Interesting and Unusual Flux Patterns of Nitrogen and Carbon in Pinus Contorta Germinants Colonized by the Ectomycorrhizal Fungus Suillus Tomentosus

Interesting and Unusual Flux Patterns of Nitrogen and Carbon in Pinus contorta Germinants Colonized by the Ectomycorrhizal Fungus Suillus tomentosus

Joshua Michael Smith

B.Sc., The University of British Columbia, 2014

A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF

MASTER OF SCIENCE

The College of Graduate Studies

(Biology)

THE UNIVERSITY OF BRITISH COLUMBIA (Okanagan)

September 2018

Interesting and Unusual Flux Patterns of Nitrogen and Carbon in Pinus contorta Germinants Colonized by the Ectomycorrhizal Fungus Suillus tomentosus

submitted by Joshua Michael Smith in partial fulfillment of the requirements of the degree of Master of Science .

Dr. Melanie Jones, Irving K. Barber School of Arts and Sciences Supervisor

Dr. Daniel Durall, Irving K. Barber School of Arts and Sciences Supervisory Committee Member

Dr. Louise Nelson, Irving K. Barber School of Arts and Sciences Supervisory Committee Member

Dr. Peter Millard, Manaaki Whenua - Landcare Research, New Zealand Supervisory Committee Member

Dr. Jeff Curtis, Irving K. Barber School of Arts and Sciences University Examiner

Abstract

The most commonly known characteristics of the ectomycorrhizal symbiosis are translocation of soil nitrogen by fungi to plant roots, and the allocation of plant carbon to the fungi. Here, I report on observations that contradict these well-known ecophysiological pathways: movement of nitrogen from the plant toward the fungus, and the fungus behaving saprotrophically while in symbiosis. In both experiments for this thesis mycorrhizal Pinus contorta and Suillus tomentosus were used.

In the first experiment, these organisms were in low-nitrogen microcosms where some nitrogen was accessible only by hyphae. After 73 days, the hyphal nutrient medium was replaced with water or fresh nitrogen solutions. Forty-eight hours after nutrient manipulations, shoot nitrogen concentrations had dropped by 70% in microcosms where nitrogen was added to the hyphal compartment. These seedlings contained only 55% of the nitrogen present in the seed. Loss of nitrogen did not occur if water was added to the hyphal compartment or if hyphal connections to the fresh nitrogen were severed prior to nitrogen addition. Because severing of hyphae prevented loss of nitrogen I speculate that seedling nitrogen was translocated to S. tomentosus. If a similar effect occurs in the field with germinants, the nitrogen would be especially beneficial to ectomycorrhizal fungi colonizing from spores after wildfire, a scenario typical for lodgepole pine.

For the second experiment I used microcosms where the substrate was amended with soil organic carbon (SOC) that was at natural abundance or 13C- enriched. After 148 days of growth, both the CO2 respired by a combination of plant plus fungus, and that respired by S. tomentosus alone, were significantly enriched in 13C

iii over natural abundance levels. An isotope mixing model was used to determine that

35% of the carbon respired by the fungus came from the SOC. This indicates that S. tomentosus possesses the ability to behave saprotrophically while in symbiosis to supplement its carbon supply. Saprotrophy by ectomycorrhizal fungi is currently a debated topic in our field.

Lay Summary

This thesis provides evidence that lodgepole pine seedlings may have the ability to give some nitrogen to beneficial fungi that associate with their roots. It also indicates that these fungi, which are thought to get their carbon from the plant, can behave like decomposer fungi and acquire carbon from the soil. Pine seedlings may be able to donate nitrogen to the fungus to help it grow when it first colonizes the roots. This would help the plant over the long-term by kick-starting the beneficial fungi. Additionally, when the fungus needs extra carbon to acquire nutrients, survive, or because the plant is not capable of providing enough carbon to the fungus, the fungus may have the ability to supplement its carbon supply with soil-carbon. Both of these effects are not well known nor understood. These capabilities may help explain why lodgepole pine regenerates so successfully after wildfires and other disturbances.

Preface

Chapter 2 is based on a similar experiment conducted by Dr. Matthew Whiteside and

Dr. Melanie Jones with phosphorus. Dr. Whiteside, Dr. Jones and I contributed to the conception and design of this experiment. I conducted the experiment with considerable assistance from Dr. Whiteside. I also analyzed the data. Dr. Jones and I wrote a manuscript based on the material presented in Chapter 2, with editing from Dr.

Whiteside. I altered the manuscript version for use in this thesis, with editing from Dr.

Jones and the rest of my committee (Dr. Daniel Durall, Dr. Louise Nelson, and Dr. Pete

Millard). Dr. Jones and I have submitted the manuscript version of this chapter to New

Phytologist. The manuscript is under revision for a new submission.

Chapter 3 is based on an experiment originally conceived with blueberries and ericoid mycorrhizal fungi. The original experiment was designed by Dr. Gwen Grelet, Dr. Jones,

Dr. Millard, Dr. Durall, and Dr. Whiteside. With assistance from Dr. Jones, and feedback from my committee, I adapted the experiment for lodgepole pine and ectomycorrhizal fungi. I conducted the experiment and analyzed the data with assistance from Dr.

Jones. I wrote Chapter 3, with editing from Dr. Jones and the rest of my committee. A manuscript version of this chapter will be submitted for publication after I complete my thesis defense.

I composed chapters 1 and 4, with editing from Dr. Jones and the rest of my committee.

Table of Contents

Abstract ...... iii

Lay Summary ...... v

Preface ...... vi

Table of Contents ...... vii

List of Tables ...... x

List of Figures ...... xi

List of Abbreviations ...... xiii

Acknowledgements ...... xiv

Dedication ...... xv

1 Introduction ...... 1

1.1 Ectomycorrhizae ...... 2 1.1.1 Evolution, taxonomy, and morphology ...... 2 1.1.2 General function and ecology ...... 3 1.2 Nitrogen and ectomycorrhizae ...... 6 1.2.1 Nitrogen use by ectomycorrhizal fungi ...... 7 1.3 Carbon and ectomycorrhizae ...... 11 1.3.1 Relevance ...... 11 1.3.2 Ectomycorrhizal carbon physiology ...... 12 1.3.3 Definition of fungal saprotrophy ...... 13 1.3.4 Evidence for fungal saprotrophy ...... 14 1.3.5 Evidence against fungal saprotrophy ...... 17 1.4 Species used in this thesis ...... 19 1.4.1 Suillus tomentosus ...... 19 1.4.2 Pinus contorta var. latifolia ...... 20 1.5 Thesis organization, objectives and hypotheses ...... 22 1.5.1 Thesis organization ...... 22 1.5.2 Objectives and hypotheses for Chapter 2 ...... 22 1.5.3 Objectives and hypotheses for Chapter 3 ...... 24

vii

2 Nitrogen Priming: Can an Ectomycorrhizal Fungus Withdraw Nitrogen from its Plant Symbiont?...... 25

2.1 Background ...... 25 2.2 Materials and methods ...... 27 2.2.1 Overview of experiment ...... 27 2.2.2 Preparation of fungal material and seedlings ...... 28 2.2.3 Assembly of microcosms ...... 29 2.2.4 Foliage and well treatments ...... 30 2.2.5 Harvest ...... 31 2.2.6 Nitrogen content of microcosm components and field seedlings ...... 32 2.2.7 Data analysis ...... 33 2.3 Results ...... 34 2.3.1 Nitrogen concentration in seedling tissues ...... 34 2.3.2 Nitrogen concentration in naturally-occurring field seedlings ...... 44 2.4 Discussion ...... 45 3 The Saprotrophic Potential of a Mycorrhizal Fungus in Symbiosis ...... 49

3.1 Background ...... 49 3.2 Methods ...... 51 3.2.1 SOC creation ...... 52 3.2.2 Microcosm construction ...... 53 3.2.2 Preparation and incorporation of organisms ...... 55 3.2.3 Growth of seedlings ...... 56

3.2.4 Analysis of respired CO2 ...... 57 3.2.5 Harvest ...... 58 3.2.6 Statistical analyses ...... 59 3.2.7 Isotope mixing model ...... 60 3.3 Results ...... 61 3.3.1 Contamination ...... 61 3.3.2 Total respiration ...... 62 3.3.3 Fungal respiration ...... 63 3.4 Discussion ...... 66 4 Conclusion ...... 73

viii

4.1 Hypothesis summary and significance ...... 73 4.1.1 Chapter 2 ...... 73 4.1.2 Chapter 3 ...... 75 4.2 Strengths and limitations ...... 76 4.3 Potential applications and future research directions ...... 79 4.3.1 Chapter 2 applications and future directions ...... 79 4.3.2 Chapter 3 applications and future directions ...... 81 Bibliography ...... 83

Appendices ...... 97

Appendix A: Ch. 2 supplementary information ...... 97 Appendix B: SOC extraction protocol ...... 101 Appendix C: Modified Ingestad’s pine medium ...... 102

List of Tables

Table 2.1 Summary of experimental treatments...... 28

Table 3.1 Experimental treatments and replicates...... 52

Table A.1 Replicates used for N% and total N analyses as well as mean final weight and 48 h biomass increase for shoots and roots of each treatment...... 97

Table A.2 Total N inputs into the microcosms...... 98

List of Figures

Figure 2.2 Nitrogen concentrations (% N; mean ± SD) of P. contorta roots and shoots in symbiosis with S. tomentosus after (a) 70 d growth followed by 3 d foliage

treatment (H2O or 4.7 mM NH4; one-factor ANOVA on Treatments 1 and 2;

shoots: F1,10 = 0.3252, p = 0.6; roots: F1,7 = 0.0416, p = 0.8), or an additional

2 d of exposure of S. tomentosus to fresh media in the hyphal well (H2O or

9.46 mM N from glycine or NH4Cl)...... 36

Figure 2.3 Final nitrogen concentrations of the solutions applied to the seedling foliage, at harvest...... 38

