Laccase Gene Composition and Relative Abundance in Oak Forest ...

Microb Ecol (2009) 57:50–57 DOI 10.1007/s00248-008-9437-0

SOIL MICROBIOLOGY

Laccase Gene Composition and Relative Abundance in Oak Forest Soil is not Affected by Short-Term Nitrogen Fertilization Christian L. Lauber & Robert L. Sinsabaugh & Donald R. Zak

Received: 26 October 2007 / Accepted: 25 July 2008 / Published online: 29 August 2008 # Springer Science + Business Media, LLC 2008

Abstract Anthropogenic nitrogen (N) deposition affects a wide range of soil processes including phenol oxidase (PO) activity and soil organic matter dynamics. Depression of phenol oxidase activity in response to N saturation is believed to be mediated by the activity of white-rot basidiomycetes, whose production of extracellular oxidative enzymes can be limited by high N availability. We examined the effect of short-term N deposition on basidiomycete laccase gene diversity and relative abundance in temperate oak forest soil in which significant decreases in phenol oxidase and increased SOM have been recorded in response to experimental N deposition. UniFrac was used to compare the composition of laccase genes between three control- and three nitrogen-fertilized (80 kg−1 ha−1 per year) oak forest soils. The relative abundance of laccase genes was determined from qPCR analysis of laccase and basidiomycete ITS gene abundances. Our results indicate that there was no significant shift in the composition of laccase genes between control- and N-fertilized soils, nor was there a significant change in the relative abundance of laccase genes. These data suggest that N deposition effects on mineral soil PO activity do not result from changes in C. L. Lauber : R. L. Sinsabaugh Department of Biology, University of New Mexico, Albuquerque, NM 87131-0001, USA D. R. Zak School of Natural Resources and Environment, University of Michigan, Ann Arbor, MI 48109-1014, USA C. L. Lauber (*) University of Colorado at Boulder CIRES, 216 UCB, Boulder, CO 80309-0216, USA e-mail: [email protected]

laccase gene diversity of white-rot basidiomycetes but are likely the result of altered microbial abundance or expression in this ecosystem type. Furthermore, laccase gene composition may be tied to factors that structure microbial communities in general, as soil laccase gene communities are more similar to other forest soils than with the corresponding litter.

Introduction Basidiomycetes are key organisms in the decomposition of plant litter because of their ability to translocate nutrients and produce extracellular oxidative enzymes that can efficiently degrade lignin [4, 33]. Their role in the C dynamics of mineral soil is less clear, particularly in ecosystems in which the dominant plants also have ectomycorrhizal symbionts. Both brown and white rot basidiomycetes produce laccases acting at various redox potentials, and in some cases, redox mediators are used to step up these potentials to enhance lignin degradation [6, 15, 25]. In addition to laccases, some taxa produce lignin and manganese peroxidases, both of which contribute to the degradation of lignin and other polyphenolic compounds, but the mechanisms of degradation vary among taxa and are not well described [33]. Ectomycorrhizal basidiomycetes as well as saprotrophs carry genes for laccases, but the contribution of ectomycorrhizal fungi to soil oxidative potentials is not known [5, 9]. The contribution of basidiomycetes to soil C dynamics is of particular interest in the context of global N enrichment [22]. Fog [18], extrapolating from culture studies of lignin peroxidase expression by white rot basidiomycetes, proposed that decreased rates of decomposition for litter of high lignin content under conditions of high N availability

