Application of 13C-labeled litter and root materials for in situ decomposition studies using phospholipid fatty acids

Soil Biology & Biochemistry 40 (2008) 2485–2493

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Application of 13C-labeled litter and root materials for in situ decomposition studies using phospholipid fatty acids Jennifer Moore-Kucera a, Richard P. Dick b, * a b

Oregon State University, United States School of Environment and Natural Resources, 2021 Coffey Road, Ohio State University, Columbus, OH 43210-1085, United States

a r t i c l e i n f o

a b s t r a c t

Article history: Received 27 March 2008 Received in revised form 28 May 2008 Accepted 2 June 2008 Available online 1 July 2008

Microorganisms play a central role in litter decomposition and partitioning C between CO2 evolution and sequestration of C into semi-permanent pools in soils. At the ecosystem level, forest stand age influences rates of litter accumulation and quality, and micro-climatology which could affect the microbial community structure and C sequestration processes. Although numerous laboratory experiments have studied the decomposition of model 13C-labeled compounds, few studies have verified these findings under field conditions. The objective of this study was to track decomposition of 13C-labeled Douglas-fir (Pseudotsuga menziesii (Mirb.) Franco) materials into the soil microbial community using 13C-phospholipids fatty acid (PLFA) analysis in three different aged forest stands. A field experiment was conducted that had three forest stand age treatments: old-growth (>500 yrs); 8-year-old clear-cut (CC8); and 25year-old clear-cut (CC25) (landscape reps of n ¼ 2). Each stand age had in situ microcosms that were amended with either 13C-labeled surface litter or root material. Microcosms were destructively sampled seven times over a 22-month period and the soil was analyzed for the relative amounts of 13C incorporated (13C%INCORP) into PLFAs and the proportional distribution of 13C incorporated into PLFAs. The 13 C%INCORP was affected by stand age and 13C source with greater 13C%INCORP in samples from CC8 than OG or CC25. Also, the level of 13C%INCORP was greater for labeled litter than root material in five out of the seven sample dates. In general, 18:1u9 and 18:2u6,9 (common fungal biomarkers) had the greatest amount of 13C incorporation throughout the study period in both clear-cut and old-growth sites, especially in plots with 13C-labeled litter. Our data showed a low fungal 13C-PLFA: bacterial 13C-PLFA ratio (0.45) 1 month after incubation was initiated compared to 5, 7 and 9 months after incubation (two of these dates were >1.0). This suggests that initially bacteria played a greater role in the decomposition of the added needles with fungi playing a more important role in subsequent sample dates. Our results illustrate that the use of 13C-labeled materials in field studies coupled with13C-PLFA profiling is a powerful tool for determining microbial dynamics during decomposition – enabling statistically significant detection of land management treatment effects on C acquisition by microbial functional groups. Ó 2008 Elsevier Ltd. All rights reserved.

Keywords: 13 C-PLFA Isotope probing Clear-cut Old-growth Douglas-fir Litter Root Decomposition

1. Introduction Soil microbial community structure is a function of the interactions between litter quality and quantity, root deposition, and the microclimate of the soil. This community is responsible for the decomposition of litter, the cycling of nutrients for plant growth and the formation of soil organic matter. Forest management practices, such as clear-cutting, have been shown to disrupt the soil microbial ecosystem via removal of organic matter, reduction in litter input, changes in root dynamics, alteration of nutrient cycles (Schmidt et al., 1996; Lindo and Visser, 2003) and increased temperature and moisture extremes (Keenan and Kimmins, 1993; * Corresponding author. Tel.: þ1 614 247 7605; fax: þ1 614 292 7432. E-mail address: [email protected] (R.P. Dick). 0038-0717/$ – see front matter Ó 2008 Elsevier Ltd. All rights reserved. doi:10.1016/j.soilbio.2008.06.002

Holmes and Zak, 1999). Although considerable progress has been made in understanding the factors affecting the rates of litter decomposition and formation of soil organic matter (Swift et al., 1979; Stevenson, 1994; Heal et al., 1997), most of what is known about the role of microbial functional groups during decomposition/C sequestration in soils is based on indirect methods that use inferences such as activity measurements, biomass C, or respiration to study the ‘‘black box’’ of decomposition. The composition and structure of the microbial community may affect degradation rates of organic substances. For example, communities with relatively high proportions of fungi have advantages over communities dominated by bacteria because fungi are able to extend hyphae into the soil to extract nutrients and water (Holland and Coleman, 1987) whereas bacteria are limited in movement. Fungi are major players in decomposition and formation of organic

