Tannins can slow-down but also speed-up soil enzymatic activity in boreal forest

Soil Biology & Biochemistry 107 (2017) 60e67

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Tannins can slow-down but also speed-up soil enzymatic activity in boreal forest € m b, Bartosz Adamczyk a, *, Maarit Karonen b, Sylwia Adamczyk a, Marica T. Engstro €a € a, Veikko Kitunen a, Aino Smolander a, Judy Simon c Tapio Laakso a, Pekka Saranpa a b c

Natural Resources Institute Finland, Vantaa Research Unit, PO Box 18, FI 01301, Vantaa, Finland Laboratory of Organic Chemistry and Chemical Biology, Department of Chemistry, University of Turku, FI 20014, Turku, Finland €tsstrasse 10 DE 78457, Konstanz, Germany Chair of Plant Physiology and Biochemistry, Department of Biology, University of Konstanz Universita

a r t i c l e i n f o

a b s t r a c t

Article history: Received 20 July 2016 Received in revised form 29 December 2016 Accepted 30 December 2016

The boreal forest ecosystem stores substantial amounts of soil organic matter (SOM), which may act either as a source or as a sink for atmospheric CO2 under climate change and that is why enzymatic SOM degradation is gaining increasing attention. The boreal forest ecosystem is rich in plant secondary compounds and in particular tannins which are seen as enzyme inhibitors. We studied changes in enzymatic activity after the addition of tannins. Our experimental design combined direct studies of the tannin effects on enzymes with laboratory soil mesocosm experiments. Our results showed that the addition of tannins directly led to both decreases and increases in the catalytic activity of enzymes, however, some differences between enzymes were observed. Overall, low concentrations of tannins increased the coiled structures of the enzymes boosting their catalytic activity. High concentrations of tannins acted in the opposite way, thereby diminishing the catalytic activity. We observed that tannins caused a similar change in enzymatic activity in soil mesocosm experiments. Tannin-enzyme synergy needs more study as these interactions can potentially play an important role in SOM decomposition of future climate, especially in the tannin-rich ecosystems. © 2016 Elsevier Ltd. All rights reserved.

Keywords: Boreal forest soil Enzymatic activity Soil organic matter Tannins

1. Introduction Soil organic matter (SOM) contains more carbon (C) than global vegetation and the atmosphere combined and therefore SOM is a crucial component in the C cycle (Farrior et al., 2015; Kaiser et al., 2015; Li et al., 2012; Schmidt et al., 2011). The enzymatic release of C from SOM and subsequent conversion into CO2 or CH4 can cause significant increase of atmospheric concentrations of these greenhouse gases, which in turn accelerate global warming (Kaiser et al., 2015; Li et al., 2012; Schmidt et al., 2011). Soil organic matter decomposition is gaining increasing attention for this reason (Hunter, 2008; Melillo et al., 2011). Ecosystems with high SOM content, like boreal forest, are of particular interest. The boreal forest soil ecosystem is characterized also by high content of plant secondary compounds of which tannins are the most abundant ~ uelas and Estiarte, 1998; Tharayil et al., 2011). Global group (Pen

* Corresponding author. Current address: University of Helsinki, Department of Food and Environmental Sciences, PO Box 27, FI 00014, Helsinki, Finland. E-mail address: Bartosz.Adamczyk@helsinki.fi (B. Adamczyk). http://dx.doi.org/10.1016/j.soilbio.2016.12.027 0038-0717/© 2016 Elsevier Ltd. All rights reserved.

