Decoupling of priming and microbial N mining during a short-term soil incubation

Accepted Manuscript Decoupling of priming and microbial N mining during a short-term soil incubation Birgit Wild, Jian Li, Johanna Pihlblad, Per Bengtson, Tobias Rütting PII:

S0038-0717(18)30397-3

DOI:

https://doi.org/10.1016/j.soilbio.2018.11.014

Reference:

SBB 7339

To appear in:

Soil Biology and Biochemistry

Received Date: 20 August 2018 Revised Date:

15 November 2018

Accepted Date: 15 November 2018

Please cite this article as: Wild, B., Li, J., Pihlblad, J., Bengtson, P., Rütting, T., Decoupling of priming and microbial N mining during a short-term soil incubation, Soil Biology and Biochemistry (2018), doi: https://doi.org/10.1016/j.soilbio.2018.11.014. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

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Decoupling of priming and microbial N mining during a short-term soil incubation

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Birgit Wilda,b,c*, Jian Lid, Johanna Pihlblada,e, Per Bengtsond, Tobias Rüttinga

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a

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b

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Sweden

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Bolin Centre for Climate Research, Stockholm University, Stockholm, Sweden

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Department of Biology, Lund University, Lund, Sweden

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Hawkesbury Institute for the Environment, Western Sydney University, Sydney, Australia

Department of Earth Sciences, University of Gothenburg, Gothenburg, Sweden

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Department of Environmental Science and Analytical Chemistry, Stockholm University, Stockholm,

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Birgit Wild, Tel: +468 674 7250, email: [email protected]

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Abstract

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Soil carbon (C) and nitrogen (N) availability depend on the breakdown of soil polymers such as lignin,

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chitin, and protein that represent the major fraction of soil C and N but are too large for immediate uptake

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by plants and microorganisms. Microorganisms may adjust the production of enzymes targeting different

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polymers to optimize the balance between C and N availability and demand, and for instance increase the

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depolymerization of N-rich compounds when C availability is high and N availability low (“microbial N

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mining”). Such a mechanism could mitigate plant N limitation but also lie behind a stimulation of soil

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respiration frequently observed in the vicinity of plant roots (“priming effect”). We here compared the

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effect of increased C and N availability on the depolymerization of native bulk soil organic matter (SOM),

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and of

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microcosm incubation experiments. A significant reduction of chitin depolymerization (described by the

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recovery of chitin-derived C in the sum of dissolved organic, microbial and respired C) upon N addition

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indicated that chitin was degraded to serve as a microbial N source under low-N conditions and replaced

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in the presence of an immediately available alternative. Protein and lignin depolymerization in contrast

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were not affected by N addition. Carbon addition enhanced microbial N demand and SOM decomposition

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rates, but significantly reduced lignin, chitin, and protein depolymerization. Our findings contrast the

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hypothesis of increased microbial N mining as a key driver behind the priming effect and rather suggest

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that C addition promoted the mobilization of other soil C pools that replaced lignin, chitin, and protein as

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microbial C sources, for instance by releasing soil compounds from mineral bonds. We conclude that

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SOM decomposition is interactively controlled by multiple mechanisms including the balance between C

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vs N availability. Disentangling these controls will be crucial for understanding C and N cycling on an

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ecosystem scale.

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C-enriched lignin, chitin, and protein added to the same soil in two complementary ten day

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Keywords: decomposition, lignin, chitin, protein, phospholipid fatty acids, extracellular enzymes

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1. Introduction

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Polymers such as lignin, chitin, and protein constitute the bulk of soil C and N (Knicker, 2011; Paul,

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2016) but are too large for immediate uptake by plants and microorganisms. Their depolymerization by

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microbial extracellular enzymes consequently controls soil C and N availability (Schimel and Bennett,

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2004; Schimel and Weintraub, 2003). Since enzyme synthesis requires an investment of C and N,

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microorganisms may adjust the production of enzymes targeting different compounds to balance C and N

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availability and demand. Activities of enzymes targeting N-rich compounds have thus been found to

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decrease after N addition (Allison and Vitousek, 2005; Geisseler and Horwath, 2009; Koranda et al., 2013;

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Wild et al., 2017), and to increase after C addition (Geisseler and Horwath, 2009; Meier et al., 2017).

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These adjustments can occur rapidly, often within a few days (Allison and Vitousek, 2005; Geisseler and

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Horwath, 2009; Koranda et al., 2013; Wild et al., 2017).

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An increased breakdown of N-rich compounds under low N and high C availability (“microbial N

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mining”) has been suggested as a key mechanism behind the rhizosphere priming effect, i.e., a stimulation

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of SOM mineralization frequently observed in the vicinity of plant roots (Craine et al., 2007; Dijkstra et

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al., 2013; Kuzyakov, 2010). Plant roots take up available N from the soil and release a range of organic

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compounds that contain little or no N (e.g., sugars, organic acids; Jones et al., 2009), potentially

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stimulating the microbial synthesis of N-targeting enzymes, and thereby the breakdown of N-rich

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polymers such as proteins or chitin. Supporting such a mechanism, previous studies have found higher

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activities of N-targeting enzymes in rhizosphere than non-rhizosphere soil (Brzostek et al., 2013; Koranda

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et al., 2011; Weintraub et al., 2007). Overall, microbial N mining has the potential to mitigate plant N

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limitation and thereby promote plant C fixation, but also soil C losses.

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Although ecosystem C and N cycling depend on breakdown rates of N-containing polymers, our

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understanding of these processes and their controls is still limited. Proteins account for the largest fraction

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of soil N (Knicker, 2011); nevertheless, gross protein depolymerization rates have shown little adjustment

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to increased soil C and N availability in a previous incubation study (Wild et al., 2017). Changes in soil N

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cycling induced by increased C availability might be rather related to the breakdown of other soil

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polymers, such as chitin, heterocyclic compounds, or lignin. Although lignin itself does not contain N, a

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negative relationship between lignin degradation and N availability has been frequently observed

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(Knicker, 2011) and possibly results from either an inhibiting effect of N on ligninolytic microorganisms

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or enzymes (Carreiro et al., 2000) or a binding of N to lignin phenols (Schmidt-Rohr et al., 2004). Lignin

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depolymerization rates might thereby affect soil N availability. While gross protein depolymerization rates

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can be quantified using an established

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method for quantifying chitin depolymerization rates has only recently become available (Hu et al., 2017),

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and suitable methods for quantifying depolymerization rates of other soil compounds are still lacking.

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Most studies on the adjustment of soil polymer breakdown to environmental conditions are consequently

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limited to consider only proteins, or use indirect measures such as enzyme activities or soil C and N

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mineralization rates.