Figure 2.4 Total nitrogen (µg N ± SD) of roots and shoots of P. contorta seedlings in symbiosis with S. tomentosus after 70 d growth, then 3 d foliage treatment,

and finally 2 d of hyphal well replenishment (H2O or 9.46 mM N from glycine

or NH4Cl)...... 40

Figure 2.5 Sources of readily available N, and the final distribution of N in shoots and roots of P. contorta seedlings at the 75-day harvest...... 42

Figure 2.6 Nitrogen concentrations (% N; mean ± SD) of root and shoots of P. contorta seedlings after 70 days of growth followed by 3 days of foliage treatment

(H2O or 4.7 mM NH4) and finally 2 days of hyphal well addition (9.46 mM N

from glycine or NH4Cl)...... 43

Figure 2.7 Frequency distribution of nitrogen concentrations in (a) shoots and (b) roots of eight 16 to 20-week-old naturally regenerating P. contorta seedlings from clearcut sites near Kelowna, British Columbia, Canada (elevation 1480 m)...... 45

Figure 3.1 Diagram of microcosm used in the labelling experiment...... 55

Figure 3.2 13C atom percent values of total respiration readings. Treatment M1 and M2 were one Pinus contorta in symbiosis with Suillus tomentosus with 1% substrate SOC content at: M1) 4.1 at% 13C or M2) 1.1 at% 13C. Some microcosms contained no organisms but still had the SOC addition (E1, 4.1 at% 13C; or E2, 1.1 at% 13C)...... 62

Figure 3.3 13C atom percent values of fungal respiration, following a 24 h dark incubation after seedling removal...... 65

Figure A.1 Shoot 15N at% values...... 99

Figure A.2 Root 15N at% values...... 100

xii

List of Abbreviations

ANOVA Analysis of Variance BSA Bovine Serum Albumin IR-MS Isotope Ratio Mass Spectrometry MMN Modified Melin-Norkrans PDA Potato Dextrose Agar REML Restricted Maximum Likelihood SD Standard Deviation SEM Standard Error of the Mean SOC Soil Organic Carbon

xiii

Acknowledgements

I would like to express my deepest appreciation for Dr. Melanie Jones, my supervisor.

She gave me the encouragement and positive reinforcement needed to pursue a much more interesting and fulfilling career. She changed the trajectory of my life for the better.

I will always be grateful for her support, kindness, keen intellectual advice and the occasionally needed reprimand.

I am very grateful for my committee, Dr. Daniel Durall, Dr. Louise Nelson, and Dr. Pete

Millard. Their insight, constructive feedback, and reassurances inspired me to always try my best and stay positive throughout this project.

A special thank you goes out to my parents, Linda and Michael Smith, and Dayna

Betsill. Words cannot express how much their love, compassion, and understanding meant to me over the years. This work would not be possible without these magnificent people.

I also thank Elise Blake, Anna Epp, Andy Midwood, Sarah McDonald, Michelle

Stephenson, Matthew Whiteside and Anton Hsu for technical assistance. This work was made possible by funding from a NSERC Discovery Grant (#170627-13) and a

Discovery Accelerator Supplement to Dr. Jones. I was grateful to receive a NSERC

Canada Graduate Scholarship, a UBC Graduate Dean’s Entrance Scholarship, and several University Graduate Fellowships. Thank you to NSERC and UBC for having faith in me.

xiv

Dedication

To my parents

1 Introduction

Mycorrhizal fungi are ubiquitous, specialized fungi that form symbiotic associations with the roots of over 90% of plant species (Malloch et al., 1980). A subset of mycorrhizal fungi, ectomycorrhizal fungi, colonize almost all boreal and many northern temperate tree species (Bahr et al., 2013). To put this in perspective, the boreal forest alone covers all continents between the latitudes of 50̊ and 60˚ N and represents roughly 25% of the global forest canopy as the world’s largest terrestrial biome (Chapin & Danell, 2001). Ectomycorrhizal fungi are a significant and integral component of ecophysiological processes in these vast biomes (Finlay & Read, 1986a;

Nehls et al., 2010; Itoo & Reshi, 2013), utilizing up to 20% of the net primary production

(Hobbie & Hobbie, 2008) of 740 billion boreal and 610 billion temperate trees (Crowther et al., 2015) through an extensive plant-fungal interface, estimated to be around

320,000 m2 ha-1 (Stögman et al., 2013).

Mycorrhizal systems are critically important for study because soil respiration is a major global carbon flux which almost balances all CO2 fixation by terrestrial photosynthesis (Vargas & Allen, 2008), and is an order of magnitude greater than all anthropogenic CO2 emissions (Högberg & Read, 2006). More than 50% of CO2 released from boreal soil can come from living plant roots, their mycorrhizal fungi, and root-associated microorganisms (Högberg & Read, 2006). Consequently, a comprehensive knowledge of the ecophysiology of ectomycorrhizal fungi is required to develop policies and create climate models for the northern temperate and boreal biomes. Additionally, ectomycorrhizal forests are frequently nitrogen limited (Hunt et al.,

1988; Attiwill & Adams, 1993; Stump & Binkley, 1993), so it is of particular importance to

understand how climate change and anthropogenic factors will affect carbon and nitrogen cycling in these systems, as this will lead to changes in global fluxes of these elements (Read, 1991; Johnson et al., 1997; Högberg & Read, 2006; Nehls et al.,

2010).

1.1 Ectomycorrhizae

1.1.1 Evolution, taxonomy, and morphology

There are four main morphotypes of mycorrhizal associations: arbuscular mycorrhizae, ectomycorrhizae, ericoid mycorrhizae, and orchid mycorrhizae (Read,

1991; Read & Perez-Moreno, 2003; Talbot et al., 2008). For the purposes of this thesis I will focus only on ectomycorrhizae. Ectomycorrhizal fungi are members of two phyla,

Basidiomycota and Ascomycota (Bidartondo, 2005; Scott, 2008). Globally there are estimated to be between 7 000 and 10 000 species of ectomycorrhizal fungi, and around 8 000 ectomycorrhizal plant species (Taylor & Alexander, 2005). Species richness in ectomycorrhizal fungal communities is generally very high; for example, examination of sporocarps in an Interior Cedar-Hemlock forest found around 75 species per 100 m2 (Durall et al., 1999), and surveys of sporocarps almost certainly underestimate ectomycorrhizal fungal diversity in soil or on roots (Gardes & Brun,

1996).

Arbuscular mycorrhizal associations between members of the Glomeromycota and plants have existed for more than 400 Ma, dating back to when the first terrestrial plants arose (Remy et al., 1994; Hibbett et al., 2000; Krings et al., 2007; Field et al.,

2015). The ectomycorrhizal morphotype has convergently evolved several times, the first known occurrence being with members of Pinaceae, 180-154 Ma (Tedersoo et al.,

2010, Kohler et al., 2015). Ectomycorrhizal plants now include many members of the

Betulaceae, Dipterocarpaceae, Fagaceae, Myrtaceae, Pinaceae, Salicaceae, and others (Brundrett, 2009; Smith & Read, 2010). Ectomycorrhizal fungi do not fully penetrate root cell walls and avoid contact with the plant cell membrane (Bending &

Read, 1995). There are two distinct morphological characteristics associated with ectomycorrhizae. The first characteristic is a labyrinthine network of hyphae between root cortex cells or root epidermal cells; this is called a Hartig net (Brundrett, 2004;

Scott, 2008). The Hartig net is the location of bidirectional exchanges of nutrients and carbon between plants and their fungal symbionts (Itoo & Reshi, 2013). A positive correlation has been found between Hartig net surface area and host growth response, supporting the above statement that the Hartig net is the primary zone of nutrient transfer (Brundrett, 2004). Ectomycorrhizae also display a fungal covering of the fine- roots, called a mantle (Scott, 2008). Hyphae fan out from these mantles and spread into the soil, thereby increasing the surface area from which the root can acquire nutrients

(Finlay & Read, 1986b; Blanke et al., 2012; Bahr et al., 2013).

1.1.2 General function and ecology

Important physiological characteristics of the ectomycorrhizal association are allocation of photosynthetic carbon from the plant to the fungus, and the transfer of mineral nutrients from the fungus to its plant partner (Abuzinadah et al., 1986;

Abuzinadah & Read, 1986; Brundrett, 2004; Bidartondo, 2005). This relationship represents a significant investment by the plant host, as up to 30% of the carbon fixed by a plant (i.e., gross primary production) can be allocated to ectomycorrhizal fungi

(Jones et al., 1991; Durall et al., 1994a; Högberg et al., 2008; Corrêa et al., 2008; Nehls

et al., 2010). With this investment, ectomycorrhizal fungi excrete a wide variety of extracellular enzymes into the soil so that they are capable of acquiring nitrogen, phosphorus, carbon, and various other micro- and macro-nutrients from soil organic matter (Tibbett & Sanders, 2002; Zhao et al., 2013). Ectomycorrhizal fungi have been found to increase the carbon-to-nitrogen ratio of colonized organic material in the soil

(Bending & Read, 1995), implying that they prioritize assimilation of soil nitrogen over soil carbon (Lindahl et al., 2007).

Despite the most studied, and commonly known, aspects of these systems being carbon transfer from plants to fungi and nutrient transfer from soil to plants via the fungi

(Abuzinadah & Read, 1986; Norton et al., 1990; Perez-Moreno & Read, 2000; Schimel

& Bennett, 2004; Treseder et al., 2004; Walder et al., 2012; Bahr et al., 2013), these symbioses influence other aspects of plant physiology. Ectomycorrhizae can significantly enhance drought, heavy-metal, pathogen, and disease resistance of plants

(Parke et al., 1983; Leyval et al., 1997; Cairney & Meharg, 2003; Meharg, 2003; van der

Heijden et al., 2015; Song et al., 2015), and can enhance seedling survival (Miller et al.,

1998; Teste et al., 2009). Also, at the ecosystem level, ectomycorrhizal fungi can reduce nitrogen loss from the soil due to leaching, aggregate soil particles, and stimulate plant diversity (Finlay, 2008; van der Heijden et al., 2015).