scale assessments and future projections point to increase in temperature, drought, and levels of greenhouse gases (IPCC, 2013; Schindler and Lee, 2010). These factors affect the production of ~ uelas plant secondary compounds (Jamieson et al., 2015, 2012; Pen and Estiarte, 1998; V€ ais€ anen et al., 2013; Zhao et al., 2016). It was recently proven that trees produce more reactive tannins in response to climatic stress (Tharayil et al., 2011). Tannins are polyphenolic compounds usually divided into hydrolysable tannins (HT), and condensed tannins (CT, proanthocyanidins). The role of tannins in boreal forest soil span numerous functions, like defense against herbivores, metal complexation, influence on C and N cycling and inhibition of microbial activity €ttenschwiler and Vitousek, 2000; Kraus et al., 2003). Tannins (Ha are seen as protein precipitating agents and potential enzyme inhibitors (Adamczyk et al., 2012; Goldstein and Swain, 1963; Hagerman, 2012; Strumeyer and Malin, 1970; Uchida et al., 1987; Upadhyay and Singh, 2011). Numerous direct biochemical studies proved that tannins decrease enzymatic activity (Adamczyk et al., 2009; Goldstein and Swain, 1963; Hagerman, 2012; Uchida et al., 1987; Upadhyay and Singh, 2011), but in some cases only a minor decrease in activity was observed (Juntheikki and Julkunen-Tiitto,

B. Adamczyk et al. / Soil Biology & Biochemistry 107 (2017) 60e67

2000; Strumeyer and Malin, 1970). There is surprisingly little information on the interference of tannins with soil enzymes (Fierer et al., 2001; Joanisse et al., 2007). There is currently no evidence that demonstrates and explains the enhancement of enzymatic activity through reaction with tannins. Here we wanted to take a first step to explore the possibility that tannins can accelerate or decelerate enzymes responsible for SOM decomposition. We studied changes in enzymatic activity of betaglucosidase, acid phosphatase, arylsulfatase and chitinase, crucial enzymes in C, P, S, N cycling (Acosta-Martínez et al., 2007). We hypothesized that (i) tannins, through changes in enzyme conformation, can increase or decrease the enzyme activity, (ii) there are differences between enzymes in response to tannins, and (iii) both, increase and decrease of enzymatic activity through reaction with tannins, occur in soil conditions, but inhibitory effect is prevalent. 2. Materials and methods 2.1. Experimental setup Our experimental design combined direct biochemical studies of the tannin effects on enzymes (no soil included) with laboratory soil mesocosm studies. In direct biochemical part, we studied the effect of tannins in different concentrations on enzyme activity, kinetics of enzyme inhibition by high tannin concentration, enzyme precipitation through very high tannin concentrations and residual activity of precipitated enzymes. We also studied changes in enzyme secondary structure due to reaction with tannins. Biochemical studies took into account condensed tannins (CT) and hydrolysable tannins (HT). In laboratory soil mesocosm studies we added different amounts of tannins to two soils differing in characteristics including the native amounts of condensed tannins (called later CTrich and CT-poor soil). Under soil conditions we studied only the effect of CT as they constitute the major source of tannins and soil concentration of HT is very low (Adamczyk et al., 2009). We used CT in 3 doses, 10 mg, 50 mg and 100 mg per soil bottle. One soil bottle contained 1.26 g or 1.46 g SOM for CT-poor and CT-rich soil, respectively. Amount of 10 mg CT was equal to the native amount of CT in CT-rich soil. Incubation lasted for 20 days. This incubation time was chosen on the basis of earlier studies (e.g. Kanerva et al., 2006) in which changes in N and C transformations occurred up to 15 days, with no difference at later time points in comparison to control (i.e. no tannin addition). Below we outline the methods, but more details can be found in Supplementary Materials. 2.2. Characterization of tannins As representative of HT we used tannic acid (TA) which was already characterized (Adamczyk et al., 2012) according to Salminen and Karonen (2011). The TA contained simple galloyl glucoses (tri-, tetra- and pentagalloyl glucoses), gallotannins (hexato tridecagalloyl glucoses), but also gallic acid and digallic acid. The average molecular mass was approximated as 1000 Da. Condensed tannins from Norway spruce needles were extracted and fractionated as previously described (Fierer et al., 2001; Kanerva et al., 2006). The CT fraction was analyzed by two €m et al., different LC-MS methods (Karonen et al., 2011; Engstro 2014). 2.3. Biochemical studies of tannin effect on enzymatic activity Activities of acid phosphatase, beta-glucosidase, arylsulfatase and chitinase were measured as described earlier (de la Mata et al.,