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matter (Stevenson, 1994; Carlile et al., 2001; Cairney and Meharg, 2002), in part because they have a wide range of extracellular enzymes that can degrade recalcitrant materials such as lignin and cellulose (Hammel et al., 1993). Consequently, with clear-cutting there is a large reduction in litter inputs and root turnover which would reduce the substrates fungi are uniquely adapted to degrade, particularly lignin (Carlile et al., 2001). Furthermore, fungi are filamentous and therefore susceptible to the disturbance and compaction that occurs with clear-cutting activities. However, structurally different microbial communities may still function at similar levels (Buyer and Drinkwater, 1997). The ability to identify specific groups of microorganisms responsible during the decomposition of organic material will provide needed information regarding the direct pathways of C movement between plants, roots, microorganisms and soils (Bromand et al., 2001; Loya et al., 2002). Traditional methods such as culturing and identification, and direct counts through microscopy are limited because a high percentage of organisms are not culturable and identification through morphological characteristics can be difficult. Plus, these methods are extremely labor intensive and nonquantitative, making field-based studies of soil microbial ecology/ diversity impractical. The use of phospholipid fatty acids (PLFA) is a community-level biochemical method that can provide information regarding shifts in the microbial community structure resulting from management or environmental changes. It is difficult, however, to link PLFA markers with community function because the overall PLFA profile is a consortia of all organisms present at sampling time (Ludvigsen et al., 1997) and any given microbial process may be carried out by diverse taxa (Hill et al., 2000). Stable isotope probing (SIP) with 13C of specific biomarkers, such as PLFAs, is a promising tool to directly link specific microbial processes with the organisms responsible; thus shedding valuable insight into the biogeochemical transformations in natural environments that are mediated by microorganisms (Boschker and Middelburg, 2002). Although the term, SIP, was first used with DNA biomarkers (Radejewski et al., 2000), the first published example of SIP (Boschker et al., 1998) used PLFAs as the biomarkers. Isotopic techniques have been used to track C allocation and remobilization in plants (Mordacq et al., 1986; Svejcar et al., 1990; Thompson, 1996) and the fate of individually labeled C sources in soil in laboratory studies (Hu and van Bruggen, 1997; Abraham et al., 1998; Chotte et al., 1998; Arao, 1999; Ekblad and Hogberg, 2000; Ekblad et al., 2002; Abraham and Hesse, 2003; Burke et al., 2003). Similarly, 13C-labeled plant residues have been used in laboratory incubations (Butler et al., 2003; Malosso et al., 2004; McMahon et al., 2005; Waldrop and Firestone, 2004) and in field incubations (Bird and Torn, 2006; Williams et al., 2006). However, no studies to date have used 13C-labeled litter material in forest ecosystems to track its decomposition in situ using 13C-PLFA techniques. Our hypothesis was that forest stand age alters C uptake by microorganisms during decomposition. Specifically, we expected the fungal community to be less active (i.e., incorporate less 13C) in samples from the CC sites compared to those from the OG sites. The objective of this study was to track the decomposition of 13C-labeled Douglas-fir (Pseudotsuga menziesii) materials into the microbial community using 13C-phospholipids fatty acid (PLFA) analysis in forest soil of old-growth stands (>500 yrs) compared to stands clear-cut eight (CC8) and 25 years (CC25) prior to sampling. 2. Methods 2.1. Site description The study site is located within the Gifford Pinchot National Forest in the southern Cascade Range of Washington State. The forests are predominately Douglas-fir (P. menziesii) stands of

varying ages and have been described in detail by Shaw et al. (2004). The climate is cool and moist with annual averages of precipitation at 222 cm (5% of falls during June–August) and temperatures of 8.7  C (Shaw et al., 2004). The three forest stand ages in our study were (1) old-growth (OG) ranging from 300 to 500 years old, (2) an 8-year-old clear-cut stand (CC8) that was cut in 1994, and (3) a 25-year-old clear-cut stand (CC25) that was cut in 1977 and have been previously described (Moore-Kucera and Dick, 2008a). Both clear-cuts were immediately replanted with Douglas-fir (P. menziesii). The CC8 and CC25 had two landscape level replications where each had an adjacent paired OG stand giving OG four replications. The two OG sites nearest each other were averaged to give a balanced design (all treatments having n ¼ 2). The O horizons were 2–4 and 0– 0.5 cm deep for OG and clear-cut sites, respectively. The soils are deep, well drained loams and silt loams, medial, mesic Entic Vitrands (Stabler series, http://ortho.ftw.nrcs.usda.gov/ cgi-bin/osd/osdname.cgi). The overall mean soil pH value (0– 10 cm) was 5.1 for the three stand ages with a range between 5.0 and 5.3. The mean C values in the O horizon were 34%, 29%, and 21% for OG, CC8, and CC25, respectively, and the mean C values in the 0– 10 cm depth were 3.7%, 3.4%, and 4.3% for OG, CC8, and CC25, respectively. All of the stands except for CC8 are located within the Wind River Experimental Forest (WREF) – a 4208-ha area where active forest research has been conducted since 1908. The CC8 stands are adjacent to the WREF on USDA Forest Service land. 2.213C-labeled Douglas-fir material A pulse-chase technique was used to label 670 Douglas-fir seedlings with 13CO2 (99 at.%) as described by Moore-Kucera and Dick (2008b). At the end of the 9-week labeling period, seedlings were harvested, rinsed with deionized water, dried, and separated into needle, stem, and root parts. Over 1.5 kg of seedling dry matter was produced with 13C signatures (relative to the Pee Dee Belemnite standard) of 415, 295, and 228& for needles, stems, and roots, respectively. This was a significant enrichment over non-labeled plants that had isotopic signatures of 21, 19, and 24.0& for needles, stems, and roots, respectively. 2.3. Microcosms Between November 3 and 7, 2002, 14 microcosms constructed of 10-cm diameter  12- to 16-cm long open-ended PVC columns were inserted into the top 10 cm of mineral soil at each of the three sub-replicate plots in each stand (extra length accounts for depth of litter layer). Half of the 14 microcosms received litterbags containing the 13C-labeled needles or needle/stem mix (13C-LITTER) and the other half received 13C-labeled roots (13C-ROOTS). The litterbags (7.2 cm2) were constructed of a polyester-mesh layer (1.0 mm) on top to allow entry of soil fauna and a nylon-mesh layer (0.5 mm) on the bottom to allow entry of fungal mycelium. We deliberately chose rates of litter addition for the microcosms based on actual litter input rates for each stand age at WREF (as reported by Klopatek, 2002) to enable realistic rates of C uptake by microorganisms – or in other words to impose true stand age comparisons of 13C flow through microbial communities. Therefore, 13 C-labeled needles were added at the rate of 175, 200, and 100 g m2 and correspond to 1.4, 1.6, and 1.0 g of 13C-labeled needles for OG, CC25, and CC8 sites, respectively. The labeled needles were placed into litterbags and sewn shut with nylon thread. Because there is virtually no stem litter in the CC8 or CC25 stands, only OG litterbags received stem material (1.0 g) (Klopatek, 2002). Root material was added at the rate of 388 g m2 for OG and 173 g m2 for CC25 plots, based on annual root production as estimated by

