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Response of soil C and N transformations to tannin fractions originating

from Scots pine and Norway spruce needles

Sanna Kanerva a,*, Veikko Kitunen a, Oili Kiikkila a, Jyrki Loponen b, Aino Smolander a

aFinnish Forest Research Institute, Vantaa Research Centre, P.O. Box 18, FI-01301 Vantaa, Finland

bDepartment of Chemistry, University of Turku, Vatselankatu 2, FI-20014 Turku, Finland

Received 17 May 2005; received in revised form 4 October 2005; accepted 13 October 2005

Available online 4 January 2006

Abstract

Tannins are polyphenolic compounds that may influence litter decomposition, humus formation, nutrient (especially N) cycling and ultimately,

plant nutrition and growth. The aim of this study was to determine the response of C and N transformations in soil to tannins of different molecular

weight from Norway spruce (Picea abies(L.) Karst) and Scots pine (Pinus sylvestrisL.) needles, tannic acid and cellulose. Arginine was added to

test whether the soil microbial community was limited by the amount of N, and arginineCtannin treatments were used to test whether the effects

of tannins could be counteracted by adding N. Soil and needle samples were taken from adjacent 70-year-old Scots pine and Norway spruce stands

located in Kivalo, northern Finland. Tannins were extracted from needles and fractioned based on molecular weight; the fractions were then

characterized by LCMS and GCMS. Light fractions contained tannin monomers and dimers as well as many other compounds, whereas heavy

fractions consisted predominantly of polymerized condensed tannins. Spruce needles contained more procyanidin than prodelphinidin units, while

in pine needles prodelphinidin units seemed to be dominant. The fractions were added to soil samples, pine fractions to pine soil and spruce

fractions to spruce soil, and incubated at 14 8C for 6 weeks. CO2 evolution was followed throughout the experiment, and the rates of net

mineralization of N and net nitrification, concentration of dissolved organic N (DON) and amounts of microbial biomass C and N were measured

at the end of the experiment. The main effects of the fractions were similar in both soils. Light fractions strongly enhanced respiration and

decreased net N mineralization, indicating higher immobilization of N in the microbial biomass. On the contrary, heavy fractions reducedrespiration and slightly increased net N mineralization, suggesting toxic or protein-precipitating effects. The effects of tannic acid and cellulose

resembled those of light fractions. DON concentrations generally decreased during incubation and were lower with heavy fractions than with light

fractions. No clear differences were detected between the effects of light and heavy fractions on microbial biomass C and N. Treatments that

included addition of arginine generally showed trends similar to treatments without it, although some differences between light and heavy

fractions became more obvious with arginine than without it. Overall, light fractions seemed to act as a labile source of C for microbes, while

heavy fractions were inhibitors.

q 2005 Elsevier Ltd. All rights reserved.

Keywords: Forest soil; Microbial activities; Mineralization; Nitrogen cycling; Norway spruce; Polyphenols; Scots pine; Tannins

1. Introduction

Tannins are polyphenolic compounds with the ability toform stable complexes with proteins and other compounds.

They can be divided in to two main classes: condensed tannins,

which are also called proanthocyanidins, and hydrolyzable

tannins. Condensed tannins can be further divided into

subclasses such as procyanidins and prodelphinidins. While

gymnosperms and monocots produce only condensed tannins,

dicots can produce either condensed or hydrolyzable tannins or

a mixture of the two (reviewed byKraus et al., 2003). In woody

species, foliar concentrations of tannins commonly range from15 to 25% dry weight (reviewed byKraus et al., 2004). Tannins

may influence litter decomposition rates, humus formation,

nutrient (especially N) cycling and ultimately, plant nutrition

and growth (e.g. Schimel et al., 1996; Bradley et al., 2000;

Fierer et al., 2001). Tannins from various plant species have

been shown to affect N mineralization, induce toxicity in

microbes and affect enzyme activities in soil (Schimel et al.,

1996; Bradley et al., 2000; Fierer et al., 2001; Kraus et al., 2003

and references therein). Hence, there is very strong evidence

that tannins play an important role in interspecific competition,

and many studies have suggested that individual plants may be

important in nutrient cycling on the ecosystem level (Schimel

Soil Biology & Biochemistry 38 (2006) 13641374

www.elsevier.com/locate/soilbio

0038-0717/$ - see front matter q 2005 Elsevier Ltd. All rights reserved.

doi:10.1016/j.soilbio.2005.10.013

*Corresponding author. Tel.: C358 10 211 2595; fax: C358 10 211 2206.

E-mail address:[email protected] (S. Kanerva).
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et al., 1996; Chen and Stark, 2000; Fierer et al., 2001; Castells

et al., 2003; Kraus et al., 2004).

Concentration and composition of phenolic compounds in

the soil seem to vary depending on the plant species growing in

it (e.g.Kuiters and Denneman, 1987; Smolander et al., 2005).

Tannin reactivity in soil has been suggested to be based on such

characteristics as condensed versus hydrolyzable tannins andprocyanidin versus prodelphinidin content of the tannins

(Kraus et al., 2004). In addition, molecular weight or degree

of polymerization of tannins or phenolic compounds seems to

be an important factor when their influence on soil nutrient

cycling is considered. Schimel et al. (1996) and Fierer et al.

(2001) demonstrated that high molecular weight phenolics

from balsam poplar acted as a general microbial inhibitor,

while the effects of lower molecular weight phenolics were less

predictable and depended on prior exposure of the soil

microbial community to related molecules; microbial commu-

nities previously exposed to smaller chain tannins were more

likely to use them as a C substrate, while in the communities

that had limited exposure to tannins they were more likely to

prove toxic.