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1993; Garzillo et al., 1996; Muller and Schmidt, 1986; Parham and Deng, 2000). The temperature during all incubations was 37
C. The conditions for enzyme measurements (time of incubation with substrate, pH, concentration of substrate) were optimized to obtain enzyme kinetics with the highest velocity. Briefly, acetic buffer was mixed with enzyme and 5 min later tannins were added. After 10 min of incubation a substrate was added and mixtures were incubated for 5 min (acid phosphatase), 10 min (beta-glucosidase and chitinase) or 15 min (arylsulfatase). Following substrates were used: for glucosidase p-nitrophenyl b-D-glucopyranoside, for acid phosphatase p-nitrophenyl phosphate, for arylsulphatase p-nitrophenyl sulfate and for chitinase p-nitrophenyl N-acetyl-b-D-glucosamidine. After the incubation, 25 ml of incubation mixture was transferred to a new tube containing 600 ml of 0.5 M NaOH. After 10 min, the absorbance was measured at 405 nm. The results are presented as residual activity of the control (100% activity, no tannin addition). All analyses were done in 5 laboratory replicates. To study kinetics of enzyme inhibition we used the same methods as for the measurements of enzyme activity with following concentrations of substrates: for glucosidase 10e60 mM, for acid phosphatase 5e40 mM, for arylsulphatase and chitinase 1e20 mM. The kinetics of enzyme inhibitions were calculated using Lineweaver-Burk and EadieeHofstee equations (Wilson and Walker, 2010). Studies of enzyme precipitation by very high tannin concentrations were conducted as earlier (Adamczyk et al., 2014). Standard curves were prepared separately for TA and CT, and for each enzyme. Measurements of residual activity in enzyme-tannin complexes were conducted on enzyme-tannin precipitates. Briefly, buffer (pH range from 3.5 to 7) was added to precipitates and substrate. After incubation at room temperature, (time of incubation as in earlier measurements of enzymatic activity), samples were centrifuged and 0.1 ml of supernatant was moved to the next tube already containing 0.3 ml 0.5 M NaOH and the absorbance was measured at 405 nm. We compared these residual activities to activities of the same amount of enzymes under the same conditions but without precipitation step. Studies were made in 5 laboratory replicates for each enzyme and each pH value. 2.3.1. Fourier transform infrared spectroscopy (FTIR) The infrared spectra of enzymes were recorded with a FTIR spectrometer (Shimadzu IRPrestige-21; Shimadzu Corporation, Japan) with IRsolution 1.40 software (Shimadzu, Kyoto, Japan). We scanned: i) enzymes without tannins, ii) tannin and enzyme ratios in which activity was inhibited and ratios in which activity was increased and iii) tannins without enzymes. Liquid samples were vacuum freeze dried to dry powder and 3 mg of sample were mixed with 300 mg of KBr and pressed into discs. All tests were done in triplicate. FTIR analysis of enzyme conformation was based on Shen et al. (2014). The IR spectra of 1700e1600 cm1 (Amide I), commonly used to study changes in protein conformation, were treated with second derivative to increase resolution (Byler and Susi, 1986; Baker et al., 2014). Band assignments were done on the basis of literature (Byler and Susi, 1986; Choi and Ma, 2005; Jackson and Mantsch, 1995; Kong and Yu, 2007; Lu et al., 2015). 2.4. Soil microcosm experiment Samples were taken from the soil organic layer of two study sites, one located in Eno (middle-eastern Finland), called CT-poor soil and the other located in Salla (northern Finland), called CTrich soil. Both sites have three replicate plots. The characteristics of the soils can be found in Table 1. For more information see Adamczyk et al. (2015), Luiro et al. (2010), Smolander et al. (2010).

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Table 1 Characteristics of the site; presented values are means, for CT mean (SD).