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We here investigated the regulation of lignin, chitin, and protein depolymerization by C and N availability

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in a forest soil. Specifically, we hypothesized that (1) C addition would promote microbial N demand and

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N-targeting enzyme activities, and thereby accelerate lignin, chitin and protein depolymerization, as well

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as SOM depolymerization in general, and that (2) N addition would have the opposite effect, decrease N-

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targeting enzyme activities, lignin, chitin, protein and SOM depolymerization. These hypotheses were

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tested using a combination of two simultaneous ten day laboratory incubation experiments. Our findings

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therefore reflect short-term responses to increases in C and N availability. In Experiment 1, forest soil was

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amended with easily available sources of C (glucose) and N (ammonium), and glucose was enriched in

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experiment. In Experiment 2, the same forest soil was mixed with lignin, chitin, or protein before addition

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of easily available C or N as above. In this second experiment, glucose was of natural 13C content, but

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lignin, chitin, and protein were enriched. This setup permitted us to trace lignin-, chitin- and protein-

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derived (i.e., 13C-enriched) C into dissolved organic C (DOC), microbial C, and respiration, and thereby to

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study changes in lignin, chitin, and protein mobilization induced by increased soil C or N availability.

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Although absolute C fluxes from lignin, chitin and protein added to the soil do not reflect absolute C

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N pool dilution assay (Wanek et al., 2010), a corresponding

C which permitted us to distinguish fluxes of SOM-derived and glucose-derived C in a classic priming

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fluxes from their native counterparts, their relative responses to C or N addition (Experiment 2) help in

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elucidating mechanisms behind changes in bulk SOM decomposition (Experiment 1).

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In order to resolve the effect of C and N addition on the utilization of polymer-derived C by different

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microbial groups, we further analyzed the incorporation of SOM-, lignin- chitin- and protein-derived C

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into phospholipid fatty acids (PLFAs) that serve as biomarkers of fungi and bacteria. Previous studies

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suggest that lignin degradation is strongly dominated by fungi (Treseder and Lennon, 2015), with less

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contribution by some bacteria (Bugg et al., 2011). Chitin as well as protein degradation in contrast can be

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performed by both fungi and bacteria (Baldrian et al., 2011; Beier and Bertilsson, 2013; Treseder and

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Lennon, 2015; Vranova et al., 2013). Although strong degradation activity of a microorganism does not

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necessarily imply strong incorporation of breakdown products, we expected to find a preferential

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incorporation of lignin-derived C into fungal markers, and a more even incorporation of chitin- and

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protein-derived C into both fungal and bacterial markers.

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2. Material and Methods

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2.1. Experimental setup

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Soils for the experiment were taken from the Svartberget experimental forest (64° 15’ N, 19° 47’ E) in

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northern Sweden. The site is characterized by a mean annual temperature of 1.8°C and a mean annual

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precipitation of 614 mm (1981-2010; Laudon et al., 2013), dominated by Pinus sylvestris and Picea abies

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with an understory of Vaccinium myrtillus and Vaccinium vitis-idaea, and underlain by a Podzol soil. Soil

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was sampled from the Podzol B horizon in May 2016, sieved to < 2 mm, homogenized by mixing in a

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plastic bag, and stored at 18°C for a few days. The soil was then again homogenized, and aliquots of 40 g

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fresh weight were weighed into plastic bags and pre-incubated at 18°C for 24 hours. Before the

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experiment, soils contained 4.56 ± 0.32% C and 0.13 ± 0.01% N (GSL Elemental Analyzer coupled to 20-

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22 Stable Isotope Mass Spectrometer, Sercon Ltd.), gravimetric C/N ratios averaged 36.3 ± 0.4, and

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gravimetric water contents 31.7 ± 0.2% fresh soil weight (means ± standard errors of five analytical

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replicates). Note that inorganic C content is negligible at the study site so that organic C is close to total C

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content.

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At the start of the experiment, the pre-weighed soil samples were randomly assigned to one of 16

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treatment combinations in five replicates. For Experiment 1 (four treatments), pre-weighed soil samples

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were filled into 250 ml glass flasks, and then received either a solution of glucose as an easily available C

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source (+C), a solution of ammonium sulfate as an easily available N source (+N), a solution of both

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glucose and ammonium sulfate (+CN) or Milli-Q water as control (+0). In this experiment, the added

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glucose was enriched in

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Cambridge Isotope Laboratories with conventional, natural abundance glucose). Substrates were added at

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concentrations of 5 mg glucose and/or 1.875 mg ammonium sulfate in 250 µl Milli-Q water at different

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locations of the sample surface, and homogenized with the soil by thorough shaking. For the +CN

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treatment, the added substrates had a gravimetric C/N ratio of 5, which is at the lower end of the range

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observed in microbial biomass (Xu et al., 2013).

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C at 5 atom% (mixture of 99 atom% universally 13C-enriched glucose from

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For Experiment 2 (twelve treatments), pre-weighed samples were first mixed with

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chitin, or protein. Lignin (Zea mays isolate, universally enriched in

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(Aspergillus niger isolate, universally enriched in

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protein (algal crude protein extract, universally enriched in

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Amounts of lignin, chitin, and protein equivalent to 4 mg C per sample were suspended in 250 µl Milli-Q

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water, added to the pre-weighed samples in the plastic bags, and homogenized with the soil before

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samples were filled into 250 ml glass flasks. Samples mixed with lignin, chitin, or protein then received

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the same treatments of C or N addition as in Experiment 1, but in this case, glucose was of natural 13C

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abundance. For both Experiments 1 and 2, flasks were closed and incubated at 18°C for ten days. For

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comparison, summer temperature at the sampling site is ca. 15°C at 10 cm depth. Addition of easily

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available C, N, CN, or water was repeated daily by adding solutions to the soil surface and shaking as

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described above. This disturbance of the soil might have induced some bias by stimulating microbial

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activity. Nevertheless, since all samples were exposed to the same disturbance, this does not limit our

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ability to test our hypotheses based on relative differences between treatments. Additions of easily

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available C and N summed up to a total of 22 mg C and 4.375 mg N after the last addition, respectively,

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which corresponds to moderate increases of the initial SOC pool by 1.8% and of the soil N pool by 12.7%.

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After the last respiration measurement, samples were destructively harvested for analysis of soil C and N

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pools, microbial community function based on potential activities of extracellular enzymes, and microbial

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community composition and substrate utilization based on phospholipid fatty acids. Samples were

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processed within a few days after harvest and kept at incubation temperature until then.

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C at 97 atom%) and chitin

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C at 98 atom%) were purchased from Isolife, and 13

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C at 98 atom%) from Sigma Aldrich.