Norway spruce (Picea abies (L.) Karst) and Scots pine

(Pinus sylvestris L.) are the dominant tree species in Finland.

Both similarities and differences have been reported in the C

and N transformations of soil under these species (Priha and

Smolander, 1997, 1999; Smolander and Kitunen, 2002). The

aim of this study was to find out the response of soil C and N

transformations to tannins of different molecular weight from

Norway spruce and Scots pine needles. Fractions prepared

from spruce and pine needles were added to spruce and pine

soils, respectively, to examine their effects on microbial

activities by measuring CO2 evolution, net mineralization ofN and net nitrification rates, concentrations of dissolved

organic nitrogen (DON) and amounts of C and N in the

microbial biomass. The availability or inhibition of these

fractions to bacteria and fungi was also assessed.

2. Materials and methods

2.1. Study site, soil and needle sampling and chemical analysis

The stands used in this study were adjacent 70-year-old

stands in Kivalo, northern Finland (668200N/268400E), which

were dominated by Scots pine, Norway spruce or silver birch

(Betula pendula Roth) growing in soil that originally was

similar in all three stands. The soil type was podzolic and

humus type mor. Three study plots (25!25 m) were placed in

each stand. The coniferous stands also contained species other

than the dominant one (the spruce stand contained 76% spruce

and 24% other species, and the pine stand contained 88% pine

and 12% tree species other than pine). For a more detailed

description of the study site and the tree stands, seeSmolander

and Kitunen (2002).

In August 2001 soil samples (2030 cores, core diameter

58 mm) were taken systematically from the humus layer of

spruce and pine plots. The samples were combined to give one

composite sample per plot, and the composite samples from

each plot were combined to give one sample that represented

one stand. After the green plant material was removed, the

samples were sieved through a 4.0 mm mesh and stored in

plastic bags at 4 8C until used. Content of soil organic matter

(o.m.) was measured as loss-on-ignition at 550 8C. The soil

characteristics have been described earlier (Smolander and

Kitunen, 2002); the pH (H2O) of both conifer soils was 4.0, andthe C-to-N ratio in spruce soil was 37 and in pine soil 32.

Undamaged bulk green needles were collected from the pine

and spruce plots in spring 2001. After collection, the needles

were freeze dried and finely ground.

2.2. Extraction, fractionation and analysis of tannin fractions

Tannins were extracted and fractioned as described in Fierer

et al. (2001) but with some modifications. The ground plant

material (1300 g spruce/1070 g pine needles) was placed in a

steel container and 5 l of hexane was added. The material was

soaked in hexane and stirred occasionally with a power drill; at

the end the solvent was decanted from the plant material. The

plant material was extracted again with 2.5 l hexane, stirred and

decanted. This procedure was repeated two more times. After

that, the remaining plant material was extracted with acetone

water (70:30) overnight. The next day the suspension was

stirred and filtered through a filter paper (S & S 5893). The

filtrate was collected in a glass bottle. Needles were extracted

again with acetonewater (70:30), stirred 30 min and filtered.

The procedure was repeated once overnight. The three

acetonewater extracts were combined and concentrated by

roto-evaporation. This concentrated extract was then extracted

with 100% ethyl acetate for 30 min at 200 rev minK1. Ethyl

acetate and water were separated with a siphon, and theprocedure was repeated three more times. The ethyl acetate

fractions were combined and roto-evaporated to dryness. This

fraction was labelled F1.

The acetonewater fraction was loaded onto a Sephadex

LH-20 column that had previously been equilibrated with

methanolwater (50:50). The column was eluted with

methanolwater (50:50) followed by acetonewater (70:30)

until the eluate was colourless. The acetonewater fraction was

concentrated by roto-evaporation.

The concentrated acetonewater fraction was loaded into a

clean Sephadex LH-20 column that had previously been

equilibrated with methanolwater (50:50). The acetonewater

fraction was eluted with 100% ethanol, and the eluate was

concentrated by roto-evaporation and labelled F2.

The extract in the LH-20 column was then eluted with 100%

methanol. The eluate was concentrated and labelled F3.

Finally, the extract in the column was extracted with

acetonewater (70:30), and the eluate was concentrated and

labelled F4.

All four fractions were analyzed using thin layer chroma-

tography (TLC) to determine their composition. The solvent

system for TLC analyses was toluene/acetone/formic acid

(3/6/1). The components in each fraction were detected by UV-

light. In addition, the compositions of the fractions, in terms of

number and types of monomer units, molecular weights of

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proanthocyanidins (up to heptamers) and identification of

anthocyanidin monomers (procyanidin/prodelphinidin), as

well as a tannic acid product (Merck, Tannic acid powder

pure, DAB7, FU, pH Helv, USP) were confirmed by reversed-

phase and normal-phase high-performance liquid chromatog-

raphy (RP- and NP-HPLC) coupled with an ultraviolet (UV)

detector and electrospray-ionization mass spectrometer (ESIMS) (for methods see Loponen et al., 2001; Karonen et al.,

2004). Soluble condensed tannins were quantified by the

modified acid-butanol assay (proanthocyanidin assay) (Porter

et al., 1986; Terrill et al., 1992; Ossipova et al., 2001). The

standard curves for the calculations were determined by using

previously purified proanthocyanidin from leaves of mountain

birch (Betula pubescens ssp. czerepanovii) (Bae et al., 1993;

Ossipova et al., 2001). The content of low molecular weight

substances other than condensed tannins in the fractions were

analysed with GCMS. A known amount of each fraction was

weighed for GCMS analysis. Samples were analysed before

and after BSTFA derivatization and 4-chlorobenzoic acid,

erytritol and heptadecanoic acid were used as internal

references.