Latitude Longitude Altitude, m a.s.l. Annual mean precipitation, mm Tree species Soil texture Humus type Soil type Organic matter content, % Site typea pH Total C, g kg1 Total N, g kg1 C/N Condensed tannins, mg g1 SOM water fraction ethanol fraction

CT-rich site

CT-poor site

67
180 N 29
140 E 260 551 Norway spruce fine sand mor podzol 79.6 EMT 3.9 501 13.3 37

62
470 N 30
90 E 130 646 Norway spruce silt mull podzol 14.0 OMaT 6.1 540 31.8 17

4.0 (0.3) 2.8 (0.2)

0 0.2 (0.1)

a Site types according to Cajander et al. (1949): EMT e Empertum nigrumVaccinium myrtillus, OMaT - Oxalis acetosella-Maianthemum bifolium. Information about sites were collected from Adamczyk et al. (2015), Smolander et al. (2010), Luiro et al. (2010).

2.5. Statistics To determine significant differences between the activities of the enzymes after incubation with CT from the control, a one way ANOVA was used, followed by Dunnett's test. The difference was reported as significant when P < 0.05. The assumption of normality was assessed using Kolmogorov-Smirnov and Shapiro-Wilk tests and homogeneity of variances using Levene's test. All statistical analyses were made using SPSS software (22.0 version; IBM Corporation, New York, U.S.A.). 3. Results 3.1. Condensed tannins characterization Condensed tannins contained 94.3% procyanidins and 5.7% prodelphinidins, with the mean degree of polymerization of 7 and molecular mass of about 2 kDa (for more results see Fig. S1). 3.2. Biochemical studies of enzyme-tannin interaction

Representative samples (20 cores, diameter 5.8 cm) were taken from the soil organic layer (Ofh), in case of CT-poor site such layer could not be distinguished; therefore the uppermost 4 cm was sampled. The samples passed through a 4.0 mm mesh. Samples from different plots were not bulked. The soil samples were then frozen until the laboratory experiments began and preincubated in þ4
C for 3 days prior to use. The dry weight (þ105
C, 16 h) was determined, and the organic matter content was measured as loss on ignition (þ550
C, 4 h). The soil pH was measured in a soil water suspension of 15 ml of soil in 25 ml of ultrapure water. Parts of the soil samples were dried at þ40
C, ground (0.5 mm mesh) and stored at 20
C until analyzed for the amount of CT with common acid-butanol assay (Adamczyk et al., 2016; Kanerva et al., 2008). We used CT extracted from Norway spruce needles as a standard.

Enzymes increased activity in response to low tannin concentrations whereas the enzyme activity diminished with higher tannin concentrations (Fig. 1). Fig. 1 shows the most important range of tannin concentration for acid phosphatase and arylsulfatase (full range at Fig. S2) and full range of tannin concentration for beta-glucosidase and chitinase. Larger increases or decreases of enzyme activity were observed for arylsulfatase, acid phosphatase and chitinase than for beta-glucosidase. Tannins in very high concentrations (0.4% for CT, and 2% for TA) substantially precipitated acid phosphatase, arylsulfatase and chitinase (more than 40%); but only 10% of beta-glucosidase (Fig. 3; see Fig. S3 for amount of precipitated enzymes, CT and TA). Inhibition kinetic studies proved that at high concentrations tannins acted as noncompetitive inhibitors on all studied enzymes (see Fig. 2 for acid phosphatase; for other enzymes see Fig. S4). As

