Physiological Changes in White-Rot Fungi in Response to Simulated Nitrogen Deposition Are Not Readily Reversed Through Experimental Evolution

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Master's Theses and Capstones Student Scholarship

Winter 2015

PHYSIOLOGICAL CHANGES IN WHITE-ROT FUNGI IN RESPONSE TO SIMULATED NITROGEN DEPOSITION ARE NOT READILY REVERSED THROUGH EXPERIMENTAL EVOLUTION

Elizabeth Amy Landis University of New Hampshire, Durham

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PHYSIOLOGICAL CHANGES IN WHITE-ROT FUNGI IN RESPONSE TO SIMULATED NITROGEN DEPOSITION ARE NOT READILY REVERSED THROUGH EXPERIMENTAL EVOLUTION

ELIZABETH LANDIS

Biology BS, Wright State University, 2013

THESIS

Submitted to the University of New Hampshire in Partial Fulfillment of the Requirements for the Degree of

Master of Science in Microbiology

December, 2015

THESIS COMMITTEE PAGE

This thesis has been examined and approved in partial fulfillment of the requirements for the degree of

Masters of Science in Microbiology by:

Thesis Director: Serita Frey, Professor, Soil Microbial Ecology

Linda T.A. van Diepen, Assistant Professor, Ecosystem Science and Management, University of Wyoming

Louis Tisa, Professor, Microbiology

On December 14th, 2015

Original approval signatures are on file with the University of New Hampshire Graduate School.

TABLE OF CONTENTS

ACKNOWLEDGEMENTS...... iii. LIST OF TABLES...... iv. LIST OF FIGURES...... v. ABSTRACT...... vi.

CHAPTER PAGE I. INTRODUCTION...... 1 II. MATERIALS AND METHODS...... 4 Fungal isolation and selection...... 4 Experimental Design………………………………………………………………………… 6 Statistical Analysis…………………………………………………………………………... 10 III. RESULTS...... 12 IV. DISCUSSION...... 16 V. SUPPLEMENTARY INFORMATION…………………………………………………. 28

REFERENCES...... 30

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ACKNOWLEDGEMENTS:

This work was produced with help from lab members at the University of New Hampshire Linda van Diepen, Sarah Griffin, Shana Whitney, and Mel Knorr, and also lab help from Theresa

Fennel, statistical help from Rich Smith at the University of New Hampshire, support from the

Boonshoft Museum of Discovery, and encouragement from friends and family. Significant editorial contributions also came from advisors Serita Frey and Linda van Diepen. This material is based upon work supported by the National Science Foundation Graduate Research

Fellowship under Grant No 13-084.

LIST OF TABLES

PAGE Table 1. Replication and origin of isolates used in the decomposition experiments...... 20 Table 2. Experiment 1: Percent change in fungal decomposition ability and enzyme activities after long-term exposure to two levels of N addition (N50 and N150)...... 21 Table 3. Experiment 2: Significance values for pair-wise comparisons between groups identified through non-metric multidimensional scaling (Figure 4)...... 22 Table 4. Experiment 2: Indicator species analysis of extracellular enzymes measured for Stereum complicatum and Irpex lacteus using isolate origin, growth conditions, and decay environment as grouping factors ..……………………………………………………………. 23

LIST OF FIGURES

PAGE Figure 1. Experimental design ...... 24 Figure 2. Experiment 1: Mass loss for oak litter decomposed for 7 weeks by Stereum complicatum, Panellus stipticuss and Irpex lacteus isolated from control or N addition plots at the Chronic N Amendment Study at Harvard Forest………………...... 25 Figure 3. Experiment 2: Mass loss for oak litter decomposed for 7 weeks by Stereum complicatum and Irpex lacteus after experimental evolution...... 26 Figure 4. Experiment 2: Nonmetric multidimensional scaling of enzyme activities associated with Stereum complicatum and Irpex lacteus isolates showing the similarity of groups based on isolate origin, growth conditions, and decay environment. ………………………………….. 27

ABSTRACT PHYSIOLOGICAL CHANGES IN WHITE-ROT FUNGI IN RESPONSE TO SIMULATED NITROGEN DEPOSITION ARE NOT READILY REVERSED THROUGH EXPERIMENTAL EVOLUTION by

Elizabeth Landis

University of New Hampshire, December, 2015

Increases in nitrogen (N) deposition have been shown to slow decomposition of organic material including lignin, particularly in forests with high lignin content. Since basidiomycete fungi are the primary decomposers of lignin, their response to increases in available N are of interest. The Chronic Nitrogen Amendment Study at the Harvard Forest Long-Term Ecological

Research (CNAS HF-LTER) site is an experimental N gradient in Petersham, MA. Since 1988, three megaplots have received varying levels of NH4NO3: ambient-only deposition (control), 50 kg N ha-1 yr-1 (N50), or 150 kg N ha-1 yr-1 (N150). In order to examine how species of lignin- degrading basidiomycetes might evolve in response to increases in available N, isolates of

Stereum complicatum, Irpex lacteus, and Panellus stipticus were reciprocally transplanted from the megaplots at CNAS HF-LTER onto a laboratory-simulated N gradient. These isolates were tested for their ability to decompose oak leaf litter, and were also subjected to an experimental evolution to test whether changes in litter decay capacity are inducible or reversible over shorter time periods. The short-term experimental evolution failed to change litter decay rates or enzyme activity profiles. However, for two of the three species, exposure to 25+ years of N addition significantly changed their decay capacities. S. complicatum isolated from the N150 plot exhibited increased decomposition capacity and N150 isolates of I. lacteus exhibited decreased decomposition capacity compared to control isolates of the same species. These finding suggest

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that N deposition may have long-lasting effects on the decay capacities of fungi that may help to explain suppression of decomposition which occurs with increases in N deposition.