Mycorrhizal symbiosis has been generally labelled as a mutualism, characterized as a balanced reciprocal parasitism (Hacskaylo, 1972), or an association between organisms to exchange goods and commodities that are relatively easy to procure by one symbiont, for ones that are more difficult to obtain or synthesize by that symbiont

(Brundrett, 2004; Bidartondo, 2005). However, classifying mycorrhizal associations as

mutualisms is somewhat incorrect because, in some circumstances, carbon and nutrient transfer between these mycorrhizal partners does not appear to be equivalent

(Wallander, 1995; Jones & Smith, 2004; Corrêa et al., 2011; Valtanen et al., 2014). A mutualism is defined as an interaction between species that results in reciprocal increase in the fitness of both species, whereas mycorrhizae can function at very different points along the parasitism-mutualism continuum (Johnson et al., 1997; Jones

& Smith, 2004). The associated difficulty with determining where any particular mycorrhizal association functions along this continuum is that mycorrhizal function is determined by morphological, phenological, and physiological characteristics of the symbionts (Johnson et al., 1997), as well as plant and fungal genotypes (Rúa et al.,

2018), and their biotic and abiotic environments (Jones & Smith, 2004). All of these characteristics, in combination with spatial, temporal, species, and nutrient changes, can influence which ectomycorrhizal fungal species are present on a plant and their associated position on the parasitism-mutualism continuum (Read, 1991; Jones &

Smith, 2004; Walker et al., 2014).

In summary, evidence contrary to the mycorrhizal association always functioning in a beneficial manner has led to the suggestion that mycorrhizal symbioses be classified on a structural or developmental basis and that the assumption of mutualism be eliminated (Jones & Smith, 2004). Mycorrhizae could be more succinctly defined as root-fungus associations where nutrient and carbon transfer occurs across an interface

(Brundrett, 2004). Keeping in mind that trees are seldom carbon-limited (Körner, 2003), one could argue that as long as long-term plant fitness is not affected by context- dependant fungal parasitism, there would be little selective pressure away from the

symbiosis. Given the prevalence and dominance of ectomycorrhizal associations in certain ecosystems, ectomycorrhizae must be a significantly beneficial association to both fungal and plant symbionts under certain conditions over their life span (Tibbett &

Sanders, 2002).

Nutrient and carbon fluxes within mycorrhizal plants are often interdependent, usually relying on other elements at critical steps, as triggers, or for feedback processes

(Högberg & Read, 2006; Fransson & Johansson, 2010; Valtanen et al., 2014; Hendriks et al., 2016). As this thesis involves nitrogen and carbon fluxes, a relevant example of nutrient cycling interdependence is the exchange of plant-carbon and fungal-nitrogen at the mycorrhizal interface. This exchange appears to be highly regulated and monitored by both partners; increased plant allocation of carbon to the fungus can increase

(Fellbaum et al., 2012) or decrease (Hasselquist et al., 2016) fungal nitrogen allocation to the plant, and some plants can detect when mycorrhizal fungi are behaving more parasitically and restrict their carbon allocation (Nehls et al., 2010; Kiers et al., 2011).

Although the next two sections discuss nitrogen and carbon individually, it is important to keep in mind that the cycling of these two elements is intricately intertwined.

1.2 Nitrogen and ectomycorrhizae

Soils in pine forests have been shown to be nitrogen-limited for both plant production and decomposition (Hunt et al., 1988). This means that, when examining micro- and macro-nutrients in these systems, nitrogen availability and accessibility has the largest proportional effect on plants, their associated microorganisms, and the carbon cycle (Högberg et al., 1998; Norton et al., 1990; Finlay et al., 1992; Averill et al.,

2014; Kieloaho et al., 2016). Given the global extent of ectomycorrhizal forests, it is

clear that an understanding of the dynamics of nitrogen allocation and partitioning in these systems is of importance when developing policies, practices, research projects or conservation efforts (Finlay & Read, 1986a,b; Högberg et al., 2007; Nehls et al.,

2010; Itoo & Reshi, 2013).

1.2.1 Nitrogen use by ectomycorrhizal fungi

Nitrogen is the only plant nutrient that originates from the atmosphere (Scott,

2008). Until the last decade or so, it was widely accepted that plants could only use inorganic nitrogen and that plants competed poorly against soil microbes for nitrogen, only being able to use microbial ‘leftovers’ (Schimel & Bennett, 2004). This is a byproduct of early research focusing on nitrifying bacteria and nitrogen mineralization, the process whereby microbes in the soil break down nitrogen-containing organic polymers and monomers into inorganic nitrogen (Ingestad & Kähr, 1985; Hunt et al.,

1988; Stump & Binkley, 1993; Bending & Read, 1995). Nitrogen mineralization is still an important component of the nitrogen cycle because microbes can release excess inorganic nitrogen while acquiring carbon during decomposition, and this nitrogen can be used by plants and other microbes. However, in ectomycorrhizal forests, inorganic nitrogen can comprise as little as 5% of soil nitrogen (Chalot & Brun, 1998). It is now clear that organic nitrogen, which constitutes the majority of soil nitrogen in ectomycorrhizal soils, can be taken up by both mycorrhizal and non-mycorrhizal roots when in simple forms such as amino acids and amino sugars (Abuzinadah et al., 1986;

Abuzinadah & Read, 1986; Finlay et al., 1992; Näsholm et al., 2000). Additionally, in ectomycorrhizal plants, uptake of organic nitrogen is equal to or greater than inorganic nitrogen use (Lipson & Näsholm, 2001; Read & Perez-Moreno, 2003; Kranabetter et al.,

2007).

In the currently accepted nitrogen model (Schimel & Bennett, 2004), nitrogen cycling in ectomycorrhizal soils is driven by depolymerization of nitrogen-containing polymers, including nucleic acids, proteins, and chitin, by extracellular enzymes of soil microbes, including mycorrhizal fungi (Hobbie & Hobbie, 2008). The resulting nitrogen- containing monomers may be utilized by plants and microbes (Read, 1991; Chalot &

Brun, 1998; Schimel & Bennett, 2004).

To acquire nutrients, ectomycorrhizal fungi possess suites of enzymes to mine nitrogen, phosphorus, and sulphur from soil organic matter (Luis et al., 2005; Pritsch &

Garbaye, 2011; Rineau et al., 2012; Zhao et al., 2013; Bödeker et al., 2014; Lindahl &

Tunlid, 2015). Nutrient-containing biopolymers are commonly occluded by structural components of plant, fungal, and bacterial cell walls (Hobbie & Hobbie, 2008; Pritsch &

Garbaye, 2011; Rineau et al., 2012). These structural components, such as cutin, lipids, waxes, pectin, cellulose, cellobiose, hemicellulose, monophenols, polyphenols, and lignin, can be degraded by various species of ectomycorrhizal fungi (see Read & Perez-

Moreno, 2003 for a review) using extracellular enzymes, such as phosphatases, proteinases, cellulases, chitinases, laccases, and phenol oxidases (Hobbie & Hobbie,

2008; Pritsch & Garbaye, 2011). Nitrogen-containing products including ammonium, oligopeptides, amino acids, and amino sugars, are taken up by the extramatrical mycelium (hyphae growing outside of plant tissues) via active and passive transporters

(Chalot & Brun, 1998; Hobbie & Hobbie, 2008; Lipson & Näsholm, 2001). It is important to note that the ability of an ectomycorrhizal fungus to depolymerize, take up and transport various nitrogen sources is species-, and even strain-specific (Finlay et al.,

1992; Wallenda & Read, 1999; Perez-Moreno & Read, 2000; Pena & Polle, 2014). For example, when Hebeloma cylindrosporum becomes mycorrhizal it upregulates the expression of di- and tripeptide transport genes (Garcia et al., 2016a). This upregulation modifies the transportome of the fungus to the extent that one of the transport genes becomes constitutively expressed (HcPTR2b) and, under nitrogen-limitation, an additional high-capacity uptake transporter gene (HcPTR2a) is also expressed. This mycorrhiza-only fungal expression confers a higher uptake rate of peptides over the non-mycorrhizal fungal transportome (Garcia et al., 2016a).

Once nitrogenous molecules have been taken up, some of this extramatrical nitrogen is transported to the plant-fungal interface. One mechanism to accomplish this is to incorporate the nitrogen into compounds such as spermine or polyamines, which are then transferred into fungal vacuoles. These low C/N ratio molecules are then transported via vacuolar transport from cell to cell (Morel et al., 2005). Another method of transporting nitrogen to the plant-fungal interface is to make use of source/sink concentration gradients by converting the nitrogen to an amino acid, usually arginine

(see Courty et al., 2014 for a review). After reaching the interface, which is an arginine sink, arginine is converted into glutamate by ornithine oxoacid transaminase to maintain the strength of the arginine sink at the interface; sometimes glutamate is deaminated to ammonium before being transferred to the plant (Courty et al., 2014).

Many key elements of the export of fungal nitrogen into the plant-fungal interface are still being determined (Courty et al., 2014; Garcia et al., 2016a). However, an

+ aquaporin in Laccaria bicolor, and a Saccharomyces cerevisiae NH4 export-protein homolog found in L. bicolor and Amanita muscaria appear to be likely candidates for

ammonium export at the symbiotic interface (Garcia et al., 2016a). The main forms of organic nitrogen transfer from the fungus into the interface appear to be the amino acids glycine, glutamate, and allantoin (Chalot & Brun, 1998; Garcia et al., 2016a). A metabolomic model of L. bicolor has even predicted that these three specific compounds may be synthesized specifically for transport to mycorrhizal plant partners

(Larsen et al., 2011). Some amino acid transporters have been functionally characterized in ectomycorrhizal fungi, but the characterization of oligopeptide transporters is incomplete (Garcia et al., 2016a). Spermine and other polyamines, indicated as possible vectors for fungal-nitrogen transport, have been found to be increased in mycorrhizal Pinus sylvestris root tissues, but the mechanism of polyamine export and import at the plant-fungal interface has not been identified (Niemi et al.,

2007).