2.4.1. Soil incubation and enzyme activity measurements The effects of CT on the enzyme activity were studied in a soil incubation experiment at constant moisture (60% WHC) and temperature (þ17
C). The soil samples (5 g of fresh soil for CT-rich and 11.7 g for CT-poor corresponding to 20 ml in volume) were placed in 125 ml glass bottles. Amount of soil organic matter per bottle was 1.26 g and 1.46 g for CT-poor and CT-rich soil, respectively. Condensed tannins were mixed with silica gel in proportion of 2:1 (w/w) and added to the soil. Silica gel is an inert carrier (Schimel et al., 1996) which aids handling and application of tannins to soils (Kanerva et al., 2006). We used CT in 3 doses, 10 mg, 50 mg and 100 mg per soil bottle. Only silica gel was added to the control bottles. The glass bottles were covered with gas-tight septa and aerated 3 times per week. All analyses were made in triplicate bottles for each treatment and each plot. Incubation was stopped after 0, 6, and 20 days and enzymatic activity was measured. Zero days of incubation means that after addition of tannins bottles were stored þ4
C overnight and activities were measured on the next day. Activities of beta-glucosidase, chitinase, acid phosphatase and arylsulfatase were measured as described previously (Adamczyk et al., 2015). We used the 40 mM buffers having the same pH value as the studied soils. For CT-rich soil acetate buffer was used, for CT-poor soil phosphate buffer. Results are presented as percent of activity of control. For values of enzyme activity of controls see Table S2.

Fig. 1. Changes in the enzyme activity after addition of tannins. Error bars indicate SD.

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Fig. 2. Enzyme inhibition kinetics on example of acid phosphatase: A) LineweaverBurk plot, B) Eadie- Hofstee plot. Other enzymes showed the same inhibition type and the results can be found in Supplementary Materials (Fig. S3).

seen from Lineweaver-Burk plots Km is not changed (X-intercept), but Vmax is decreased (Y-intercept representing 1/Vmax), which point to noncompetitive inhibition (Fig. 2A). Also Eadie-Hofstee plots point to non-competitive inhibition as lines on plots are parallel (Fig. 2B). No enzyme precipitation was observed at the tannin concentrations tested in this experiment (up to 0.2%). Enzyme-tannin complexes exerted up to 20% of the activity of the control for chitinase, beta-glucosidase and acid phosphatase, but only 10% for arylsulfatase (Fig. 4, for precise values see Table S1). Activity of complexes was detectable throughout the entire studied pH range, but it was especially high close to pH optima of the enzymes. The shapes of activity curves of enzyme-tannin complexes reflected those of control. Tannins affected the secondary structure of the enzymes, i.e. patterns in H-bonding and geometric orientations of amide bonds in alfa-helixes, beta-sheets beta-turns and random coil structures (Yang et al., 2015). Infrared spectra showed that in high concentrations tannins increased unstructured parts of enzymes (peak of 1646 cm1 assigned as random coil) at the expense of coiled structures (peak of 1655 cm1 assigned as a-helix; Fig. 5; more precise results and band assignment can be found at Fig. S5-S7). As studies of secondary structure with FTIR require relatively high enzyme concentrations, chitinase was not analyzed due to low solubility.

Fig. 4. Residual activity of CT-enzyme precipitates at different pH (in red) in comparison with control (in blue), PNP e paranitrophenyl. Error bars indicate SD. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

amounts of condensed tannins to CT-rich and CT-poor soil inhibited enzymatic activity in a concentration dependent manner (Fig. 6). The highest inhibition was observed in case of arylsulfatase in CTrich soil. The addition of the lowest dose of condensed tannins to the CTrich soil enhanced the enzyme activities, but not significantly for chitinase (Fig. 6A). Both acceleration and inhibition of enzyme

3.3. Mesocosm soil experiment Soil mesocosm studies were based on a laboratory incubation experiment in which we added different amounts of tannins to two soils differing in native amounts of condensed tannins (called later CT-rich and CT-poor soil). The addition of medium and high

Fig. 3. Amount of precipitated enzymes by very high tannin concentration (0.4% for CT, light-green columns, 2% TA, dark-green columns). Error bars indicate SD. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

Fig. 5. Infrared spectra of enzyme secondary structure presented as stacked plot of second derivative. Black lines represent enzyme without tannins and blue lines enzymes with low tannin concentrations. Red lines show enzymes with high tannin concentration. Spectras are smoothed by 8 points. Region of alfa-helix marked in yellow. We used following concentrations of tannins: 0.2% and 0.001% TA and 0.1% and 0.0025% CT (for acid phosphatase and arylsulfatase) and 0.2% and 0.01% TA and 0.1% and 0.01% CT (for beta-glucosidase), as these concentrations gave significant changes in enzyme activities (see Fig. 1). Spectras of tannins without enzymes did not provide a peak within Amide I region. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