The fractions and the tannic acid product were mixed with

silica gel in a ratio of 1:2 (w:w), slurried in acetone or acetone

water and dried with rotary evaporation. The dry mixture was

ground in a mortar. This procedure binds tannin components to

the silica gel, which aids handling and application to soils.

2.3. Incubation experiment

Soil (corresponding to a volume of 20 ml) from both

coniferous stands was weighed into 120 ml glass bottles. The

treatments consisted of the four needle fractions (see above),tannic acid and cellulose, 45 mg gK1 soil o.m. each, and

control. All treatments were done in three replicates. Tannic

acid was added to test how the soils respond to hydrolyzable

tannins, since birch, which is often mixed with conifers in

natural forests, contains both hydrolyzable and condensed

tannins (Ossipova et al., 2001). Cellulose was used as a

compound that would supply C to the soil microbia without

having any specific physiological effects (such as toxicity). All

the same treatments were also done with arginine addition

(1 mg arg-N gK1 o.m.). Arginine was added to test whether the

microbial community in the soil was limited by the amount of

N, and adding arginineCneedle fractions tested whether the

effects of the fractions could be counteracted by adding N.

Fraction-silica mixture and cellulose were added to soil dry,

while arginine was added in solution. To determine how

addition of silica gel affected soil processes, soil with and

without addition of silica was also incubated. Since the changes

in the soil properties measured were affected not much by silica

gel alone, we conclude that adding silica gel with tannin

fractions did not distort the results of this experiment, and

therefore silica gel alone is not discussed later in this article.

The four fractions isolated from spruce needles were added

to the spruce soil and those from pine needles were added to the

pine soil. After the additions, the soils were adjusted to 60%

water saturation.

The samples were incubated at 14 8C for 42 or 43 days, and

their moisture content was adjusted weekly. CO2 evolution

bottles were sealed with rubber septa, and the other bottles

were capped with aluminium foil. Some samples of pure soil

and controls had been frozen immediately before incubation

and were extracted concurrently with the incubated samples.

The reason for this was to determine the initial concentrationsof NHC4N, NO

K

2CNOK

3 N and total N in the soils.

2.4. Chemical and microbiological analyses

During the 6-week incubation, CO2evolution was measured

10 times by sampling the headspace and analyzing the amount

of CO2 on a gas chromatograph (Priha and Smolander, 1997).

At the beginning of incubation, CO2 was analyzed more

frequently than in the end of the run. The bottles were aerated

24 h before each sampling. To determine net N mineralization

rates over the 6-week incubation period, samples were

extracted at time zero and after the incubation with 40 ml

1 M KCl (2 h 200 rev minK1) and filtered through S & S 5893

filter papers. Concentrations of NHC4N, NOK

2CNOK

3 N and

total N were measured using a flow injection analyzer (FIAstar

5012 analyzer C5042 detector, Tecator). To calculate net

ammonification and nitrification, initial concentrations of NHC4N and NOK2CNO

K

3 N from non-incubated samples were

subtracted from the final (post-incubation) NHC4N and NOK

2

CNOK3 N concentrations. DON was calculated as the

difference between total N and inorganic N.

At the end of the incubation period, the amounts of C (Cmic)

and N (Nmic) in the microbial biomass were measured using the

chloroform fumigation-extraction method (Priha and Smo-

lander, 1997). Carbon and nitrogen contents in the microbialbiomass were calculated by subtracting the total C and N in

unfumigated samples from that in fumigated samples. Because

the results represent a flush of C and N instead of the total

microbial biomass, flushes were converted to microbial

biomass with the formulas of Martikainen and Palojarvi

(1990), which are for Cmic in humus samples (1.9 CfC428)

mg gK1 dry mass (d.m.) and for Nmic(1.86NfC74.82) mg gK1

d.m (Cf and Nf represent amounts of C and N measured in

fumigation-extraction).

In addition to the incubation experiment, water extracts of

the needle fractions (F1F4) and tannic acid were used to

assess the availability or inhibition of these amendments to soil

bacteria and fungi. Fractions (45350 mg) were shaken

(250 rev minK1) for 1 h with 20 ml of water and filtered

through a 0.45 mm PES membrane. The concentrations of C

were measured, and two concentrations of C were chosen for

testing the conditions where the extracts of the fractions could

act as substrates (30 mg C lK1) for or inhibitors (700 mg C lK1)

to bacterial and fungal growth. A preliminary experiment was

done with F3 and F4, which were expected to inhibit microbes

(Schimel et al., 1996; Fierer et al., 2001). Based on the results

of that experiment, 700 mg C lK1 was chosen as an inhibitory

concentration, being near the 50% inhibition of bacterial

growth compared to growth in water solution. As an inoculum

of bacteria and fungi we used a mixture of humus layer

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collected from the same study site and also including a birch

stand (Kiikkilaet al., 2005).

The rate of bacterial growth was measured using the 3H-

thymidine incorporation technique on bacteria extracted from

soil (Baath, 1992), with the modifications introduced byBaath

et al. (2001) and Kiikkilaet al. (2005).Inoculum was prepared

by shaking a soil sample (3 g fresh weight, fw) with 100 ml ofwater, followed by low-speed centrifugation and filtration

through quarz wool. Two replicates of the mixture of bacterial

suspension (8.6 ml) and the extracts of the fractions (11.4 ml)

were prepared, and after pre-incubation of the suspensions

(shaken in 80 rev minK1) at 20 8C for 1, 2, 3, 7 and 8 days, the

rate of thymidine incorporation was measured. Then 1.4 ml of

the suspension was transferred to Eppendorf tubes, 3.5 ml of

[methyl-3H]thymidine (925 GBq mmolK1) was added, and the

samples were incubated for 2 h. Washing of excess tracer and

measurement of radioactivity are described in detail byBaath

et al. (2001).