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INTRODUCTION

Fungi play important roles as mediators of soil ecosystem processes, for instance, governing rates of soil carbon (C) cycling. The rate of fungal C cycling may be altered in the face of global changes in climate and nutrient availability. Atmospheric nitrogen (N) deposition, created through fixation of N2 for fertilizer and as a byproduct of fossil fuel combustion, is projected to increase globally but heterogeneously, with some areas expected to experience up to

100 times their natural rate of N deposition by 2050 (Galloway et al., 2008). Simulated N deposition has been shown to suppress decomposition rates and activities of fungal lignin- degrading enzymes in high lignin, low quality (high C:N) environments (Fog et al., 1988;

Carreiro et al., 2000; Knorr et al., 2005; Janssens et al., 2010; Frey et al., 2014). However, the mechanism by which fungal enzyme activities are reduced in these environments is unknown.

Decomposition could be slowed due to changes in microbial community structure, transient effects on microbial physiology, non-plastic evolutionary changes to decomposer microbes, or some combination of these mechanisms.

If changes are evolutionary and non-plastic, their impact on microbial processes may be long-lasting. In order to predict how fungi might evolve in response to global change, one option is to examine physiological and genetic variation within fungal species along environmental gradients that simulate global change. A complementary approach to examining environmental variation is to subject organisms to experimental evolution, where they are exposed to a stimulus over some time frame. Experimental evolution can provide information about rates of evolutionary adaptation seen in environmental gradients and can also confirm that the

environmental stimulus in question is the cause of evolution (Van Dooerslaer et al., 2007; Bell &

Collins 2008; Lohbeck et al., 2012; Reusch et al., 2013).

In traditional experimental evolution studies, rates of evolution are measured over generations. However, concurrent asexual and sexual reproduction in fungi confounds attempts at quantifying a single generation. Therefore, evolutionary studies with fungi often use time or growth as a denominator for rates of evolution rather than generations (Ratcliff et al., 2011; Jeon et al., 2013; Voyles et al., 2014; Wartenberg et al., 2014). Research on fungal evolution has been focused on ascomycetes and some chytridiomycetes, where evolution has been shown to occur over relatively short time periods in laboratory experiments lasting weeks to months

(Ratcliff et al., 2011; Jeon et al., 2013; Voyles et al., 2014; Wartenberg et al., 2014).

Experimental evolution of basidiomycetes, however, is not well documented. Because of their role mediating decomposition processes through secretion of extracellular lignin-degrading enzymes (Floudas et al., 2012, Thevenot et al., 2010), it is critical to understand how basidiomycetes might evolve in response to global change, particularly in terms of enzyme production. In order to assess the direction, extent, and plasticity of the effects of soil N enrichment on the capacity of white-rot basidiomycetes, I utilized a microbial common garden approach.

Common-garden techniques are derived from plant ecology where they have historically served as an indirect way to measure genetic variation between populations of plants from diverse environments (Linhart & Grant, 1996). In plant and animal studies, common gardens are still widely used to measure phenotypic plasticity or the relative contributions of genetic versus environmental influences on phenotypes (Cordell et al., 1998, Nygren et al., 2008, Stillwell &

Fox, 2009). If representatives of the same species are placed in a common garden and exhibit

different phenotypes, then those differences are considered non-plastic and are a function of evolutionary differences rather than a response to environmental stimuli. In a microbial context, common-garden experiments have been used previously to measure genetic variation and plasticity of fungi along temperature and altitudinal gradients (Desprez-Loustau et al., 2010,

Stefansson et al., 2013).

Here, I utilized the common-garden approach to examine differences in decay rates among white-rot basidiomycetes isolated from a soil N gradient. My primary objectives were to determine (1) the effect of long-term simulated N deposition on the decay capacity of these fungi, (2) whether responses to N enrichment are plastic in nature, and (3) whether changes in decay capacity in response to N enrichment could be induced or reversed. I hypothesized that when placed in a common garden, fungal isolates which had been exposed to long-term simulated N deposition would exhibit lower decay rates and lower activities of lignin-degrading enzymes compared to isolates from control plots. Additionally, I predicted that N-induced changes in fungal physiology would be inducible or reversible by growing fungal isolates in control or high N environments in the laboratory.