J. Moore-Kucera, R.P. Dick / Soil Biology & Biochemistry 40 (2008) 2485–2493

Klopatek, (2002). The rate of root production for the CC8 was assumed to be 75% that of the CC25 site. Thus, root material was added at the rate of 130 g m2 for these sites. These rates correspond to the addition of 1.2, 0.5, and 0.4 g of 13C-labeled root material to the soils within OG, CC25, and CC8 plots, respectively. To install microcosms, a custom-made steel soil corer with beveled edges (10-cm diameter by 16-cm) and an internal floating plate (Scotty’s Welding, Wenatchee, WA) was used to extract intact soil cores. The PVC column was placed in the hole and a coupler connected the column to the corer. The soil was transferred from the corer to the column by lightly tapping the internal plate. This ensured that minimal disturbance occurred and soil layers remained intact. Immediately in the field, cores were amended with labeled plant materials. For the labeled litter, the top O horizon was removed, set aside, and a litterbag containing the 13 C-LITTER was placed on the soil surface and the O horizon repositioned on top of the litterbag. Columns that received 13CROOTS were established in a similar manner except the 13C-ROOTS were freely incorporated (i.e., no bags) into the top 10 cm of mineral soil before the O horizon was replaced. 2.4. Sample collection On December 16, 2002 (M1), April 25 (M5), June 20 (M7), August 31 (M9), and November 23 (M12), 2003, and June 14 (M19) and September 1 (M22), 2004, one pair of microcosms (one containing 13 C-LITTER and one containing 13C-ROOTS material) was randomly chosen from each sub-replicate plot for PLFA analysis. Abbreviations in parentheses correspond to months (M) of incubation from 1 to 22. Control soil samples adjacent to microcosms (>0.5 m) were collected in April 2003. For control samples, five soil samples (0– 5 cm) were collected using a soil corer and bulked to give one composite sample for each site. Samples were stored at 4  C and processed within 3 days. One of the three sets was analyzed for PLFA concentration and the 13C value for each PLFA by GC–C–IRMS. Each sample was separated into litter layer, 0–5 cm soil below the litterbag or 0–5 cm mineral soil (root cores) and sieved through a 4.75 mm mesh to remove wood pieces, roots, or stones and homogenized. Sub-samples were dried at 65  C for 24 h to calculate moisture content. 2.5. Tissue and phospholipid fatty acid (PLFA) analysis Needle, stem and root tissue were analyzed for total C and N on Carlo Erba Elemental Analyzer (EA1108, Carlo Erba, Milan, Italy). PLFAs were extracted in three steps using a modified Bligh and Dyer (1959) procedure. In brief, lipids were extracted from soils using a solution of phosphate buffer, chloroform, and methanol at a ratio of 0.8:1:2 and then fractionated into neutral- glycol- and phospholipids on a silicic acid column (Supelco, Inc., Bellfonte, PA). The phospholipids were then subjected to alkaline methanolysis and analyzed via gas chromatography (Agilent 6890) equipped with a 30-m Hewlett Packard Innowax 2 column (0.25 mm internal diameter; 0.25 mm film thickness) connected to a Europa ORCHID on-line combustion interface attached to a Europa 20–20 mass spectrometer. The carrier gas was He, and the oven temperature was ramped from 120 to 260  C at a rate of 5  C per min with a 5min hold at 260  C. Carbon dioxide of known isotopic composition was injected at the beginning and end of each run. Individual fatty acids (FA) were identified using the following standards: 37 FAMEs mixture (FAME 37 47885-4; Supelco, Inc.), 24 bacterial FAMEs mixture (P-BAME 24 47080-U; Supelco, Inc.), and MIDI standards (Microbial ID, Inc., Newark, DE). Quantification of FAMEs was accomplished by comparing peaks to analytical standards containing 19– 458 pmol ml1 tridecanoic FAME (Supelco, Inc.). Of the 23 FAs identified, 15 could

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not be used because of GC–C–MS limitations where the separation of FAs during the GC stage was not maintained during the combustion and MS analysis stages so that these CO2 pulses had mixed d13C value. The nine peaks we did use, however, accounted for between 60 and 70% of total PLFAs that we found in a subsequent study at the same site. Here we used the other two sub-replicate samples and identified 43 individual peaks (Kucera, 2006). The peaks labeled 16:1u7 and 18:1u7 co-eluted with PLFAs 10Me16:0 and 10Me18:0, respectively and hence are designated as 16:1u7þ and 18:1u7þ, respectively. 2.6. Calculations PLFA d13C values were corrected for the addition of the extra C atom introduced during derivatization with methanol of known d13C value (44.6&) as described by Pelz et al. (1997). Following the equation used by Boschker et al. (2001), the percentage of added 13C (LITTER or ROOTS) incorporated into a specific PLFA (%13CINCORP) was calculated as:

  % 13 C ¼ 100 ðFtx  Ft0 Þ½PLFAi tx =½13 C  added ;

(1)

13

with the fraction of C before (t0) and after (tx) incubation as: F ¼ 13C/(13C þ 12C) ¼ R/(R þ 1), the concentration of PLFA in mg C kg1 soil, and the concentration of 13C added in mg 13C kg1 soil. The carbon isotope ratio (R) was derived from the measured d13C values (&) relative to the Pee Dee Belemnite (PDB) standard as: R ¼ (d13C/1000 þ 1)*RPDB, with RPDB ¼ 0.0112. This equation accounts for the difference in total mass of added materials to the three stand age groups. The proportion of 13C incorporated into an individual PLFA 13 ( CPLFA) of the microbial community was expressed relative to the 13 C incorporated in all PLFAs (%13CM-DIST) as:

% 13 C ¼

13

C  PLFAi =

X

13

C  PLFAi ;

13

where CPLFA is the amount of PLFA and is calculated as: 13

C ¼ ðFtx  Ft0 Þ  ½PLFAi tx ;

13

(2)

C incorporated into an individual

(3)

13

with the fraction of C before (t0) and after (tx) incubation and [PLFA]tx described in Eq. (1). The %13CM-DIST in bacterial and fungal PLFAs was calculated using the sum of i15:0, 16:1u7 þ 10Me16:0, and 18:1u7 þ 10Me18:0, cy17:0, and cy19:0 PLFAs for bacterial PLFAs and the sum of 18:1u9 and 18:2u6,9 for fungal PLFAs (Federle et al., 1986; Phillips et al., 2002). Unfortunately for interpretation purposes, the peaks that coeluted represent two different microbial groups and thus cannot be used together as a general taxonomic category. The PLFAs 16:1u7 and 18:1u7 are Gram-negative biomarkers whereas 10Me16:0 and 10Me18:0 are characteristic of the actinomycetes, a specialized group of Gram-positive biomarkers (Zelles and Bai, 1994; Zelles, 1999). Using data from a separate study that investigated soils from the same stand ages, we were able to determine that on average, 66% of the 16:1u7 þ 10Me16:0 peak was attributed to 16:1u7 (Kucera, 2006). Similarly, 18:1u7 made up about 89% of the 18:1u7 þ 10Me18:0 peak concentration. Although we were able to account for mass differences, we were not able to determine if 66 and 89% of the 13C signature is due to 16:1u7 and 18:1u7, respectively, because a large signature can come from a peak with a small area. 2.7. Statistical analysis All statistical analyses were conducted using SAS (SAS Institute, Inc., Cary, NC, version 8) and PC-ORD (MJM Software Design,

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Gleneden Beach, OR, version 5). The stand by age treatment effects were analyzed by standard ANOVA. To determine how the structure and composition of soil microbial communities differed between stand age (AGE), 13C-SOURCE, and sampling date (DATE), a threeway repeated measures analysis of variances was conducted. The compound symmetry variance–covariance matrix was used because it does not require equal spacing between data sampled through time to meet convergence criteria. Tukey’s honestly significantly different tests were used for pair-wise comparisons to control for the experiment-wise error rate with significant differences defined at P  0.05. Changes in the distribution of 13C in individual PLFAs (%13CM13 C-SOURCE, or DATE DIST) resulting from the main effects of AGE, were assessed using a non-metric multidimensional scaling (NMS) plot. Prior to NMS construction, a monotonic transformation (square root) of all PLFAs was also conducted to create a more normally distributed data set and to reduce the coefficient of variation among PLFAs. The square root transformation is similar in effect to the log transform but is less drastic and is commonly used in ecological studies (McCune and Mefford, 1999). A second matrix containing the original %13CM-DIST values was used to construct a joint plot overlaid on the NMS. This joint plot is used to visualize the relationship between a set of variables (in this case, %13CM-DIST of individual PLFAs or the sum of all bacterial PLFAs (15:0i, 16:1u9þ, 18:1u7þ, cy17:0, and cy19:0), fungal PLFAs (18:2u6,9 and 18:1u9, and the fungal:bacteria ratio (F:B))) and NMS scores. The angle and length of a line indicate the direction and strength of the relationship (McCune and Grace, 2002). 3. Results 3.1. Needle decomposition rates Except for M9, the differences in mass loss of needles between M1 and M19 were significantly greater in samples from old-growth sites compared to those from the clear-cut sites (Fig. 1). By M22, there was little to no litter remaining in any of the samples making precise weight measurements difficult as a result of encroachment of soil into the bags, a common limitation in litterbag studies. There were no significant differences (P < 0.05) in the percent litter mass loss between CC8 and CC25 at any sampling date. The stem materials that were added to OG remained intact and lost less than 5%