Nitrogen Amendment and Laccase Gene Diversity

was the result of reduced oxidative enzyme expression by basidiomycetes. Subsequent research has demonstrated that not all white rot basidiomycetes exhibit N-dependent expression, and at least some white rot fungi use laccase rather than peroxidase as their primary lignolytic agent [5, 25]. Nevertheless, Carreiro et al. [10] showed that the effects of N availability on litter decomposition were correlated with changes in phenol oxidase (e.g. laccase) activity. The relationship between N amendment and changes in phenol oxidase activity was extended to the ecosystem scale in studies at experimental N deposition sites in northern Michigan that showed that high N availability promotes increased cellulolytic and phenol oxidase activity in labile organic matter fractions and depresses phenol oxidase and peroxidase activities in recalcitrant fractions [14, 20, 36, 37]. As a result of these enzymatic responses, organic matter accumulated in N-amended ecosystems dominated by oak (Quercus velutina–Quercus alba) and declined in N-amended ecosystems dominated by sugar maple (Acer saccharum) [38]. Blackwood et al. [7], sampling the same experimental sites, found that basidiomycete diversity in the litter was similar for oak- and maple-dominated ecosystems, but basidiomycetes were an order of magnitude more abundant at the oak sites, in which the greatest declines in phenol oxidase activity have been observed in response to experimental N deposition. Despite the declines, neither the abundance of basidiomycetes nor the diversity of laccase sequences significantly changed in response to 2 years experimental N deposition [7]. In contrast, fungal biomass, ectomycorrhizal fungal diversity, and phenol oxidase activities all declined in hardwood and pine forest stands at the Harvard Forest, where experimental N deposition had been in progress for 15 years [19]. These observations suggest that the mechanisms underlying short-term and long-term losses of soil oxidative activity may differ. Short-term responses, which appear after a single N application, are probably the result of altered enzyme expression [20, 21]. It is plausible that, as treatment is extended over time, the varied effects of N saturation may begin to alter microbial community composition. Neither the short-term nor long-term effects have been resolved at the level of microbial community composition. This resolution is necessary for understanding soil carbon sequestration trends across ecosystems. In this study, we have used a molecular-based approach to characterize the effect of nitrogen fertilization on the diversity and abundance of basidiomycete laccase genes in mineral soil. The sites, established in 2001, are part of the long-term N deposition study described above. Our hypothesis was that the reduced phenol oxidase activity was the product of shifts in the laccase gene ‘community’

51

or selection against basidiomycetes likely to contain laccase genes (e.g., relative abundance), indicating changes in basidiomycete community composition are driving PO activity. Our rationale was that sustained suppression of oxidative enzyme expression in fungi that use environmental N availability to regulate expression would be a selective disadvantage, and such organisms might ultimately be competitively excluded from the community. In addition, we compared the laccase gene communities in the mineral soil to those in the corresponding litter previously described by Blackwood et al. [7] and to those described by Luis et al. [28, 29] in a German beech–oak forest soil.

Materials and Methods Site Description The three experimental sites, located in the Manistee National Forest (MNF) of northern Michigan (approximately 44°17′ N, 86° W), have an overstory of black and white oak (Quercus velutina–Quercus alba) on entic haplorthods of the Rubicon soil series. The sites are a subset of those examined by Zak et al. [41], and have been used for several studies [31, 41, 42], including a recent publication by Blackwood et al. [7], who examined the sequence diversity of laccase genes in the litter; and Gallo et al. [20], who examined oxidative enzyme activity. Beginning in 2001, these sites have been part of a large-scale N deposition experiment. Each site has three experimental plots (10× 30 m) that are randomly assigned to three levels of N deposition (ambient/control, +30 and +80 kg N ha−1 per year). The N is applied as NaNO−3 pellets in six equal increments over the growing season. Mean ambient N deposition is approximately 17 kg ha−1 per year [32]. Mean annual precipitation for the study area is 81 cm, mean annual temperature is 7.2°C, and the growing season varies from 100 to 150 days [1]. At each site, the overstory canopy is composed entirely of oak trees, about 90 years in age. The soil (0–10 cm) has a sandy texture (72% sand), a pH of 4.0, and 2.2% organic matter. Analyses of phospholipid fatty acids show that fungal biomass in the upper 20 cm of mineral soils are twice that of bacteria [20]. Descriptive data on soil and nutrient pools for these sites have been compiled by Gallo et al. [20]. Sample Collection Soil samples were collected in June 2003 from three replicate black oak–white oak stands (sites 3, 58, and 59) in which each stand had a control- (C) and N- (N) fertilized (+80 kg NaNO−3 ha−1 per year) plot, resulting in six samples for analysis. The plots were visually divided into

