The Role and Contribution of Saprotrophic Fungi During Standing

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Master's Theses

Spring 5-2018

The Role and Contribution of Saprotrophic Fungi during Standing Litter Decomposition of Two Perennial Grass Species, Schizachyrium scoparium and Schizachyrium tenerum

Matthew Lodato University of Southern Mississippi

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Recommended Citation Lodato, Matthew, "The Role and Contribution of Saprotrophic Fungi during Standing Litter Decomposition of Two Perennial Grass Species, Schizachyrium scoparium and Schizachyrium tenerum" (2018). Master's Theses. 342. https://aquila.usm.edu/masters_theses/342

This Masters Thesis is brought to you for free and open access by The Aquila Digital Community. It has been accepted for inclusion in Master's Theses by an authorized administrator of The Aquila Digital Community. For more information, please contact [email protected]. THE ROLE AND CONTRIBUTION OF SAPROTROPHIC FUNGI DURING STANDING LITTER DECOMPOSITION OF TWO PERENNIAL GRASS SPECIES, SCHIZACHYRIUM SCOPARIUM AND SCHIZACHYRIUM TENERUM.

Matthew Baine Lodato

A Thesis Submitted to the Graduate School, the College of Science and Technology and the Department of Biological Sciences at The University of Southern Mississippi in Partial Fulfillment of the Requirements for the Degree of Master of Science

Approved by:

Dr. Kevin A. Kuehn, Committee Chair Dr. Micheal A. Davis Dr. Carl P. Qualls

______Dr. Kevin A. Kuehn Dr. Janet R. Donaldson Dr. Karen S. Coats Committee Chair Department Chair Dean of the Graduate School

May 2018

Matthew Baine Lodato

2018

Published by the Graduate School

ABSTRACT

In terrestrial ecosystems, most of the plant biomass produced enters the detrital pool, where microbial decomposers colonize, enzymatically degrade, and assimilate plant litter carbon and nutrients in amounts sufficient to bring about the decomposition of plant litter. Here, I estimated the biomass and production of fungi and microbial respiration associated with decaying Schizachyrium scoparium and Schizachyrium tenerum leaf litter, and constructed a partial organic matter budget estimating C flow into and through fungal decomposers. Significant losses in S. scoparium (57%) and S. tenerum (68%) leaf mass was observed during litter decomposition along with concomitant increases in fungal biomass, which reached a maximum of 36 and 33 mgC/g detrital C in S. scoparium and S. tenerum, respectively. Cumulative fungal production during leaf decay totaled 96 mgC/g initial detrital C in S. scoparium and 71 mgC/g initial detrital C in S. tenerum, indicating that 17 and 11% of the leaf litter C was converted into fungal biomass, respectively. Next generation sequencing (Illumina) of fungal ITS regions identified several fungal taxa associated with decaying S. scoparium and S. tenerum leaf litter, respectively, with the majority of sequences belonging to the Ascomycota

(Dothideomycetes and Sordariomycetes). These findings extend our current understanding of fungal processes in grasslands, which should be incorporated into existing models depicting major biogeochemical pathways.

ACKNOWLEDGMENTS

I would like to thank my major advisor, Dr. Kevin A. Kuehn, my committee members,

Drs. Micheal A. Davis and Dr. Carl P. Qualls, as well as Dr. Halvor M. Halvorson for their guidance and advice during this research project. In addition, I would like to extend my gratitude to Dr. Gene Saucier for allowing me access to his property and Jerrid

Boyette, Rachel Smilo and Stephanie Koury for providing assistance in both the field and laboratory. Lastly, I would like to thank my parents, Darryl and Karen Lodato, for all their support during my academic studies. This study would not have been possible without funding from the National Science Foundation (DBI 0923063) and Mississippi

Computational Biology Consortia (EPS-0556308) and the University of Southern

Mississippi

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DEDICATION

This thesis is dedicated to my family, friends, and dogs Belle and the late Jazz.

TABLE OF CONTENTS

ABSTRACT ...... ii

ACKNOWLEDGMENTS ...... iii

DEDICATION ...... iv

LIST OF TABLES ...... vii

LIST OF ILLUSTRATIONS ...... viii

CHAPTER I - INTRODUCTION ...... 1

CHAPTER II – MATERIALS AND METHODS ...... 5

Study Site and Field Procedures ...... 5

Litter Mass Loss Patterns ...... 5

Fungal Biomass and Production ...... 7

Glucosamine ...... 9

Fungal Community Composition ...... 9

Microbial Respiration ...... 10

Daily and Cumulative Fungal Production and Respiration ...... 11

Data Analyses ...... 12

CHAPTER III – RESULTS ...... 14

Litter mass loss and nutrient dynamics ...... 14

Fungal biomass and glucosamine ...... 16

Fungal production and microbial community respiration ...... 19 v

Cumulative fungal production and respiration rates ...... 19

Fungal Community Composition ...... 21

CHAPTER IV - DISCUSSION ...... 31

REFERENCES ...... 37

LIST OF TABLES

(cumulative production + respiration) by the total leaf C mass loss. Values for total leaf mass loss and fungal biomass and the means±1SD. Values for cumulative fungal production and respiration are the mean±1SD, as estimated using Monte Carlo

Simulation Analysis. Transformed variables, such as P:B ratio, turnover time, fungal growth efficiency, fungal yield coefficient and fungal contribution are the mean±1SD. . 21

vii

LIST OF ILLUSTRATIONS

Figure 1. Changes in maximum (solid line) and minimum (dashed line) (A) air temperatures, and (B) relative humidity and daily total precipitation (dark vertical bars) at the Schizachyrium grass study site during the annual study period ...... 6

Figure 2. Percent leaf litter C remaining of standing S. scoparium and S. tenerum leaves during the study period. Symbols indicate the mean±1SE ...... 14

Figure 3. Changes in (A) carbon:nitrogen and (B) carbon:phosphorus ratios in S. scoparium and S. tenerum leaf litter during standing-litter decomposition. Symbols indicate the mean±1SE...... 17

Figure 4. Patterns of (A) fungal biomass and (B) glucosamine concentrations in S. scoparium and S. tenerum leaf litter during standing-litter decomposition. Symbols indicate the mean±1SE ...... 18

Figure 5. Rates of (A) fungal production and (B) microbial respiration associated with S. scoparium and S. tenerum leaf litter during standing-litter decomposition. Symbols indicate the mean±1SE...... 20

Figure 6. Relationship between (A) cumulative fungal production and (B) cumulative microbial respiration (CO2 flux) with cumulative leaf litter carbon loss in standing S. scoparium and S. tenerum leaf litter during the study period. Symbols indicate the mean±1SE...... 22

Figure 7. Relative abundance of major fungal phylotypes identified in (A) S. scoparium and (B) S. tenerum leaf litter based on ITS rRNA gene sequence reads from Illumina

MiSeq high-throughput sequencing. Values are expressed as percentages of 174,268 sequences...... 23

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Figure 8. Non-metric multidimensional scaling (NMDS) ordinations of community dissimilarity patterns in (A) S. scoparium and (B) S. tenerum leaf litter during standing litter decomposition as determined from relative abundance based sequence metrics. .... 25

CHAPTER I - INTRODUCTION

The production and decomposition of plant matter dominates the flow of carbon and nutrients in most terrestrial and aquatic ecosystems (Swift et al. 1979; Moore et al.