Plant root cells in the interface have well-documented active and passive nitrate and ammonium transporters. They also have active protein symporters with broad substrate specificity, including for amino acids (Chalot & Brun, 1998; Wallenda & Read,

1999; Javelle et al., 2001; Lipson & Näsholm, 2001; Scott, 2008). To avoid the toxicity effects associated with ammonium, plants rapidly convert ammonium to amino acids.

This is done through the glutamine synthesis pathway, where ammonium and glutamate are converted to glutamine by glutamine synthetase in the plastids of root or leaf cells

(Scott, 2008). Glutamine is then used to produce other amino acids via transamination reactions. Amino acids can then be moved throughout the plant according to demand, and source-sink gradients (Morel et al., 2005; Scott, 2008).

It is important to note that, while ectomycorrhizal fungi translocate nitrogen to their plant partners, these fungi still need to take up nitrogen to meet their own physiological needs as well. Many factors can alter how much fungal nitrogen is exported to the plant symbiont. Both the availability of soil nitrogen and the allocation of plant carbon to ectomycorrhizal fungi can greatly influence nitrogen translocation from the fungi to their plant hosts (Perez-Moreno & Read, 2000). Different fungal species have been found to allocate varied amounts of nitrogen when provided with identical nitrogen sources and quantities (Colpaert et al., 1996). Even fungal physiology has been found to influence allocation of nitrogen to the plant. For example, increased allocation of photosynthate to a nitrogen-limited fungus can result in reduced nitrogen allocation to the plant (Hasselquist et al., 2016). This is likely because the fungus was using the increased carbon allocation to explore, colonize and metabolize new nutrients.

1.3 Carbon and ectomycorrhizae

1.3.1 Relevance

Soil contains more carbon than the atmosphere and all terrestrial vegetation combined (Averill et al., 2014; Averill, 2016). The boreal and temperate forest regions of

North America alone sequester 1.3 ± 0.5 PgC per year; this constitutes roughly 10% of the global net ecosystem productivity (Dolman et al., 2010). Ectomycorrhizal soils in particular contain 70% more carbon per unit of nitrogen than arbuscular mycorrhiza- dominated soils (Averill et al., 2014), and can sequester up to 320 kg C ha-1 yr-1 (Bahr et al., 2013). Much of this stored carbon is linked to plant roots and their associated biosphere. For example, in a boreal island chronosequence, 50 - 70% of all stored carbon was derived from roots and root-associated organisms (Clemmensen et al.,

2013). Consequently, in boreal forest soils, turnover of extraradical mycelia (hyphae growing outside of root tissues) is the main factor regulating soil biomass (Hagenbo et al., 2017). An increased understanding of the interaction between ectomycorrhizal fungi and soil carbon will therefore aid in predictions of the effects of climate change on global carbon cycles (Jakobson & Rosendahl, 1990; Bahr et al., 2013; Lehmann & Kleber,

2015).

1.3.2 Ectomycorrhizal carbon physiology

The carbon flux between plant and fungal symbionts is highly controlled and maintained by both organisms through the regulation of transporters, and conversion or sequestration of common carbon molecules into alternative forms to create concentration gradients (Hughes & Mitchell, 1995; Nehls et al., 2001; Nehls et al., 2010;

Martinez-Garcia et al., 2013). Nehls et al. (2010) reviewed carbon flux at the plant- fungal interface of ectomycorrhizae and concluded that carbon compounds excreted by the plant consist primarily of soluble sugars, such as sucrose, glucose, and fructose, but also carboxylic and amino acids. Plants have been observed to control carbon loss by active regulation of sucrose export from the phloem in roots; fungi, in turn, generate a strong carbon diffusion potential by rapid metabolism or conversion of fructose and glucose into fungal-specific sugar molecules or insoluble storage molecules (Nehls et al., 2001; Nehls et al., 2010; Martinez-Garcia et al., 2013). Most ectomycorrhizal fungi lack transporters for the disaccharide sucrose and, with the known exception of Tuber melanosporum (Ceccaroli et al., 2011), have to rely on host-plant invertases in the plant-fungal interface to hydrolyze the disaccharide into monosaccharides for carbon uptake (Nehls et al., 2001; Nehls et al., 2010; Martinez-Garcia et al., 2013; Hupperts et

al., 2017). After monosaccharides are taken up by fungal tissues the mycorrhizal fungus must rapidly process the carbon to maintain the concentration gradient and carbon-sink strength of its tissues. The monosaccharides are (i) directly shuttled into growth and maintenance metabolism, (ii) converted into insoluble long-term carbon storage, such as glycogen, or (iii) converted into fungus-specific sugars such as trehalose, and sugar alcohols such as mannitol, which are very mobile and utilized as long-distance transport molecules and short-term carbon storage (Hughes & Mitchell, 1995; Nehls et al., 2001;

Nehls et al., 2010).

1.3.3 Definition of fungal saprotrophy

The term saprotrophy is used frequently in this thesis. Recently, Lindahl and

Tunlid (2015) attempted to define the separation between fungal saprotrophs and biotrophs. They defined saprotrophs as “heterotrophic organisms that obtain a major fraction of their metabolic carbon from dead organic matter” (i.e. free-living, litter- decomposing fungi) and biotrophs as “heterotrophic organisms that obtain a major fraction of their metabolic carbon via interaction with the living cells of a host organism”

(e.g. mycorrhizal fungi and parasites). However, I would argue that this classification is not as binary as Lindahl and Tunlid (2015) suggest. Fungi are not just saprotrophs or biotrophs; many of these fungi behave differently depending on their host species, and their biotic and abiotic environments (Johnson et al., 1997; Jones & Smith, 2004; Koide et al., 2008; Cullings & Courty, 2009). In order to more accurately describe the carbon acquisition method of a fungus, modifiers such as obligate (by necessity), facultative

(occurring optionally in response to circumstances), and opportunistic (exploiting chances offered by immediate circumstances; Koide et al., 2008; Baldrian, 2009;

Cullings & Courty, 2009) can be used. Furthermore, if a fungus possesses degrees of saprotrophic and biotrophic behaviour, it should be defined with as much detail as possible, rather than being classified according to the trophic type that provides > 50% of their carbon (Koide et al., 2008). For example, in order to asexually reproduce, an arbuscular mycorrhizal fungus relies on a plant host, thus the fungus is defined as an obligate biotroph (Gui et al., 2017). By contrast, free-living, decomposer fungi are typically defined as obligate saprotrophs. Throughout this thesis I use the term

‘saprotrophy’, as meaning ‘any fraction of metabolic carbon being derived from dead organic matter’.

1.3.4 Evidence for fungal saprotrophy

Most research on mycorrhizal associations has focused on the unidirectional flow of carbon from plant to fungus to soil. However, mycorrhizal fungi excrete oxidative and hydrolytic enzymes that break down soil organic matter (Grelet et al., 2009; Lindahl &

Tunlid, 2015; Shah et al., 2016), which release soluble nutrients, including carbon, that can be taken up by the hyphae (Talbot et al., 2008; Lindahl & Tunlid, 2015). Many of these enzymes are active against ligno-cellulose components in plant cell walls because these form a major component in litter. For example, ectomycorrhizal fungi degrade cellulose via endoglucanases and assimilate the products using a cellodextrin transporter (T. melanosporum; Hacquard et al., 2013). They also express enzymes that can partially break down lignins, including highly variable, substrate-specific laccases

(Luis et al., 2005), and manganese-dependent peroxidases. The latter use a similar mechanism to white-rot saprotrophic fungi (Cortinarius glaucopus; Bödeker et al., 2014), which are typically considered the only soil fungi capable of fully degrading lignin. By

contrast, Paxillus involutus employs an oxidative enzyme that uses a similar chemical mechanism to that of brown-rot saprotrophic fungi (i.e., Fenton reactions; Rineau et al.,

2012; Tunlid et al., 2016). It is important to note that the evolution of ectomycorrhizal/biotrophic capabilities in fungi has been generally associated with an overall reduction in the genes encoding for saprotrophic plant cell wall-degrading enzymes (Kohler et al., 2015; Tunlid et al., 2016). Nevertheless, many ectomycorrhizal fungi retain a diverse set of these enzymes, despite the reduction in overall number of enzyme-encoding genes (Kohler et al., 2015).

The view that ectomycorrhizal fungi can act saprotrophically is supported by evidence that the oxidative and hydrolytic enzyme activities of ectomycorrhizal fungi are at least comparable to, or even higher than those of saprotrophic fungi. For example,

Phillips et al. (2014) found that sterilized soil organic matter subsequently colonized primarily by ectomycorrhizal fungi in the field had similar or higher enzyme activities as those colonized primarily by saprotrophs. Courty et al. (2007) found that the saprotrophic enzyme activity of Lactarius quietus was significantly increased during bud-break of their Quercus spp. hosts. The authors hypothesized that the fungus increased its enzyme activity because carbon allocation from the plant had slowed down as a result of the large amounts of carbon required for bud reactivation. Cullings et al. (2008) also hypothesized that the production of wood-degrading enzymes by ectomycorrhizal fungi would be induced by a reduction in host photosynthetic potential.

To test this, they removed 50% of the needles from trees mycorrhizal with Suillus granulatus. In response to defoliation, the excretion of endocellulase, D-glucosidase,

laccase, Mn-peroxidase, lignin peroxidase, phosphatase, and protease by the fungus increased.