MATERIALS AND METHODS

Fungal isolation and selection

Known lignin-degraders were cultured from soil and fruiting bodies collected from the Chronic

Nitrogen Amendment Study at the Harvard Forest Long-Term Ecological Research (CNAS HF-

LTER) site in Petersham, MA, USA (Table 1). This experiment was established in the spring of

1988 to examine ecosystem responses to simulated atmospheric N deposition (Magill et al.,

1998). Three 30 x 30 m megaplots were established in a mixed hardwood forest dominated by

Quercus rubra and Quercus velutina (red and black oak). Each megaplot receives one of three N treatments: no N addition (control, N0); 50 kg N ha-1 yr-1 (N50), representative of atmospheric N deposition levels expected by 2050 in some parts of the world (Galloway et al., 2008), or 150 kg

N ha-1 yr-1 (N150), a space for time substitution meant to push the ecosystem to N saturation

(Aber et al. 1993). The current rate of ambient N deposition at the site is ~8 kg N ha-1 per yr-1

(Ollinger et al., 1993). The N amendment plots are fertilized with an aqueous solution of

NH4NO3 applied monthly in six equal doses during the growing season (May to October).

Basidiomycete fungi from the Chronic N Addition Study were initially isolated on modified

Melin-Norkrans (MMN) media (Marx, 1969) from sporocarps found growing on decaying wood or were isolated from soil by dilution plating using methods outlined by Thorn et al. (1996)

(Table 1). A subculture was then grown on potato dextrose agar (PDA) (Sigma-Aldrich 70139,

1.5%) to create a stock culture for use in decomposition experiments. Species identity was confirmed by DNA extraction following procedures outlined by James et al. (2008). In order to easily extract DNA from mycelia, isolates were grown on cellophane placed on the surface of

PDA in a 9 cm Petri dish. Mycelium was scraped off the cellophane and ground in 2x

cetyltrimethylammonium bromide (CTAB) buffer using glass beads (3 mm diameter)

(Winnepenninckx et al., 1993). The internal transcribed spacer (ITS) region (Schoch et al.,

2012) was PCR amplified using the primer pair ITS1f and ITS4 (Gardes & Bruns, 1993, White et al., 1990) and Sanger sequenced using the ITS1f primer at the University of New Hampshire’s

Hubbard Center for Genome Studies sequencing Core Facility (Durham, New Hampshire, USA).

ITS sequences were identified using the Basic Local Alignment Search Tool (BLAST, Altschul et al. 1990) against the NCBI database.

The criteria by which species were selected for this study were that they were found commonly in both the control and N addition plots and were known lignin decomposers.

Biological replicates (details on replication can be found in table 1) were isolated from each treatment plot, with the goal of minimizing the risk of obtaining isolates of the same individual.

To that end, isolates were collected from subplots that were not adjacent to one another and that were at least 5 m apart. Of the ~200 isolates cultured, 46 belonged to species which were found across all three plots. After sequencing, three species met the above outlined criteria: Stereum complicatum, Irpex lacteus, and Panellus stipticus. Anecdotally, these three species were relatively common on decaying wood across the field plots, but they were only found in very low abundance in a sequence dataset of decomposing litter from a litterbag experiment implemented at the CNAS site from 2010-2012. Using 95% sequence similarity as a cutoff, only S. complicatum was found at less than 0.01 percent abundance in the decomposing litter of the litterbag experiment (unpublished).

Stereum spp., I. lacteus, and P. stipticus have been shown to degrade lignin (Lingle 1993,

Lee et al., 2007, Song et al., 2012, Manavalan et al., 2014) and all three sporulate sexually from basidiospores located on their fruiting bodies. The selected species are of diverse lineages, with

S. complicatum, I. lacteus, and P. stipticus belonging to orders Russulales, Polyporales, and

Agaricales, respectively. S. complicatum is found commonly growing on oak. I. lacteus grows on both hard and softwoods (Cejp, 1931). P. stipticus, of the family Mycenaceae, is well- characterized in respect to its ability to bioluminesce (Shimomura, 1989, Weitz, 2001). Although its ecology is less well-documented, it is generally found on dead hardwood (Lewis & McGraw,

1984).

Experimental design

Two laboratory experiments were conducted to address my outlined objectives. Experiment 1 was designed to address objectives 1 and 2 — to assess (1) the direction and extent of each species’ response to long term-elevated N addition in terms of decay capacity, and (2) — to measure the plasticity of these decay capacities in response to N enrichment. Plasticity here is defined as a change in fungal physiology (decay capacity) when isolates are exposed to environments with novel levels of available N compared to their home environments.

Experiment 2 was designed to address objective 3 — to determine whether changes in decay capacity observed in the field in response to simulated N enrichment could be induced or reversed experimentally.

Experiment 1: Effect of long-term N enrichment on the decay capacity of isolates.

Fungal isolates of each species collected from the three N treatments in the CNAS field experiment (Table 1) were reciprocally transplanted onto simulated “decay environments” in the laboratory—Petri dishes intended to simulate the field N environments at Harvard Forest (figure

1). More specifically, the Petri dishes contained agar with concentrations of available inorganic

N comparable to those observed in the top 10 cm of soil in each N treatment (N0, N50, or N150).