2004; Hagen et al. 2012). Hence, investigating the natural fate of vascular plant matter is a critical step to our understanding of detrital pathways that are important to carbon and nutrient cycling. In ecosystems, the decomposition of plant matter involves a wide range of abiotic and biotic transformations that result in the growth and biomass production of detrital consumers, the formation of CO2 and other mineral substances via mineralization of organic matter, as well as the formation of intermediate breakdown products such as dissolved and particulate organic matter (see Gessner et al. 1999; Gessner et al. 2007;

Kuehn 2016). From a microbial perspective, the rates of these processes are strongly influenced by the types and response of microbial decomposers to the prevailing environmental decay conditions, such as temperature, water and nutrient availability, as well as the intrinsic chemical quality of plant litter substrate (e.g., nutrient concentrations).

When investigating plant litter decomposition, an important phenologic attribute to consider is the presence or absence of leaf or shoot abscission (e.g., Newell 1993;

Bärlocher 1997). In many grass and grass-like plants, abscission and collapse of plant organs (e.g., leaf blades) to the soil surface does not typically occur following senescence and death of the plant shoot. As a result, significant quantities of standing-dead plant litter can accumulate (Risser et al. 1981; Seastedt 1988; Asaeda et al. 2002), where it begins initial decomposition in an upright standing position. Several recent studies in arid and semi-arid grassland systems have documented that increases in nighttime relative

humidity and subsequent dew formation stimulate microbial decomposers and accelerate rates of litter decomposition within standing plant litter (Dirks et al. 2010; Jacobson et al.

2015; Gliksman et al. 2017; Wang et al. 2017), which underscores the episodic pulse- dynamics of microbial processes within these types of ecosystems (Collins et al. 2008;

McHugh et al. 2015). For example, Wang et al. (2017) reported that standing litter of

Cleistogenes squarrosa experienced much higher nighttime relative humidity in comparison to litter at the soil surface, resulting in greater levels of microbial biomass and activity and higher rates of standing litter decomposition in comparison to soil surface litter (see also Liu et al. 2015). Furthermore, microbial preconditioning of C. squarrosa litter during the standing dead phase also led to a more rapid rate of litter decomposition following litter collapse to the soil surface, thus increasing the efficiency of soil organic matter formation. Earlier research by Jacobson et al. (2015) reported that

Ascomycete fungi were likely a key microbial decomposer in standing litter of the Namib dune grass, Stipagrostis sabulicola. Fungal communities inhabiting standing litter responded rapidly to periods of increased water availability and were able to survive the extreme daily thermal and desiccation stress experienced in this arid landscape.

Furthermore, they also observed that fungal colonization and growth significantly decreased the C:N ratio of standing litter and that the termite detritivore, Psammotermes allocerus, showed a clear preference for litter colonized by fungi.

These recent findings are not particularly surprising to researchers investigating standing litter decay processes in other habitats such as wetlands, where strikingly similar spatial and temporal microbial decay patterns have been known for several decades. A number of studies conducted in both salt and freshwater emergent marshes have firmly

established that microbial communities, particularly fungi, are well adapted to life in the standing litter environment, where they rapidly shift their metabolism from an inactive to a fully active state when water becomes available (Gallagher et al. 1984; Newell et al.

1985; Kuehn and Suberkropp 1998; Kuehn et al. 1998; Welsch and Yavitt 2003; Kuehn et al. 2004). Additional studies also documented that a sizable fraction of the plant litter

C is channeled into and through litter inhabiting fungal decomposers, with cumulative fungal biomass production and respiratory losses, which is likely fungal (see Jacobson et al. 2015), accounting for a significant portion of the observed losses in standing litter mass (Newell et al. 1996; Gessner 2001; Kuehn et al. 2011; Su et al. 2015). Collectively, this evidence together with more recent findings from arid grassland systems suggests that fungi may play a key role in standing litter decomposition (Kuehn 2016).

Although litter decomposition in grasslands has been extensively studied at the soil surface and subsurface environment (e.g., Hossain et al. 2010; Fraser and Hockin

2013; Allison et al. 2013; Suseela et al. 2014), the patterns, controls and overall importance of decomposition processes in standing litter have yet to be fully explored.

Furthermore, standing litter decomposition within these ecosystems has rarely been examined in relation to the growth dynamics of the microbial communities that are central to carbon and nutrient cycling pathways. This is particularly true for litter- associated fungal decomposers, which are widely considered as important decomposers in terrestrial ecosystems, yet detailed knowledge of their dynamics and overall contribution to litter decomposition and other ecosystem processes is noticeably scarce

(van der Heijden et al. 2008; van der Wal et al. 2013).

In the present investigation, the decomposition and fungal decay dynamics associated with naturally standing leaves of two perennial grasses, Schizachyrium scoparium and Schyzachirum tenerum, were examined in a subtropical longleaf Pine savanna ecosystem. Here, I quantified changes in leaf mass (carbon), fungal taxa, fungal biomass and production rates, and rates of microbial respiration during leaf senescence and initial standing litter decomposition. These data were used to construct a partial organic matter decay budget estimating C flow into and through litter associated fungal decomposers within the standing litter compartment. I also examined changes in standing litter C:N and C:P contents in relation to fungal growth dynamics in order to assess the potential role of fungi in detrital nutrient cycling.

CHAPTER II – MATERIALS AND METHODS

Study Site and Field Procedures

This study was conducted in a longleaf Pine savanna ecosystem located near

Hattiesburg, Mississippi (31°22'8.32"N, 89°25'18.18"W). During each sampling period,

10 naturally standing leaves of S. scoparium and S. tenerum were randomly collected from plants in each of 4 established plots. Leaf collection began in October while leaves were living (peak growth) and continued periodically during leaf senescence and standing litter decomposition for 261 days. Collected samples were placed individually into clean zip-lock bags, placed on ice in a cooler, and immediately returned to the laboratory and processed to determine losses in leaf carbon mass, litter nutrient concentrations (nitrogen and phosphorus), litter-associated fungal biomass (ergosterol) and glucosamine concentrations, and rates of fungal production and microbial respiration (see below). Air temperatures and relative humidity were continuously monitored and recorded at 15 min intervals throughout the study period using two Onset Hobo H8 Pro series data loggers that were placed ~12 cm above ground level (Figure 1). Daily precipitation data were obtained via a permanent NOAA meteorological data station located near the study site.