It is surprising that there continues to be debate about ectomycorrhizal saprotrophy, given that it was documented almost 25 years ago. Durall et al. (1994b) noted 14C-enriched fungal respiration when 14C-labelled hemicellulose, cellulose, humic polymers, and conifer needles were provided to several different species of ectomycorrhizal fungi in symbiosis with Pseudotsuga menziesii, indicating that the fungi had taken up and metabolized the carbon. Furthermore, ectomycorrhizal fungi assimilate soil carbon when they acquire nitrogen as amino acids or amino sugars

(Abuzinadah et al., 1986; Abuzinadah & Read, 1986; Leake & Read, 1991; Talbot et al.,

2008; Baldrian, 2009). Amino acids contain a minimum of two carbon atoms, with most of the common nitrogen-containing amino acids containing four to five carbon atoms.

Labelling studies have confirmed that soil-derived, amino-acid carbon is not only assimilated by the fungus, but also translocated to plant shoots (Näsholm et al., 1998;

Jones et al., 2009). By the definition laid out in 1.3.3, this constitutes saprotrophy by ectomycorrhizal fungi (Hobbie et al., 2001; Talbot et al., 2008; Hobbie et al., 2013;

Hobbie et al., 2014).

After an extensive review of the evidence of ectomycorrhizal saprotrophy, Talbot et al. (2008) proposed three hypothetical scenarios under which mycorrhizal fungi might metabolize soil carbon: (i) ‘Plan B’: mycorrhizal fungi metabolize soil carbon as an alternative carbon source when supplies of photosynthate from the host plant are low;

(ii) ‘Coincidental Decomposer’: mycorrhizal fungi decompose soil carbon as a byproduct of mining the soil for organic nutrients; and (iii) ‘Priming Effect’: mycorrhizal

fungi decompose soil carbon when allocation of plant photosynthate to mycorrhizal roots is high, such that plant carbon ‘primes’ the growth and activity of mycorrhizal fungi.

They proposed a new model for soil carbon cycling that incorporated the decomposition of organic matter by ectomycorrhizal fungi. Several recent studies since then have supported the ‘Coincidental Decomposer’ hypothesis put forth by Talbot et al. in 2008

(Phillips et al., 2014; de Vries & Caruso, 2016; Fernandez & Kennedy, 2016; Hupperts et al., 2017).

1.3.5 Evidence against fungal saprotrophy

Despite the evidence described above, there is debate about whether the soluble carbon released by mycorrhizal fungi, while mining litter, contributes in a substantial way to the metabolic carbon of the fungus (Talbot et al., 2008; Clemmensen et al.,

2013; Lindahl & Tunlid, 2015). Those that disagree with ectomycorrhizal saprotrophy being incorporated into a new carbon cycling paradigm usually cite Treseder et al.

(2006), who found no evidence for uptake of carbon from 14C–labelled litter by ectomycorrhizal fungi associated with Quercus alba in situ. Although no saprotrophic behaviour by ectomycorrhizal fungi was observed, the authors acknowledge that the detection range of their equipment was limited to > 5% of fungal carbon being from the litter. While isotopic labeling has been used successfully in studying ectomycorrhizal systems (Hobbie, 2001; Mayor et al., 2008), the low precision of the study conducted by

Treseder et al. (2006) means that a significant contribution of mycorrhizal fungi to the ecosystem carbon cycle could have gone undetected (Cullings & Courty, 2009).

In an important paper, Baldrian (2009) argued against the conclusions of Courty et al. (2007), Cullings et al. (2008), and Talbot et al. (2008) that ectomycorrhizal fungi

could function saprotrophically. Baldrian (2009) highlighted that ectomycorrhizal fungi generally have higher proportional abundance in lower soil horizons where the availability of compounds with positive carbon energy values is very low, and that the lignolytic enzymes and cellulases produced by ectomycorrhizal fungi are much weaker than those of saprotrophs. Baldrian (2009) speculated that the increased enzyme activity observed by Courty et al. (2007) and Cullings et al. (2008) was because, during high carbon-demand/low carbon-supply periods, ectomycorrhizal root tissues begin to die, and the ectomycorrhizal fungi begin to break down these root tissues to fuel new hyphal growth through soil. He was also highly critical of the enzyme assay methods used in Courty et al. (2007) and Cullings et al. (2008). In response, Cullings and Courty

(2009) pointed out that many of Baldrian’s arguments still indicated a saprotrophic potential for ectomycorrhizal fungi. Specifically, ‘weaker’ enzymes are still functional and breakdown of senescent root tissues is still saprotrophy. Ultimately, they posed the question: ‘How much breakdown is significant?’ and pointed out that even if 1% of the carbon supply of an ectomycorrhizal fungus came from soil organic matter, it would represent millions of tons of soil carbon released as carbon dioxide via respiration. In conclusion, Cullings and Courty (2009) state that although the data are still preliminary and often conflicting, the abundance of recent enzymatic data cannot be ignored and must be pursued to shift the old paradigms in our understanding of mycorrhizal fungi.

Clearly the topic of mycorrhizal saprotrophy deserves to be studied in more detail, and at the simplest of scales, to provide evidence to help settle this debate.

1.4 Species used in this thesis

1.4.1 Suillus tomentosus

Suillus is an ectomycorrhizal genus, belonging to the Class Agaricomycetes within the Phylum Basidiomycota; it contains approximately 100 species from all over the globe. These species are widely distributed and can be very similar. Suillus spp. are well-documented ectomycorrhizal symbionts of pine, with Suillus tomentosus and its closely-related, European sister-species S. variegatus, occurring with Pinus contorta

(Douglas) and other two-needle pines (Dahlberg & Finlay, 1999; Durall et al., 1999;

Kranabetter et al., 2006; Paul et al., 2007). Suillus spp. have been observed to form roughly 2% of the ectomycorrhizae identified on two-year-old P. contorta seedlings in an

Interior Cedar-Hemlock forest study in British Columbia (Durall et al., 1999), and Suillus tomentosus Singer constituted a disproportionally large amount of sporocarp, or fruiting body, production in conifer forests in North America (Dahlberg & Finlay, 1999).

Sporocarps of Suillus spp. can have up to 40% higher nitrogen concentrations than sporocarps of other ectomycorrhizal fungi (Kranabetter et al., 2006), suggesting that these species are especially effective in acquiring nitrogen, even in nitrogen-limited forests. Additionally, S. tomentosus has been implicated as harboring nitrogenase- producing bacteria in P. contorta tubercules (Paul et al., 2007). These bacteria may be essential for the survival of P. contorta in nitrogen-deficient soils (Chapman & Paul,

2012). For both experiments in this thesis I chose S. tomentosus because of its common occurrence on lodgepole pine seedlings, and because the Jones lab had a local isolate in culture.

1.4.2 Pinus contorta var. latifolia

Pinus contorta is a member of the Pinaceae, commonly found in the coastal, montane, subalpine, and boreal forests of western North America, spanning 33˚ of latitude and 3900 m of elevation (Rehfeldt et al., 1999; Despain, 2001). Pinus contorta

Dougl. var. latifolia Engelm. is the most extensive subspecies of lodgepole pine, ranging from New Mexico, USA to the Yukon Territory, Canada (Despain, 2001). Together with its sister-species, jack pine (Pinus banksiana), P. contorta has dominated broad areas of montane and boreal forests in northern and western North America since the

Pleistocene (Despain, 2001). Pinus contorta is capable of growing in nitrogen-poor environments, and is a shade-intolerant, early successional species (Miller et al., 1998;

Despain, 2001). Lodgepole pines produce serotinous or non-serotinous cones, depending on their genotype; it is common to find both genotypes in a single stand

(Anderson, 2003). Serotinous cones require heat from forest fires to open, so while lodgepole pine is not reliant on fire for dispersal, it regenerates very effectively after a low-medium strength fire (Miller et al., 1998; Anderson, 2003). A laboratory study of growth rates of several coniferous tree species along a nitrogen addition gradient found that P. contorta had a relatively fast growth rate and had the highest nitrogen productivity, or growth rate per unit of nitrogen (Ingestad & Kähr, 1985). Because of this rapid, whole-plant growth rate, even on poor soils, P. contorta is a popular tree species for commercial reforestation; it is planted on 53% of forested areas in British Columbia

(3.8 million ha; Reid et al., 2016). Studying P. contorta is of importance in British

Columbia due to our reliance on pine forests for lumber and its significant contribution to our resource-based economy. The forestry and logging sector contributes $1.65 billion

in GDP to British Columbia’s economy (2012 statistics, Corbett et al., 2016), and the forest-related goods sector produced more than $16 billion of revenue in 2012 (Corbett et al., 2016).

Because it is easy to cultivate, P. contorta is commonly used as a host plant when studying ectomycorrhizae. Classic studies involving ectomycorrhizal P. contorta have examined most aspects of the symbiosis, from the utilization of proteins as an organic nitrogen source (Abuzinadah et al., 1986; Abuzinadah & Read, 1986) and contrasting the nitrogen-scavenging capabilities of ectomycorrhizal fungi when in pure culture vs symbiotic (Finlay et al., 1992), to tracing carbon flow in the rhizosphere of ectomycorrhizal seedlings (Norton et al., 1990) and determining how forestry practices affect ectomycorrhizal community diversity (Jones et al., 2012). An examination of ectomycorrhizal morphotypes on 133 two-year-old P. contorta seedlings found a mean of around six ectomycorrhizal fungi associated with each seedling (Durall et al., 1999), and a genetic study of the ectomycorrhizal community, using single-core soil samples from twenty-six P. contorta trees in Southern Oregon identified 125 distinct operational taxonomic units, indicating up to 125 species were found from a relatively small sample size (Garcia et al., 2016b). With the information provided in this, and 1.4.1, I feel that

Suillus tomentosus and Pinus contorta var. latifolia ectomycorrhizae are a relevant and realistic symbiosis with which to examine host-fungus carbon and nitrogen dynamics for ectomycorrhizal forest ecosystems of British Columbia.

1.5 Thesis organization, objectives and hypotheses

1.5.1 Thesis organization

This thesis is organized into four chapters. This first chapter covers the background to the research questions and lays out the objectives of the research. The next two chapters describe experiments on nitrogen and carbon nutrition in the mycorrhizal symbiosis between Pinus contorta and Suillus tomentosus. Specifically,

Chapter 2 builds on an experiment initiated during my honours program. The objective of my honours research project was to determine if differential allocation of nitrogen occurred within a mycorrhizal network if the plant hosts had differing nitrogen statuses.