52

quadrants; two soil cores (2 cm diameter×20 cm deep), excluding litter and roots, were randomly taken from each quadrant. The eight cores for each plot were combined into a composite sample and passed through a 2-mm mesh. After removing sub-samples for analyses of moisture, organic matter, and enzyme activity, the remaining material was frozen at −80°C for subsequent molecular analyses. Enzyme activity data can be found in Gallo et al. and Sinsabaugh et al. [20, 37]. The diversity of laccase genes for litter in these plots was previously described by Blackwood et al. [7]. DNA Extraction and Amplification Genomic DNA was isolated from 0.25-g sub-samples of soil stored at −80°C using the Mobio PowerSoil™ DNA Isolation Kit (Carlsbad, CA, USA). Amplification of laccase genes was accomplished using degenerate primers describe by Luis et al. [28]: Cu1F 5′ CA(T/C) TGG CA(T/ C) GGN TT(T/C) TT(T/C) CA-3 and Cu2R 5′ G G(A)CT GTG GTA CCA GAA NGT NCC-3′. Final concentration of polymerase chain reaction (PCR) reagents are as follows, 1× Promega Taq DNA polymerase buffer (Promega Life Sciences, Madison, WI, USA), 0.04% bovine serum albumin, 2 mM MgCl2, 200 μM dNTP mix, 400 μM each forward and reverse primers, 1.25 U Taq DNA polymerase, and 2.0 μl genomic DNA; PCR grade water was used to bring the reaction volume to 50 μl. Thermocycling parameters were a modification of conventional touchdown PCR method. Initial denaturation was for 5 min at 94°C: stage 1, 94°C 60 s, 55°C 60 s, 72°C 90 s for seven cycles; stage 2, 94°C 60 s, 55–45°C 90 (−0.5°C per cycle), 72°C 90 s for 21 cycles; stage 3, 94°C 60 s, 40°C 60 s, 72°C 90 s for seven cycles. A final primer extension of 10 min was added after stage 3. Each sample was amplified in triplicate and run on 2.0% NuSieve GTG low melting point agarose (Cambrex Life Science, East Rutherford, NJ, USA) to isolate PCR bands of the expected size (≈200 bp). The bands were excised from the gel and purified using the Qiagen Gel Extraction Kit (Qiagen, Valencia, CA, USA). The bands were pooled and treated to ensure the addition of T residues at the end of the PCR fragments for efficient cloning. Cloning and Amplification of PCR Products PCR products were cloned using the TOPO TA Cloning Kit for Sequencing (Invitrogen, Carlsbad) and transformed into TOP10 chemically competent E. coli and grown overnight at 37°C on L-agar supplemented with 50 μg ml−1 kanamycin. Template DNA for sequencing was generated by direct amplification of cloned laccase genes or by Phi 29 rolling circle amplification. Direct amplification of laccase