Litter Mass Loss Patterns

Mass loss of standing S. scoparium and S. tenerum leaves was estimated by losses in leaf area-specific carbon mass (Gessner 2001; Kuehn et al. 2011; Su et al. 2015). Upon return to the laboratory, collected leaves were immediately scanned using a LiCor LI-

3100 Area Meter to determine total leaf surface area. After scanning, 2 leaves per plot for each species were randomly selected to determine litter-associated fungal biomass

Figure 1. Changes in maximum (solid line) and minimum (dashed line) (A) air temperatures, and (B) relative humidity and daily total precipitation

(dark vertical bars) at the Schizachyrium grass study site during the annual study period

(ergosterol), rates of fungal production, and rates of microbial respiration. The remaining

8 leaves collected per plot for each species were immediately frozen (-20˚C) and later lyophilized to dryness and weighed. Leaves for each species were then pooled together per plot and ground to 40-mesh using a Wiley mill. Subsamples of ground leaf litter were

then analyzed for litter carbon, nitrogen, and phosphorus concentrations. Additional subsamples of ground leaf litter were also used to assess fungal community structure via

Illumina MiSeq next generation sequencing and analyzed for glucosamine (chitin) concentrations as a secondary indicator molecule of fungal colonization (see below). Leaf litter carbon and nitrogen concentrations were determined using a Costech elemental CN analyzer. Phosphorus concentrations were determined using a SEAL AA3 nutrient analyzer (molybdate-ascorbic acid method) following combustion (500˚C) and hot HCl extraction and dilution of the ashed ground litter subsamples.

The initial leaf area-specific mass of S. scoparium and S. tenerum leaves was determined by dividing the leaf carbon mass at the initial sampling date when leaves were still living by the corresponding scanned leaf area. Subsequently, the percent of leaf carbon mass remaining during later sampling dates was estimated as changes in mean leaf area-specific carbon mass per plot (n= 10 leaves) relative to the initial mean leaf area-specific carbon mass. Mass loss rates (k) of S. scoparium and S. tenerum leaf blades

-kt were calculated for each plot using a negative exponential decay model (Nt =Noe ), where Nt is the percent of leaf area-specific carbon mass remaining at time (t) in days, No is the estimated initial leaf area-specific carbon mass. Mass loss rates (k) and estimated initial leaf area-specific carbon mass were then averaged across plots for both species

(mean±SD, N=4).

Fungal Biomass and Production

Living biomass and instantaneous growth rates of fungi associated within standing leaf litter were estimated from concentrations of the fungal membrane sterol ergosterol and from rates of [1-14C]acetate incorporation into ergosterol, respectively (Gessner

2005; Suberkropp and Gessner 2005). One of the two selected leaves of S. scoparium and

S. tenerum was cut into ~2-cm long sections, placed individually into sterile 20 mL glass scintillation vials containing 3.95 ml of sterile water, and allowed to hydrate for 2 hours at 15˚C. Afterwards, an aliquot of a Na[1-14C]acetate solution was added to each sample

(final concentration of 5 mmol L-1 Na[1-14C]acetate, specific activity = 48.5 MBq mmol-

1) and samples incubated in darkness for 5 h at 15˚C. Two additional vials containing leaf sections of S. scoparium and S. tenerum, respectively, were preserved with formalin (2% v:v final concentration) prior to the addition of the [1-14C]acetate radiolabel (i.e., killed control). After incubation, incorporation of [1-14C] acetate was stopped by placing sample vials on ice and immediately filtering (0.7 µm Whatman GF/F) and washing the sample litter pieces with sterile water. Filters and litter pieces were placed back into glass scintillation vials and stored frozen at -20˚C until analyzed for ergosterol.

Frozen litter pieces were lyophilized to dryness, weighed, and ergosterol extracted in alcoholic KOH (0.8% KOH in HPLC grade methanol, total extraction volume 10 ml) for

30 min at 80°C in tightly capped digestion tubes. The resultant crude extract was partitioned into n-pentane, evaporated to dryness with nitrogen gas, and dried ergosterol residues were dissolved by sonication in 1 ml of methanol. Dissolved ergosterol samples were transferred to 2 ml screw cap HPLC vials and stored at -20° C until analyzed.

Separation and analysis of ergosterol was performed using a Shimadzu high-pressure liquid chromatography (HPLC) system, with ergosterol fractions being identified and quantified based on comparison with ergosterol standards (see Kuehn et al. 2011; Su et al. 2015). Ergosterol fractions eluting from the HPLC were collected in 20 mL scintillation vials using an automated Advantec (SF-3120) fraction collector, mixed with

10 mL of scintillation fluid (Ecolume, MP Biomedicals), and radioactivity assayed by using a Beckman LS6500 Scintillation Counter. Ergosterol concentrations and incorporated radioactivity within ergosterol fractions were determined as the average of two HPLC sample injections per sample. To estimate fungal biomass in standing litter, ergosterol concentrations were converted to fungal C assuming a conversion factor of 5

µg ergosterol mg-1 fungal dry mass, and 43% C in fungal dry mass (Kuehn et al. 2011).

Rates of acetate incorporation into ergosterol were multiplied by 12.6 µg fungal biomass nmole-1 acetate incorporated to obtain a fungal growth rate (µ, % h-1) (Gessner and

Suberkropp 2005).

Glucosamine

Glucosamine (chitin) concentrations in decaying standing litter were determined following protocols described earlier by Su 2014 and Su et al. 2015. Briefly, subsamples of lyophilized, ground plant litter were initially extracted in 0.2M NaOH to deproteinate samples and convert litter-containing chitin to chitosan. The resulting ground litter pellets were washed 4 times with sterile water and the resulting chitosan hydrolyzed to individual glucosamine residues in 8M HCI at 100˚C. Glucosamine residues in hydrolyzed samples were then converted to 9-fluorenylmethylchloroformate (FMOC-Cl) derivatives and analyzed by HPLC using fluorescence detection. Glucosamine residues were identified and quantified on the basis of comparison with known FMOC derivatized glucosamine standards (Sigma Chemical).

Fungal Community Composition

Fungal community phylotypes associated with Schizachyrium scoparium and

Schizachyrium tenerum litter samples were determined on three dates (November, March

and July) during decomposition using Illumina MiSeq next generation sequencing of fungal ITS rRNA gene regions. Briefly, DNA from ground litter subsamples were extracted using a PowerSoil extraction kit (MoBio, Carlsbad, CA) following procedures outlined by the manufacturer. Sample DNA was amplified and analyzed using a dual index barcoding approach, with primers BITS (5’ ACCTGCGGARGGATCA-3’) and

B58S3 (5’GAGATCCRTTGYTRAAAGTT-3’) targeting fungal ITS rRNA regions

(Bokulich and Mills 2013). Amplification products were then sequenced using the

Illumina MiSeq platform at the Molecular and Genomics Core Facility at the University of Mississippi Medical Center (UMMC). Raw data files (FASTQ) were processed using

Mothur bioinformatics pipeline (Schloss et al. 2009) following procedures outlined in

Kozich et al. (2009). All diversity analyses were conducted using fungal phylotypes (e.g.,

OTU’s) defined by 97% sequence similarity.