mass at M1 and no more than 30% by M22. Thus, we assumed that 13 C transfer from stems to PLFAs would be negligible during the early stages of decomposition (e.g., during the first 9 months of incubation). 3.2. Total amount and percent 13C incorporated into PLFAs as a function of stand age and litter source The total amount of 13C incorporated into all of the PLFAs (13CPLFATOT) averaged over AGE and 13C-SOURCE was greater for the first 9 months of incubation compared to 19–22 months after incubation. This value peaked at M7 (3.8 mg 13C-PLFATOT kg1 soil) and was significantly greater at this sampling date than all other dates. There was no significant difference in 13C-PLFATOT from M9 until the end of the incubation (M22) (0.6 mg 13C-PLFATOT kg1 soil) (Table 1). In order to compare the amount of 13C incorporated into PLFAs across AGE, it was necessary to account for the difference in total mass of added materials to the three stand age groups. Thus, the percentage of added 13C (LITTER or ROOTS) that was incorporated into PLFAs (%13CINCORP) was calculated. The %13CINCORP in PLFAs ranged from 0 (e.g., several PLFAs at M1) to 0.02 % (18:2u6,9 for CC8 LITTER in M5 and M7) (data not shown). The total %13CINCORP into all PLFAs was significantly greater in CC8 than OG and CC25 for M1 and both clear-cuts had significantly greater total %13CINCORP for M7 (Fig. 2). Separating the %13CINCORP of the bacterial PLFAs or fungal PLFAs showed the same trend as illustrated in Fig. 2 with significantly greater %13CINCORP for CC8 than OG and CC25 (data not shown). Averaged over the entire study period, %13CINCORP into all PLFAs for CC8 was highest (0.032%), CC25 intermediate (0.020%), and OG the least (0.011%). There was also a significant difference due to 13C-SOURCE with a greater %13CINCORP into all PLFAs for LITTER (0.026%) compared to ROOTS (0.016%). 3.3. Stand age, litter source and temporal effects on the distribution of 13C incorporated into PLFAs Based on the %13CM-DIST for individual PLFAs, there was a significant effect of AGE for %13CM-DIST only for the fungal PLFA (18:1u9) and the mixed PLFA 18:1u7þ (Fig. 3). The %13CM-DIST in 18:1u9 was significantly greater in OG (16.1%) compared with CC25 (8.8%) and CC8 (12.4%). There was a significant effect of SOURCE on the %13CM-DIST only for one of the fungal PLFAs (18:1u9) with greater %13CM-DIST of 18:1u9 in LITTER (14%) compared with ROOTS (11%). Using the sum of %13CM-DIST for the five bacterial PLFAs, AGE was a significant effect. The %13CM-DIST for the five bacterial PLFAs was significantly greater for CC25 (53%) than either OG (46%) or CC8 (47%) whereas the %13CM-DIST of the F:B ratio was significantly lower for CC25 (0.59) than either OG (0.79) or CC8 (0.80). Table 1 Absolute amount of 13C incorporated into all PLFAs (mg 13C-PLFATOT kg1 soil) for each of the seven sample dates averaged over stand age and 13C source (standard error in parentheses, n ¼ 12)

Fig. 1. Needle mass lost (%) for samples from OG, CC25, and CC8 sites during a 22month incubation (M1 through M22) (stem material was removed by hand before determining needle mass on OG samples). Error bars represent standard error (n ¼ 6; three sub-replicates per replicate). Data for M7 was lost. Within a given date, means with the same letter were not significantly different at P  0.05.

Sampling date

Sum of 13C-PLFA incorporation (mg 13C-PLFATOT kg1 soil)

November 3–7, 2002 (T0) December 16, 2002 (T1) April 20, 2003 (T2) June 14, 2003 (T3) August 31, 2003 (T4) November 23, 2003 (T5) June 14, 2004 (T6) September 1, 2004 (T7)

– 2.16b (0.58) 1.78b (0.37) 3.97a (0.64) 0.92bc (0.12) 1.15bc (0.19) 0.69c (0.11) 0.60c (0.08)

Means followed by the same letter are not significantly different at P  0.05.

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Fig. 2. The percent of 13C incorporated in total PLFAs relative to the total amount of 13C added (%13CINCORP) averaged over 13C source for each stand age (OG, CC25, and CC8) and sample date (M1–M22). Error bars represent standard error (n ¼ 2). Within a given date, means with the same letter or ns were not significantly different at P  0.05.

The NMS plot of %13CM-DIST revealed changes in the relative 13C enrichment of the PLFA biomarkers over time (Fig. 4a). The lines depicted on the joint plot overlay show %13CM-DIST for individual PLFAs having a strong correlation along the axes (Fig. 4b). For example, the fungal PLFAs, 18:1u9 and the F:B ratio increases from top to bottom along axis 2 (i.e., M19 and M22 have the lowest values, M5 and M9 have the highest values, and M1, M7, and M12 have similar mid-range values of %13CM-DIST for these PLFA markers). In contrast, the %13CM-DIST for the sum of all bacterial PLFAs (BAC_sum) increases from bottom to top along axis 2 left with greater %13CMDIST for M19 and M22 compared to M5 and M9. Additionally, the %13CM-DIST for cy19:0 increases from right to left along axis 1. However, joint plot overlays only provide information on general trends along a given axis. Differences of least squares means with Tukey adjustment resulted in a significant effect of DATE for all of the PLFAs biomarkers except for cy17:0. (Table 2). The lowest %13CM-DIST among four out of the five bacterial PLFAs occurred at M5 or M9. The %13CM-DIST of i15:0, a Gram-positive bacterial biomarker, was significantly lower at M9 than M1, M7, or M22. The %13CM-DIST

Fig. 3. Relative distribution of 13C in individual PLFAs (%13CM-DIST) for each of the three stand ages averaged over the 22-month incubation and two 13C sources. Error bars represent standard error (n ¼ 14). Within a PLFA, means with the same letter were not significantly different at P  0.05.