These simulated N environments consisted of 9 cm Petri dishes including a base of 30 mL agar, adjusted N levels, and Hoagland's micronutrient solution (per L of media: 2.5 mL KSO4 (5 M),

1.25 mL CaCL2 (1 M), 250 uL KH2PO4 (1 M), 250 uL MgSO4 (1 M), 1 mL EDTA-Fe (0.6557 g/100mL), 2.86 mg H3BO4 , 1.81 mg MnCl2 · 4H20, 0.22 mg ZnSO4 7 H20, 0.08 mg

CuSO4 · 5H20, 0.02 mg MoO3 (85%)). NH4NO3 was added to the agar at levels designed to match concentrations found in the organic horizon at CNAS HF-LTER: 0.18 mg mL-1 (N0), 0.45 mg mL-1 (N50), and 0.67 mg mL-1 (N150). Field levels were determined using KCl extractable

N (Harvard Forest Data Archive, http://harvardforest.fas.harvard.edu/data-archive). Agar pH was slightly but significantly (p < 0.01) different according to the levels NH4NO3 added to the media, with averages of 5.74 (N0), 5.68 (N50), and 5.64 (N150) (three independent pH measurements were taken during Petri plate preparation).

Oak leaf litter (~0.7 g) was added on top of the agar medium. Oak litter was selected because it is the dominant tree species in the field plots, representing ~85% of the total litter fall.

The litter for each decay environment was collected from litter baskets located at the corresponding N treatment plot (N0, N50, or N150). Litter was air dried, destemmed, cut to uniform size (~2 x 2 cm pieces), and weighed. All litter was sterilized by autoclaving three times at 121°C for 30 min prior to being placed in the simulated N environments. Inoculum plugs of each fungal isolate were taken from the stock culture grown on 1.5% PDA. Inoculum plugs were taken from the edge of the growth front using the blunt end of a sterile glass pipette, creating equal-sized round plugs (7 mm diameter) for use in subsequent trials. The plugs were placed on the litter in the center of the plate and the sealed Petri dishes were incubated in

darkness at 25°C and 25-30% relative humidity. After seven weeks of incubation, all litter remaining was removed from the surface of the Petri dish, well mixed, and weighed. A litter subsample (~0.5 g) was taken for enzyme analysis and the remaining litter was lyophized and weighed for determination of moisture content. Moisture content was calculated for each sample

((wet weight – dry weight) / wet weight), and an average moisture content was calculated for each species. Litter mass loss was determined by subtracting the dry weight of lyophized leaf litter from the dry weight of initial litter. Control Petri dishes, which contained leaf litter but no fungal plug, were also incubated along with each species and no mass loss was detected.

Extracellular enzyme activities were measured colorimetrically using a slurry of sodium acetate buffer solution and the decomposed leaf litter using the methods previously described by

Saiya-Cork et al. (2002) and DeForest (2009). Briefly, litter (~0.5 g) was blended for 30 seconds in 125 ml sodium acetate buffer (50 mM) at the average pH (4.7) of litter from the field environments, using a Magic Bullet (Homeland Housewares LLC). Sample homogenate (200 µl) was transferred to a 96-well microplate and 50 µl substrate was added. Six enzymes were assessed: Peroxidase (PER), phenol oxidase (OX), cellobiohydrolase (CBH), phosphatase

(PHOS), N-acetyl-ß-glucosaminidase(NAG), ɑ-glucosidase (AG), and β-d-glucosidase (BG).

PER activity was measured using the substrate 3,3',5,5'-Tetramethylbenzidine (Johnsen and

Jacobsen 2008), while OX activity was measured using azino-bis(3-ethylbenzothiazoline-6- sulphonic acid) (ABTS). Hydrolytic enzymes CBH, PHOS, NAG, AG, and BG were assayed using the methylumbelliferyl-(MUB) linked substrates ß-D-cellobioside, phosphate, N-acetyl-ß-

D-glucosaminide, and β-D-glucopyranoside, respectively. Tests for each species were conducted to determine incubation times as well as substrate concentrations to exceed enzyme saturation

(Table S1). Microplates were incubated for 15 min to 5 hours, depending on the enzyme

substrate, to achieve peak fluorescence or absorbance (Table S1). After incubation, fluorescence was measured for hydrolytic enzymes at an excitation wavelength of 360 nm and an emission wavelength of 450 nm, and absorbance was measured for oxidative enzymes at 420 nm (OX) and

450 nm (PER) on a Biotek HT plate reader (Biotek, Winooski, USA). All enzyme assays were done with sixteen replicate wells per sample and corrected for background fluorescence or absorbance of substrate (negative control). Conversion of PER (TMB substrate) and OX (ABTS substrate) was determined based on an empirically determined extinction coefficient of ε450=

-1 -1 -1 -1 59,000 M cm (Josephy et al. 1982) and ε420=36,000 M cm (Bourbonnais and Paice 1990), respectively. Final enzyme activities were calculated using formulas outlined in DeForest (2009) and expressed as µmol of substrate converted per hour per g litter dry mass (µmol h-1 g-1).

Experiment 2: Experimental evolution of isolates

Isolates were experimentally evolved by serially transfer under the simulated N conditions described above (Figure 1), except that leaf litter was not added to the Petri dishes. The Petri dishes for the experimental evolution consisted of agar (1.5 % PDA) containing three levels of N and Hoagland’s nutrient solution. For each “generation”, fungi were grown to the edge of the 9 cm Petri dishes and a 7 mm plug from the edge of growth was transferred to new Petri dishes.