Microbial Respiration

Collected plant litter samples were also used to determine rates of litter- associated microbial respiration. One of the two selected leaves of S. scoparium and S. tenerum was cut (~10-cm long subsamples), placed into sterile Petri dishes containing sterile filter paper and wetted with sterile distilled water (see Kuehn et al. 2004; Su et al. 2015). Samples were allowed to hydrate for 2 hours at ambient laboratory temperatures (~15˚C). Afterwards, rates of microbial respiration (CO2 evolution) from plant litter were monitored using a Licor LI-6400 Infrared Gas Analyzer with a coupled

LI-6400-89 Insect Respiration Chamber. All measurements were conducted in darkness.

Following respiration measurements, litter samples were stored frozen (-20˚C) and later lyophilized and weighed. Lyophilized, weighed leaves were also subsampled and

ergosterol concentrations determined as additional sample estimate of fungal biomass within collected leaf material.

Daily and Cumulative Fungal Production and Respiration

Estimates of daily fungal production and respiration during S. scoparium and S. tenerum litter decay were calculated using procedures described earlier by Kuehn et al.

(2011) and Su et al. (2015). Since the conditions for radiolabelled incorporation and respiration assays require submergence/wetting of standing litter samples, estimates of daily fungal production were corrected for diel changes in water availability as the major controlling variable affecting microbial activities in standing litter (see Kuehn and

Suberkropp 1998; Kuehn et al. 2004). Hobo Onset temperature and relative humidity loggers were deployed at the study site to continuously monitor and estimate daily changes in water availability and temperature. These data revealed daily time periods in which standing litter microbial assemblages are likely metabolically active (i.e., 100% relative humidity – dew formation). Consequently, estimates of daily fungal growth and microbial respiration rates were calculated by multiplying the hourly fungal growth or respiration rate, as determined in the laboratory, by the hours per day in which standing S. scoparium and S. tenerum litter was exposed to ~100% relative humidity. Both fungal growth and respiration rates were also temperature adjusted (assumed Q10 = 2) to reflect in situ field temperatures (see Kuehn et al. 2004; Kuehn et al. 2011). Rates of daily fungal production were calculated by multiplying the daily growth rate (µ) by the litter- associated fungal biomass (B).

Cumulative litter-associated fungal production and microbial respiration over the entire study period was estimated followed procedures described by Suberkropp et al.

(2010) and Kuehn et al. (2011). Briefly, to determine daily fungal production or respiration values for days between sampling dates the following criteria was assumed and calculated: 1) the average hourly fungal growth (µ) and microbial respiration rate and litter fungal biomass for the first half of the days during the sampling interval were assumed to be equal to the values obtained on the preceding sampling date, and the hourly fungal growth and microbial respiration rate and litter fungal biomass for the latter half of days in the sampling interval were assumed to be equal to the corresponding values observed on the next sampling date, 2) Daily fungal growth and respiration rates were then determined (as above) by multiplying the hourly fungal growth and microbial respiration rate by the specific number of hours in that day where the relative humidity was 100%. As above, fungal growth and respiration rates were temperature adjusted

(assumed Q10 = 2) to account for observed daily changes in in situ temperatures (see

Kuehn et al. 2004, Kuehn et al. 2011), 3) Estimates of daily fungal production were subsequently calculated by multiplying the daily growth rate by the litter-associated fungal biomass. This raw data set for the entire sampling period was subsequently used to estimate cumulative fungal production using a Monte Carlo Simulation (see below).

Data Analyses

Statistical analyses of the collected data was conducted using R 3.1.2 (2015 R

Foundation for Statistical Computing), with differences at the p<0.05 level being considered significant. If necessary, data were transformed prior to analysis to ensure normality and reduce heteroscedasticity. Data were analyzed using repeated measure

ANOVA’s to test the effects of time (within plots) and grass species (across plots), on changes in leaf-area specific mass, litter nutrients dynamics, fungal biomass and

production, and microbial respiration. Pearson’s correlation analysis was used to determine relationships between selected variables. Nonmetric multidimensional scaling

(NMDS) ordinations were used to visualize any community similarities or differences between fungal phylotypes detected from both grass species. Analysis of similarity

(ANOSIM) was performed to test the effect of decay stage (a priori grouping: early, middle, late) on fungal community assemblage.

Monte Carlo Simulation Analysis using Microsoft Excel PopTools add-ins was used to estimate both cumulative fungal production and microbial respiration. The raw data set of estimated daily rates of fungal production and microbial respiration over the study period was resampled with replacement to produce 10,000 sets, from which the mean±1SD and 95% confidence intervals were calculated. Variance of transformed variables, such as production:biomass ratio and fungal yield, were estimated using the delta method (Salkind 2007).

CHAPTER III – RESULTS

Litter mass loss and nutrient dynamics

Significant decreases in S. scoparium and S. tenerum leaf carbon mass were observed during senescence and standing litter decomposition (p<0.001, Table 1), with 57 and 68% of the initial leaf carbon lost over the 261 day sampling period, respectively (Figure 2).

Initial leaf area-specific carbon mass of S. scoparium and S. tenerum leaves in October

(day 0, peak living green mass) averaged 3.78±0.40 and 5.18±0.50 mgC/cm2 (mean1SE, n=4), respectively. By the end of the study period (July), leaf area-specific carbon mass of S. scoparium and S. tenerum leaves had decreased to similar values, averaging

1.63±0.07 and 1.67±1.00 mgC/cm2, respectively. The resulting mean±SD leaf litter decay rate (-k) for S. scoparium was -0.0027±0.0009 (r2 = 0.84±0.06) and

Figure 2. Percent leaf litter C remaining of standing S. scoparium and S. tenerum leaves during the study period. Symbols indicate the mean±1SE