Fig. 4. NMS plot (a) with joint plot overlay (b) of relative distribution of 13C in total PLFAs (%13CM-DIST for each of the seven sample dates averaged over stand age and 13C source). Error bars represent standard error (n ¼ 12). Joint plot overlay showings vectors based on %13CM-DIST for individual PLFAs. The angle and length of the vector indicate the direction and strength of the variable and the NMS axis. Axes are arbitrary; the closer points are on the plot, the more similar they are in SMC composition.

of 16:1u7þ was significantly lower at earlier dates (M5 and M9) compared to the end of the incubation (M12, M19, and M22). The %13CM-DIST of 18:1u7þ was highest at M1 (24%) and significantly higher than all other dates except at M19 (M5–M12 and M22 average ¼ 13%). The %13CM-DIST of the cyclopropyl PLFAs, cy17:0 and cy19:0, often indicators of stress, showed different patterns: cy17:0 remained constant over the sampling period, whereas cy19:0 was low at the beginning of the incubation and increased through M9. The %13CM-DIST for cy19:0 was significantly lower at M1 (4.8%) than M9 and M12 (average 14%). The effect of DATE also was significant for the %13CM-DIST in the fungal PLFAs 18:2u6,9 and 18:1u9 (Fig. 5). The %13CM-DIST in 18:2u6,9 was lowest at M1 (13%) and lower than the next three sample dates (representing the first 9 months of incubation). The greatest %13CM-DIST was at M9 (24%) and this was significantly greater than M1 and M22 (16%). For 18:1u9, the %13CM-DIST was significantly greater at M5 and M9 than M1, M19, and M22. A significant effect of DATE was found for the arbuscular mycorrhizal PLFA, 16:1u5 (P ¼ 0.042), however, further separation between dates was not found using the Tukey honestly significantly different tests. The proportion of the active fungal decomposers to bacterial decomposers (%13CM-DIST-F:B) was lowest at M1 (0.45) (Fig. 6). This ratio was highest at M5 (1.1) and M9 (1.0) and was significantly greater for these dates compared to M1, M12, M19, and M22.

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Table 2 Relative distribution of microbial-13C (%13CM-DIST) for the general biomarker, 16:0, and the five bacterial PLFAs, and the sum of the bacterial PLFAs (BAC_sum) for each of the seven sample dates averaged over stand age and 13C source Date

%13CM-DIST 16:0

i15:0

16:1u7þ

18:u17þ

cy17:0

cy19:0

BAC_sum

T1 T2 T3 T4 T5 T6 T7

20.3a (2.9) 14.5b (1.4) 13.3b (1.1) 13.1b (1.4) 14.8b (1.3) 12.7b (1.5) 16.0ab (1.2)

7.5a (1.5) 6.8ab (1.4) 8.8a (1.5) 2.1b (0.7) 7.1ab (0.9) 5.8ab (1.3) 8.0a (1.6)

10.1ab (1.7) 7.0b (1.0) 11.6ab (0.6) 7.7b (1.2) 13.2a (1.0) 14.4a (2.6) 16.0a (1.1)

24.0a (4.0) 13.3b (2.3) 12.4b (0.8) 12.3b (1.0) 12.3b (1.6) 19.2ab (2.8) 13.5b (2.3)

6.2a (1.8) 2.9a (1.0) 5.0a (0.6) 4.6a (0.8) 3.9a (0.8) 7.2a (1.8) 4.5a (1.0)

4.8a (1.8) 7.7ab (1.4) 10.3ab (0.8) 15.2b (1.2) 12.9b (0.9) 10.6ab (3.3) 10.8ab (1.2)

52.6a (2.1) 37.6c (1.8) 48.0ab (1.9) 42.1bc (2.3) 49.4ab (2.3) 57.1a (3.2) 52.8a (3.3)

Standard error is in parentheses (n ¼ 12). Within a column (individual PLFAs), means followed by the same letter are not significantly different at P  0.05.

4. Discussion 4.1. In situ

13

C-PLFA stable isotope probing methodology

With regular GC analysis 40 þ PLFA peaks can be identified. However, in our study only nine PLFAs had a clear d13C signature. In part this was due to GC–C–MS limitations where peaks or PLFAs separated by the GC were combined in the combustion chamber and entered the MS as a single pulse of 13CO2. Thus, this makes separation of d13C signatures impossible for these overlapping fatty acids. With newer GC–C–MS instruments that have reduced the combustion chamber size, combined with optimized columns and run conditions, this problem has been greatly reduced (R. Dick, personal communication) – so that most peaks separated on the GC can obtain a d13C signature value on the MS. Regardless, our results are very promising for further labeling studies in the field because statistically significant treatment effects on 13C-PLFA measurements could be detected. This was feasible in spite of: the added variability of field incubation conditions; use of a more depleted 13C material; and use of a complex plant source. This is in contrast to the highly enriched and simple model compounds used in most other studies. For example, Boschker et al. (1998) reported less than 0.1% of 13C-enriched acetate (99% 13C) in individual PLFAs of sulfatereducing bacteria from aquatic sediments. We reported a maximum of 0.11% of total PLFA-C derived from labeled Douglas-fir material and a maximum of 0.022% of any one individual PLFA. We were able to find only two published studies that utilized 13 C-labeled plant residues in situ; one study investigated the incorporation of straw or clover residue into PLFAs over time

Fig. 5. Relative distribution of 13C in total PLFAs (%13CM-DIST) for the three fungal PLFAs for each of the seven sample dates averaged over stand age and 13C source. Error bars represent standard error (n ¼ 12). Within a given PLFA, means with the same letter were not significantly different at P  0.05.