The data from that experiment arrived too late to be included in my honours thesis.

Furthermore, when the data were analyzed, they revealed very little effect of plant nitrogen status on distribution of nitrogen from the fungus to the plant. In Chapter 2, the results from that experiment were investigated for evidence that, contrary to expectations, nitrogen moved from pine seedlings to the mycorrhizal fungus. I performed additional analyses to construct a total nitrogen budget for the experimental system as part of my MSc research. Chapter 3 presents an experiment to test for saprotrophic behavior in an ectomycorrhizal fungus when in symbiosis. Chapter 4 integrates material from the two experimental chapters and discusses future directions for this kind of research.

1.5.2 Objectives and hypotheses for Chapter 2

With over 25% of the global forest canopy (Chapin & Danell, 2001) growing in nitrogen-limited soil (Hunt et al., 1988; Attiwill & Adams, 1993; Stump & Binkley, 1993), it is important to examine nitrogen flux in ectomycorrhizal plants. The objective of the

first experiment of my thesis was to determine whether plant nitrogen status, which was altered via foliar nitrogen application, or exposure of the hyphae to an altered nutrient environment, affected the amount and proportion of nitrogen movement between S. tomentosus and two P. contorta seedlings in microcosms containing the pine seedlings associated with the same fungal mycelium.

Nitrogen Priming: Can an Ectomycorrhizal Fungus Withdraw Nitrogen from its Plant

Symbiont?

H10 = Nitrogen transfer between symbionts in an ectomycorrhizal network containing

two pine seedlings associated with a single ectomycorrhizal mycelium is

unaffected by host nutrient status.

H1a = Nitrogen transfer between symbionts in an ectomycorrhizal network containing

two pine seedlings associated with a single ectomycorrhizal mycelium shows

preferential allocation from the fungus to the seedling that did not receive foliar

nitrogen.

H20 = Nitrogen transfer between symbionts in an ectomycorrhizal network containing

two pine seedlings associated with a single ectomycorrhizal mycelium is

unaffected by the form and amount of nitrogen available to the fungus.

H2a = Nitrogen transfer between symbionts in an ectomycorrhizal network containing

two pine seedlings associated with a single ectomycorrhizal mycelium shows

increased nitrogen transfer to the host when the fungus has access to inorganic

nitrogen.

1.5.3 Objectives and hypotheses for Chapter 3

With mycorrhizal associations observed in over 90% of plant species (Malloch et al., 1980), up to 30% of plant carbon allocated to ectomycorrhizal fungi (Jones et al.,

1991; Durall et al., 1994a; Högberg et al., 2008), and surface areas of plant-fungal interfaces reaching up to 320,000 m2 ha-1 (Norway spruce forest; Stögman et al., 2013), even a small proportion of carbon mineralization by ectomycorrhizal fungi represents a significant, but unattributed, amount of carbon being metabolised from the terrestrial soil pool. For the second experiment of my thesis, the objective was to determine whether

S. tomentosus, growing symbiotically, could break down and metabolize organic carbon in the substrate, under laboratory conditions.

The Saprotrophic Potential of a Mycorrhizal Fungus in Symbiosis

H30 = No carbon is metabolized from labelled soil organic carbon by S. tomentosus

when growing symbiotically.

H3a = A detectable amount of carbon is metabolized from labelled soil organic carbon

by S. tomentosus when growing symbiotically.

2 Nitrogen Priming: Can an Ectomycorrhizal Fungus

Withdraw Nitrogen from its Plant Symbiont?

2.1 Background

Ectomycorrhizae are ancient symbiotic relationships between fungi and plant roots, which evolved first with members of the Pinaceae, 180-154 Ma, and later with angiosperms (Tedersoo et al., 2010; Kohler et al., 2015). Ectomycorrhizal plants, including many members of the Betulaceae, Diptocarpaceae, Fagaceae, Myrtaceae,

Pinaceae, Salicaceae, and others, depend heavily on their fungal partner for adequate uptake of mineral nutrients from soil (Smith & Read, 2010; Brundrett, 2009). In boreal and montane forests of the Northern Hemisphere, where nitrogen is typically the limiting nutrient (Hunt et al., 1988; Högberg et al., 1998; Averill et al., 2014), ectomycorrhizal fungi mineralize organic forms of nitrogen and translocate a portion of the solubilized nitrogen to their plant partners (Finlay et al., 1992; Bending & Read, 1995; Högberg et al., 2007). Many ecologically and commercially important trees from temperate forests require ectomycorrhizae to develop normally (Hatch, 1936; Nuñez et al., 2009) because they cannot access nutrients from complex soil organic matter on their own

(Abuzinadah et al., 1986; Bending & Read, 1995). In particular, members of the

Pinaceae are considered obligately ectomycorrhizal (Hayward et al., 2015).

Lodgepole pine, (Pinus contorta Dougl.), together with its sister species jack pine

(Pinus banksiana Lamb.), are ectomycorrhizal trees that dominate broad areas of montane and boreal forests in northern and western North America. These early- successional species often form monospecific forests after stand-clearing fires or

clearcutting. Because of their rapid early growth, lodgepole pine seedlings are currently planted on 53% (3.8 million ha) of all second-growth forests in British Columbia, Canada

(Reid et al., 2016). The ability of lodgepole pine to grow rapidly, even under low soil nitrogen (Weetman et al., 1998; Miller et al., 1998), has been attributed to nitrogen supplied by ectomycorrhizal fungi and other root symbionts (Bothwell et al., 2001;

Kranabetter et al., 2006; Chapman & Paul, 2012). To examine the ecophysiological movement of nitrogen in mycorrhizal systems, I set up a laboratory experiment to study nitrogen uptake and allocation by Suillus tomentosus (Kauffman), a fungus that is common on lodgepole pine roots in lower nitrogen sites (Kranabetter et al., 2006; Jones et al., 2012), to two P. contorta var. latifolia Engelm. seedlings differing in their nitrogen status. The purpose of this was to determine if S. tomentosus preferentially allocated nitrogen to ectomycorrhizal seedlings with a lower nitrogen status.

2.2 Materials and methods

2.2.1 Overview of experiment

Figure 2.1 Diagram of microcosms containing two Pinus contorta seedlings growing in association with a single Suillus tomentosus colony on a low-N agar medium overlain with a layer of perlite. The hyphal well contained high-N medium overlain with perlite and was accessible by only hyphae.

For this experiment, pairs of Pinus contorta Dougl. var. latifolia Engelm. seedlings were grown together with Suillus tomentosus (Kauffman) in Petri plate microcosms containing solid medium low in nitrogen (Figure 2.1). Each microcosm contained a well, which was accessible only to the hyphae and was enriched in nitrogen. Once the fungus had colonized the pine roots and hyphae had explored the well for about 4 weeks, two treatments were applied to the foliage: (i) one shoot received soluble nitrogen and H2O, and the other received only H2O (Table 2.1), or (ii)

both shoots received H2O. Three days later, these foliar treatments were crossed with three well treatments: replacement of the original well medium with (i) glycine, (ii) ammonium, or (iii) H2O. To quantify seedling nitrogen content prior to addition of fresh nitrogen to the wells, some microcosms were harvested just before the hyphal well treatments were administered (Table 2.1). Some microcosms received both foliar and nitrogen well treatments, but had any hyphae crossing into the well manually severed just prior to nitrogen addition to the wells (referred to as the severing controls).

Table 2.1 Summary of experimental treatments.

Treatment Foliar Hyphal Harvest Hyphae Number Treatment Well3 Time (d)

1 Differential1 N/A 73 Intact 2 2 H2O N/A 73 Intact 1 3 Differential NH4 75 Intact 4 Differential1 Glycine 75 Intact 1 5 Differential H2O 75 Intact 2 6 H2O NH4 75 Intact 2 7 H2O Glycine 75 Intact 2 8 H2O H2O 75 Intact 1 9 Differential NH4 75 Severed 1 Glycine 75 Severed 10 Differential 1Microcosms where the foliage of one of the two P. contorta seedlings was treated with 4.7 mM- N NH4Cl and the other received deionized H2O. 2 Microcosms where the foliage of both seedlings received H2O. 3 + 15 + 15 At 73 d, 9.46 mM-N NH4Cl (98 at% N), 9.46 mM-N glycine (99 at% N), or deionized H2O were added to wells accessible by hyphae but not roots; N/A indicates treatments that were harvested before N additions were made to the hyphal wells. 2.2.2 Preparation of fungal material and seedlings

Suillus tomentosus was grown on solid 1/10 Modified Melin-Norkrans (MMN) medium (Marx & Bryan, 1975) for one month and then eight ~1 cm3 pieces of medium,

containing a distal edge portion of the hyphae, were placed individually on eight 1 x 8 cm pieces of charcoal-infused filter paper (#728, Macherey-Nagel GmbH & Co. KG), plated on 1/10 MMN agar. Fungi were allowed to grow for one month after which they were used in the microcosms. Pinus contorta seeds (Seedlot 30556, British Columbia

Ministry of Forests, Lands and Natural Resource Operations, Tree Seed Center, Surrey,

British Columbia, Canada, [email protected]) were surface sterilized in aerated 30% hydrogen peroxide for 15 min and then rinsed in aerated sterile deionized water overnight. The seeds were then plated on sterile water agar (8%) and kept at a constant 22°C, with a 12 h photoperiod of 370 μmol m-2 s-1 photosynthetically active radiation (GC-20, Biochambers Inc., Winnipeg, MB) for one month.