The rate of fungal growth was measured using the techniqueof 14C-acetate incorporation into ergosterol devised byNewell

and Fallon (1991)and modified byBaath (2001)for use in the

soil habitat. Extracts of fractions (30 mg C lK1, 30 ml) were

transferred into Erlenmeyer flasks, and 0.03 g fw of soil was

added. To diminish the bacterial growth, streptomycin and

ampicillin (50 mg lK1) were added. The bottles were shaken at

80 rev minK1 in the dark at 20 8C for 1, 2, 7 and 8 days. After

this pre-incubation, the suspension was filtered through a

Whatman GF/D glass fiber filter. The soil and the filter were

transferred to a test tube with 1.5 ml of the extract. Samples of

700 mg C lK1 were used only after one-day pre-incubation in

test tubes (0.03 g fw soil and 1.5 ml extract). After pre-incubation, 14C-acetate solution (0.05 ml, 1,2,-[14C]acetic acid,

sodium salt, 2.07 GBq mmolK1) and 1 mM non-radioactive

acetate (0.45 ml) were added to the test tubes. After the mixture

was incubated for 20 h at 20 8C, 1 ml of 5% formalin was

added, the test tubes were centrifuged and the supernatant was

discarded. The ergosterol was then extracted (Baath, 2001) and

measured with HPLC (Hitachi, Merck), and 14C-ergosterol was

determined with the HPLC radioactivity monitor (Berthold, LB

506 C-1). The proportion of radioactivity of the total ergosterol

was calculated.

The mean of three samplings (30 mg C lK1) was calculated

(bacterial growth: after 1, 2, and 3 days pre-incubation; fungal

growth: after 1, 7 and 8 days pre-incubation). For 700 mg C lK1

the sampling after one-day pre-incubation was used so that

the inhibition effect could be measured before the microbesadapted. Relative availability (30 mg C lK1) or inhibition

(700 mg C lK1) of fractions to bacteria (TdR 30, TdR 700) or

fungi (Ac-erg 30, Ac-erg 700) was calculated by dividing the

incorporation of the sample by the incorporation in water

solution (valueZ1). Thus the higher the TdR 30 or Ac-erg 30

value the better the fraction is as substrate. Low TdR 700 or

Ac-erg 700 values mean that the fraction inhibited growth of

bacteria.

2.5. Statistical analysis

Differences in the measured characteristics betweentreatments were compared with one-way analysis of variance.

ANOVA was performed separately for the two soil-treatment

combinations. When needed, transformations were made to

fulfill the assumptions of the analysis of variance. Significant

differences of the means by treatments were determined by

Tukeys test using a significance level ofP!0.05.

3. Results

Chromatographic and mass spectrometric results from

TLC, RP- and NP-HPLC-UV/ESI-MS analyses revealed that

both Norway spruce and Scots pine needle fractions F3 andF4 contained polymers of condensed tannins that were

longer than those in the F1 and F2 fractions. Acid-butanol

assay showed that fractions F3 and F4 consisted mainly of

condensed tannins (5587%) while F1 and F2 contained

only 1.75.5% condensed tannins (Table 1). Minor amounts

of several compounds other than condensed tannins were

found in light fractions, especially in F1 (Table 2), the rest

containing other needle constituents.

Table 1

Qualitative and quantitative data for fractions of condensed tannins from spruce and pine needles

Fraction Polymeric composition of fractions Molecular weight

of fractions

Estimation of the

most abundantCT unit

CT concentration

in fractions(g kgK1)

Spruce F1 Mono-dimers 290610 PC 39

F2 Mono-trimers 290914 PC 55

F3 Tetra-heptamers and higher tannin polymers, minor

amounts of mono-trimers

11542130 and

higher

PC 622

F4 Tetra-hexamers and higher tannin polymers, trace

amounts of mono-trimers

11541826 and

higher

PC 853

Pine F1 Mono-dimers 290610 PD 20

F2 Mono-dimers 290610 PD 17

F3 Tri-tetramers and higher tannin polymers, minor

amounts of mono-dimers

8661218 and

higher

PD 551

F4 Tetramers, pentamers and higher tannin polymers,

trace amounts of mono-trimers

11541522 and

higher

PD 870

CT, condensed tannins; PC, procyanidin; PD, prodelphinidin.


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Spruce needles contained more procyanidin than prodel-

phinidin units while in pine needles prodelphinidin units were

dominant. In addition, HPLC-ESI-MS analysis confirmed that

the tannic acid product contained a mixture of galloylglucoses

of different molecular sizes (see also Hagerman and Butler,

1989). Acid-butanol assay with tannic acid gave no indication

of condensed tannins.

In both soils, and with and without arginine, fractions F1

and F2 and tannic acid caused a sharp increase in CO2

production during the first days after addition; but after that C

mineralization settled to the same level as the control ( Fig. 1).

In contrast, throughout the incubation experiment both spruce

and pine F3 and F4 fractions decreased C mineralization rates

relative to the control. Cellulose showed slightly higher CO2production in the spruce soil during last weeks of the

incubation than the other treatments did.

Net N mineralization rates measured over the 6-week

incubation were mostly negative in the absence of arginine

Table 2

Compounds other than condensed tannins in spruce and pine needle fractions analysed after BSTFA derivatization

Compound (mg kgK1) Spruce Pine

F1 F2 F3 F4 F1 F2 F3 F4

Low molecular weight

phenols

518 4.5

Sesqui-and diterpenes 170 10Resin acids 108 148 2.7

Glucose 34 14 7.9 7.6 6.7

Phenolic glucosides 45 21 1.1

Stilbene glucosides 41 292

Flavonoid monomers 228 117 6.3 4.3 233 80 38 4.0

Sterols 6.4 5.6

Cyclitols 44 23

Fatty acids 23 4.6

Fig. 1. Respiration rates during 6-week incubation for spruce and pine control soils and soils treated with spruce or pine needle fractions F1F4, respectively, tannic

acid or cellulose and the corresponding treatments with arginine addition. Mean of three replicates.