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C-enriched lignin,

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2.2. Respiration rates

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Respiration rate and isotopic composition of respired CO2 were measured immediately after the start of

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the experiment, and after 1, 2, 3, 4, 6, 8, and 10 days of incubation, after the daily addition of easily

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available C and N. Flasks were closed with caps equipped with septa, evacuated four times and refilled

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with pressurized air of quantified, atmospheric concentration and isotopic composition of CO2. Flasks 7

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were then incubated for 7-8 hours (day 0) or 4 hours (all other days) at 18°C before analysis. Empty flasks

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were processed with the samples as blanks. Concentration and isotopic composition of accumulated CO2

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were analyzed with an Isotope Ratio Infrared Spectrometer (Delta Ray, Thermo Scientific) and calibrated

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against two CO2 standards of different isotopic composition. Concentration and isotopic composition of

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CO2 were corrected for the initially present CO2 using the values derived from the blank bottles. We then

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distinguished CO2 originating from different organic carbon sources using Equations 1 and 2.

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(Equ. 1) (Equ. 2)

The abbreviations CTotal, C1, and C2 represent concentrations of total CO2, CO2 from source 1, and CO2

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from source 2, respectively, and atom%Total, atom%1 and atom%2 the corresponding

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atom% 13C) of respired CO2. In Experiment 1, source 1 was the added 13C enriched glucose (+C and +CN

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treatment), and source 2 the native, natural abundance soil organic C. In Experiment 2, source 1 was the

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added 13C enriched polymer (lignin, chitin, or protein), and source 2 included both native soil organic C

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and added glucose (+C and +CN treatments) that were of natural

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distinguished here. The development of cumulative respiration from different sources is shown in

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Supplementary Fig. 1, and total cumulative respiration in Supplementary Table 1.

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C contents (in

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C abundance and cannot be

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2.3. Carbon and nitrogen pools

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Concentration and 13C content of DOC were measured in 0.5 M K2SO4 extracts of fresh soil with an LC

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Isolink IRMS system (Thermo Scientific) at the Stable Isotope Service Lab, Department of Biology, Lund

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University, Sweden. The system was operated in flow injection analysis mode (flow rate 400 µl min-1),

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and samples were analyzed by direct injection of 10 µl extract. Within the LC Isolink interface, organic C

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contained in the extract was oxidized to CO2 at 99.9°C using Na2SO2O7 as oxidant and H3PO4 as acid

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reagent, both supplied at flow rates of 50 µl min-1. Concentration and isotopic composition were

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quantified against IAEA sucrose as well as glycine standards, and blank corrected.

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Concentration and

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(modified after Vance et al., 1987). Aliquots of fresh soil were fumigated for 48 hours in an atmosphere of

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ethanol-free chloroform, extracted with 0.5 M K2SO4 and analyzed with LC Isolink IRMS as described

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above. Microbial C was calculated as the difference between organic C in fumigated and non-fumigated

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extracts, and the 13C content of microbial C using Equation 3.

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C content of microbial C were measured using chloroform fumigation extraction

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(Equ. 3)

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The abbreviations CFum, CDOC, and CMic stand for the organic C concentrations of fumigated extracts, DOC

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and microbial C, respectively, and atom%Fum, atom%DOC, and atom%Mic for the corresponding

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contents. We did not apply a correction factor to account for incomplete extraction of microbial C

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considering the large variability of correction factors reported in the literature, and the likely location of C

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recently taken up that is the focus of this study in the easily extractable cytoplasm. Dissolved organic C

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and microbial C from different sources were distinguished as described above using Equations 1 and 2.

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Total DOC and total microbial C are presented in Supplementary Table 1.

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Concentrations of ammonium and nitrate were measured in 0.5 M K2SO4 extracts using the photometric

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assays described by Kandeler and Gerber (1988) and Miranda et al. (2001), respectively.

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2.4. Extracellular enzyme activities

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We measured potential activities of the six extracellular enzymes cellobiohydrolase (CBH; targeting

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cellulose), β-glucosidase (BG; targeting cellulose and cellobiose), N-acetyl-β-D-glucosaminidase (NAG;

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targeting chitin and peptidoglycan), leucine-aminopeptidase (LAP; targeting proteins and peptides),

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phenoloxidase (POX; non-specific) and peroxidase (PER; non-specific) using a combination of

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fluorometic (hydrolytic enzymes CBH, BG, NAG, LAP) and photometric (oxidative enzymes POX, PER)

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assays (Kaiser et al., 2010b; Marx et al., 2001; Saiya-Cork et al., 2002). Briefly, aliquots of 2 g fresh soil

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were suspended in 100 ml 100 mM sodium acetate buffer at pH 5.5. For analysis of the hydrolytic

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enzymes, 200 µl aliquots of the soil suspensions were amended with fluorescence labeled substrates

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(CBH: 4-methylumbelliferyl-β-D-cellobioside; BG: 4-methylumbelliferyl-β-D-glucopyranoside; NAG: 4-

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methylumbelliferyl-N-acetyl-β-D-glucosaminide;

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incubated for 140 min at room temperature. Fluorescence was measured at 365 nm excitation and 450 nm

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emission, and concentrations of released 4-methylumbelliferone (MUF) and 7-amido-4-methylcoumarine

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(AMC), respectively, were calibrated against external standard curves. To prepare standards, target

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compounds were mixed at different concentrations with the soil suspension to account for possible

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quenching effects. Potential activities of cellobiohydrolase, β-glucosidase, and N-acetyl-β-D-

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glucosaminidase were expressed in nmol MUF g-1 dry soil h-1, and activities of leucine-aminopeptidase in

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nmol AMC g-1 dry soil h-1.

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For analysis of the oxidative enzymes, 1 ml aliquots of the soil suspensions were mixed with 1 ml 20 mM

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L-3,4-dihydroxyphenylalanine (DOPA) as substrate, and samples for PER additionally with 10 µl 0.3%

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H2O2. Samples were shaken, centrifuged, and measured photometrically at 450 nm absorbance

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immediately, and after 20 hours of incubation at room temperature. Activities of POX were calculated

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from the difference in absorbance of the DOPA amended samples between the two time points, and

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activities of PER from the difference in absorbance of the DOPA+H2O2 amended samples between the

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two time points, subtracting the difference between DOPA amended samples. Potential activities of both

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enzymes were expressed in nmol DOPA g-1 dry soil h-1.

and

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L-leucine-7-amido-4-methylcoumarin)

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LAP:

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2.5. Phospholipid fatty acids (PLFAs)

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Phospholipid fatty acids were analyzed as described in Ehtesham and Bengtson (2017). Briefly, 2 g freeze

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dried soil was extracted using a 0.8:1:2 (v/v/v) solution of citrate buffer, chloroform and methanol

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(Frostegård et al., 1991), extracts were centrifuged (3000 × g, 10 min), and the supernatant was transferred 10

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to a new set of test tubes and split into two phases by adding equal amounts of CHCl3 and citrate buffer.