C. L. Lauber et al.

genes were performed on overnight cultures of single colonies (37°C, L-broth with 50 μg μl−1 kanamycin) according to Sambrook and Russel [35] with M13 forward and reverse primers. Amplification was carried out at 94°C 5 min, then 30 cycles of 94°C 15 s, 55°C 30 s, and 72°C 30 s. PCR reactions were purified using Eppendorf Perfectprep® PCR Cleanup kit for 96-well plates (Eppendorf, Hamburg, Germany). Phi 29 rolling circle amplification was accomplished by incubating a small portion of a single colony in 1.25 μl annealing buffer (800 μl 25 mM MgCl2, 12.5 μl 1-M Tris–HCl at pH 8.0, and 187.5 μl sterile deionized water), 1 μl exonuclease-resistant random hexamer primers (500 μM, Fermentas Life Science, Hanover, MD, USA), and 3.25 μl sterile deionized water. The cell suspension was incubated for 3 min at 95°C then immediately placed on ice. Phi 29 polymerase was added in 5 μl aliquots of a solution containing 1.0 μl 10× Phi 29 buffer, 0.4 μl 10 mM dNTPs (Promega Life Science), 0.2 μl (2 U) Phi 29 DNA polymerase (Fermentas Life Science), and 3.4 μl sterile deionized water per reaction. Samples were incubated for 12 h at 30°C followed by 20 min at 65°C to deactivate the enzyme. The reaction was diluted with 10 μl Tris–HCl (10 mM, pH 8.2). All samples were stored at −20°C until needed. Sequencing Sequencing was accomplished using the BigDye Terminator Cycle Sequencing Kit v 1.1 (Applied Biosystems, Foster City, CA). Sequencing reactions were composed of 2 μl of a 1:5 dilution (2 μl BDT with 8 μl 5× reaction buffer) of the BigDye Terminator mix, 0.5 μl purified laccase gene fragment, 1 μl (20 pmol) of M13 forward primer, and 3.5 μl DNA/DNase-free molecular biology grade water. The samples were initially denatured at 95°C for 5 min then subjected to 99 cycles of 95°C for 20 s, 50°C for 20 s, 60°C for 4 min, with a 4°C hold at the end of the program [34]. Extension products were precipitated using the ethanol/EDTA method described in the sequencing kit manual and air dried for 5 min. The volume of each reagent was scaled to match the 7-μl volume of the sequencing reaction. The air dried samples were resuspended in 10 μl Hi Dye Formamide (Applied Biosystems), vortexed for 3 min then denatured for 5 min at 95°C and run on an ABI 3100 automated genetic analyzer. Quantitative PCR The relative abundance of laccase genes was determined from the ratio of laccase gene abundance to basidiomycete ITS gene abundance. Basidiomycete abundance was determined using 5.8sr and ITS4-B primers [17], while laccase


Sequence Analysis Introns were identified by aligning the laccase fragments in this study with an alignment kindly provided by Dr. H. Kellner [28, 29]. The exons were translated in the +1 reading frame and compared to GenBank sequences to ensure putative laccase genes were analyzed. Similarity to other laccase sequences was done by grouping sequences into operational taxonomic units (OTU) using FastGroup II [40] at 77% similarity [7]. The relationship of OTU to other laccase sequences was done using the BLAST algorithm [3] against a set of laccase genes from cultured fungi. Laccase gene sequences were deposited in GenBank under the accession numbers EF116965–EF117199. The aligned exons were used to produce a neighborjoining phylogenetic tree with the probable metallooxidoreductase gene of Pseudomonas aeruginosa PAO1 (GI 9949938) in Phylip 3.6 as the out group [16]. The NJ tree was then sent to UniFrac to determine the phylogenetic distance between each sample using the weightednormalized options to account for differences in sample size. The resulting phylogenetic distances were then analyzed for N fertilization effects using fertilization as the main factor and visualized by principle coordinate analysis (PCA) [26, 27]. We also compared the phylogenetic relationship of laccase genes in different soil, litter, and forest types of previous studies to those reported here. Using similar methodology as outlined above, we compiled laccase genes from Blackwood et al. and Luis et al. [7, 28, 29] and visualized the phylogenetic distance between each ‘community’ of sequences by PCA. Sequences reported by Blackwood et al. [7] represent litter from the same plots

sampled in this study as well as laccase genes from sugar maple–basswood (SMB; Acer saccharum–Tilia Americana) forest sites at MNF. The Luis et al. [28, 29] data set is representative of sequences across O, A, and B horizons of a 20-cm core from a German beech–oak (Fagus sylvatica– Quercus robur) forest soil. Sequences in this study are from homogenized samples of O and A horizon mineral soils. Statistical Analysis Statistical analyses were performed to determine the effect of N fertilization on the phylogenetic distance between laccase gene communities and the relative abundance of laccase genes between the control- and N-amended soils. The UniFrac phylogenetic distance matrix was analyzed using the analysis of similarity matrix (ANOSIM) in Plymouth Routines in Multivariate Ecological Research v. 5 (Primer v. 5, Lutton, UK) with fertilization as the main effect. qPCR data was tested using the Kolmogorov– Smirnoff test for non-parametric data in Systat 11.0. In all cases, the effect of N fertilization was considered significant when P0.7) on the relative abundance of laccase genes

OTU grouping showed the majority of laccases in this study (55%) were most closely related to laccase genes of ectomycorrhizal fungi belonging to the genus Russula (Table 1). Laccase gene communities were spatially heterogeneous and were qualitatively assessed by comparing sequences reported by Blackwood et al. and Luis et al. to the data presented in this study [7, 28, 29]. The PCA plot (Fig. 1) indicates that laccase gene communities of mineral soils tend to be more similar, while laccase sequences derived from litter appear to be phylogenetically distinct between litter types as well as from soil. Laccase genes reported by Luis et al. [28, 29] for A and B soil horizons clustered with the samples in this study than with O horizon or litter sequences. Litter laccase gene communities appear to be unique to each litter type and those from oak litter bear little resemblance to the underlying soil communities (Fig. 1).