(Williams et al., 2006) and the other studied C and N dynamics using 13C-labeled Ponderosa pine needles and roots (Bird and Torn, 2006). Neither of these reported the actual percent of 13C incorporated into the microbial biomass relative to the amount added and thus, we were not able to compare our results. Nonetheless, Williams et al. (2006) report much higher recovery of 13C in PLFAs ranging between 40 and 200 nmol C g1 soil of root-derived PLFA-C. Using the same units as Williams et al. (2006), our recovery was at most, 12.4 nmol 13C-PLFA g1 soil (data not shown). Likely, the high recovery reported by Williams et al. (2006) resulted from the incorporation of relatively high-quality, 13C-labeled compounds that were released into the rhizosphere during the multiple pulselabeling events over the growing season. Bird and Torn (2006) also reported a relatively high total 13C recovery of approximately 30% in bulk soil but did not indicate how much was recovered in the microbial biomass.

4.2. Effect of stand age We hypothesized that the dominant microbial members actively decomposing would be different as a function of stand age and the associated levels of disturbance caused by clear-cutting. Specifically, we expected the fungal community to be less active (i.e., incorporate less 13C) in samples from the CC sites compared to those from the OG sites. This hypothesis was supported by data reported by Moore-Kucera and Dick (2008a) who found significant differences in the soil microbial community structure (based on mol %) between the recent clear-cut (CC8) and OG as well as a significant

Fig. 6. The relative distribution of 13C (%13CM-DIST of fungal:bacterial PLFAs (F:B)) for each of the seven sample dates averaged over stand age and 13C source. Error bars represent standard error (n ¼ 12). Means with the same letter were not significantly different at P  0.05.

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reduction in bacterial and fungal biomass (based on PLFA concentrations) in CC8 compared to OG. However, we report that the total PLFAs and specifically, the fungal PLFAs 18:2u6,9 and 18:1u9, in CC8 were more highly labeled with 13C (i.e., greater %13CINCORP) than that found in the OG sites. Reduced 13C incorporation in the OG sites, despite a greater rate of needle mass loss and greater microbial biomass, may be the result of dilution of the 13C signature from microbial assimilation of alternate (and more abundant) C sources. For example, OG sites had a more complex understory and a litter layer containing 58% more organic C (on an area basis) (Moore-Kucera and Dick, 2008a). This would result in a microbial population at the OG sites that has greater access to more C sources for metabolism compared to the population found at the CC sites. Alternatively, a lower amount of 13 C incorporation in OG samples may occur when the microorganisms are actively mineralizing C but are not necessarily synthesizing lipids, especially when N is limiting (Williams et al., 2006). As the three forest stands are equally N limited (MooreKucera and Dick, 2008a), mineralization of C without lipid synthesis is likely occurring at similar levels across all three ages and not a contributing factor to the AGE effect on %13CINCORP. The distribution of 13C among PLFAs (%13CDIST) better describes how C was distributed among the ‘active’ microbial population. We are uncertain why there was an AGE effect for the %13CDIST of the fungal marker 18:1u9 but not for 18:2u6,9. Monounsaturated fatty acids are also characteristic of Gram-negative bacteria so one cannot be certain the signature for the 18:1u9 is from fungi. However, the %13CDIST of these two fungal markers tracked similarly across time suggesting they were behaving in a similar fashion. A third fungal biomarker, the 16:1u5, which is more specifically used as an arbuscular mycorrhizal (AM) marker, was found to be equally labeled across all three stand ages. This was in spite of the fact that only OG and CC8 sites had understory plants capable of forming arbuscular associations. For example, vine maple (Acer circinatum) was a major understory plant at OG and various grasses were found at CC8, whereas little to no vegetative growth was observed at CC25 (Moore-Kucera and Dick, 2008a). Again, this monounsaturated fatty acid has been shown to characterize some Gram-negative bacteria (Bradley et al., 2006). 4.3. Effect of

13

C-source

We expected, at least at the early stages of decomposition, a greater %13CINCORP in PLFAs from ROOTS compared to LITTER because of the close contact between added root material and soil but this was not the case. Although there were no differences in SMC structure between ROOT and LITTER treatments (Moore-Kucera and Dick, 2008a), rates of decomposition were faster for LITTER (based solely on the amount of 13C incorporated in PLFAs); a trend most prevalent in CC8 samples and especially so for the fungal PLFA, 18:1u9. This is consistent with Bird and Torn (2006) who showed that needles turned over 52% faster than roots in a Ponderosa pine system over a 1.5 yr study. They attributed this difference to the chemical quality of the pine material with a greater proportion of acid-resistant compounds and a higher C:N ratio for roots compared to needles. However, our results showed that C:N ratios were quite similar (w19) between roots and needles. Unfortunately, too little material remained at the end of the experiment to get a reliable estimate of lignin content but root materials are expected to be higher in lignin than above ground plant parts and may, in part, explain the slower decomposition of root material. We report no difference in SMC structure between samples from ROOT compared to LITTER treatment. Other studies have shown a residue-associated microbial community whose labeling patterns were influenced by residue quality over time (Williams et al., 2006; Lu et al., 2003). The lack of a community shift in our samples was