Table1. Repeated-measures analysis of variance (ANOVA) summary table indicating the effects of time and grass species on losses in leaf-area specific mass, % nitrogen, % phosphorus, carbon:nitrogen (C:N), carbon:phosphorus (C:P), nitrogen:phosphorus (N:P) ratios, litter fungal biomass and glucosamine concentrations, and rates of fungal production and microbial respiration. Within-Subjects (plots) Across-Subjects (plots) Response Factora F-value P-valueb Factora F-value P-valueb Leaf-area specific mass T 76.52 <0.001 S 46.57 <0.001 SxT 8.39 <0.001 % Nitrogen T 9.29 <0.001 S 0.05 0.829 SxT 3.51 <0.01 % Phosphorus T 37.14 <0.001 S 5.78 0.053 SxT 11.01 <0.001 C:N T 10.48 <0.001 S 0.05 0.838 SxT 3.75 <0.01 C:P T 28.52 <0.001 S 2.78 0.146 SxT 7.87 <0.001 Fungal Biomass T 36.601 <0.001 S 22.47 <0.01 SxT 1.963 0.062 Glucosamine T 33.55 <0.001 S 11.65 0.014 SxT 1.27 0.275 Fungal Production T 46.74 <0.001 S 0.67 0.451 SxT 0.93 0.509 Microbial Respiration T 73.19 <0.001 S 11.16 0.016 SxT 5.41 <0.001 a Factors S = species (S. scoparium and S. tenerum) and T = time (days) b P values, bolded values indicate significance at p<0.05

2 -0.0051±0.0007 (r = 0.85±0.06) for S. tenerum. The estimated initial leaf masses (NO) of

S. scoparium and S. tenerum leaves were 88±11 and 96±5%, respectively. Decay rates (k) were significantly different between the Schizachyrium species (t = 8.2, p =0.0037), with

S. tenerum decaying at a much faster rate. Mass loss patterns and decay rates reported above are most likely a conservative estimate, as these values were based on only losses in leaf-area specific mass. Any additional losses in leaf mass due to fragmentation were not taken into account.

Nitrogen and phosphorus concentrations (%) of Schizachyrium leaves decreased significantly during plant senescence (p<0.001, Table 1). Initial N and P concentrations of living S. scoparium and S. tenerum leaf blades were similar, averaging 0.65±0.06 and

0.67±0.10% N and 0.034±0.002 and 0.031±0.003% P, respectively. Following senescence, N and P concentrations of S. scoparium and S. tenerum leaf blades decreased by roughly ~50 and ~30%, respectively, resulting in a corresponding significant increase in leaf litter C:N and C:P ratios (p<0.001, Table 1, Figure 3). After this decrease, leaf litter N and P concentrations remained relatively stable until the end of the study period when both litter N and P concentrations increased, leading to a decrease in litter C:N and

C:P ratios (Figure 3). No significant differences were observed in the dynamics of N and

P concentrations between S. scoparium and S. tenerum (p>0.05, Table 1).

Fungal biomass and glucosamine

Fungal biomass, as measured by ergosterol concentrations, increased significantly in

S. scoparium and S. tenerum leaves during plant senescence and standing litter decomposition (p<0.001, Table 1), with peak fungal biomass accounting for 3.6 and 3.3% of the total detrital mass, respectively (Figure 4). Patterns of fungal biomass accrual in standing leaf litter were significantly different between S. scoparium and S. tenerum

(p<0.001, Table 1), with S. tenerum accumulating more fungal biomass during the early phases of leaf decomposition. Glucosamine (chitin) concentrations followed a similar pattern as fungal biomass (ergosterol) concentrations in both S. scoparium and S. tenerum

Figure 3. Changes in (A) carbon:nitrogen and (B) carbon:phosphorus ratios in S. scoparium and S. tenerum leaf litter during standing-litter decomposition. Symbols indicate the mean±1SE.

leaf litter (r = +0.86 and +0.85, respectively, p<0.001, Pearson), with glucosamine increasing significantly during standing litter decomposition (p<0.001, Table 1, Figure 4) and exhibiting greater accrual on S. tenerum litter (p=0.014, Table 1). Changes in mean fungal biomass and glucosamine concentrations within S. scoparium and S. tenerum

Figure 4. Patterns of (A) fungal biomass and (B) glucosamine concentrations in S. scoparium and S. tenerum leaf litter during standing-litter decomposition. Symbols indicate the mean±1SE

leaves were negatively correlated with changes in mean area-specific leaf carbon mass (r

≥ -0.86 , p<0.001, Pearson), providing evidence that increases in fungal biomass and glucosamine within standing leaf litter were consistent with concomitant losses in leaf litter carbon mass.

Fungal production and microbial community respiration

Significant increases in rates of fungal production associated with S. scoparium and S. tenerum leaves were observed during the study period (p<0.001, Table 1), with a large increase in fungal production occurring shortly after leaf senescence in February (111 days) (Figure 5). Rates then declined and remained lower until increasing by the end of the study period in July. No significant differences in rates of fungal production were observed between S. scoparium and S. tenerum (p>0.05, Table 1). Corresponding rates of microbial respiration from S. scoparium and S. tenerum leaf litter, measured from post leaf senescence onward, increased significantly during the study period (p<0.001, Table

1) and followed a strikingly similar pattern to observed rates in fungal production (Figure

5). Rates of microbial respiration were significantly correlated with rates of fungal production in both S. scoparium and S. tenerum (r = +0.82 and +0.96, respectively, p<0.001, Pearson), suggesting that most of the respiratory activity from decaying litter originated from fungi. Throughout decomposition, respiration rates from S. tenerum litter were significantly greater compared to those from S. scoparium litter (P<0.001, Table 1).

Cumulative fungal production and respiration rates

When integrated over the entire study period, estimated cumulative fungal production using Monte Carlo Simulation Analysis totaled 96±6 and 71±4 mgC/g initial detrital C in

S. scoparium and S. tenerum (Table 2), which equated to a fungal yield of 17±2% and

11±1%, respectively. Corresponding estimates of cumulative microbial respiration from

S. scoparium and S. tenerum totaled 235±16 and 301±19 mgC/g initial detrital C, respectively, indicating that a significant portion of Schizachyrium leaf litter carbon is

Figure 5. Rates of (A) fungal production and (B) microbial respiration associated with S. scoparium and S. tenerum leaf litter during standing- litter decomposition. Symbols indicate the mean±1SE.

likely mineralized by microbial decomposers during the standing litter phase (Table 2). If respiratory activities are assumed to be fungal, then total fungal assimilation (fungal production + respiration) could account for 58±5 and 55±5% of the total C losses observed during standing S. scoparium and S. tenerum litter decay (Table 2), with the remaining C losses occurring through other decay processes, such as plant senescence, 20

leaching, or through the decay activities of other microbial decomposers (e.g., bacterial).

Cumulative increases in both fungal production and microbial respiration were significantly related to cumulative losses in S. scoparium and S. tenerum leaf C (Figure

6), providing compelling evidence that fungal metabolism of grass litter is an important pathway of carbon flow during the standing litter phase.