This transfer occurred 10 times before the isolates were placed on decay environments prepared as described above for experiment 1, with agar and oak litter (Figure 1). After seven weeks of incubation at 25ºC and 25-30% relative humidity in darkness, litter mass loss and enzyme activity were determined as previously described for Experiment 1. Each experimental condition was replicated three times for all fungal isolates. Several replicates of P. stipticus isolates were lost due to contamination during the experimental evolution phase in experiment 2, so P. stipticus was omitted from Experiment 2.

Statistical Analyses

I assessed the effect of long-term simulated N deposition on the decay dynamics (litter mass loss and enzyme activities) of each species using two-way analysis of variance (ANOVA) with isolate origin (N0, N50, or N150) and simulated decay environment as main factors

(Experiment 1). In order to make pairwise comparisons, separate ANOVAs were conducted for

N50 vs control isolates and N150 vs control isolates. Prior to conducting ANOVAs, the data were tested for and met the assumptions of normal distribution of residuals using a Shapiro

Wilke’s test and normal distribution of variance using a Bartlett’s test. ANOVAs and tests of normality and variance were done in R version 3.0.2 (R core team, 2013).

For Experiment 2, nonmetric tests were performed to assess the impacts of 25+ years of

N addition, experimental evolution, and decay environment. These nonmetric tests included

Nonmetric Multidimensional Scaling (NMDS), Multiresponse Permutation Procedure (MRPP), and Indicator Species Analysis (ISA). NMDS was used to measure the effects of isolate origin, experimental evolution, and decay environment on enzyme patterns with a Bray-Curtis distance measure in PC-ORD version 6.0 (Mather, 1976, Kruskal, 1964) using 250 permutations with real data and random starting configurations. Dimensionality of the final solution was in accordance with the recommendation of PC-ORD and was based on least final stress. For I. lacteus, 75 iterations were used for the final solution, and a final stress of 6.257 was reached with a final instability of 0.0. For S. complicatum, the final stress was 4.334 and the final instability was 0.0 after 87 iterations.

MRPP is a nonmetric test of variance similar to a MANOVA (Mielke & Berry, 2001) and was used to determine how groups in Experiment 2 separated according to isolate origin, growth conditions, or decay environment. Because MRPP is a quantitative complement to NMDS, the same distance measure (Bray-Curtis) was employed, and analyses were done in PC-ORD v. 6.0.

ISA is typically used to compare the populations of individual species when grouped by an environmental factor (McCune & Grace, 2002). Here, an ISA was used to contrast the performance of enzymes across N treatments within my reciprocal transfer design in Experiment

2. I used this analysis to calculate both the abundance of enzyme activity within a group and the abundance relative to other groups (McCune & Grace, 2002). These two terms were multiplied and expressed as an indicator value. Only the N treatment with the highest indicator value for any given enzyme is reported. ISA was conducted using PC-ORD version 6.0, using 4,999 randomizations in a Monte Carlo test.

RESULTS

Experiment 1: Effect of long-term N enrichment on the decay capacity of fungal isolates.

Litter mass loss ranged from 24 to 60 percent and was dependent on species (S. complicatum, P. stipticus, or I. lacteus), isolate origin (the N addition treatment from which fungi were isolated), and the decay environment (the simulated conditions in which isolates were placed in the lab) (Fig. 2). Isolates exposed to the lower N level for 25 years (i.e., isolates cultured from the N50 treatment) were not significantly different in their decay capacity from the control (N0) isolates (Fig. 2a,c). In contrast, litter decay rates were significantly impacted for S. complicatum and I. lacteus isolates cultured from the highest N addition (N150) plots, with the direction of response differing between the two species. Litter decay was significantly higher

(+33.4%) for S. complicatum isolates from the N150 plot compared to control isolates (Figure

2b, Table 2). Litter decay by I. lacteus, however, was significantly lower (-16.5%) for N150 isolates compared to control isolates (Figure 2e, Table 2). There was no significant effect of isolate origin on the decay capacity of P. stipticus. For decay capacity, there was no significant interaction between isolate origin and decay environment.

Increases in decay rates for S. complicatum and decreases in decay rates for I. lacteus were accompanied by respective increases and decreases in the potential activities of lignin- degrading enzymes. S. complicatum isolated from the N addition plots (N50 and N150) showed significantly higher levels of PER (+59.8% for N50, + 52.7% for N150) and OX enzyme activities (+15.6% for N50, + 101% for N150) than those isolated from control plots. N150 isolates of I. lacteus showed a significant decrease in PER activity (-37.7%) and a nonsignificant decrease in OX activity (-13.3%). P. stipticus showed no significant change in oxidative enzyme

activity levels. Significant shifts in hydrolytic enzymes also occurred for all three species but had no clear pattern following decay rates (Table 2).

The simulated decay environment in which isolates were placed in the lab significantly influenced litter decay rates, particularly at the highest N concentration. Although none of the species showed changes in decomposition rate under the lower level of N addition (mid N), all isolates showed significantly accelerated litter decay under high N enrichment, regardless of isolate origin. That is, isolates cultured from both control (N0) and high N (N150) treatment plots showed enhanced ability to decay oak litter in the high N decay environment, with increases ranging from 41.7 to 75.9 percent (Figure 2b,d,e and Table 2). These increases in litter decay were accompanied by changes in the activities of certain enzymes. When decaying litter in this high N environment, CBH activity was increased significantly by S. complicatum and P. stipticus (Table 2). P. stipticus also showed a significant increase in NAG enzyme activity and a significant decrease in OX activity in the high N decay environment. S. complicatum had significantly increased BG and OX enzyme activities when decaying litter in a high N environment. I. lacteus had no significant changes in hydrolytic enzyme activities when decaying litter in a high N environment (Table 2). Any interactions between origin and decay environment for the enzymes?