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The lipid fraction was loaded on silica based sorbent cartridges (Bond Elut LRC-SI, Agilent), and

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fractionated into neutral lipids, glycolipids and phospholipids by elution with chloroform, acetone and

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methanol, respectively. The phospholipid fraction was amended with a known amount of methyl

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nonadecanoate (19:0) as internal standard, and lipids were transmethylated to their fatty acid methyl esters

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(FAMEs) using mild alkaline methanolysis at 37ºC for 15 min in a temperature controlled water bath.

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Following the alkaline methanolysis, FAMEs were flash evaporated under N2 gas, re-dissolved and stored

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in 200 µl hexane at -20°C until analysis. Concentration and isotopic composition of individual FAMEs

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were determined at the Stable Isotope Service Lab, Department of Biology, Lund University, Sweden

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using a GC-C-IRMS system consisting of a Trace GC Ultra connected to a Delta V Plus isotope-ratio

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mass spectrometer via the GC Isolink II preparation device and ConFlow IV interface (Thermo Scientific).

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Samples were injected in splitless mode at 250°C and individual FAMEs were separated on a HP-5MS UI

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column (60 m × 0.25 mm, 0.25 µm stationary phase, Hewlett Packard) using Helium as a carrier gas (flow

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rate 1.5 ml min-1) and the temperature program described in Ehtesham and Bengtson (2017). Individual

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PLFAs were identified using a combination of previously established mass spectra, retention times

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relative to the internal standard 19:0, and an external bacterial acid methyl ester standard (BAME, Sigma

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Aldrich, 47080-0). The PLFAs 18:1ω9 and 18:2ω6 were considered as markers for fungi (Kaiser et al.,

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2010a), 16:1ω7, 18:1ω7 and cy19:0 as markers for gram negative bacteria, i15:0, a15:0, i16:0 and

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10Me18:0 as markers for gram positive bacteria (Zelles, 1997; Zogg et al., 1997), and 15:0, 16:0, 18:0,

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16:1ω5 as unspecific. The contribution of C from different sources to individual PLFAs was calculated

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using Equations 1 and 2. The total amount of C and the contribution of C from different sources in

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individual PLFAs are presented in Supplementary Tables 2 and 3, respectively.

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2.6. Statistical analyses

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Statistical analyses were performed in R 3.3.1 (R Development Core Team, 2016) with the additional

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packages vegan (Oksanen et al., 2016) and GenABEL (Aulchenko et al., 2007), on C and N pools, enzyme 11

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activities and PLFAs measured upon harvest after ten days of incubation, and on cumulative respiration

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over the full incubation period. To describe the total mobilization of SOM-, lignin-, chitin-, and protein-

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derived C, we additionally defined a “mobile C pool” that was calculated as the sum of DOC and

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microbial C at the end of the incubation, as well as cumulative respiration over the incubation period that

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was derived from SOM, lignin, chitin, and protein, respectively. We used one-way ANOVA followed by

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Tukey’s HSD test to assess differences in absolute and relative (percentage of total) concentrations of

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PLFAs associated with specific microbial groups, as well as in enzyme activities between samples with no

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additional polymer, lignin, chitin, and protein, separately for +0, +C, +N, and +CN treatments. We then

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applied two-way ANOVA to test for significant main effects of C and N addition on measured parameters

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as well as their interaction separately for each polymer treatment, after log- or rank-transformation where

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necessary to meet the conditions of homogenous variances and normality. Significant interactive effects of

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CxN indicate that the effect of one factor (C or N addition) depends on the level of the other. P-values

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below 0.05 were considered significant. We further used Spearman’s rank sum correlations to test for

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monotonic relationships between parameters, and Pielou’s Evenness Index to compare the distribution of

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lignin-, chitin-, and protein-derived C among individual PLFAs. Pielou’s Evenness Index was calculated

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by dividing the Shannon Diversity Index (diversity function in vegan) of the relative contribution of

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polymer-derived C to individual PLFAs by the natural logarithm of the number of individual PLFAs. High

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values of Pielou’s Evenness indicate a similar contribution of polymer-derived C to all individual PLFAs,

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and low values the preferential incorporation of polymer-derived C into specific PLFAs.

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3. Results

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3.1. Effect of carbon and nitrogen addition on nitrogen availability

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The addition of easily available C significantly reduced soil ammonium concentrations irrespective of

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polymer amendment (Table 1), reflecting enhanced microbial N demand that increased microbial

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ammonium uptake, decreased microbial ammonium release by N mineralization or both. Not surprisingly,

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the addition of easily available N in the form of ammonium sulfate strongly increased ammonium

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concentrations; nevertheless, a decrease in ammonium concentrations by C addition was also observed

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under these N-enriched conditions. Although C addition significantly increased nitrate concentrations in

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all polymer treatments, the overall effect on dissolved inorganic N was strongly negative as nitrate

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accounted for only a small fraction of the inorganic N pool.

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3.2. Effect of carbon and nitrogen addition on fluxes of SOM-, lignin-, chitin-, and protein-derived carbon

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To assess the effect of C and N addition on the mobilization of C from SOM in Experiment 1, and from

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lignin, chitin and protein in Experiment 2, we defined a mobile C pool that was calculated as the sum of

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DOC, microbial C and cumulative respiration from SOM, lignin, chitin, or protein, respectively. The

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mobile C pool thus serves as a proxy for C that is not bound in the form of large polymers but is

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immediately available for microbial uptake (DOC) or has already been taken up (microbial C, respiration).

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Comparing the recovery of SOM-, lignin-, chitin-, and protein-derived C in the mobile C pool reveals

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contrasting effects of C addition on C mobilization from native bulk SOM in Experiment 1, and from

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lignin, chitin, and protein mixed to the soil in Experiment 2 (Fig. 1). Although C addition consistently and

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significantly increased the recovery of SOM-, lignin-, and protein-derived C in DOC, and reduced the

295

recovery of SOM-, lignin-, chitin-, and protein-derived C in the microbial C pool, effects on respiration

296

and the overall mobile C pool differed between SOM-derived and lignin-, chitin-, as well as protein-

297

derived C. Carbon addition increased the recovery of SOM-derived C in respiration and thereby in the

298

total mobile C pool, by + 31% without additional N (+C vs +0 treatment), and by +28% with additional N

299

(+CN vs +N treatment). The patterns observed for native bulk SOM in Experiment 1 thus suggest both the

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translocation of SOM-derived microbial C to DOC and respiration, as well as the accelerated

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depolymerization of bulk SOM, i.e., a positive priming effect. In contrast, C addition decreased the

302

recovery of lignin-, chitin-, and protein-derived C in respiration and the total mobile C pool, indicating

303

reduced lignin, chitin, and protein depolymerization. The effect was strongest for chitin (-67% without

304

additional N; -53% with additional N), followed by lignin (-41% and -38%, respectively). In the case of

305

protein, C addition slightly increased the recovery in the mobile C pool when no N was added (+2%), but

306

decreased the recovery when additional N was added (-16%). Nevertheless, the overall effect was a

307

significant reduction by C addition (p = 0.03).