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(initial concentrations of NH4N were in the spruce soil

without and with arginine 442 and 450 mg kgK1 o.m.,respectively, and in the pine soil without and with arginine

229 and 264 mg kgK1 o.m., respectively). In the spruce soil, F2

and especially F1 showed significantly lower rate of net N

mineralization whereas F3 and F4 showed significantly higher

rates of net N mineralization than the control did (Fig. 2).

Tannic acid and cellulose also showed significantly lower net N

mineralization than the control did. In the pine soil the trends

were similar, but the differences were smaller and F3 did not

differ from the control (Fig. 2). With arginine addition, the

trends observed in both soils were similar and net N

mineralization rates were positive, with the exception of

cellulose added to spruce soil (Fig. 2). In the spruce soil the rateof net nitrification was very low while in the pine soil there was

no clearly measurable NOK3 production (results not shown).

In both of the soils in the absence of arginine, all needle

fraction and tannic acid treatments showed lower DON

concentrations than the control did (pine F1 and F2 not signif.)

(Fig. 3). Cellulose had no effect on concentration of DON.

Similar trends were observed with arginine addition (Fig. 3),

but in both soils only the fractions F3 and F4 clearly decreased

DON and in the pine soil the F1 fraction increased DON.

In neither of the soils in the absence of arginine did the

treatments significantly affect the amount of microbial biomass

C, except for pine F3, which decreased it significantly (Fig. 4).

With arginine addition, F1 in spruce soil and F2 in pine soil

increased Cmic significantly during the incubation while theother treatments had no effect (Fig. 4).

In the spruce soil in the absence of arginine, addition of

tannic acid resulted in significantly higher Nmicvalues than in

the control, while the other treatments had no effect (Fig. 5). In

the pine soil, F1 and cellulose increased Nmic slightly but

significantly and F4 decreased it significantly, while the other

treatments had no effect (Fig. 5). In the spruce soil, with

arginine addition all treatments (except F3) showed signifi-

cantly lower Nmicvalues than the control did, while in the pine

soil none of the treatments differed from the control.

The relative availability of the fractions to bacteria and

fungi was studied by 3H-thymidine and 14C-acetate incorpor-ation using concentrations of 30 and 700 mg C lK1. The

highest relative availability to bacteria at the concentration of

30 mg C lK1 (TdR 30) was in pine F1 and F2, while all other

fractions and tannic acid were less available to bacteria. At the

concentration of 700 mg C lK1 (TdR 700) all fractions

inhibited bacterial growth; F1 and tannic acid showed the

highest inhibition (Fig. 6). When assessed by Ac-erg 700

(valuesO1), none of the treatments, except for tannic acid,

inhibited fungi (Fig. 6). However, if growth at the higher

concentration (Ac-erg 700) is lower than at the lower

concentration (Ac-erg 30), inhibition is also possible. In spruce

and pine F3, Ac-erg 700 was clearly lower than Ac-erg 30, thus

Fig. 2. Rate of net N mineralization during 6-week incubation for spruce and pine control soils and soils treated with spruce or pine needle fractions F1F4,

respectively, tannic acid or cellulose and the corresponding treatments with arginine addition. ANOVA was performed separately for the two soil-treatment

combinations. Significant differences (P!0.05) between the means of one soil-treatment combination are marked with different letters. Mean and SE for three

replicates.


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indicating inhibition. In pine F4 and in spruce F1, however, Ac-

erg 700 was very high and Ac-erg 30 was very low; but the

variation was high. In spruce F2 and pine F1, Ac-erg 700 was

nearly the same as Ac-erg 30.

4. Discussion

The procedure used here to extract and fractionate tannins

was similar to that ofFierer et al. (2001) with balsam poplar.

Light fractions F1 and F2 from spruce and pine needles alsocontained compounds other than tannins. In addition to the

analyzed compounds (Table 2), both spruce and pine F1

probably contained chemically neutral and also higher

molecular weight compounds like waxes, chlorophyll and

terpenoids, which are soluble in organic solvents. F2 probably

contained more polar compounds, such as phenols and

oligomeric phenols. Therefore, we cannot specify all the

effects of fractions F1 and F2 treatments as tannin effects, and

therefore we find it more justified to discuss only effects of

light fractions.

Our results indicated mainly parallel effects of spruce- and

pine-needle fractions in spruce and pine soils, respectively; butsome differences, mostly in magnitude, were also detected. The

effects caused by the lighter fractions F1 and F2 were mainly

Fig. 3. Concentrations of DON after 6-week incubation for spruce and pine control soils and soils treated with spruce or pine needle fractions F1F4, respectively,tannic acid or cellulose and the corresponding treatments with arginine addition. ANOVA was performed separately for the two soil-treatment combinations.

Significant differences (P!0.05) between the means of one soil-treatment combination are marked with different letters. Mean and SE for three replicates.

Fig. 4. C in microbial biomass after 6-week incubation for spruce and pine control soils and soils treated with spruce or pine needle fractions F1F4, respectively,

tannic acid or cellulose and the corresponding treatments with arginine addition. ANOVA was performed separately for the two soil-treatment combinations.



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contrary to those caused by the heavier fractions F3 and F4, as

was also observed Fierer et al. (2001) with Populus

balsamiferaleaf fractions.