Experiment 2: Experimental evolution of isolates

Experiment 2 was initiated to determine whether the decay capacity of isolates could be altered by exposing them to control or N-enriched conditions in the laboratory for 10 transfer generations. Results are only shown for I. lacteus and S. complicatum due to contamination issues with P. stipticus. Isolates cultured from a given field N treatment were experimentally

evolved in a novel N environment and then placed back into their “home” decay environment.

Evolution in decay capacity was tested by comparing isolates which were experimentally evolved under different levels of experimental N. For example, control N isolates of I. lacteus evolved under control conditions and subsequently decaying litter in a control environment were compared with control isolates evolved under high N conditions and subsequently decaying litter in control N environments. None of the isolates tested were affected by the experimental evolution. That is, decay capacity was not significantly altered following 10 transfer generations in the novel environment (Figure 3). Exposing isolates to experimental evolution in different N environments also did not affect enzyme activity. Nonmetric multidimensional scaling (NMDS) and complementary multiresponse permutation procedure (MRPP) of potential enzyme activities showed that the enzyme profiles of S. complicatum and I. lacteus grouped both by isolate origin and by decay environment but not by growth conditions (Figure 4, Table 3), indicating that the experimental evolution did not alter an isolate’s enzyme profile.

Nonmetric tests revealed patterns among isolates of each species across the N gradient, i.e. enzyme profiles were significantly different based on isolate origin according to the NMDS and MRPP, with S. complicatum isolates from control plots showing more variation (broader distributions) in enzyme activities than those from the N-addition plots. The reverse was shown in I. lacteus, with more variation in enzyme activities for N-isolates than control isolates (Figure

4).

The level of N addition in the decay environment also strongly influenced enzyme activity profiles. Pairwise MRPP comparisons revealed that decay environments for both S. complicatum and I. lacteus resulted in significantly distinct enzyme activity profiles (Figure 4 e,f, Table 3). For S. complicatum, enzymes did not seem to follow clear patterns, but for I.

lacteus, oxidative enzymes were associated with control N decay conditions, while hydrolytic enzymes were associated with high N decay environment.

The patterns in enzyme activities for Irpex were confirmed by indicator species analysis

(ISA), which revealed significant enzyme indicators for both isolate origin and decay environment, but not for the experimental evolution “growth conditions”. OX was an indicator of S. complicatum control isolates and also S. complicatum decaying litter in high N decay environments. BG and PHOS were indicators of S. complicatum isolated from N50 plots. For I. lacteus OX and PER enzymes were indicators of control N decay environments, while hydrolytic enzymes BG, CBH, and PHOS were indicators of the high N decay environment (Table 4).

DISCUSSION

The ability of the selected basidiomycete species to decompose litter was impacted both by long-term exposure to increased N and the presence of N in the decay environment. Chronic

(25+ years) exposure to moderate levels of N (N50 plot), meant to simulate future atmospheric N deposition rates in some parts of the world, did not influence the decay capacity of either S. complicatum or P. stipticus. This finding suggests that moderate levels of N addition, even over long timespans, do not influence the overall inherent physiological capabilities of these species.

This result is in contrast to previous observations showing that litter decay is slowed across ascomycetes in response to chronic exposure to moderate levels of N deposition in similar lab environments (unpublished), and that organic matter decay is suppressed in N50 treatment plots at the CNAS site (Frey et al., 2014). This difference in the direction of response could be attributed to the fact that in my study, isolates were cultured from wood rather than litter. There has no data collected on the retention of N in wood at the HF-CNAS plots, and as a result, it remains unclear how much N wood-decaying fungi are exposed to in the field. The finding that these fungi were not abundant in litter decomposing at HF-CNAS supports the idea that the microbial ecology of decomposing wood may be very different than that of the forest floor.

Isolates cultured from the highest N addition treatment (N150) did exhibit significant changes in decay rate after 25+ years of N addition, with S. complicatum increasing its decay rate, I. lacteus exhibiting a decrease in decay rate, and no effect for P. stipticus. The lack of interaction between the factors “isolate origin” and “decay environment” in determining decomposition rate suggests that chronic N addition has a uniform effect on isolates, regardless of the level of N in the decay environment. Differences in litter decay capacity and enzyme

activity in response to chronic N additions may represent genetic changes to these species that are not readily reversed. According to the generic interpretation of a common garden experiment, phenotypes are considered genetic if they persist in the absence of environmental differences.

High levels of N addition in the decay environment (which was designed to simulate the

N150 field treatment) significantly increased decomposition rates for the basidiomycete species I studied. This suggests that N may be stimulating production or efficiency of enzymes involved in decomposition. Indeed, we did see increases in activities for some enzymes, including CBH and

OX which may be driving this effect. It is surprising that high levels of N in the decay environment did not strongly downregulate the production of oxidative enzymes, as this is the typical pattern at simulated N experiments (van Diepen et al. 2015), and seen with basidiomycetes in response to increases in N, especially when isolates originated from low N environments (Berg & McClaugherty, 2014).