Table 2. Total leaf carbon loss, cumulative fungal production, cumulative respiration, mean fungal biomass, P/B ratio, turnover time and estimated contribution of fungal decomposers during standing leaf decay of S. scoparium and S. tenerum. Fungal growth efficiency was determined by dividing cumulative fungal production by (cumulative production+respiration) the total leaf C mass loss Fungal yield coefficient (%) = total net fungal production / total leaf mass loss x 100. The contribution of fungi to overall carbon loss from standing-dead leaf litter was determined by dividing total fungal assimilation (cumulative production + respiration) by the total leaf C mass loss. Values for total leaf mass loss and fungal biomass and the means±1SD. Values for cumulative fungal production and respiration are the mean±1SD, as estimated using Monte Carlo Simulation Analysis. Transformed variables, such as P:B ratio, turnover time, fungal growth efficiency, fungal yield coefficient and fungal contribution are the mean±1SD. ______

Parameter S. scoparium S. tenerum ______Total leaf mass loss (mgC/g initial leaf C) 569±36 677±38 Cumulative fungal production (mgC/g initial leaf C) 96±6 71±4 Cumulative respiration (mgC/g initial leaf C) 235±16 301±19 Mean fungal biomass (mgC/g initial leaf C) 18±1 22±1 P/B ratio 5.2±0.3 3.2±0.2 Turnover time (d)* 50±3 81±5 Fungal growth efficiency* 29±2 19±2 Fungal yield coefficient (%)* 17±2 11±1 Fungal contribution to overall leaf C loss (%)*‡ 58±5 55±4 *Error estimates determined using the delta method (Salkind 2007) ‡Assumes that respiratory activity is entirely due to fungal organisms

Fungal Community Composition

A total of 174,268 sequence reads were obtained from Illumina MiSeq high- throughput sequencing. Across all samples, a total of 587 fungal phylotypes were 21

detected, with most sequences attributed to the phyla Ascomycota and Basidiomycota

(Table 3). Following leaf senescence in November, unclassified ascomycetes and those classified within the Dothideomycetes (orders Capnodiales and Pleosporales) and the

Figure 7. Relative abundance of major fungal phylotypes identified in (A) S. scoparium and (B) S. tenerum leaf litter based on ITS rRNA gene

sequence reads from Illumina MiSeq high-throughput sequencing. Values are expressed as percentages of 174,268 sequences.

Sordariomycetes (orders Glomerellales and Hypocreales) were among the most abundant phylotypes detected from both S. scoparium and S. tenerum leaf litter (Figure 7). As standing leaf litter decomposition commenced, the abundance of ascomycete phylotypes decreased and the abundance of basidiomycete phylotypes in the Agaricomycetes 23

(orders Agaricales and Polyporales) increased. Based on the total number of sequence reads (174,268) across all samples, the major ascomycete phylotypes that were identified were Catenulostroma hermanusense, Alternaria sp., Colletotrichum sp., Septophoma sacchari, Toxicocladosporium strelitziae and Uwebraunia musae. Likewise, the major basidiomycete phylotypes identified from leaf litter were Mycena sp., Xenasmatella christiansenii, Marchandiomyces corallinus, Gymnopus androsaceus and Clitocybula sp.

Of the 587 phylotypes detected, 350 were represented by only 5 or less sequence reads.

In addition, many of the detected phylotypes were not taxonomically resolved (i.e., unclassified) and could only be partially assigned into a broader fungal lineage (i.e., order or family). Table 3 lists the most abundant phylotypes observed from S. scoparium and S. tenerum leaves during early (post-senescence – November), mid (March) and late (July) stages of litter decomposition

Nonmetric multidimensional scaling (NMDS) was used to ordinate sequence data from each sample to evaluate community similarity. Two dimensional NMDS ordinations based on Bray-Curtis dissimilarity scores were sufficient to account for phylotype community differences in S. scoparium and S. tenerum (stress = 0.12, r2 = 0.94), yielding a consistent pattern showing temporal changes in the fungal phylotype community during early, mid and late stages of litter decomposition (Figure 8). Analysis of similarity

(ANOSIM) confirmed that there was an overall statistically significant difference in the fungal phylotypes detected in S. scoparium and S. tenerum (R-value = 0.984, p = 0.023).

However, subsequent comparisons examining fungal phylotypes between specific dates

(November, March and July) were not statistically significant when alpha <0.05 was

Bonferroni corrected for multiple comparison tests.

Table 3. Relative abundance of fungal phylotypes identified in S. scoparium and S. tenerum leaf litter based on ITS rRNA gene sequence reads from Illumina MiSeq high-throughput sequencing. Only dominant taxa with >100 identified sequence reads are noted. % Relative Abundance S. scoparium S. tenerum November March July November March July ______Unclassified Fungi 14.3 9.5 10.1 6.6 11.4 25.4

Phylum Ascomycota Unclassified Class Unclassified Order Unclassified Family Microcyclospora quercina 1.8 0.0 0.0 1.3 0.0 0.0 Unclassified sp. 11.7 9.6 8.2 7.4 13.0 9.4 Dothideomycetes Unclassified Order Unclassified Family Septophoma sacchari 7.5* 1.7 0.0 0.0 0.0 0.0 Unclassified sp. 0.0 0.9 0.6 0.0 0.7 0.4 Botryosphaeriales Unclassified Family Camarosporium sp. 0.0 0.0 0.0 0.3 0.0 0.0 Capnodiales Unclassified Family Toxicocladosporium strelitziae 4.1 1.5 0.0 0.4 0.0 0.0 Unclassified sp. 4.1 8.6 8.1 4.8 10.0 7.3 Davidiellaceae Cladosporium grevilleae 0.0 0.0 0.0 0.3 0.0 0.0 Davidiella tassiana 0.0 0.0 0.0 0.8 0.0 0.0 Unclassified sp. 3.8* 1.1 0.0 6.3* 2.2 0.4

Mycosphaerellaceae Uwebraunia musae 4.8* 0.0 0.0 1.3 0.0 0.0 Unclassified sp. 1.5 1.3 0.5 2.1 1.7 0.5 Teratosphaeriaceae Catenulostroma hermanusense 1.9 2.7 1.9 1.9 9.9 3.2 Devriesia thermodurans 0.0 0.0 0.0 0.0 0.0 0.4 Teratosphaeria sp. 2.1 0.0 0.0 0.3 0.0 0.0 Unclassified sp. 1.8 0.7 0.7 0.9 1.2 0.9 Pleosporales Unclassified Family Unclassified sp. 4.9 15.5 6.0 3.3 7.0 4.4 Unclassified sp. 0.0 0.0 0.0 2.8 0.9 0.0 Massarinaceae Keissleriella poagena 0.0 0.7 0.0 0.0 0.0 0.4 Montagnulaceae Unclassified sp. 0.0 0.0 0.0 0.5 0.0 0.0 Pleosporaceae Alternaria sp. 2.7 2.6 0.4 5.4* 1.9 0.0 Bipolaris microstegii 0.0 0.0 0.0 0.2 0.0 0.0 Curvularia lunata 0.7 0.0 0.0 0.6 0.0 0.0 Pleospora herbarum 0.0 0.0 0.0 0.4 0.0 0.0 Unidentified sp. 0.0 0.4 0.5 0.0 0.0 0.0 Unidentified sp. 0.0 0.0 0.0 1.8 0.0 0.0 Phaeosphaeriaceae Stagonospora sp. 0.0 0.0 0.0 0.3 0.0 0.0 Unidentified sp. 0.0 0.8 0.0 1.0 0.0 0.0 Dothideales Dothioraceae Aureobasidium pullulans 1.4 0.0 0.0 0.4 0.0 0.0