Discussion (ANOSIM, P>0.1). The relative abundance of laccase genes was similarly not affected by N fertilization. Although laccase genes were slightly more abundant in control than fertilized plots (Fig. 2), Kolmogorov–Smirnoff testing indicated the difference was not significant (P>0.7).

Nitrogen fertilization reduces PO activity and has been shown to slow the decomposition and mineralization of plant detritus [10, 38]. One possible explanation is that N selects for specific taxa, of which some may be inefficient

Table 1 Distribution of operational taxonomic units (OTU) among the sample sites OTU

n

3C

3N

58C

58N

59C

59N

EF117137 EF117095 EF117166 EF117027 EF116969 EF117117 EF116989 EF116990 EF117032 EF116995 EF117176 EF117015 EF117017 EF117104 EF116973 EF116986 EF116998 EF117035 EF117069 EF117134 EF117135 EF117181 EF117193

49 29 27 26 23 13 12 11 9 8 7 6 3 3 1 1 1 1 1 1 1 1 1

1 2 11 3 5 5 0 3 0 0 1 1 2 0 0 0 0 0 0 1 1 0 0

12 8 16 1 4 2 0 2 1 1 2 0 0 0 0 0 0 0 0 0 0 1 1

0 0 0 6 4 0 12 3 1 6 0 0 0 0 1 1 1 0 0 0 0 0 0

5 1 0 6 6 1 0 3 2 0 0 3 1 0 0 0 0 0 0 0 0 0 0

27 5 0 4 1 2 0 0 4 1 0 0 0 0 0 0 0 1 1 0 0 0 0

4 13 0 6 3 3 0 0 1 0 4 2 0 3 0 0 0 0 0 0 0 0 0

Closet Match

Accession

Russula nigricans Russula nigricans Pleurotus cornucopiae Russula mairei Lepista nuda Russula nigricans Macrolepiota procera Mycena crocata Russula nigricans Mycena crocata Russula mairei Marasmius alliaceus Stropharia squamosa Mycena crocata Mycena galopus Russula mairei Mycena crocata Lentinula edodes Lentinula edodes Russula mairei Russula nigricans Russula mairei Russula mairei

AJ542622 AJ542621 AJ420174 AJ542625 AJ542593 AJ542621 AJ542611 AJ542586 AJ542621 AJ542587 AJ542625 AJ542603 AJ542599 AJ542587 AJ542640 AJ542625 AJ542587 AJ420172 AJ420171 AJ542625 AJ542621 AJ542625 AJ542625

An OTU is defined as 77% sequence homology within the putative coding region, the accession number for the representative sequence is shown. n is the number of sequences detected for each OTU. Sites are designated by location (3, 58, 59) and by treatment (C control, N nitrogen amended). Accession numbers represent laccase sequences from cultured fungi