The observed differences in the direction of evolution after 25+ years of N addition between S. complicatum and I. lacteus may be due in part to changes in the degree of plasticity within these species, which were demonstrated in Experiment 2. I. lacteus increased its plasticity, or broadened its range of enzyme activity in response to environmental N after 25+ years of N150 addition, whereas, S. complicatum restricted its range. The evolution of plasticity has been linked in several studies across systems to the ability to adapt to novel environments

(Ghalambor et al., 2015, Lande, 2009, Murren et al., 2015). It is possible that plasticity was actively selected against in S. complicatum and that it was selected for in I. lacteus in response to chronic N addition.

Changes in litter decay capacity which were evolved over long time periods at Harvard

Forest were not induced or reverted over short lab incubations, as experimental evolution failed

to influence either decay rates or enzyme activity levels. Though the experimental evolution failed to definitively give a timeframe over which evolution occurs, the presence of 10 transfer generations validated the results of the simple common garden (Experiment 1). These transfers minimized any residual effect of “maternal” traits – traits influenced by the environment of the parent organism which are heritable but not genetic (Riggs & Porter, 1996). Differences found between isolates in Experiment 2 are less prone to be confounded by factors other than N, such as stress of the parent fungi during reproduction. Buffer generations such as these 10 transfers are often included in common garden experiments as a means of purging heritable but non- genetic traits (Barrett et al., 2010, Elgersma et al., 2013, Mosseau & Roff, 1995).

It is possible that the experimental evolution did not alter decay capacity because of artifacts from the experimental design. First, fungi were not evolved on Petri dishes containing litter. Since they were not actively decomposing lignin at the time of their supposed evolution, the selective pressure to utilize these compounds may have been inadvertently removed.

Second, transfers occurred over relatively short time periods and cultures were non-sporulating.

Exclusive non-sporulation is a condition which does not occur in nature for basidiomycetes and which does not promote rapid evolution. Non-sporulating fungi have been shown to evolve considerably more slowly than fungi which are allowed to sporulate (Zhang et al., 2015), presumably because sporulation spreads haploid cells whose phenotypes cannot be suppressed through heterozygosity (Verweij et al., 2009). So, although species may not have evolved in the

10-transfer time period under which they were measured, that is not to say that they are unable to evolve in short time periods in field conditions.

In summary, high levels of long-term simulated N deposition in the field were shown to affect both the decomposition capabilities and enzyme profiles of S. complicatum and I. lacteus.

Further, these changes were not reversed following 10 transfer generations in the experimental evolution experiment. These results suggest that long-term exposure to high levels of soil N enrichment can alter fungal physiology in such a way that it is not readily reversible. The use of microbial common garden experiments can provide insight into the influence of long-term simulated global change on the capacity of microorganisms to provide important ecosystem services.

Table 1. Replication and origin of isolates used in the decomposition experiments. Panellus stipticus isolates were not used in Experiment 2. Here, n is equal to the number of replicates obtained from each source. .

Table 2. Experiment 1: Percent change in fungal decomposition capacity and enzyme activities after long-term exposure to two levels of N addition (N50 and N150). A negative value indicates a decline in litter mass loss or enzyme activity with N addition relative to the control isolate, while a positive value indicates an increase. Significant differences are indicated by asterisks with * = P ≤ 0.05, ** = P ≤ 0.01 *** = P ≤ 0.001. Absence of data is indicated by “---“.

ⱶ -1 -1 -1 -1 Isolate origin abbreviations: 50 kg nitrogen ha y (N50) and 150 kg nitrogen ha y (N150) ɸ -1 Growth conditions and decay environment abbreviations: 0.45 mg nitrogen mL (mid N), and 0.67 mg nitrogen -1 mL (high N)

Table 3. Experiment 2: Significance values for pair-wise comparisons between groups identified through non-metric multidimensional scaling (Figure 4). Values were calculated using the multiresponse permutation procedure. Significant differences (P<0.05) are bolded. Statistic “T” describes separation between groups, and “A” describes within-group homogeneity.

Table 4. Experiment 2: Indicator species analysis of extracellular enzymes measured for Stereum complicatum and Irpex lacteus using isolate origin, growth conditions, and decay environment as grouping factors. Indicator value (IV) represents the degree to which the enzyme is associated with an N addition level (CN/N0, LN/N50, or HN/N150) within each experimental treatment (isolate origin, growth conditions, or decay environment). Significant differences (P<0.05) indicator values are bolded.

Figure 1: Experimental design. Isolate origin represents the experimental field treatment plot (N0, N50, or N150) from which a fungal isolate was cultured. Isolates were tested for their capacity to decay oak litter in either their “home” or “away” environment to determine whether differences in decay rates and enzyme activities were plastic or whether decay rates and enzyme activities differed due to non-plastic variation between N addition treatments. In Experiment 2, isolates were experimentally evolved under simulated “home” or “away” environments (growth conditions) before testing their decay capacity to determine whether a change in decay capacity could be induced based on the N environment in which the fungi were grown.