Leotiomycetes Unclassified Order Unclassified Family Unclassified sp. 0.0 0.0 0.4 0.0 0.0 0.0 Helotiales Unclassified Family Unclassified sp. 0.0 0.5 0.7 0.0 0.0 1.2 Helotiaceae Hymenoscyphus sp. 0.0 0.7 0.6 0.0 0.0 0.7 Sordariomycetes Unclassified Order Unclassified Family Unclassified sp. 1.9 0.8 0.0 4.6* 0.0 0.0 Hypocreales Unclassified Family Myrothecium cinctum 0.0 0.0 0.0 0.4 0.0 0.0 Unclassified sp. A 0.0 1.2* 0.0 12.2* 3.4 0.0 Unclassified sp. B 0.0 0.0 0.6 3.0 0.9 0.0 Unclassified sp. C 0.0 0.0 0.0 2.1 0.0 0.0 Glomerellales Unclassified Family Colletotrichum sp. 3.5* 0.0 0.0 6.3* 0.7 0.0 Xylariales Amphisphaeriaceae Pestalotiopsis rhododendri 0.0 0.0 0.0 0.2 0.0 0.0 Apiosporaceae Arthrinium sp. 0.4 0.0 0.0 0.0 0.0 0.0 Unclassified Family Dinemasporium trichophoricola 0.0 0.0 0.0 1.4 0.0 0.0

Phylum Basidiomycota Unclassified Class Unclassified Order Unclassified Family Unclassified sp. 0.0 1.1 1.0* 0.7 1.0 3.2 Agaricomyetes Unclassified Order Unclassified Family Unclassified sp. 0.0 0.0 5.5* 3.2 0.0 10.5 Agaricales Unclassified Family Unclassified sp. 0.0 1.2 1.0 0.0 0.0 0.8 Boletales Rhizopogonaceae Rhizopogon occidentalis 0.0 0.0 0.0 1.2 0.0 0.0 Mycenaceae Mycena sp. A 5.6 10.6 19.0* 2.1 8.4 9.6* Mycena sp. B 0.0 0.4 0.4 0.4 0.0 0.0 Unclassified sp. 0.0 1.0 0.6 0.0 0.0 0.5 Marasmiaceae Clitocybula sp. 0.0 1.1 1.2 0.0 0.9 1.0* Gymnopus sp. A 0.0 0.8 0.0 0.0 0.0 0.4 Gymnopus androsaceus 0.0 2.1 0.4 0.0 0.8 0.6 Marasmiellus opacus 0.0 0.6 0.0 0.9 0.0 0.0 Unclassified sp. A 1.6 5.5 3.2 0.7 2.7 2.9* Atheliales Unclassified Family Athelia sp. 0.6 0.0 0.0 0.0 0.0 0.0 Unclassified 0.0 0.0 0.0 0.0 5.5 2.1

Corticiales Corticiaceae Marchandiomyces corallinus 0.0 0.0 1.4* 0.0 0.0 2.3 Marchandiomyces sp. 0.0 0.0 0.0 0.0 0.0 0.3 Hymenochaetales Hymenochaetaceae Hymenochaete sp. 0.0 0.8 0.9 0.0 0.7 1.0 Polyporales Polyporaceae Perenniporia subacida 0.0 0.0 0.0 0.3 0.0 0.0 Xenasmataceae Xenasmatella christiansenii 0.0 0.0 15.0* 0.0 0.0 2.1*

Unclassified Family Unclassified sp. 0.0 0.0 0.9 0.4 0.0 0.0 Thelephorales Thelephoraceae Tomentella sp. 0.0 1.1 1.0 0.3 0.9 0.9 Trechisporales Hydnodontaceae Luellia recondita 0.0 1.0* 0.0 0.0 0.0 0.0 Russulales Peniophoraceae Unidentified sp. 0.0 0.0 0.0 0.3 0.0 0.0 Unclassified Family Unidentified sp. 0.0 0.0 0.0 0.6 0.0 0.0

CHAPTER IV - DISCUSSION

In the present study, significant changes in Schizachyrium leaf carbon and nutrients (N & P) were observed during the study period, which concurs with prior research findings that appreciable transformation and decomposition of plant litter can occur in the aerial standing-dead phase, prior to its detachment and collapse from the parent plant. In Schizachyrium, rapid losses in leaf carbon mass and nutrients were observed during the transition of plant leaves from a living green to fully brown (dead) condition. During this plant senescence phase, fungal colonization and growth within plant leaves were noticeably low, suggesting that the initial declines in Schizachyrium leaf carbon mass were likely a result of plant translocation and reabsorption to belowground rhizomes or through leaching losses (Aesaeda et al. 2008; Tylova et al.

2008; Su et al. 2015; Wang et al. 2017). Losses in leaf carbon mass continued following senescence and were accompanied by a concomitant increase in the production and biomass accumulation of fungal decomposers and the respiration rates of the collective litter-associated microbial community. Moreover, high-throughput sequencing analysis of the fungal community associated with standing Schizachyrium leaves established the presence of a diverse assemblage of decomposer fungi, which exhibited a distinct temporal successional pattern during standing litter decomposition.

The observation of notable carbon and nutrient loss in standing Schizachyrium leaf litter along with a concomitant increase in fungal colonization and growth should not be considered surprising in Schizachyrium or any other grassland system known to possess a standing-dead litter component. The observed patterns of leaf mass loss, nutrient loss and fungal colonization in both S. scoparium and S. tenerum standing leaves

were remarkably consistent with the sequential senescence, death and fungal decay reported previously in standing litter of emergent macrophytes in both freshwater and salt marsh ecosystems. For example, recent studies by Su et al. (2015) estimated the contribution of fungal decomposers to standing litter decomposition of Typha domingensis leaves in a subtropical freshwater marsh. Significant increases in both fungal ergosterol and chitin (glucosamine) concentrations were observed in Typha leaves during senescence and standing litter decay, which revealed the rapid colonization, growth and assimilation of Typha leaf carbon by inhabitant fungal decomposers. During that study, estimated cumulative biomass production of fungi associated with T. domingensis leaf litter totaled 39 mg fungal C/g initial detrital C, indicating that 11% the observed leaf carbon lost (37%, 370 mg C/g initial detrital C) was transformed into fungal biomass

(i.e., fungal yield, see also Kuehn et al. 2011). Corresponding estimates of cumulative microbial respiration from decaying T. domingensis leaves totaled 133 mg C/g initial detrital C, indicating that a significant fraction of the standing Typha litter (~36%) was also mineralized by litter inhabiting microbial communities, most likely by fungal decomposers. When integrated over the entire study period, cumulative increases in both fungal production and microbial respiration were significantly related to cumulative losses in T. domingensis leaf carbon mass, providing evidence that a significant fraction of the plant detrital carbon is channeled into and through litter-associated fungi during standing litter decomposition.