PO producers or may not produce laccase for the purpose of C mineralization [4, 13]. However, in this study, we have shown that N fertilization does not affect the diversity of laccase genes (Fig. 1), indicating that N has not selected for any one group of laccase gene containing fungi in MNF oak forest soil. Previous work in these plots also demonstrates that N fertilization has little effect on microbial community composition. Blackwood et al. and Hofmockel et al. [7, 21] have reported no significant changes in laccase gene diversity within the litter, while Gallo et al. [20] reported only slight differences in the PLFA profiles of mineral soil shortly after N fertilization, even though PO activity was significantly reduced. Although N fertilization has been linked to declines in fungal diversity, microbial biomass and PO activity in long-term studies, these sites have received additional N for longer periods than MNF and have had significant changes in some edaphic properties [8, 14, 19, 39]. Collectively, these data indicate that the effects of short-term N fertilization on PO activity are not conspicuously associated with changes in the diversity of laccase genes and are likely the result of reduced expression. The phylogenetic examination of laccase genes in this study was designed to identify shifts in specific laccase gene communities that may be related to PO activity. Alternatively, we tested the possibility that the effect of additional N was non-specific and reduced the relative abundance of laccase genes in N-fertilized soils, thus providing an explanation for reduced PO activity. qPCR analysis, however, showed that there was no significant difference in the relative abundance of laccase genes (Fig. 2), indicating that N fertilization is not likely affecting PO activity by excluding laccase gene containing basidiomycetes. The lack of N fertilization effect on the relative abundance of laccase genes, though, raises the question as to why there is no selection against fungi believed to be sensitive to N availability, which participate in lignocellulose mineralization [13]. A recent report by Allison et al. [2] showed that short-term N fertilization (2 years) had little effect on the relative abundance of most active fungi, suggesting that competitive exclusion by N amendment may not occur immediately after the commencement of N fertilization. This observation is concomitant with the results in this study, suggesting that the relative abundance of laccase genes may follow a general pattern of fungal community dynamics in response to short-term N fertilization, which is independent of nutrient availability. We were curious as to why there was no selection for any particular group of laccase genes and looked to previous reports to identify possible factors that may contribute to structuring laccase gene ‘communities.’ Luis et al. [29] have observed that the distribution of laccase genes was spatially diverse, suggesting that niche selection

55

may be an important factor in determining the composition of laccase genes. Principle component analysis of laccase sequence data shows that laccase gene communities between soil and litter from the same plot are more phylogenetically distant than mineral soils from different locations (Fig. 1). One possible explanation for this observation is that the laccase gene composition is defined by the types of fungi found in soil and litter. For instance, litter typically contains more saprotrophs, while soils tend to have more ECM fungi. BLAST analysis of our data indicates that approximately 55% of the laccase sequences could be linked to ECM fungi; similar results were reported by Luis et al. [29]. This is in contrast to Blackwood et al. [7] who found that a majority of laccase sequences in litter were related to saprotrophic fungi. The data suggest that laccase gene composition is directly related to the overall composition of the fungal community rather than a product of specific selection. The use of culture-independent techniques to assay microbial communities has expanded greatly in recent years. Yet, relationships between microbial function and specific groups of organisms may be occluded depending on the strategy used to amplify sequences from environmental samples. For instance, the use of DNA, rather than RNA, may be contributing to the results presented in this study. Basidiomycetes may contain multiple laccase genes, of which a subset may be expressed for the mineralization of polyphenolic substrates [30]. What is more, DNA preparations from soil may also include sequences from inactive populations, which would further limit attempts to link function with phylogeny [21]. Minimizing these factors should be taken into account in future studies attempting to link functional gene diversity with microbial community function. Much attention has been given to the role of fungi as degraders of aromatic C. However, a number of bacteria have the ability to produce powerful oxidative enzymes that mediate the mineralization of polycyclic aromatic substrates [11, 12, 23]. Yet, there have been few studies that link the diversity of these genes to soil C cycles [24]. What is more, the contribution of bacteria to total PO activity is undefined and may be an important facet of litter decomposition and mineralization. The results presented here and in other studies would suggest that examining shifts in bacterial community composition as well as bacterial functional genes may be warranted when attempting to link N effects to PO activity. Even with large step increases in N deposition, it apparently takes several years for the relatively rapid physiological responses by the soil microbial community to be mirrored by significant changes in microbial community composition. Several studies have shown that laccase gene diversity is not affected by N fertilization and

56

responses may be constrained by edaphic factors that control microbial community composition. Changes in the edaphic characteristics of the microbial environment appear to have a strong influence on the composition of laccase genes, so much so, that laccase gene ‘communities’ across a similar depth profile tend to be more similar than between litter and mineral soil of the same site. Available evidence suggests that microbial enzymatic responses to increased rates of N deposition, even after years of treatment, are the result of altered expression by microbial communities whose composition is edaphically constrained. Acknowledgment This research was supported by the Office of Science (BER), US Department of Energy, grant no. DE-FG0203ER63591.