Based on the results for soil respiration, the lighter fractions

seemed to act as a C source for microbes, while the heavier

fractions were inhibitors. Results for the net N mineralization

also indicated the difference between light and heavy fractions;

in both soils net N mineralization was clearly lower with F1

and F2 than with F3 and F4. Effects of tannic acid and cellulose

resembled the effects of light fractions more than those ofheavy fractions.

One reason for inhibition of respiration by heavy fractions

may be the same asSchimel et al. (1996)suggested for balsam

poplar tannins: inhibition of exoenzyme activity or complexa-

tion of proteinaceous substrates. Bradley et al. (2000) also

expected that decreases in mineral N cycling would be a result

of the ability of Kalmia angustifolia and balsam fir tannins to

bind and sequester organic sources of N. Moreover,Kumar and

Horigome (1986)showed that in black locust (Robinia pseudo

acacia) the protein-precipitating capacity and the percentage of

protein-precipitable phenolics increased with increasing degree

of polymerization of the tannin fractions, and Kraus et al.

(2003)reported that high molecular weight tannins precipitate

more protein than low molecular weight tannins do. In our

study, fractions F3 and F4 contained longer tannin polymers

than fractions F1 and F2 did (Table 1). Therefore it is likely,

that protein precipitation could have played an important role

in inhibiting respiration in soils treated with heavy fractions.

Most fractions seemed to slightly reduce DON, but heavy

fraction treatments slightly more than light fraction treatments.

This points to protein precipitation by the heavy fractions,

since probably most of protein-tannin complexes do not appear

in DON, due to their weak extractability. According toNorthup

et al. (1995), the soil DON:mineral-N ratio correlates with soil

tannin concentration; but our soils did not generally show

increased DON:mineral-N ratios (results not shown) due to

addition of the needle fractions. However, it is possible that

added tannins do not affect soil in the same way as natural soil

tannins do.

Inhibition of soil respiration by heavy fractions may also

have been due to toxic effects. Since heavy-fraction treatments

with addition of arginine showed similar results for soil

respiration as treatments without added arginine, it is possible

that N was not a limiting factor in our soils, but more probably,

arginine addition was not large enough to overcome the

Fig. 5. N in microbial biomass after 6-week incubation for spruce and pine control soils and soils treated with spruce or pine needle fractions F1F4, respectively,

tannic acid or cellulose and the corresponding treatments with arginine addition. ANOVA was performed separately for the two soil-treatment combinations.


Fig. 6. Relative availability (30 mg C lK1) and inhibition (700 mg C lK1) of

spruce and pine needle fractions F1F4 and tannic acid (TA) to bacteria (TdR)

and fungi (Ac-erg). Bars represent SE for three replicates.


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negative effects of the precipitation of organic N compounds or

that the inhibition mechanism of heavy fractions was other than

substrate complexation, e.g. toxicity. This conclusion is also

supported by C in microbial biomass, since in some cases Cmicwas slightly decreased by heavy fractions, although not

significantly, which could indicate direct toxic effects of

tannins on the microbial community or decreased enzymeactivities (Kraus et al., 2004). Therefore slight increases in net

N mineralization by heavy fractions relative to the control were

likely a consequence of reduced microbial activity and N

uptake rather than the result of gross mineralization of N

becoming more effective. The rate of N mineralization may not

have decreased as much as the rate of N immobilization, which

would result in accumulation of mineral N in the soil.

Decreased net mineralization of N by light fractions does

not necessarily indicate a reduction in gross mineralization of

N. More probably, mineralized N was immobilizated by soil

microbes since soil respiration increased shortly after addition

of the fractions. This indicates that the compounds in those

fractions were easily metabolized, as has also been seen in

other studies with different plant species (Basaraba, 1964;

Schimel et al., 1996; Fierer et al., 2001; Castells et al., 2003;

Kraus et al., 2004). However, microbial biomass C and N were

not significantly increased with additions of light fractions,

except only Nmic with pine F1.Kraus et al. (2004)also found

no increase in the amounts of C and N in the microbial biomass

with additions of purified tannins from different plant species.

Fierer et al. (2001)observed an increased C-to-N ratio in the

microbial biomass due to addition of tannins from balsam

poplar, and especially additions of the lighter fractions. This

was also observed in our study, but only in spruce soil with F1,

and when arginine was added, in spruce soil with both lightfractions and in pine soil with F2 (results not shown). In spruce

soil with arginine the addition of tannic acid and cellulose

clearly increased the C-to-N ratio in the microbial biomass, but

in pine soil with added arginine increased it only slightly. In

general, while the trends with arginine were mainly similar to

those without it, in pine soil, arginine addition made the

differences in net N mineralization and DON concentrations

between light and heavy fractions more obvious, which

indicates that for determining the maximal effects of the

fractions, nitrogen addition was essential. However, better

understanding of the N flux in both soils could be obtained

using 15N labelling.

In some studies, nitrification has been found to be inhibited

by phenolic compounds (Basaraba, 1964; Thibault et al., 1982;

Baldwin et al., 1983), but it is unclear, is the mechanism

specific or only dependent on the change in net ammonifica-

tion. In this study, however, nitrate concentrations were always

very low, close to the limit of determination; therefore, it is not

relevant to make conclusions about net nitrification based on

them.

Some differences were found between spruce soil with

spruce-needle fractions and pine soil with pine-needle

fractions, but in both soils the main trends were similar.

Spruce needles contained more procyanidin units, while in

pine needles prodelphinidin units were dominant (Table 1).