Similar findings were observed in the present study, with S. scoparium and S. tenerum leaf litter losing 58% and 55% of their total leaf carbon during standing decomposition, respectively. During this time, rates of fungal production associated with

S. scoparium and S. tenerum leaves increased, averaging 1.52 mgC/gC/d and 1.21 mgC/gC/d, respectively, which were similar to rates of fungal production reported from standing emergent macrophyte litter in both tropical and subtropical wetlands systems

(Newell 1993; Newell et al. 1989, 1995; Kuehn et al. 2000, 2011; Su et al. 2015 see references therein). When integrated over the 261day study period, estimated rates of cumulative fungal production associated with S. scoparium and S. tenerum averaged 96 and 71 mgC/g initial leaf C, respectively, which translated into 17% and 11% of the total leaf C being transformed and converted to fungal biomass. Likewise, estimates of cumulative respiration from S. scoparium and S. tenerum leaf litter also indicated that a sizable fraction of the leaf carbon was also respired by litter-inhabiting microorganisms, again most likely by fungi. Similar to Su et al. (2015), when integrated over the entire study period, cumulative increases in both fungal production and microbial respiration were significantly related to cumulative losses in S. scoparium and S. tenerum leaf carbon.

In addition to temporal patterns in litter-associated fungal biomass and production, this study also observed a change in fungal community composition during standing litter decomposition, which significantly differed between S. scoparium and S. tenerum. During the early and mid-stages of litter decomposition, Ascomycetes were the most prevalent taxa associated with standing litter of both Schizachyrium species, likely due to the greater availability of labile compounds (Koide et al. 2005a; Gessner et al.

2010). With more advanced stages of litter decomposition towards the end of the study period, I observed a shift in fungal taxa towards a higher proportion of Basidiomycetes, possibly due to their ability to degrade more recalcitrant plant material, such as cellulose,

hemicellulose, and lignin (Lindeberg 1946; Mikola 1956; Satio 1960; Hering 1967; Tuor et al. 1995; Leonowicz et al. 1999; Miyamaoto et al. 2000; Lynd et al. 2002; Osono and

Takeda 2002; Ososno et al 2003; Boer et al. 2005). These observed patterns of fungal community succession have also been reported within collapsed litter at the soil surface

(Tokumasu 1996, 1998; Osono 2002; Virzo de Sant et al. 2002; Koide et al. 2005a,

2005b), implying that the fungal decomposers identified in standing litter may be shared between the standing and soil litter environment.

These findings indicate that paradigms of senescence, microbial colonization and decomposition of plant litter are not restricted to just standing plant litter within humid wetland ecosystems, but may be a typical decomposition process observed in standing litter within a variety of terrestrial ecosystems whereby fungi are likely a key microbial participant in litter decomposition (Newell et al. 1995; Kuehn et al. 1999; Gessner et al.

2001; Kuehn et al. 2011; Jacobson et al. 2015; Wang et al. 2017). A large proportion of grassland litter studies have focused primarily on abiotic drivers of decomposition, such as photodegradation (i.e., UV photolysis), or have only presumed but neglected to quantify the activities of microbial decomposers during standing litter decay (Dirks et al.

2010; King et al. 2012; Barnes et al. 2015; Liu et al. 2015; Austin et al. 2016). Prior research on microbial colonization and activity within standing litter in terrestrial grasslands are sparse (but see Jacobson et al. 2015; Glicksman et al. 2017; Wang et al.

2017), and to our knowledge, this is the first study to investigate standing litter decomposition in relation to microbial growth dynamics in these systems.

As in most terrestrial ecosystems, water availability is frequently identified as the critical factor influencing microbial activities in both plant litter and soils (Borken and

Matzner 2009). A growing number of studies have now established that microbial decomposers, particularly fungi, are well-adapted for survival and growth in the harsh aerial standing litter environment, where they can rapidly shift their metabolic decay activities when water becomes available (Kuehn et al. 1998, 1999, 2004; Kuehn and

Suberkropp 1998a; Welsch and Yavitt 2003). Early studies by a number of investigators

(Newell et al. 1985; Kuehn et al. 1998) observed that rates of microbial respiration from standing plant litter of emergent macrophytes increased rapidly following exposure to wetting conditions. For example, under field conditions and in the absence of precipitation, Kuehn et al. (1998) observed that rates of microbial respiration from standing Juncus effusus litter exhibited a pronounced diel periodicity, with the highest rates coinciding with nighttime increases in relative humidity and subsequent dew formation on the surfaces of standing plant litter. In contrast, rates of microbial respiration virtually ceased during the day as a result of increased daytime temperatures, litter drying and ensuing microbial desiccation stress. Additional research established that fungi inhabiting standing Juncus litter likely survive these cyclical periods of desiccation stress by actively adjusting their intracellular compatible solute concentrations in response to fluctuations in water availability (Kuehn et al. 1998). Similar patterns of microbial respiratory activity observed in terrestrial grassland systems (Dirks et al. 2010;

Jacobson et al. 2015; Glicksman et al. 2017; Wang et al. 2017) suggest that temperature- driven increases in nighttime relative humidity and dew formation on standing litter may be a key physiological strategy for fungal survival and growth in the standing litter environment, whereby fungi rapidly take advantage of even short-term periods of moisture availability to exploit detrital resources (Kuehn 2016).

During this study the metabolic and growth activities of diverse communities of saprotrophic fungi had profound impacts on the decomposition of Schizachyrium leaf litter during the standing dead phase. Over half of Schizachyrium litter carbon was assimilated and likely respired by fungi, indicating that the standing dead phase of decomposition is a major pathway of detrital carbon transformation. Furthermore, I observed significant changes in standing litter C:N and C:P molar ratios implying that fungi may also be a key player in nutrient cycling processes within grassland ecosystems.

These data highlight that fungal-mediated pathways of decomposition are not only restricted to wetlands, but likely occur in standing litter within various terrestrial ecosystems. Therefore, the data obtained from this study should be integrated into conceptual carbon and nutrient cycling models.

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