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Fig. 6. Effect of tannin addition to soil: (A) to CT-rich soil, (B), to CT-poor soil. The results are presented as a percentage of residual activity. Mean and SD for 3 replicates. Statistically significant differences (P < 0.05) to the control are marked with an * (decrease in activity) or # (increase in activity). Condensed tannin additions were 10, 50 and 100 mg per bottle. One bottle of CT-poor soil contained 1.26 g SOM and CT-rich soil contained 1.46 g SOM. First incubation time (0 days) means that after addition of tannins bottles were stored þ4
C overnight and activities were measured on the next day.

activity by CT were weakened with time of incubation. Overall, the activities were especially affected at first measurement (0 days) and/or at second measurement (6 days), but after 20 days enzyme activities recovered to the level of control. This recovery trend was especially visible for additions of 100 mg CT (Fig. 6, darkest columns). 4. Discussion 4.1. Tannin-enzyme interaction in studies without soil It is well established that tannins can decrease enzymatic activity (Hagerman, 2012), although sometimes the inhibition effect is negligible (Juntheikki and Julkunen-Tiitto, 2000; Strumeyer and Malin, 1970). Our study provides first time evidence for an acceleration of enzyme activity through direct reaction with tannins. However, two earlier studies suggested similar result. In first of them some phenolic compounds enhanced the activity of pepsin (Tagliazucchi et al., 2005), but it was explained by phenolicinduced changes in substrate protein (Mole and Waterman, 1985; Oh and Hoff, 1986; Tantoush et al., 2012) and not directly by tannin reactions with enzyme. The second study, conducted under highly heterogenic conditions with tannin-rich leaves in the rumen, showed increased capacity of glutamate ammonia ligase, but no mechanical evidence was provided (Makkar et al., 1988). In our study, tannins affected enzyme activities to a different extent. We observed the weakest increases and decreases for the activity of beta-glucosidase, which could emerge from low affinity of this enzyme to tannins. This was confirmed by enzyme precipitation studies since not more than 10% of beta-glucosidase was precipitated by very high tannin concentrations. Different affinities of proteins, including enzymes, to tannins were observed previously (Adamczyk et al., 2011a, b; Hagerman, 2012; Juntheikki and

Julkunen-Tiitto, 2000). One can assume that precipitated enzymes are completely inactive. However, in our study, enzyme-tannin complexes exerted residual activity up to 20% of the control. To understand the mechanism underlying the shift in enzyme activity triggered by tannins, we studied inhibition kinetics. Our results showed that at high concentrations, tannins act as noncompetitive inhibitors, which is in accordance with earlier studies (Tamir and Alumot, 1969; Uchida et al., 1987; Wu et al., 2013). Noncompetitive inhibitors bind to an enzyme at sites other than the active site, possibly altering the conformation of the enzyme (Berg et al., 2010; Wu et al., 2013). On the other hand, competitive inhibitors bind to active center of an enzyme, blocking activity (Berg et al., 2010), so enhancement of activity by competitive inhibitor would be impossible. As inhibition kinetic studies suggested that tannins affect enzyme activity through changes in conformation, we used Fourier transform infrared spectroscopy (FTIR), a well-established tool in determining secondary structure of proteins including enzymes lu et al., 2015; Byler and Susi, 1986; Sytina et al., 2008; (Baltacıog Vonhoff et al., 2010). Infrared spectra showed that tannins in high concentration increased unstructured parts of enzymes at the expense of coiled structures. Conversely, tannins in low concentrations caused the adverse effect, decreasing unstructured parts, but increasing coiled structures. These changes in enzyme conformation explain both increase and decrease in enzymatic activity. 4.2. Tannin-enzyme interaction in soil conditions and its possible ecological significance To extrapolate our results to soil conditions we continued with mesocosm experiments. Effects observed in biochemical studies, if also present in soil, may potentially accelerate decomposition (if tannins would increase soil enzymatic activity) or decelerate