References 1. Albert DA, Denton SR, Barnes BV (1986) Regional landscape ecosystems of Michigan. School of Natural Resources, University of Michigan, Ann Arbor 2. Allison SD, Hanson CA, Treseder KK (2007) Nitrogen fertilization reduces diversity and alters community structure of active fungi in boreal ecosystems. Soil Biol Biochem 39:1878–1887 3. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignment tool. J Mol Biol 215:403–410 4. Baldrian P (2006) Fungal laccases—occurrence and properties. FEMS Microbiol Rev 30:215–242 5. Bending GD, Read DJ (1997) Lignin and soluble phenolic degradation by ectomycorrhizal and ericoid mycorrhizal fungi. Mycol Res 101:1348–1354 6. Bermek H, Li K, Eriksson KE (1998) Laccase-less mutants of the white rot fungus Pycnoporus cinnabarinus cannot delignify kraft pulp. J Biotechnol 66:117–124 7. Blackwood CB, Waldrop MP, Zak DR, Sinsabaugh RL (2007) Molecular analysis of fungal communities and laccase genes in decomposing litter reveals differences among forest types but no impact of nitrogen deposition. Environ Microbiol 9:1306–1316 8. Bowden RD, Davidson E, Savage K, Arabia C, Steudler P (2004) Chronic nitrogen additions reduce total soil respiration and microbial respiration in temperate forest soils at the Harvard Forest. Forest Ecol Manag 196:43–56 9. Burke RM, Cairney JWG (2002) Laccase and other polyphenoloxidases in ecto- and ericoid mycorrhizal fungi. Mycorrhiza 12:105–116 10. Carreiro MM, Sinsabaugh RL, Repert DA, Parkhurst DF (2000) Microbial enzyme shifts explain litter decay responses to simulated nitrogen deposition. Ecology 81:2359–2365 11. Cenci G, Caldini G (1997) Catechol dioxygenase expression in a Pseudomonas fluorescens strain exposed to different aromatic compounds. Appl Microbiol Biotechnol 47:306–308 12. Cerniglia CE (1992) Biodegradation of polycyclic aromatic hydrocarbons. Biodegradation 3:351–368 13. Chen DM, Bastias BA, Taylor AFS, Cairney JWG (2003) Identification of laccase-like genes in ectomycorrhizal basidiomycetes and transcriptional regulation by nitrogen in Piloderma byssinum. New Phytol 157:547–554 14. DeForest JL, Zak DR, Pregitzer KS, Burton AJ (2004) Anthropogenic NO3− deposition alters microbial community function in northern hardwood forests. Soil Sci Soc Am J 68:132–138