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Nitrogen addition had no significant effect on C mobilization from bulk SOM, lignin, and protein, but

309

significantly reduced chitin-derived C in DOC and respiration, and in the mobile C pool. Carbon and N

310

effects on chitin were not additive as indicated by a significant CxN interaction term, with a much

311

stronger reduction when N was added without C (-34%) than with C (-7%).

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3.3. Effect of carbon and nitrogen addition on extracellular enzyme activities

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In contrast to our hypothesis, C addition had no significant effect on activities of the N-acquiring enzymes

315

NAG and LAP (Table 2). The effect of N addition on NAG and LAP depended on polymer amendment,

316

resulting in an increase in NAG in the lignin and chitin amended samples, an increase in LAP in the

317

protein amended samples, and a decrease in LAP in the chitin amended samples. Activities of the C-

318

acquiring enzymes CBH and BG were unaffected by C addition except for CBH in protein amended

319

samples, but generally increased by N addition, in particular when C was added together with N. Effects

320

on the oxidative enzymes POX and PER depended on polymer amendment. Carbon addition reduced POX

321

in chitin amended samples and increased both POX and PER in protein amended samples, whereas N

322

addition reduced POX and PER in lignin amended samples.

323

We further tested for relationships between activities of lignin, chitin, and protein degrading enzymes and

324

the amount of lignin-, chitin-, and protein-derived C, respectively, recovered in the mobile C pool.

325

Correlations between the chitinolytic enzyme NAG and the amount of chitin-derived C, as well as

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between the proteolytic enzyme LAP and the amount of protein-derived C were not significant (p > 0.1;

327

Fig. 2). Correlations between the unspecific, oxidative enzymes POX and PER that have been linked to

328

lignin degradation (Sinsabaugh, 2010) and the amount of lignin-derived C in the mobile C pool were not

329

significant when the full dataset was used, but were significant when one value was removed that might be

330

considered an outlier (POX: p = 0.089, Spearman’s rho = 0.457; PER: p = 0.049, rho = 0.521).

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3.4. Effect of carbon and nitrogen addition on microbial community composition and substrate utilization

333

Different distributions of lignin-, chitin-, and protein-derived C among PLFAs suggest substrate

334

preferences of microbial groups. Without additional C or N (+0 treatments), fungal markers accounted for

335

18-21% of total C in PLFAs (depending on polymer amendment), and contained 21% of the total lignin-

336

derived C recovered in PLFAs, but only 13% of protein-derived C, and 4% of chitin-derived C (Fig. 3a).

337

The incorporation of lignin-derived C into fungi thus matched their contribution to the total community,

338

whereas the incorporation of protein- and chitin-derived C was disproportionally low. Markers for gram

339

negative bacteria accounted for 33-35% of total C in PLFAs, and contained 34% of chitin-derived C, 32%

340

of protein-derived C, but only 22% of lignin-derived C. Markers for gram positive bacteria accounted for

341

14-19% of total C in PLFAs, and contained 15% of lignin-derived C, 17% of chitin-derived C, and 12% of

342

protein-derived C. Pielou’s Evenness Index of the fraction of polymer-derived C compared to total C in

343

individual PLFAs was significantly lower for chitin than lignin and protein (+0 treatments; Fig. 3b),

344

reflecting a more selective incorporation of chitin-derived C in PLFAs.

345

The addition of easily available C reduced the amount of lignin-, chitin-, and protein-derived C in PLFAs,

346

suggesting the preferential use of glucose as a microbial C source. This reduction was stronger for lignin

347

(total PLFAs: -46% without additional N; -43% with additional N) and chitin (-72% and -37%,

348

respectively) than protein (-26% and -1%), but significant for all polymers and microbial groups (Fig. 4),

349

and for most individual PLFAs (Supplementary Table 3).

350

Nitrogen addition significantly reduced the amount of protein-derived C (-50% without additional C; -

351

33% with additional C) and chitin-derived C (-73% and -39%, respectively) in total PLFAs, suggesting the

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preferential use of ammonium as a microbial N source. This reduction was significant for all microbial

353

groups and all individual PLFAs in the case of protein, but only for gram negative bacteria and a set of

354

individual markers in the case of chitin (Supplementary Table 3). Nitrogen addition had no significant

355

effect on the amount of lignin-derived C in total PLFAs or PLFAs associated with certain microbial

356

groups, but changed the recovery of lignin-derived C in some individual PLFAs (Supplementary Table 3).

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3.5. Differences between polymer treatments and the control

359

Amending soils with lignin, chitin or protein itself also significantly affected microbial community

360

composition and function compared to the control (Supplementary Table 4). In +0 treatments without

361

addition of easily available C or N, lignin addition significantly increased the absolute and relative

362

abundance of PLFA markers for fungi, gram positive as well as gram negative bacteria. Lignin further

363

reduced activities of the cellulolytic enzymes CBH and the chitinolytic enzyme NAG, and increased

364

activities of the ligninolytic enzymes POX and PER. Chitin addition had little effect on microbial

365

community composition, but decreased activities of NAG and increased activities of the proteolytic

366

enzyme LAP, as well as of POX and PER. Protein addition increased the absolute and relative abundance

367

of fungi and gram positive bacteria, and decreased activities of BG as well as NAG. Similar differences

368

between polymer amendments were observed in +C, +N, and +CN treatments (Supplementary Table 4).

369

Overall, these findings indicate an adjustment of microorganisms to increased lignin, chitin, and protein

370

availability.

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4. Discussion

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Soil microorganisms may adjust to high C and low N availability by accelerating the breakdown of N-

373

containing polymers into assimilable units (“microbial N mining”), a mechanism that could mitigate N

374

limitation in the vicinity of plant roots, but also enhance soil C losses by respiration (“priming effect”).

375

We here show that N addition significantly reduced the recovery of chitin-derived C in the mobile C pool

376

over the course of a ten day laboratory incubation (Fig. 1), indicating reduced chitin depolymerization in

377

the presence of a more easily available alternative (the added ammonium), and thereby supporting the N

378

mining hypothesis. We expected that C addition would in turn stimulate the breakdown of polymers that

379

serve as microbial N sources such as chitin, by enhancing the microbial investment into N-targeting

380

enzymes. We found that C addition increased microbial N demand and the mobilization of SOM-derived

381

C, but not its incorporation into microbial C, which might indicate that microorganisms accelerated the

382

breakdown of N-containing polymers to access available N and respire excess C. Nevertheless,

383

depolymerization rates of lignin, chitin, and protein were significantly reduced. Although it is possible that

384

C addition promoted the extracellular mineralization of chitin- and protein-bound N to ammonium to meet

385

the microbial N demand, for instance via the Fenton reaction (Op De Beeck et al., 2018), this effect would

386

not explain the observed increase in SOM-derived respiration rates. The contrasting effects of C addition

387

on bulk SOM as well as lignin, chitin, and protein depolymerization thus challenge the concept of N

388

mining as a key driver behind the priming effect, and shed light on gaps in our current understanding of

389

SOM decomposition.