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decomposition (if inhibitory effect would be prevalent). The addition of CT in medium and high amounts to CT-rich and CT-poor soil, inhibited enzymatic activity, but the addition of the lowest dose of CT to the CT-rich soil enhanced the enzyme activities. Joanisse et al. (2007) also observed that addition of CT to soil in high amounts suppressed beta-glucosidase and acid phosphatase, but low amounts of CT increased enzymatic activity. However, the authors explained this result as the binding of CT to enzyme inhibitors other than tannins, which diminished the overall amount of inhibitors thus increasing soil enzyme activity. The results of our biochemical assays provided another explanation: low concentration of tannins modified enzyme conformation, boosting the activity. As soil is highly heterogenic environment, other reasons for changes in enzyme activity cannot be excluded, especially after longer incubation period. Increase in soil enzyme activity could emerge also from adaptation of soil microbial populations which secreted more de novo synthesized enzymes over time. Moreover, for some microbes CT may cause toxic effect (observed by e.g. Kanerva et al., 2008) thus these dead cells provided easy-available nutrients causing a positive feedback to microbes well-adjusted to CT. In addition, inactivation of added CT by soil constituents over time could also take place. Tannins could be also partially digested by soil microorganisms, which could result in decrease of inhibitory effect, but such effect cannot be substantial as CT are recalcitrant compounds (Kraus et al., 2003; Smolander et al., 2012, 2010). After 20 days of incubation, enzyme activity from treatments with CT were equal to controls, indicating the recovery of soil enzymes in response to CT-input. Regardless of the mechanism, an ability to recover the enzyme machinery after high CT levels seems to be necessary to sustain the productivity of tannin-rich boreal forest soil. However, soil is exposed to constant deposits of tannins provided from above- and belowground litter (Xia et al., 2015). Thus, an increase or decrease in enzyme activity due to reactions with tannins may be observed continuously during the vegetation period. Our soil study was performed under controlled laboratory conditions and we used CT amounts corresponding to natural soil concentrations. The lowest addition (10 mg) was equal to the native amount of CT in CT-rich soil. High CT flux to soil from litter after heavy rain may outweigh even the highest CT amounts. We can therefore assume that the effects of CT observed in our study can potentially also occur under field conditions, especially in CT-rich boreal forest ecosystems. However, as under field conditions soil enzymatic activity is subjected to numerous factors, including a mixture of enzyme inhibitors and enhancers, their combined effect may be weakened or strengthened. In addition, abiotic factors, especially moisture and temperature modify the influence of tannins on decomposition processes. Moreover, site-specific effects may play significant role. In our mesocosm experiment, the addition of CT increased enzyme activities in CT-rich soil, in which the amount of native CT was higher than those found in CT-poor soil. Differences between the level of soil enzyme inhibition by tannins in different soils may emerge from “isoenzyme acclimatization” to high native amounts of CT (Triebwasser et al., 2012). Our first experiments without soil proved that enzyme-tannin complexes may retain up to 20% of residual activity. This phenomenon, if present in soil conditions, may additionally explain complicated changes in enzymatic activity due to interaction with tannins. Ecologically, precipitation of enzymes by CT can have positive effect on soil productivity in a long-term, as enzymes may be stabilized in the soil after reaction with tannins (Burns et al., 2013). Furthermore, these bound enzymes represent a significant reservoir of activity and function as the first catalytic response to changing the fluxes of substrates and are a potential source of substrate turnover during periods when microbial biomass is low