This is in agreement with the results of Maie et al. (2003)

where the procyanidin/prodelphinidin ratio was 70:30 in

Norway spruce needles, and those of Kraus et al. (2004)

with bishop pine needles in which the percentage of

procyanidin monomer units versus prodelphinidin monomer

units was 22%. Procyanidins and prodelphinidins differ from

each other in terms of the hydroxylation pattern in theB-ring, which in terms of reactivity is a critical factor for

condensed tannins (Kraus et al., 2004). Hernes et al. (2001)

reported that, compared to procyanidin units, prodelphinidin

units may be structurally less stable and thus more prone to

chemical transformation by abiotic processes. In addition,

different protein-binding capacity for the procyanidin-type

and the prodelphinidin-type condensed tannins have been

suggested to affect the total amount of extractable free

condensed tannins in forest soils (Hernes et al., 2001; Maie

et al., 2003).

When the control, tannic acid and cellulose treatments

were compared in order to ascertain the differences betweenspruce and pine soils, spruce soil seemed to be somewhat

more active than the pine soil, because it showed higher

respiration rates than pine soil did. This difference was to be

expected because spruce soil from this particular stand has

already previously been shown to have higher rates of C

mineralization and net N mineralization than the soil under

pine (Smolander and Kitunen, 2002). No toxic effects of

tannins in light fractions were observed in either of the soils,

which was probably due to the addition of fractions only in

their natural soils where the microbes were adapted to use the

compounds contained as C substrates. On the other hand,

concentrations of condensed tannins found in the light

fractions were so low that it may not be possible to detect

their potential toxic effects.

At the concentration of 700 mg C lK1, none of the spruce or

pine needle fractions was clearly inhibitory to fungi. It is

possible, however, that heavy fractions (except pine F4) may

have been somewhat inhibitory to fungi because fungal growth

rate was lower at the concentration of 700 mg C lK1 than at the

concentration of 30 mg C lK1. In contrast, at the concentration

of 700 mg C lK1, all fractions seemed to be inhibitory to

bacteria and at the concentration of 30 mg C lK1, available to

bacteria. These results did not clearly support the results of the

soil incubation experiment. The soil incubation experiment,

where fungi may out-compete the bacteria, may indicate morefungal than bacterial activity. In addition, bacteria and fungi

may play different roles in mineralization of soil organic

matter; fungi and bacteria have been shown to be specialized to

grow on different types of phenolic compounds (Blum and

Shafer, 1988; Ley and Schmidt, 2002). It is also possible that

the concentration of 700 mg C lK1 was not enough to inhibit

fungi, which are suggested to degrade the most refractory

compounds (Mller et al., 1999). These results must, however,

be interpreted and compared to those from the soil incubation

experiment with caution, because only the water-extractable

part of tannin fractions was studied and the application of the

used methods was novel.


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4.1. Conclusions

Spruce and pine needle fractions seem to play an important

role in controlling transformations of C and N in spruce and

pine soils. Light fractions, consisting of tannin mono- and

dimers and also many other compounds, were used as C

substrates, while heavy fractions, consisting mainly of thepolymerised condensed tannins, inhibited growth. However,

these results cannot be applied directly to natural forest soil,

since in nature, light and heavy fractions are introduced to soil

concurrently through litter and leachates, addition is more or

less continuous and it is not known whether in natural

conditions the effects of light fractions overcome the effects

of heavy fractions or vice versa. In the future, it would be

reasonable to find out the differences between the effects of

spruce and pine needle fractions, and to explore whether

differences between spruce and pine soils can be explained by

those compounds.

Acknowledgements

We are grateful to Anneli Rautiainen and Pauli Karppinen

for excellent laboratory work, to Anne Siika for making the

figures and to Dr Joann von Weissenberg for checking the

English language of this paper. Many thanks to Prof Erland

Baath for discussions and comments on the results of 14C-

acetate-in-ergosterol method. This study was supported by the

Academy of Finland.

References

Baath, E., 1992. Thymidine incorporation into macromolecules of bacteria

extracted from soil by homogenizationcentrifugation. Soil Biology &

Biochemistry 24, 11571165.

Baath, E., 2001. Estimation of fungal growth rates in soil using 14C-acetate

incorporation into ergosterol. Soil Biology & Biochemistry 33, 20112018.

Baath, E., Pettersson, M., Soderberg, K.H., 2001. Adaptation of a rapid and

economical microcentrifugation method to measure thymidine and leucine

incorporation by soil bacteria. Soil Biology & Biochemistry 33,

15711574.

Bae, H.-D., McAllister, T.A., Muir, A.D., Yanke, L.J., Bassendowski, K.A.,

Cheng, K.-J., 1993. Selection of a method for condenced tannin analysis for

studies with rumen bacteria. Journal of Agricultural and Food Chemistry

41, 12561260.

Baldwin, I.T., Olson, R.K., Reiners, W.A., 1983. Protein binding phenolics and

the inhibition of nitrification in subalpine balsam fir soils. Soil Biology &Biochemistry 15, 419423.

Basaraba, J., 1964. Influence of vegetable tannins on nitrification in soil. Plant

and Soil 21, 816.

Blum, U., Shafer, S.R., 1988. Microbial populations and phenolic acids in soil.

Soil Biology & Biochemistry 20, 793800.

Bradley, R.L., Titus, B.D., Preston, C.P., 2000. Changes to mineral N cycling

and microbial communities in black spruce humus after additions of

(NH4)2SO4and condensed tannins extracted from Kalmia angustifolia and

balsam fir. Soil Biology & Biochemistry 32, 12271240.

Castells, E., Penuelas, J., Valentine, D.W., 2003. Influence of the phenolic

compound bearing species Ledum palustre on soil N cycling in boreal

hardwood forest. Plant and Soil 251, 155166.

