DOI: 10.5061/dryad.prr4xgxm4
title: Low-quality carbon and lack of nutrients result in a stronger fungal than bacterial home-field advantage during the decomposition of leaf litter

description: Decomposition of litter is a key biochemical process that regulates the
 rate and magnitude of CO2 fluxes from biosphere to atmosphere and
 determines soil nutrient availability. Although several studies have shown
 that plant litter decomposes faster in their native compared to a foreign
 environment, i.e. a home field advantage (HFA) for litter degradation, to
 date HFA has only been considered in terms of respiration or litter mass
 loss. The ecological success will be determined by the decomposer
 microorganism’s ability to transform used OM into population growth, and
 therefore we hypothesized that HFA for microbial growth would be more
 pronounced than for decomposition, by driving the feedback between
 community alignment and environmental resource conditions. We also
 expected that HFA for microbial processes would increase with lower
 quality litter, that the fungal role in litter decomposition would be more
 dominant than that of bacteria, and that HFA effects would increase with
 more pronounced environmental differences between sites. We designed a
 2-month microcosm reciprocal transplant experiment with litter from two
 sites with contrasting climates (Atlantic and Sub-Mediterranean climates)
 and including 3 tree species (Quercus robur, Pinus sylvestris and Fagus
 sylvatica). We found a stronger HFA for microbial growth than for
 decomposition, that the nutrient content and C-quality of litter
 influenced the microbial HFAs, and that interactions between bacterial and
 fungal communities during litter decomposition modulated the HFA for
 litter degradation. Low litter nutrient content, strong nutrient
 limitations and low C-qualities all favoured fungal over bacterial
 decomposers, and our results suggest a dominant functional role of the
 fungal community and gave rise to HFA for fungal growth but that
 translated to only marginal implications for overall decomposition of
 litter.
description: <b>2.1 Experimental design and sample
 collection </b> We designed a 2-month microcosm
 reciprocal transplant study with litter and soil from 2 sites with
 contrasting climatic conditions (M-site was in a Sub-Mediterranean climate
 and A-site was in an Atlantic climate) and including 3 different tree
 species (<i>Quercus robur </i>L., <i>Pinus sylvestris
 </i>L. and <i>F</i><i>agus sylvatica
 </i>L.<i>)</i> at each of the two sites<i>.
 </i>The laboratory microcosm study was carried out at controlled
 temperature and moisture to standardise any direct influence by climate on
 the litter decomposition processes. We measured respiration, bacterial
 growth and fungal growth throughout the experiment to determine the
 responses of the microbial community to different treatments and estimated
 the HFA effect for each process. To elucidate putative factors that could
 drive HFA effect, we determined the litter quality of all studied litters,
 and the physiochemistry of the studied soils. In addition, to link HFA to
 the perceived nutrient quality of the studied litters, enzymatic
 activities related with C, N and P cycles were also measured. Finally,
 PLFA analysis were performed to screen for differences between microbial
 community structures resulting from different combinations of soils with
 litter. The O layer (approx. 5-10 cm deep) and leaves
 for the microcosm experiment were collected within the forest at Artikutza
 (43°13'N 1°48'W, Goizueta) and Montoria (42°38'N,
 2°44'W, Urizaharra), located in the northern and southern Basque
 County, respectively, in north Spain. Both study sites have acid soils,
 with pH-H<sub>2</sub>O 3.9 in Artikutza (“A-site”) and 4.4 in
 Montoria (“M-site”). Soil sampled from the A site has a sandy clay loam
 soil texture with 13% of organic matter and that sampled from the M site
 also had a sandy loam soil texture, with 9.4% of organic matter. Both
 soils were classified to be Umbisols (WRB, 2015). These two study sites
 differ in climatic conditions. While the northern site (Artikutza,
 orA-site), is located in an Atlantic climate with annual mean
 precipitations about 2500 mm, 16<sup>◦</sup>C annual mean
 temperature and annual 31 frost days, the southern M-site is in a
 sub-Mediterranean climate in which mean annual precipitation is
 approximately 650 mm, i.e. 4 times lower than in A site, and the mean
 annual temperature is 10.5<sup>◦</sup>C, i.e. about 5.5 °C
 cooler than the A-site, and the frost days are approximately double than
 in A-site (58 days). We selected three different tree species, similar in
 age, density and structure, which were present in the two areas: common
 oak (<i>Quercus robur </i>L.), scots pine (<i>Pinus
 sylvestris </i>L.) and European beech (<i>Fagus sylvatica
 </i>L.). Naturally senesced leaves were
 collected with litter traps (composited from 4 litter traps per forest
 type) between late October 2017 and early January 2018 from oak, pine and
 beech stands within each study site. They were oven-dried (40˚C) until dry
 (&gt;48 h) and ground. O layer samples (top 5-10 cm) were collected in
 early January 2018 during frost free conditions in both sites. Composite
 samples were collected in each stand around each litter trap. O layer
 samples were sieved (4 mm) and kept at 4˚C for a week until microcosm
 experiment started. Different microcosms were
 prepared for a reciprocal transplant experiment of litter from different
 study sites for each of the three tree species (Figure 1). Litter from the
 A-site was combined with its native soil (‘home’), but also with soil from
 the other site (‘away’), and <i>vice versa</i>, resulting in 4
 different treatments per tree species, and thus a total of 12 treatments.