65

(Burns et al., 2013). 5. Conclusions The ability of tannins to modify enzyme activity furthers our current understanding on how tannins may potentially regulate SOM decomposition processes and nutrient fluxes in boreal forest soil. The present study showed that tannin-enzyme interaction led to decreases but also increases of enzymatic activity due to changes in enzyme conformation and that tannin-enzyme precipitates can exert residual enzyme activity. Moreover, the enzyme susceptibility to tannins was a function of enzyme type and in soil, site-specific effects modified enzyme response to tannins. As hypothesized, decrease of enzymatic activity was prevalent in soil studies. Ecologically, decrease of decomposition through inhibition of enzyme activity and building-up of SOM through retention of organic N could be a mechanism for buffering soil CO2 emissions in a changing climate, however, this issue needs more study with special attention to field experiments. Overall, our results suggest that tannin-enzyme interactions can potentially play important role in SOM decomposition of the tannin-rich ecosystems. Acknowledgements We thank Anneli Rautiainen and Irmeli Luovula for her help with laboratory work. This work was funded by The Academy of Finland (grant no. 126667) and Emil Aaltonen Foundation (grants no. 140006 and 130007). Appendix A. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.soilbio.2016.12.027. References rez-Alegría, L., 2007. Acosta-Martínez, V., Cruz, L., Sotomayor-Ramírez, D., Pe Enzyme activities as affected by soil properties and land use in a tropical watershed. Applied Soil Ecology 35, 35e45. Adamczyk, B., Adamczyk, S., Kukkola, M., Tamminen, P., Smolander, A., 2015. Logging residue harvest may decrease enzymatic activity of boreal forest soils. Soil Biology and Biochemistry 82, 74e80. Adamczyk, B., Adamczyk, S., Smolander, A., Kitunen, V., 2011a. Tannic acid and Norway spruce condensed tannins can precipitate various organic nitrogen compounds. Soil Biology and Biochemistry 43, 628e637. €, O.-M., Kanerva, S., Kieloaho, A.-J., Adamczyk, B., Ahvenainen, A., Sietio €, P., Laakso, T., Strakov Smolander, A., Kitunen, V., Saranp€ aa a, P., Heinonsalo, J., 2016. The contribution of ericoid plants to soil nitrogen chemistry and organic matter decomposition in boreal forest soil. Soil Biology and Biochemistry 103, 394e404. €, O., Kitunen, V., Smolander, A., 2014. Can we measure Adamczyk, B., Kiikkila condensed tannins from tannin-protein complexes? - A case study with acidbutanol assay in boreal forest soil organic layer. European Journal of Soil Biology 64, 40e45. Adamczyk, B., Kitunen, V., Smolander, A., 2009. Polyphenol oxidase, tannase and proteolytic activity in relation to tannin concentration in the soil organic horizon under silver birch and Norway spruce. Soil Biology and Biochemistry 41, 2085e2093. Adamczyk, B., Salminen, J.-P., Smolander, A., Kitunen, V., 2012. Precipitation of proteins by tannins: effects of concentration, protein/tannin ratio and pH. International Journal of Food Science and Technology 47, 875e878. Adamczyk, S., Adamczyk, B., Kitunen, V., Smolander, A., 2011b. Influence of diterpenes (colophony and abietic acid) and a triterpene (beta-sitosterol) on net N mineralization, net nitrification, soil respiration, and microbial biomass in birch soil. Biology and Fertility of Soils 47, 715e720. Baker, M.J., Trevisan, J., Bassan, P., Bhargava, R., Butler, H.J., Dorling, K.M., Fielden, P.R., Fogarty, S.W., Fullwood, N.J., Heys, K.A., Hughes, C., Lasch, P., -Suso, J., Strong, R.J., Martin-Hirsch, P.L., Obinaju, B., Sockalingum, G.D., Sule Walsh, M.J., Wood, B.R., Gardner, P., Martin, F.L., 2014. Using Fourier transform IR spectroscopy to analyze biological materials. Nature Protocols 9, 1771e1791. lu, H., Bayındırlı, A., Severcan, M., Severcan, F., 2015. Effect of thermal Baltacıog treatment on secondary structure and conformational change of mushroom polyphenol oxidase (PPO) as food quality related enzyme: a FTIR study. Food Chemistry 187, 263e269.

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