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not surprising because we added materials in levels that were consistent with normal annual litter fall or root turnover rates. Moreover, we did not exclude materials already present and thus, it is reasonable that the native microbial populations would not be altered by our additions; this approach was intentional as we sought to mimic the true C inputs of each stand age. 4.4. Temporal dynamics Destructive sampling over a 22-month period showed there was seasonal variation in the degree of 13C enrichment of individual PLFAs and enrichment was highest when PLFA biomass was highest during June (M7) when environmental conditions are best for supporting microbial activity. The seasonal variability of C acquisition by microorganisms showed the importance of capturing seasonal dynamics to understand the fate of C and to correctly identify the microbial players. This supports the previous work of Lipson et al. (2002), Brant et al. (2006), and Moore-Kucera and Dick (2008a) who showed strong changes in microbial communities, especially of fungi, on a seasonal basis. The rapid loss of litter and labeling of lipids after only 1 month (M1), especially for OG, was surprising, despite the relatively cold and wet conditions of the soil during this time (November–December). We applied the material in late fall because this is the time when most of the litter fall occurs in PNW forests. The soil temperature at 10 cm, averaged over 3 days prior to sample collection at M1 was 7.3  C (Moore-Kucera and Dick, 2008a) and mean total precipitation in November typically is over 350 mm (Shaw et al., 2004). However, our results fall in line with recent studies by Lipson et al. (2002) and Schadt et al. (2003) who have shown high microbial biomass and activity during the winter, largely due to seasonal shifts in fungal species and diversity. Our results suggest that frequent sampling is needed at early stages of decomposition for these forest residues, even if initiated in the winter for this region. A potentially interesting outcome that needs further investigation is the rapid labeling of 13C-litter in fungal biomass after only 1 month incubation. This and our visual observations showing fungal hyphae growing into the litter from the mineral soil layer provide further evidence of the possibility of fungal transport of residue nutrients and C to the mineral soil which has been suggested by isotopic studies of Frey et al. (2003) and Williams et al. (2007). Alternatively, we cannot reject the possibility that water-soluble 13C had moved from the litterbag into the mineral soil below and then taken up by fungi. In the latter scenario, since water-soluble C is a relatively labile C source, it would be reasonable for bacterial biomarkers to have greater 13C uptake than the fungal biomarkers at the early stages of decomposition. This is supported by considering the fungal:bacterial %13CM-DIST ratio, which was lower at M1 than the next three sample dates (representing the first 9 months of incubation) indicating that bacteria were more actively acquiring labeled material compared to fungal populations at the early stages of decomposition; a trend found for both samples under LITTER and ROOT materials. It may well be that both leaching and hyphal transport of litter C to mineral soil happen simultaneously. More controlled studies that isolate the effect of leaching are needed to distinguish these two mechanisms. Given the relatively low fungal:bacterial 13C-PLFA ratio (0.45) for M1 compared to the next three sample dates (M5 and M9 > 1.0), it appears that bacteria had greater access to the added (13C labeled) needle or root material at M1 than fungi at later dates. This scenario is in accordance with general decomposition models where a twostage decomposition process occurs with bacteria (primarily Gram negative) predominating at early stages of decomposition, followed by Gram-positive bacteria and then fungi at later stages (Minderman, 1968; Chen et al., 2002). It is possible that the disturbance to

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the site at the study onset including the addition of fresh litter stimulated the bacterial population. However, our comparisons of unlabeled PLFA inside and outside the microcosms showed no significant differences in microbial community structure (MooreKucera and Dick, 2008a), indicating minimal disturbance effects due to microcosm installation. An inherent challenge of SIP at later stages of decomposition is determining whether 13C microbial uptake is secondary feeding on labeled primary microbial biomass rather than the original labeled substrate. Our study provides indirect evidence that secondary feeding of the isotopically labeled biomass was minimal. First, we showed a decrease in the total 13C-incorporation in all PLFAs as the incubation proceeded. Second, the temporal enrichment of functional groups fits the general conceptual model of microbial succession during decomposition as described above. 5. Conclusions In this study we tracked 13C from labeled Douglas-fir litter or root material into the microbial phospholipid fatty acids during decomposition over a 22-month incubation under field conditions. Our data showed that the SMC structure shift (diversity) was much greater for seasonal effects (DATE) than stand AGE or residue SOURCE. This suggests that environmental controls take precedence for driving the differences in community structure over clear-cutting. Nonetheless there were nuances in microbial 13C acquisition, within dates, as a function of stand age. For example, there was greater uptake of 13C by fungi in the CC8 stand than the other treatments at nearly all sampling dates. Furthermore, C sources (i.e., ROOTS vs. LITTER) were degraded at different rates but overall a similar suite of soil microbial community members were found to be taking up 13C regardless of C source. The high variability of 13C in lipids can be attributed to the high micro-scale spatial variability of the isotope technique relative to microbial uptake of C. From an experimental perspective this would suggest there is a need to have smaller litter pieces that are highly homogenized if incorporated in soils or evenly distributed in the litter layer for litter layer placement of labeled materials. In spite of high spatial variability and the rapid rates of decomposition, we were able to detect significant stand age and substrate treatment effects on 13C enrichment of PLFAs – particularly for the fungal biomarker throughout the 22-month period in the clear-cut sites. Our work shows that the 13C-PLFA method is a robust tool that can substantially aid soil ecologists at the field scale. Acknowledgments This research was supported by the Office of Science (BER), U.S. Department of Energy, through the Western Regional Center of the National Institute for Global Environmental Change under Cooperative Agreement No. DE-FC02-03ER63613 and No. DE-FC0390ER-90ER61010. The authors would also like to thank the following people: Dr. Rockie Yarwood for technical assistance and for running 13C samples; Plum Creek Timber Toledo, OR for donating the seedlings; Joan Sandeno for field assistance and editing the manuscript; and Scotty’s Welding Wenatchee, WA for assistance designing and manufacturing the soil corer. We appreciate the thoughtful comments and suggestions of the anonymous reviewers. References Abraham, W.R., Hesse, C., 2003. Isotope fractionations in the biosynthesis of cell components by different fungi: a basis for environmental carbon flux studies. FEMS Microbiology Ecology 46, 121–128.

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