390

Carbon and N effects on SOM depolymerization were determined without addition of lignin, chitin, or

391

protein. Differences in microbial community composition and function between samples with and without

392

additional polymers (Table 2, Supplementary Table 4) suggest an adaptation of the microbial community

393

to polymer amendment that could favor a positive response of polymer breakdown to changes in

394

environmental conditions in samples that received additional polymers. The observed reduction in lignin,

395

chitin, and protein depolymerization upon C addition, however, rather points to increased availability of

396

other, native SOM compounds that replaced chitin, lignin, and protein as microbial C sources. Carbon

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addition might have enhanced the depolymerization of other soil compounds, with an overall positive

398

effect in spite of reduced lignin, chitin, and protein depolymerization rates. The amount of 13C-enriched

399

lignin, chitin and protein added to the soil corresponded to only 1.8% of the native soil organic C and C

400

fluxes from added compounds were consequently low compared to those from bulk SOM (Figs. 1 and 4).

401

Nevertheless, native, non-enriched lignin, chitin and protein have been shown to account for a large

402

fraction of SOM in a wide range of soils (Cécillon et al., 2012; Grandy et al., 2007; Haddix et al., 2011)

403

and their depolymerization might respond in a similar way to C and N addition than that of the added

404

compounds. Alternatively, C addition might have promoted not the breakdown of specific chemical

405

structures, but released organic compounds from mineral or physical protection. Lignin, chitin, and protein

406

artificially added in this study are chemically similar to their native counterparts, but likely less affected

407

by physical protection or the formation of organo-mineral complexes due to limited interaction time with

408

native soil minerals. The increased mobilization of bulk SOM in the presence of an easily available C

409

source might therefore result from reduced protection of native soil C, for instance a release from mineral

410

bonds mediated by organic acids as suggested in a previous study (Keiluweit et al., 2015). Organic acids

411

can be excreted by plants as well as microorganisms and produced during organic matter decomposition

412

(Jones et al., 2003).

413

The mobilization of lignin-, chitin-, and protein-derived C was only partly connected to activities of the

414

lignin-, chitin-, and protein-degrading enzymes measured in this study. The recovery of chitin- and

415

protein-derived C in the mobile pool was not correlated with the corresponding hydrolytic enzymes, i.e.,

416

the chitinolytic enzyme NAG and the proteolytic enzyme LAP. Previous studies have observed a similar

417

disconnection between gross protein depolymerization rates (measured using 15N pool dilution) and LAP

418

activities (Wanek et al., 2010; Wild et al., 2017), and have suggested that individual hydrolytic enzymes

419

might not be representative for the diversity of enzymes active in soil (see e.g., Vranova et al., 2013). In

420

contrast, the observed changes in lignin depolymerization to at least some extent matched the changes in

421

the oxidative enzymes POX and PER (Fig. 2) that have been connected to lignin degradation (Sinsabaugh,

422

2010). Oxidative enzyme activities might thus be more representative for oxidative depolymerization

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reactions in the soil than hydrolytic enzyme activities for hydrolytic reactions, possibly due to the higher

424

specificity of hydrolytic than oxidative enzymes.

425

Although proteins represent the largest fraction of soil N (Knicker, 2011) and amino acids are key N

426

sources for plants especially in boreal forests (Inselsbacher and Näsholm, 2012; Näsholm et al., 2009), we

427

found that the mobilization of protein-derived C was only minimally affected by changing C and N

428

availability. Carbon input significantly, but only weakly reduced protein-derived C mobilization, and N

429

input had no significant effect (Fig. 1). Nevertheless, the reduced microbial incorporation not only of

430

chitin-, but also protein-derived C into PLFAs upon N addition (Fig. 4) suggests that not only amino

431

sugars derived from chitin depolymerization, but also amino acids derived from protein depolymerization

432

served as microbial N sources under low-N conditions and were replaced by ammonium in the N addition

433

treatments. A weak response of protein depolymerization to changes in environmental conditions has also

434

been observed in previous incubation and field studies that quantified gross protein depolymerization rates

435

using

436

depolymerization rates by C and N input (Wild et al., 2017), elevated CO2, temperature and drought

437

frequency (Mooshammer et al., 2017; Wild et al., 2018). Similarly, a study quantifying 14CO2 evolution

438

from

439

(Jan et al., 2009). Our study suggests a higher flexibility of chitin than protein depolymerization under

440

changing environmental conditions, possibly related to a lower functional redundancy of chitinolytic than

441

proteolytic microbial communities. Supporting a more specific chitin-utilizing microbial community, we

442

observed a less even distribution of C from chitin than protein and lignin among individual PLFAs (Fig.

443

3b). Nevertheless, overall both chitin- and protein-derived C were preferentially incorporated into

444

bacterial compared to fungal PLFAs, while the opposite was the case for lignin-derived C (Fig. 3a). We

445

note however that microbial community composition and function might differ between laboratory

446

incubations and field conditions, and that in particular mycorrhizal fungi that have been linked to the

447

breakdown of N-containing soil polymers (Read and Perez-Moreno, 2003) are likely underrepresented in

448

our study. Microbial communities might further adjust to environmental conditions with time, and

14

N pool dilution assays. These studies reported only minor changes in gross protein

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C-labelled protein added to soil found no significant effect of ammonium or nitrate amendment

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microbial community functions such as SOM decomposition might change on time frames beyond our ten

450

day experiment, as shown in response to repeated additions of both C and N in previous more long-term

451

studies (Hamer and Marschner, 2005; Lavoie et al., 2011; Mau et al., 2018; Ohm et al., 2007).

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5. Conclusions

453

Based on two complementary microcosm incubation experiments, we here show that C addition

454

stimulated the degradation of bulk SOM over the course of ten days but reduced the degradation of lignin,

455

chitin and protein. Degradation products of the latter two are likely key N sources for soil microorganisms

456

under low-N conditions; we therefore reject the hypothesis that the observed positive priming effect was

457

driven by an increased microbial breakdown of N-containing polymers under increasingly N-limited

458

conditions. Our findings rather indicate that C input increased the mobilization of other SOM pools that

459

replaced lignin, chitin, and protein as microbial C sources, for instance by releasing SOM compounds

460

from mineral protection. We conclude that SOM decomposition is interactively controlled by multiple

461

mechanisms including the balance between C vs N availability. Dissecting these controls will be crucial

462

for understanding C and N cycling on an ecosystem scale.