2.2.3 Assembly of microcosms

For the hyphal wells (Figure 2.1), caps were removed from autoclaved 1.5 ml microcentrifuge tubes and secured to the bases of empty Petri dishes (10 cm diameter)

2 cm away from the plate edge, by gently melting the cap with a soldering iron. Each plate was then filled with 20 mL ¼-strength solid MMN medium containing no glucose or malt and with 1/40 the concentration of nitrogen (0.095 mM NH4Cl; 26 µg total N) in full

MMN. Each well was filled with 300 μL of ¼-strength MMN medium containing no glucose or malt, with 2.5 X the concentration of nitrogen (9.46 mM NH4Cl; 38 µg total N) in full MMN, and solidified with Phytagel™ (15 g L-1; Sigma Chemical Co., St. Louis).

The relative concentrations of nitrogen in the main plate compartment versus the well were chosen to encourage hyphal proliferation in the well and to make seedlings more dependent on the ectomycorrhizal fungus for access to nitrogen. The medium in the

plate was below the level of the hyphal well rim, creating an air gap between the plate medium and the well medium, which prevented diffusion from the well to the rest of the media.

Autoclaved 6 x 8 cm rectangles of charcoal-infused filter paper were placed on the plate medium, and then sieved Perlite™ (0.25 – 0.5 cm, The Scotts Company,

Mississauga, Canada) was spread, one granule thick, over the exposed medium in the plate to provide an aerated growth surface for S. tomentosus, as per Crahay et al.

(2013). A S. tomentosus inoculum strip (described above) was placed face-up in the middle of the filter paper rectangle (Figure 2.1). Then two one-month-old P. contorta seedlings were placed on each plate so that their roots were in contact with the inoculum (Figure 2.1). A second 6 x 8 cm rectangle of autoclaved filter paper was placed on top of the P. contorta roots and inoculum strip so that it overlapped with the bottom rectangular piece. Sterile perlite was carefully added to the hyphal well, one granule thick, while ensuring that no contact was formed between the well medium and the plate medium. The plates were then sealed with Parafilm™ and the root section wrapped in aluminum foil. The microcosms were placed in the growth cabinet at a 45° angle under the same conditions described above.

2.2.4 Foliage and well treatments

At intervals, plates were removed from the growth cabinet and visually inspected to verify the presence of hyphae in the hyphal wells and mycorrhizae on the root systems. Both were present at about 40 days. After 70 days, treatments were applied to the foliage. Autoclaved 1.5 mL microcentrifuge tubes were filled with 800 μL of either sterilized deionized water or 4.7 mM NH4Cl (53 µg total N), depending on the treatment

(Table 2.1). The foliage of each plant was gently placed inside the microcentrifuge tube until it was approximately 75% submerged in the solution.

After three days, the microcosms were removed from the growth cabinet and six microcosms were harvested (Treatments 1 and 2 in Table 2.1; see 2.2.5 for procedure).

The Phytagel™ medium in the hyphal wells of the remainder of the microcosms was carefully dissolved with 50 mM citric acid and removed by pipette under a dissecting microscope, with as little disruption to hyphae as possible. Then 300 μL of 9.46 mM 99+ at% 15N-glycine (38 µg total N; Isotech Inc., Miamisburg, OH), 9.46 mM, 98+ at%

15 NH4Cl (38 µg total N; Cambridge Isotope Laboratories Inc., Andover, MA) or deionized water were added to each well (Table 2.1). The hyphae in four microcosms each of the glycine and ammonium well treatments were manually severed with hot forceps where they entered the well. The microcosms were then placed back in the growth cabinet.

2.2.5 Harvest

After 48 h, the microcosms were harvested. The microcentrifuge tubes, intact seedlings, and microcosms were frozen separately at -80°C. Later, seedlings were removed from the freezer and the shoots were separated from the roots. Plant tissues were dried at 75°C for 5 days, weighed, and ground to 20 mesh using a Mini Wiley Mill

(Thomas Scientific, Swedesboro, NJ) prior to determination of 15N-14N isotope ratio and total nitrogen via isotope-ratio mass spectrometry (PDZ Europa ANCA-GSL elemental analyzer interfaced to a PDZ Europa 20-20 isotope ratio mass spectrometer [IR-MS],

Sercon Ltd., Cheshire, UK) at The University of California Davis, Stable Isotope Facility.

In some instances, roots of individual seedlings were too small for mass spectrometry, so some replicates were combined (Appendix A, Table A.1). Consequently, replication

was too low to compare total nitrogen content among seedlings in the early harvest treatments (Treatments 1, 2) and the differential foliar application treatment that received a water well addition (Treatment 5). The amount of nitrogen remaining in the microcentrifuge tubes applied to foliage was determined using the salicylate/nitroprusside colorimetric method of Kempers & Zweers (1986).

2.2.6 Nitrogen content of microcosm components and field seedlings

Nitrogen contents of all components of the microcosms were determined so that a nitrogen budget could be calculated. This included: (a) eight replicates of the charcoal filter paper, the high number of replicates to account for structural heterogeneity observed using confocal microscopy (J. Smith, pers. obs.); (b) four replicates each of the Perlite™ and agar medium, and (c) six replicates of the Pinus contorta seeds used in the experiment. Sample preparation was identical to that described for experimental seedlings. Nitrogen was quantified with a Flash 2000 Elemental Analyzer (Thermo

Fisher Scientific Inc.) at the British Columbia Ministry of Environment Technical

Services Laboratory, Victoria, Canada (http://www2.gov.bc.ca/gov/content/environm ent/research-monitoring-reporting/research/analytical-lab).

Additionally, to compare the tissue nitrogen concentrations observed in this study with those of first-season, naturally-establishing lodgepole pine seedlings, I sampled eight, 16 to 20 week-old, P. contorta from two recently-disturbed clear-cut sites

(50.09934°N 119.22346°W and 50.10418°N 119.22709°W, Beaver Lake Rd. near

Kelowna, Canada). Sample preparation of root and shoot tissues was identical to that described for experimental seedlings. Tissue nitrogen was analyzed in the same sample run as the microcosm components above.

2.2.7 Data analysis

One-factor ANOVAs, with shoot and root %N as the response variable, were conducted on (a) the two early harvest treatments (Treatments 1, 2); (b) the experimental treatments that did not receive differential foliar nitrogen (Treatments 6, 7,

8); and (c) the two severing control treatments (Treatments 9, 10). One-factor ANOVAs were also conducted with (i) shoot and root biomass (dry weight) as the response variable on (a) the experimental treatments that did not receive differential foliar nitrogen (Treatments 6, 7, 8), and (b) Day 73 and Day 75 seedlings to compare the two harvesting times, as well as with (ii) total shoot and root nitrogen as the response variable on (a) the early harvest treatments (Treatments 1, 2), and (b) the severing control treatments (Treatments 9, 10). Two-factor ANOVAs, with hyphal well treatment and foliar treatment as fixed factors, were used to compare shoot and root 15N at% and

%N of the treatments that received differential foliar nitrogen and renewed well nitrogen

(Treatments 3, 4), as well as shoot and root biomass, %N, and total nitrogen of all three differential foliar nitrogen treatments (Treatments 3, 4, 5). Two-factor ANOVAs were also used with severed hyphae and foliar treatments as fixed factors to examine the effect of hyphal severing further among (a) the early harvest and severing treatments

(Treatments 1, 2, 9, 10), and (b) the experimental treatments that received foliar H2O and renewed well nitrogen, and the severing treatments (Treatments 3, 4, 9, 10).

ANOVAs were two-tailed, unless stated. Due to small sample sizes, some replicates were combined for IR-MS. Because combined samples can be used as a single replicate for %N, but not total tissue nitrogen, the number of replicates for each treatment differed somewhat between %N and total nitrogen calculations (Appendix A,

Table A.1). A one-tailed t-test was used to examine whether there was net absorbance of nitrogen from the NH4 solutions applied to the foliage, or whether there was net release of nitrogen by foliage into the tubes of water, for all treatments except the early harvest and severing treatments (Treatments 1, 2, 9, 10 excluded), at time of harvest.

All data were subjected to a Shapiro-Wilk test for normality; where necessary, data were log-transformed. Due to differing replicate numbers in almost all analyses, homogeneity of variance was first verified via multiple tests: O’Brien’s, Brown-Forsythe, and Levene’s. A one-factor Welch’s ANOVA was performed on total shoot nitrogen in the foliar H2O, non-early harvest, treatments (Treatments 6, 7, 8) because variances could not be transformed to be homogeneous. Similarly, the total root nitrogen of the differential foliar experimental treatments (Treatments 6, 7, 8) could not be transformed to meet ANOVA assumptions of normality, so a non-parametric one-factor Kruskal-

Wallis rank-sum ANOVA was used to compare these treatments. When ANOVAs detected significant treatment effects (α = 0.05), differences among means were determined using Tukey’s Honest Significant Difference Tests. All analyses were conducted using JMP (V10, SAS Institute Inc., Cary, USA) and all results are shown with standard deviation unless otherwise noted.

2.3 Results

2.3.1 Nitrogen concentration in seedling tissues

The original intent of this experiment had been to examine whether an ectomycorrhizal fungus associated with more than one seedling would allocate nitrogen to those seedlings based on their nitrogen status. For that reason, the nitrogen added to

the wells was labeled with 15N. At the final harvest, many shoots from treatments with intact hyphae and nitrogen additions to the wells (Treatments 3, 4, 6 and 7, Table 2.1) were enriched in 15N compared to seedlings that received water in the wells (65% of shoots and 93% of colonized roots were in excess of the upper 95% CI for Treatments 5 and 8; Appendix A, Figures A.1, A.2), but the foliar treatments had no effect on 15N enrichment (two-factor ANOVA for foliage effect of enriched tissues from Treatments 3

15 and 4: shoots F1,17 = 1.281, p = 0.3; roots F1,22 = 0.128 p = 0.7). N enrichment of most shoots demonstrates that functional ectomycorrhizae had been established in which nitrogen absorbed from the wells was passed by hyphae to their plant partners.