 The treatments were replicated in three (n=3), resulting in 36 microcosms
 that were monitored over time. As we wanted to investigate the soil
 environment’s role in affecting the decomposition of litter by microbes,
 capturing this necessitates a sufficiently large proportion of soil to
 litter. Without it, the environments in the microcosm environments would
 be dominated by the litter only. Therefore, in each microcosm, 60 g of
 soil (O-layer) were adjusted to optimal moisture, at 50% of water holding
 capacity, after which 12 g of dried and ground litter were added. This
 combination of O layer with litter was used to simulate incoming litter
 into the top of a forest floor. Unsterilized litter will bring with it a
 small amount of microbes that could contribute to seed the soil
 environment below. However, the dried litter samples should have carried
 only a negligible microbial community compared to that already active in
 the O-horizon sample used, which we argue natural conditions of litter
 fall on a forest floor well. The microcosms were incubated at 16˚C, thus
 falling within a range of summer soil temperatures normal for both sites,
 and constant moisture (adjusted as needed) for 2 months, and respiration,
 bacterial growth and fungal growth were monitored (10 sampling times: 1,
 3, 5, 8, 12, 21, 28, 35, 42, and 55 days). In addition, biomarkers for the
 microbial community size and composition (PLFA) were determined 3 times (1
 day, 28 days and 55 days) over the experiment. Basic soil chemical
 properties, pH and electrical conductivity, and enzymatic activities were
 measured in the first and in the last sampling times to capture the
 differences in microbial nutrient availability.
 <b>2.2 Microcosm microbial parameters</b>
 Bacterial growth was measured by the incorporation of
 <sup>3</sup>H-Leucine (Leu) into bacteria (Bååth et al.,
 2001). At each time point, 1 g of soil was mixed with 20 ml demineralized
 water by vortexing for 3 min. The supernatant with a bacterial suspension
 was sampled after low-speed centrifugation (1000 <i>g</i> for
 10 min) and the incorporation of Leu was measured in 1.5 ml aliquots of
 the bacterial suspension. A combination of nonradioactive and tritiated
 Leu (<sup>3</sup>H-Leu, 37 MBq ml 1, 5.74 TBq mmol 1, Perkin
 Elmer, USA) were added to yield a final concentration of 275 nM in the
 bacterial suspension. The extracted bacteria were incubated for 1 h at
 16˚C. The samples were washed (Bååth et al., 2001) and the radioactivity
 of the incorporated Leu was measured on a liquid scintillator. Bacterial
 growth was expressed as the amount of Leu that was incorporated in the
 extracted bacteria per g dry soil and per h (Rousk and Bååth,
 2011). Fungal growth was
 assessed using the acetate-into-ergosterol-incorporation (Bååth, 2001;
 Rousk et al., 2009). A radioactively labeled acetate solution (20 µl
 1-[14C] acetic acid (sodium salt), 37 MBq/ml, 2.10 GBq/mmol, Perkin
 Elmer), was combined with 30 µl unlabelled 16 mM acetate and 1.95 ml water
 was added to 0.5 g of soil, yielding a final concentration in the soil
 mixture of 0.2 mM, and incubated for approximately 4 hours at 16˚C. Growth
 was terminated by adding 500 µl 10% formalin, and ergosterol was extracted
 and separated using high-performance liquid chromatography and a fraction
 collector (Rousk and Bååth 2007). The incorporated
 <sup>14</sup>C-acetate into ergosterol was measured using
 liquid scintillation and used as a proxy for fungal growth (Rousk
 & Bååth, 2011). Soil respiration was
 measured using a gas chromatograph equipped with a methanizer and a flame
 ionization detector. For each sampling time, 0.5 g of soil was put into a
 20 ml glass vial, which were purged with air, sealed and incubated at 16˚C
 (microcosm incubation temperature) for 2-6 hours (Silva-Sanchez et al.,
 2019). The concentration in the air used for purging was subtracted from
 the headspace concentrations, which was then divided by the incubation
 time to estimate the respiration rate. The respiration rates were
 expressed as µg CO<sub>2</sub> per g dry soil and per
 h. <b>2.3. PLFA
 analysis</b> The microbial phospholipids were
 extracted in a one-phase mixture of chloroform, methanol, and citrate
 buffer (Frostegård et al., 1991) from frozen subsamples (see above). Then,
 phospholipids were fractionated and methanolyzed into free fatty acid
 methyl esters (FAMEs) with FAME 19:0 added as an internal standard
 (Frostegård et al., 1993; Cruz Paredes et al., 2017). The resulting
 phospholipid fatty acids (PLFAs) were separated and quantified on a GC
 with a flame ionization detector (Frostegård et al., 1993). A total of 18
 PLFAs were selected and used to indicate the microbial community
 composition, and to estimate total, bacterial and fungal biomass
 concentrations.  Fungal biomass was quantified based on 18:2w6,9 content
 (PLFAfun), and bacterial biomass (PLFAbact) was quantified as the sum of
 i15:0, a15:0, i16:0, 16:1w7c, i17:0 cy17:0, 18:1w7, 18Me18:0 and cy19:0
 (Frostegård and Bååth, 1996; Ruess and Chamberlain, 2010), and the total
 biomass (PLFAtot) was calculated as the sum of the 18 PLFAs.