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Acknowledgements

465

This study was funded by the Swedish Research Councils Formas (project number 229-2011-716 to T.R.)

466

and Vetenskapsrådet (project number 2016-04710 to P.B.), as well as the Strategic Research Area

467

Biodiversity and Ecosystem Services in a Changing Climate (BECC; www.becc.lu.se). Soil samples were

468

provided by the Svartberget Experimental Forest Research Station of the Swedish University of

469

Agricultural Sciences within the framework of the Swedish Infrastructure for Ecosystem Science (SITES).

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organic nitrogen acquisition by an ectomycorrhizal fungus. New Phytologist 218, 335–343.

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Paul, E.A., 2016. The nature and dynamics of soil organic matter: Plant inputs, microbial transformations,

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Read, D.J., Perez-Moreno, J., 2003. Mycorrhizas and nutrient cycling in ecosystems - A journey towards relevance? New Phytologist 157, 475–492. doi:10.1046/j.1469-8137.2003.00704.x Saiya-Cork, K.R., Sinsabaugh, R.L., Zak, D.R., 2002. The effects of long term nitrogen deposition on

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extracellular enzyme activity in an Acer saccharum forest soil. Soil Biology & Biochemistry 34, 1309–

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1315. doi:10.1016/S0038-0717(02)00074-3

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Schimel, J.P., Bennett, J., 2004. Nitrogen mineralization: challenges of a changing paradigm. Ecology 85, 591–602. doi:10.1890/03-8002

Schimel, J.P., Weintraub, M.N., 2003. The implications of exoenzyme activity on microbial carbon and

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Schmidt-Rohr, K., Mao, J.-D., Olk, D.C., 2004. Nitrogen-bonded aromatics in soil organic matter and

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their implications for a yield decline in intensive rice cropping. Proceedings of the National Academy

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of Sciences 101, 6351–6354. doi:10.1073/pnas.0401349101

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Sinsabaugh, R.L., 2010. Phenol oxidase, peroxidase and organic matter dynamics of soil. Soil Biology & Biochemistry 42, 391–404. doi:10.1016/j.soilbio.2009.10.014 27

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Treseder, K.K., Lennon, J.T., 2015. Fungal traits that drive ecosystem dynamics on land. Microbiology and Molecular Biology Reviews 79, 243–262. doi:10.1128/MMBR.00001-15 Vance, E.D., Brookes, P.C., Jenkinson, D.S., 1987. An extraction method for measuring microbial

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biomass C. Soil Biology & Biochemistry 19, 703–707. doi:10.1016/0038-0717(87)90052-6

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Vranova, V., Rejsek, K., Formanek, P., 2013. Proteolytic activity in soil: A review. Applied Soil Ecology 70, 23–32. doi:10.1016/j.apsoil.2013.04.003

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Short-term carbon input increases microbial nitrogen demand, but not microbial nitrogen mining, in a

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Wild, B., Ambus, P., Reinsch, S., Richter, A., 2018. Resistance of soil protein depolymerization rates to eight years of elevated CO2, warming, and summer drought in a temperate heathland. Biogeochemistry

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and phosphorus in terrestrial ecosystems. Global Ecology and Biogeography 22, 737–749.

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Zelles, L., 1997. Phospholipid fatty acid profiles in selected members of soil microbial communities. Chemosphere 35, 275–294. doi:10.1016/S0045-6535(97)00155-0 Zogg, G.P., Zak, D.R., Ringelberg, D.B., White, D.C., MacDonald, N.W., Pregitzer, K.S., 1997.

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Compositional and functional shifts in microbial communities due to soil warming. Soil Science

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Figure Captions

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Fig. 1. Amount of C from native SOM (Experiment 1) and from added lignin, chitin, and protein

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(Experiment 2) in DOC, microbial C, and cumulative respiration after ten days of incubation. Samples

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were additionally amended with easily available sources of C (+C), N (+N), or both (+CN) compared to a

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control (+0). Bars are means ± standard errors. Significant effects of C and N addition as well as their

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interaction (CxN) on the amount of SOM- or polymer-derived C, respectively, in DOC, microbial C and

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respiration, as well as their sum (“mobile C pool”) were analyzed using two-way ANOVA; significance

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levels (***, p < 0.001; **, p < 0.01; *, p < 0.05; n.s., not significant at p < 0.05) and effect directions are

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indicated. See Supplementary Table 1 for total DOC, microbial C, and respiration.

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Fig. 2. Relationships between soil enzyme activities and the amount of C from added lignin, chitin, and

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protein in the mobile C pool (sum of DOC, microbial C, and respiration) after ten days of incubation: (a,

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b) Ligninolytic enzymes phenoloxidase (POX) and peroxidase (PER) compared to lignin-derived C, (c)

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chitinolytic enzyme N-acetyl-β-D-glucosaminidase (NAG) compared to chitin-derived C, (d) proteolytic

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enzyme leucine-aminopeptidase (LAP) compared to protein-derived C. Correlations were tested using

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Spearman’s rank sum correlations; p-values and Spearman’s correlation coefficient rho are presented for

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correlations significant at p < 0.1. Correlations of POX and PER with lignin C in the mobile pool were not

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significant when the full dataset was used, but were significant when one outlying value was removed.

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Fig. 3. (a) Contribution of PLFAs associated with fungi, gram negative, and gram positive bacteria to the

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total amount of C in PLFAs, as well as to the amount of C from added lignin, chitin, and protein in

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PLFAs. Bars are means ± standard errors of microbial groups. (b) Pielou’s Evenness Index of the fraction

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of PLFA C derived from lignin, chitin, or protein among individual PLFAs. Boxplots show medians with

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25th and 75th percentiles as box limits, and significant differences between polymers are indicated by

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different letters (1-way ANOVA with Tukey’s HSD post hoc test; p < 0.001). Only treatments without

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addition of easily available C or N are shown here (+0 treatments); see Supplementary Tables 2 and 3 for

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other treatments.

651 Fig. 4. Amount of C from native SOM (Experiment 1) and from added lignin, chitin, and protein

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(Experiment 2) in total PLFAs (T), as well as in markers for fungi (F), gram negative (G-) and gram

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positive (G+) bacteria, after amendment with easily available sources of C (+C), N (+N), or both (+CN)

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compared to a control (+0). Values are means ± standard errors. Significant effects of C and N addition as

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well as their interaction (CxN) were analyzed using two-way ANOVA (***, p < 0.001; **, p < 0.01; *, p

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< 0.05; n.s., not significant at p < 0.05). See Supplementary Table 3 for individual PLFAs.