C. L. Lauber et al. 15. Eggert C, Temp U, Dean JDF, Eriksson KEL (1996) A fungal metabolite mediates degradation of non-phenolic lignin structures and synthetic lignin by laccase. FEBS Lett 391:144–148 16. Felsenstein J (2005) PHYLIP (Phylogeny Inference Package) version 3.6, distributed by the author. Department of Genome Sciences, University of Washington, Seattle 17. Fierer N, Jackson JA, Vilgalys R, Jackson RB (2004) Assessment of soil microbial community structure by use of taxon-specific quantitative PCR assays. Appl Environ Microbiol 71:4117–4120 18. Fog K (1988) The effect of added nitrogen on the rate of decomposition organic matter. Biol Rev 63:433–462 19. Frey SD, Knorr M, Parrent JL, Simpson RT (2004) Chronic nitrogen enrichment affects the structure and function of the soil microbial community in temperate hardwood and pine forests. Forest Ecol Manag 196:159–171 20. Gallo M, Amonette R, Lauber C, Sinsabaugh RL, Zak DR (2004) Microbial community structure and oxidative enzyme activity in nitrogen-amended north temperate forest soils. Microb Ecol 48:218–229 21. Hofmockel KS, Zak DR, Blackwood CB (2007) Does atmospheric NO3− deposition alter the abundance and activity of lignolytic fungi in forest soils? Ecosystems 10:1278–1286 22. Holmes WE, Zak DR (1999) Soil microbial control of nitrogen loss following clearcut harvest in northern hardwood ecosystems. Ecol Appl 9:202–215 23. Kanaly RA, Harayama S (2000) Biodegradation of high-molecularweigh polycyclic aromatic hydrocarbons by bacteria. J Bacteriol 182:2059–2067 24. Kellner H, Luis P, Zimdars B, Kiesel B, Buscot F (2008) Diversity of bacterial laccase-like multicopper oxidase genes in forest and grassland Cambisol soil samples. Soil Biol Biochem 40:638–648 25. Li K, Xu F, Ericksson KEL (1999) Comparison of fungal laccases and redox mediators in oxidation of a nonphenolic lignin model compound. Appl Environ Microb 65:2654–2660 26. Lozupone C, Knight R (2005) UniFrac: a new phylogenetic method for comparing microbial communities. Appl Environ Microb 71:8228–8235 27. Lozupone C, Hamady M, Knight R (2006) UniFrac—an online tool for comparing microbial community diversity in a phylogenetic context. BMC Bioinformatics 7:371 28. Luis P, Walther G, Kellner H, Martin F, Buscot F (2004) Diversity of laccase genes from basidiomycetes in a forest soil. Soil Biol Biochem 36:1025–1036 29. Luis P, Kellner H, Zimdars B, Langer U, Martin F, Buscot F (2005) Patchiness and spatial distribution of laccase genes of ectomycorrhizal and saprotrophic basdiomycetes in the upper horizons of a mixed forest cambisol. Microb Ecol 4:570–579 30. Luis P, Kellner H, Martin F, Buscot F (2005) A molecular method to evaluate basidiomycete laccase gene expression. Geoderma 128:18–27 31. Myers RT, Zak DR, White DC, Peacock A (2001) Landscapelevel patterns of microbial community composition and substrate use in upland forest ecosystems. Soil Sci Soc Am J 65:359–367 32. National Atmospheric Deposition Program (NRSP-3) (2008) NADP Program Office, Illinois State Water Survey 2204 Griffith Dr. Champaign, IL 61280 33. Rabinovich ML, Bolobova AV, Vasil’chenko LG (2004) Fungal decomposition of natural aromatic structures and xenobiotics: a review. Appl Biochem Microb 40:1–17 34. Roe B (2005) http://www.genome.ou.edu/proto.html 35. Sambrook JD, Russell W (2001) Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory Press, Plainview, NY 36. Sinsabaugh RL, Zak DR, Gallo ME, Lauber C, Amonette R (2004) Nitrogen deposition and dissolved organic carbon production in northern temperate forests. Soil Biol Biochem 36:1509–1515

Nitrogen Amendment and Laccase Gene Diversity 37. Sinsabaugh RL, Gallo ME, Lauber C, Waldrop MP, Zak DR (2005) Extracellular enzyme activities and soil organic matter dynamics for northern hardwood forests receiving simulated nitrogen deposition. Biogeochemistry 75:201–215 38. Waldrop MP, Zak DR, Sinsabaugh RL, Gallo ME, Lauber C (2004) Nitrogen deposition modifies soil carbon storage through changes in microbial enzymatic activity. Ecol Appl 14:1172–1177 39. Wallenstein MD, McNulty S, Fernandez IJ, Boggs J, Schlesinger WH (2006) Nitrogen fertilization decreases forest soil fungal and

57 bacterial biomass in three long-term experiments. Forest Ecol Manag 222:459–468 40. Yu YN, Breibart M, McNairnie P, Rowher F (2006) FastGroupII: a web-based bioinformatics platform for analyses of large 16S rDNA libraries. BMC Bioinformatics 7:57 41. Zak DR, Host GE, Pregitzer KS (1989) Regional variability in nitrogen mineralization, nitrification, and overstory biomass in northern lower Michigan. Can J Forest Res 19:1521–1526 42. Zak DR, Pregitzer KS (1990) Spatial and temporal variability of nitrogen cycling in northern lower Michigan. Forest Sci 36:367–380