (2008) in a natural abundance labelling study with rye and maize, and could be verified for woody plants according to results from this study. According to Soederberg et al. (2004), fast generation times under conditions of sufficient nutrient supply like in the rhizosphere are characteristic for Gram-negative bacteria. Due to fast generation times, root exudates, that were rapidly incorporated by Gramnegatives might again be rapidly released into soil and initiate food web development (Kindler et al., 2006; Lueders et al., 2006). This might be a reason also for increasing isotopic signatures in PLFA of Gram-positive origin (a14:0, i/a15:0, i/a16:0) and hence for a fast turnover of carbon within the rhizosphere, although relative growth of Gram-positive bacteria was not detected. 4.5. Conclusions In the present study, during the winter period, (i) the lack of photosynthetic activity and carbon allocation created nutrient limited conditions in the rhizosphere of young beech trees. Stress conditions have been faced by an increased formation of cyclopropyl fatty acids to increase membrane stability. During the experiment, when carbon allocation to belowground was present, nutrient stress was continuously lowered and stress indicators decreased. (ii) Photosynthetically assimilated carbon could be traced into individual plant parts, where need was given at the flowering stage. Labelling could be detected in DOC and total microbial biomass, indicating plant exudation into the rhizosphere. Among rhizosphere organisms, (iii) Gram-negative bacteria and (mycorrhizal) fungi preferentially utilized plant exudates, resulting in an enhanced Gram-negative population. As in other studies (Butler et al., 2003; Lu et al., 2004; Paterson et al., 2007) assimilated carbon was not evenly distributed within PLFA, indicating a different utilization of exudates within microbial communities.

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¨ tz et al. / Soil Biology & Biochemistry 41 (2009) 1976–1985 J. Esperschu

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