Chen, J., Stark, J.M., 2000. Plant species effects and carbon and nitrogen

cycling in a sagebrush-crested wheatgrass soil. Soil Biology &


Fierer, N., Schimel, J.P., Cates, R.G., Zou, J., 2001. Influence of balsam poplar

tannin fractions on carbon and nitrogen dynamics in Alaskan taiga

floodplain soils. Soil Biology & Biochemistry 33, 18271839.

Hagerman, A.E., Butler, L.G., 1989. Choosing appropriate methods and

standards for assaying tannin. Journal of Chemical Ecology 15, 17951810.

Hernes, P.J., Benner, R., Cowie, G.L., Goni, M.A., Bergamaschi, B.A.,

Hedges, J.I., 2001. Tannin diagenesis in mangrove leaves from a tropical

estuary: a novel molecular approach. Geochimica et Cosmochimica Acta

65, 31093122.

Karonen, M., Loponen, J., Ossipov, V., Pihlaja, K., 2004. Analysis of

procyanidins in pine bark with reversed-phase and normal-phase high-

performance liquid chromatography-electrospray ionisation-mass spec-

trometry. Analytica Chimica Acta 522, 105112.

Kiikkila, O., Kitunen, V., Smolander A., 2005. Degradability of dissolved

organic carbon and nitrogen in relation to tree species. FEMS Microbiology

Ecology 53, 3340.

Kraus, T.E.C., Dahlgren, R.A., Zasoski, R.J., 2003. Tannins in nutrient

dynamics of forest ecosystemsa review. Plant and Soil 256, 4166.

Kraus, T.E.C., Zasoski, R.J., Dahlgren, R.A., Horwath, W.T., Preston, C.M.,

2004. Carbon and nitrogen dynamics in a forest soil amended with purified

tannins from different plant species. Soil Biology & Biochemistry 36,

309321.

Kuiters, A.T., Denneman, C.A.J., 1987. Water-soluble phenolic substances insoils under several coniferous and deciduous tree species. Soil Biology &


Kumar, R., Horigome, T., 1986. Fractionation, characterization, and protein-

precipitating capacity of the condensedtannins fromRobinia pseudo acacia

L. leaves. Journal of Agricultural and Food Chemistry 34, 487489.

Ley, R.E., Schmidt, S.K., 2002. Fungal and bacterial responses to phenolic

compounds and amino acids in high altitude barren soils. Soil Biology &


Loponen, J., Lempa, K., Ossipov, V., Kozlov, M.V., Girs, A., Hangasmaa, K.,

Haukioja, E., Pihlaja, K., 2001. Patterns in content of phenolic compounds

in leaves of mountain birches along a strong pollution gradient. Chemo-

sphere 45, 291301.

Maie, N., Behrens, A., Knicker, H., Kogel-Knabner, I., 2003. Changes in the

structure and protein binding ability of condensed tannins during

decomposition of fresh needles and leaves. Soil Biology & Biochemistry35, 577589.

Martikainen, P.J., Palojarvi, A., 1990. Evaluation of the fumigationextraction

method for the determination of microbial C and N in a range of forest soils.


Mller, J., Miller, M., Kjller, A., 1999. Fungal-bacterial interaction on beech

leaves: influence on decomposition and dissolved organic carbon quality.


Newell, S.Y., Fallon, R.D., 1991. Towards a method for measuring

instantaneous fungal growth rates in field samples. Ecology 72,

15471559.

Northup, R.R., Yu, Z., Dahlgren, R.A., Vogt, K.A., 1995. Polyphenol control of

nitrogen release from pine litter. Nature 377, 227229.

Ossipova, S., Ossipov, V., Haukioja, E., Loponen, J., Pihlaja, K., 2001.

Proanthocyanidins of mountain birch leaves: quantification and properties.

Phytochemical Analysis 12, 128133.Porter, L.J., Hrstich, L.N., Chan, B.C., 1986. The conversion of procyanidins

and prodelphinidins to cyanidin and delphinidin. Phytochemistry 25,

223230.

Priha, O., Smolander, A., 1997. Microbial biomass and activity in soil and

litter under Pinus sylvestris, Picea abies and Betula pendula at

originally similar field afforestation sites. Biology and Fertility of

Soils 24, 4551.

Priha, O., Smolander, A., 1999. Nitrogen transformations in soil under Pinus

sylvestris, Picea abiesand Betula pendula at two forest sites. Soil Biology

& Biochemistry 31, 965977.

Schimel, J.P., Van Cleve, K., Cates, R.G., Clausen, T.P., Reichardt, P.B., 1996.

Effects of balsam poplar (Populus balsamifera) tannins and low molecular

weight phenolics on microbial activity in taiga floodplain soil: implications

for changes in N cycling during succession. Canadian Journal of Botany 74,

8490.


8/11/2019 1-s2.0-S0038071705004013-main pino

11/11

Smolander, A., Kitunen, V., 2002. Soil microbial activities and characteristics

of dissolved organic C and N in relation to tree species. Soil Biology &


Smolander, A., Loponen, J., Suominen, K., Kitunen, V., 2005. Organic matter

characteristics and C and N transformations in the humus layer under two

tree species,Betula pendulaandPicea abies. Soil Biology & Biochemistry

37, 13091318.

Terrill, T.H., Rowan, A.M., Douglas, G.B., Barry, T.N., 1992. Determination of

extractable and bound condensed tannin concentrations in forage plants,

protein concentrate meals and cereal grains. Journal of the Science of Food

and Agriculture 58, 321329.

Thibault, J.-R., Fortin, J.-A., Smirnoff, W.A., 1982. In vitro allelopathic

inhibition of nitrification by balsam poplar and balsam fir. American

Journal of Botany 69, 676679.


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