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Table 1. Ammonium and nitrate concentrations in forest soil after ten days of incubation. Samples were amended with no additional polymer, lignin, chitin, or protein, as well as with easily available sources of C (+C), N (+N), or both (+CN) compared to a control (+0). Values are means ± standard errors, and significant effects of C and N addition as well as their interaction (CxN) are indicated (two-way ANOVA; ***, p < 0.001; **, p < 0.01; *, p < 0.05; n.s., not significant at p < 0.05). Individual treatment data (means ± standard errors) Two-way ANOVA +0 +C +N +CN C N CxN Ammonium (nmol g-1 d.s.) No polymer 563.2 ± 33.6 58.0 ± 20.0 11025.4 ± 204.2 8450.6 ± 101.1 *** (-) *** (+) *** Lignin 600.4 ± 34.9 53.3 ± 10.4 10981.6 ± 45.8 8672.7 ± 222.6 *** (-) *** (+) *** Chitin 539.5 ± 20.7 23.1 ± 4.6 11386.8 ± 140.7 9692.0 ± 257.7 *** (-) *** (+) n.s. Protein 1097.9 ± 26.1 51.0 ± 11.6 12189.6 ± 602.3 9552.7 ± 271.9 *** (-) *** (+) * Nitrate (nmol g-1 d.s.) No polymer 4.9 ± 0.3 12.7 ± 4.7 4.1 ± 0.6 5.9 ± 0.5 *** (+) ** (-) n.s. Lignin 5.0 ± 0.5 8.6 ± 1.2 5.3 ± 0.5 5.8 ± 0.6 * (+) n.s. n.s. 4.8 20.9 ± 7.9 6.6 ± 0.5 21.8 ± 9.0 * (+) n.s. n.s. Chitin 11.8 ± Protein 8.7 ± 0.7 24.9 ± 8.9 4.1 ± 0.3 22.0 ± 10.7 *** (+) * (-) n.s.

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Table 2. Potential activities of the extracellular enzymes cellobiohydrolase (CBH), β-glucosidase (BG), N-acetyl-β-Dglucosaminidase (NAG), leucine-aminopeptidase (LAP), phenoloxidase (POX), and peroxidase (PER) in forest soil after ten days of incubation. Samples were amended with no additional polymer, lignin, chitin, or protein, as well as with easily available sources of C (+C), N (+N), or both (+CN) compared to a control (+0). Values are means ± standard errors, and significant effects of C and N addition as well as their interaction (CxN) are indicated (two-way ANOVA; ***, p < 0.001; **, p < 0.01; *, p < 0.05; n.s., not significant at p < 0.05). Individual treatment data (means ± standard errors) Two-way ANOVA +0 +C +N +CN C N CxN CBH (nmol MUF g-1 d.s. h-1) No polymer 48.5 ± 4.3 48.6 ± 2.9 79.8 ± 7.7 91.8 ± 15.4 n.s. *** (+) n.s. Lignin 41.7 ± 4.4 36.6 ± 1.4 71.3 ± 9.4 103.5 ± 11.9 n.s. *** (+) * Chitin 48.8 ± 3.0 36.5 ± 1.2 48.9 ± 4.4 66.4 ± 5.8 n.s. ** (+) ** 42.4 ± 2.6 50.1 ± 1.9 77.8 ± 6.4 * (+) *** (+) *** Protein 51.3 ± 3.1 BG (nmol MUF g-1 d.s. h-1) No polymer 331.2 ± 19.5 266.3 ± 19.6 339.2 ± 25.0 535.8 ± 51.0 n.s. *** (+) *** 201.0 ± 7.1 331.2 ± 23.2 417.5 ± 23.4 n.s. *** (+) *** Lignin 260.0 ± 13.9 Chitin 268.5 ± 27.2 216.2 ± 20.1 233.2 ± 6.6 381.9 ± 27.8 n.s. ** (+) *** Protein 249.2 ± 13.8 187.6 ± 16.3 278.4 ± 25.6 426.2 ± 37.3 n.s. *** (+) *** NAG (nmol MUF g-1 d.s. h-1) No polymer 580.5 ± 16.3 580.5 ± 40.8 651.8 ± 63.1 617.0 ± 26.9 n.s. n.s. n.s. Lignin 426.4 ± 26.4 399.7 ± 25.8 528.1 ± 38.3 607.5 ± 27.3 n.s. *** (+) n.s. Chitin 389.1 ± 31.3 360.5 ± 22.8 444.1 ± 25.8 547.5 ± 50.1 n.s. ** (+) n.s. Protein 403.5 ± 29.7 378.3 ± 28.5 405.5 ± 26.3 458.9 ± 31.0 n.s. n.s. n.s. LAP (nmol AMC g-1 d.s. h-1) No polymer 101.2 ± 2.4 97.1 ± 1.4 96.7 ± 2.1 100.2 ± 1.9 n.s. n.s. n.s. Lignin 102.1 ± 1.6 101.9 ± 0.9 104.0 ± 0.8 104.9 ± 1.1 n.s. n.s. n.s. Chitin 115.2 ± 1.3 110.9 ± 1.0 107.3 ± 0.7 110.7 ± 1.6 n.s. ** (-) ** Protein 104.2 ± 2.7 101.5 ± 2.4 106.9 ± 1.5 111.8 ± 1.8 n.s. ** (+) n.s. POX (nmol DOPA g-1 d.s. h-1) No polymer 280.1 ± 10.1 255.9 ± 9.4 227.2 ± 11.2 283.4 ± 12.5 n.s. n.s. ** Lignin 459.1 ± 16.1 399.5 ± 10.7 366.4 ± 7.5 387.5 ± 21.2 n.s. ** (-) * Chitin 379.0 ± 44.2 362.7 ± 8.5 329.5 ± 8.5 362.8 ± 9.7 * (+) n.s. n.s. Protein 229.3 ± 7.8 246.9 ± 3.0 225.1 ± 4.8 246.2 ± 6.9 ** (+) n.s. n.s. -1 -1 PER (nmol DOPA g d.s. h ) No polymer 1204.4 ± 48.7 1201.7 ± 61.8 1023.6 ± 96.6 1249.5 ± 57.6 n.s. n.s. n.s. Lignin 1998.1 ± 105.7 1676.5 ± 38.7 1417.8 ± 37.1 1610.0 ± 88.2 n.s. *** (-) ** Chitin 1598.7 ± 87.7 1464.0 ± 33.3 1310.3 ± 51.1 1508.4 ± 80.8 n.s. n.s. * Protein 1025.5 ± 33.6 1138.0 ± 55.6 889.2 ± 21.3 1225.5 ± 45.8 *** (+) n.s. **

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Highlights Low N and high C availability might boost soil N-polymer breakdown (“N mining”).

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N mining might thus lie behind higher soil respiration near plant roots (“priming”).

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We tested the effect of C and N input on the degradation of SOM and added polymers.

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C input increased SOM degradation but reduced that of chitin, lignin and protein.

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Our findings challenge the hypothesis of N mining as a key driver of priming.

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