Figure 2. Experiment 1: Mass loss for oak litter decomposed for 7 weeks by Stereum complicatum, Panellus stipticus and Irpex lacteus isolated from control or N addition plots at the Chronic N Amendment Study at Harvard Forest (Experiment 1). No N50 isolates of I. lacteus were successfully cultured.

Figure 3. Experiment 2: Mass loss for oak litter decomposed for 7 weeks by Stereum complicatum and Irpex lacteus after experimental evolution. Isolates were experimentally evolved in a nitrogen gradient, and then placed back into their home environments. “Isolate origin/decay environment” on the x axis represents the N levels of both where they were cultured from, and the environment in which they decomposed litter. The color of the bars represents the level of N addition in the experimental evolution. Significant effects are bolded.

Figure 4. Experiment 2: Nonmetric multidimensional scaling of enzyme activities of Stereum complicatum and Irpex lacteus isolates grouped by isolate origin (a,b), experimental evolution (c,d), and decay environment (e,f). For Stereum complicatum, Axis 1 represents 60.2 percent of variance and axis 2 represents 37.7 percent of variance in the final solution; together they represent 97.8 of the variation in the matrix. For Irpex lacteus, axis 1 represents 95.2 percent of variance and axis 2 represents 3.3 percent of variance; together the axes represent 98.4 percent of variance.

Supplemental Table 1. Incubations times and substrate concentrations used for enzyme analysis. Peroxidase enzyme activity was measured using the substrate 3,3',5,5'-Tetramethylbenzidine (TMB), while oxidase activity was measured using azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS). Hydrolytic enzymes cellobiohydrolase (CBH), phosphatase (PHOS), N-acetyl-ß- glucosaminidase(NAG), ɑ-glucosidase (AG), and β-d-glucosidase (BG) were assayed using the methylumbelliferyl-(MUB) linked substrates ß-D-cellobioside, phosphate, N-acetyl-ß-D- glucosaminide, and β-D-glucopyranoside, respectively. Tests for each species were conducted to determine incubation times as well as substrate concentrations to exceed enzyme saturation, and those concentrations and incubation periods were used for enzyme assays. Absorbance assay substrates (TMB and ABTS) were used at standard concentrations.

Substrate Incubation concentration period Substrate Species (mM) (hrs) TMB S. complicatum 5.0 0.33 I. lacteus 5.0 0.33 P. stipticus 5.0 0.33 ABTS S. complicatum 3.0 0.33 I. lacteus 3.0 0.25 P. stipticus 3.0 0.33 BG S. complicatum 1.0 1.0 I. lacteus 0.20 1.0 P. stipticus 1.0 1.5 CBH S. complicatum 1.0 4.0 I. lacteus 1.0 4.0 P. stipticus 1.0 5.0 PHOS S. complicatum 4.0 0.5 I. lacteus 4.0 0.5 P. stipticus 1.0 0.5 NAG S. complicatum 1.0 3.0 I. lacteus 2.0 3.0 P. stipticus 1.0 3.0 AG S. complicatum 1.0 3.0 I. lacteus 4.0 3.0 P. stipticus 1.0 3.0

Supplemental 2. Average percent decomposition and enzyme activity levels for Experiment 1. Each reported value is an average of replicates for a given treatment. Decomposition values represent the percent of leaf litter that was decomposed during the 7 week-long incubation. All enzyme activities are reported in units of nmol h-1 g-1.

Decomp- Isolate Decay osition Species origin environment (%) BG CBH NAG PHOS ABTS TMB+H S. complicatum N0 control N 12.97 16383 5517.1 7562.5 63835 1712.4 148.55 S. complicatum N0 mid N 14.42 33077 10867.1 11812 101540 1124.1 118.72 S. complicatum N0 high N 22.67 28758 8134.5 10165 132600 3321.8 309.73 S. complicatum N50 control N 14.41 19627 4534.0 11472 79011 3099.6 417.79 S. complicatum N50 mid N 15.63 24817 8112.2 18283 10802 1721.87 200.90 S. complicatum N150 control N 16.31 18649 4405.5 14171 11438 3373.8 227.34 S. complicatum N150 high N 28.8 22522 4739.1 12386 5381 5321.3 402.63 I. lacteus N0 control N 29.5 3315.5 1074.3 3246.8 17116 55.730 973.39 I. lacteus N0 high N 37.01 3744.5 1599.9 2462.0 28219 61.980 471.16 I. lacteus N150 control N 21.89 3160.6 1508.4 2873.2 27385 31.100 517.76 I. lacteus N150 high N 34.38 3185.5 1294.9 2383.4 21398 51.340 177.42 P. stipticus N0 control N 28.71 10868 5292.1 7451.2 33689 155.37 11172 P. stipticus N0 mid N 25.95 11302 5955.3 9529.2 44775 46.700 577.44 P. stipticus N0 high N 31.95 12961 5089.6 10551 36430 140.65 295.02 P. stipticus N50 control N 21.95 7424.5 2310.1 4475.4 15271 161.96 86.050 P. stipticus N50 mid N 24.31 8545.6 2857.1 5269.7 21597 165.91 720.39 P. stipticus N150 control N 27.52 10686 5851.9 6486.1 14048 161.80 517.28 P. stipticus N150 high N 38.94 8637.9 8731.3 8131.7 18376 117.88 474.43

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