Figure 2.2 Nitrogen concentrations (%N; mean ± SD) of P. contorta roots and shoots in symbiosis with S. tomentosus after (a) 70 d growth followed by 3 d foliage treatment (H2O or 4.7 mM NH4; one-factor ANOVA on Treatments 1 and 2; shoots: F1,10 = 0.3252, p = 0.6; roots: F1,7 = 0.0416, p = 0.8), or an additional 2 d of exposure of S. tomentosus to fresh media in the hyphal well (H2O or 9.46 mM N from glycine or NH4Cl). Microcosms harvested at 75 d had either (b) different foliar N treatments to each seedling (two-factor ANOVA on Treatments 3-5; shoots: foliage effect F1,34 = 0.2592 p = 0.6, well effect F2,34 = 115.24 p < 0.0001, well x foliage F2,34 = 0.1661 p = 0.8; roots: foliage effect F1,27 = 4.97 p < 0.0001, well effect F2,27 = 33.22 p = 0.015, well x foliage F2,27 = 10.53 p = 0.0004) or (c) H2O applied to both seedlings (one-factor ANOVA on Treatments 6-8; shoots: F2,21 = 54.56 p < 0.0001; roots: F2,15 = 76.16 p < 0.0001). Within shoots and roots, figures shown in black are significantly different from those shown in white within treatment groups (Tukey’s HSD where applicable). Treatment numbers are in the top left corner of each box. See Table A.1 for replicate numbers and Table A.2 for nitrogen inputs.

While the 15N results confirmed that some nitrogen had passed from fungus to plant over the two days after the wells were replenished, seedling nitrogen concentrations dropped dramatically over the same period in some treatments. On Day

73, when seedlings had received the foliar nitrogen or H2O treatments for 72 h but the hyphal wells had not yet been refreshed, seedlings had high concentrations of nitrogen in both shoots and roots: approximately 4% (Figure 2.2a). Tissue nitrogen concentrations at that time were not affected by whether the foliage had been exposed to ammonium or water (one-factor ANOVA among the three seedling types in

Treatments 1 and 2; shoots: F2,9 = 1.33, p = 0.3; roots: F2,6 = 0.0182, p = 1.0). By Day

75, 48 hours after solutions in the hyphal wells were refreshed, nitrogen concentrations in shoots had dropped by approximately 70% in microcosms where nitrogen had been added to the well (Treatments 3, 4, 6, 7; Figure 2.2b,c). This occurred regardless of whether nitrogen was added as ammonium or glycine (Figure 2.2b; two-factor ANOVA on Treatments 3-5, well effect: F2,34 = 115.2, p < 0.0001, foliar effect: F1,34 = 0.259 p =

0.6, well x foliar: F2,34 = 0.166 p = 0.8; Figure 2.2c, one-factor ANOVA on Treatments 6-

8, well effect: F2,21 = 54.56 p < 0.0001). Shoots of seedlings in microcosms that received water in the wells, rather than nitrogen, had no detectable change in nitrogen concentration (Treatments 5, 8; Figure 2.2b,c). Furthermore, pretreatment of the foliage with nitrogen did not affect the reduction in nitrogen concentration in the shoots (Figure

2.2b, two-factor ANOVA on Treatments 3 and 4, foliage effect F1,28 = 0.478, p = 0.5), even though nitrogen was absorbed by the foliage from the nitrogen-containing microcentrifuge tubes in some treatments (Figure 2.3). As with shoots, nitrogen concentrations in roots decreased rapidly in response to nitrogen addition to wells, but

only in seedlings that did not have nitrogen applied to the foliage (Figure 2.2b, two- factor ANOVA on Treatments 3-5, well effect: F2,27 = 33.22 p = 0.015, foliar effect: F1,27

= 4.97 p < 0.0001, well x foliar: F2,27 = 10.53 p < 0.001; Figure 2.2c, one-factor ANOVA on Treatments 6-8, well effect: F2,15 = 76.16 p < 0.0001).

Figure 2.3 Final nitrogen concentrations of the solutions applied to the seedling foliage, at harvest. The red line represents the original N concentration of the solutions (foliar N = 4.7 mM, foliar H2O = 0 mM; 5 d before harvest, 3 d before well treatments applied). Significant differences from the original concentrations were tested by one-tailed t-test or non-parametric Wilcoxon Signed Rank t-test, where applicable (* = p < 0.05, ** = p < 0.01). Box plots show median (middle line), 1st and 3rd quartiles (box outlines) and minimum and maximum values (whiskers).

Overall, Day 75 shoots were 13% (SD: 50%) and roots were 63% (SD: 90%) larger than Day 73 seedlings (Appendix A, Table A.1), however this was not significantly different than the experimental seedlings. Additionally, no significant differences in % biomass increase were observed among the experimental treatments (two-factor

ANOVA on Treatments 3-5, shoots: foliage effect F1,34 = 0.3632 p = 0.6, well effect F2,34

= 1.838 p = 0.2, well x foliage F2,34 = 0.1214 p = 0.9, roots: foliage effect F1,20 = 2.500 p

= 0.13, well effect F2,20 = 0.9742 p = 1.0, well x foliage F2,20 = 0.0728 p = 0.9; one-factor

ANOVA on Treatments 6-8, shoots: F2,21 = 0.577 p = 0.6, roots: F2,10 = 1.959 p = 0.2).

Therefore, there was no evidence that differences in seedling growth among well treatments over the 48 h resulted in the change in nitrogen concentration.

This was confirmed by comparing total nitrogen content of the seedlings among treatments. Total tissue nitrogen content could not be calculated for some treatments (1,

2, 5) because some seedlings were too small to analyze separately. For the remaining treatments, the response to nitrogen addition to hyphal wells was the same as that observed for tissue nitrogen concentrations. Shoot nitrogen in microcosms where hyphal wells were refreshed with nitrogen were approximately 60% lower than those where water was added to wells (Treatments 3,4, 6, and 7 versus Treatment 8 in Figure

2.4). Except if their foliage had been treated with nitrogen, roots of plants in microcosms that received renewed fungal nitrogen had 77% lower nitrogen contents than ones treated with water. Treatment of the foliage with nitrogen appeared to prevent a reduction of nitrogen in roots, even though it was still lost from shoots.

Figure 2.4 Total nitrogen (µg N ± SD) of roots and shoots of P. contorta seedlings in symbiosis with S. tomentosus after 70 d growth, then 3 d foliage treatment, and finally 2d of hyphal well replenishment (H2O or 9.46 mM N from glycine or NH4Cl). Treatments are grouped into microcosms where (a) seedlings received differing foliar N treatments (H2O or 4.7 mM NH4), (b) both seedlings received H2O, and (c) hyphae entering the well were severed at the time of N addition to the wells. (a) Treatment 5 not shown due to a low number of root replicates after sample processing, NH4 foliage was 271 ± 137 µg N and H2O foliage was 312 ± 190 µg N. Two- factor ANOVA on: Treatments 3-5 shoots, foliage effect F1,39 = 0.3100 p = 0.6, well effect F2,39 = 29.96 p < 0.0001, well x foliage F2,39 = 0.7650 p = 0.8; Treatments 3 and 4 roots, foliage effect F1,21 = 16.75 p < 0.001, well effect F1,21 = 0.4383 p = 0.5, well x foliage F1,21 = 0.3312 p = 0.6). (b) Treatments 6-8; one-factor Welch’s ANOVA on shoots: F2,22 = 7.625 p < 0.01; Kruskal-Wallis non-parametric one-factor ANOVA roots: p = 0.04. (c) Treatments 9 and 10; one-factor ANOVA on well treatment; shoots: F1,8 = 0.0465 p = 0.8; roots: F1,7 = 0.4965, p = 0.5. Within shoots and roots, figures shown in black are significantly different from those shown in white within treatment groups (Tukey’s HSD where applicable). Treatments 1 and 2, as well as Treatment 5 roots, were excluded from this analysis due to combined sampling for IR-MS. Treatment numbers are in the top left corner of the boxes. See Appendix A, Table A.1 for replicate numbers and Table A.2 for nitrogen inputs.

A nitrogen budget for the microcosms indicated that the seed was the main source of nitrogen for seedlings (Figure 2.5). Seed nitrogen, combined with nitrogen originally present in the well and in the medium in the plate, accounted for approximately 90% of the nitrogen assimilated by Day 75 by seedlings not exposed to foliar nitrogen (W-W in Figure 2.5). When the 9 µg of nitrogen absorbed, on average, by the foliage (Figure 2.3) were added, the four main inputs of soluble nitrogen accounted for approximately 85% of nitrogen in seedlings receiving foliar nitrogen (N-W in Figure

2.5). The remaining 10-15% appeared to come from less available sources, such as agar or the charcoal-infused filter papers (Appendix A, Table A.2). A very small amount of nitrogen was lost from foliage treated with water (2.8 ± 6.9 µg N) (Figure 2.3, Figure

2.5). Ignoring any uptake of nitrogen from the second application of nitrogen to the hyphal wells, only 37% to 56% of the readily-available nitrogen supplied to the seedlings remained in seedlings in treatments where the wells were refreshed (Figure 2.5). An average of 8 µg of the 19 µg 15N added to the wells was detected in shoot and roots.

Although this is likely an underestimate of the total absorbed from the wells, if 8 µg are added as an additional source of readily available nitrogen, the percentage remaining in seedlings was 36% (for treatments where the foliage was exposed to water) to 54% (for treatments where the foliage was exposed to ammonium). This represents 43% (for treatments where the foliage was exposed to water) to 66% (for treatments where the foliage was exposed to ammonium) of the nitrogen originally present in the seed.

Figure 2.5 Sources of readily available N, and the final distribution of N in shoots and roots of P. contorta seedlings at the 75-day harvest. The first letter represents the foliage treatment (W = water; N = NH4); the second letter represents the well treatment (W = water; N = NH4 or glycine). For inputs, the foliage tube compartment was set at 9 µg because that was the amount, on average, taken up by the shoots. An additional 2.8 µg N was added to the W-W and W-N final distribution bars to account for N lost to the water applied to the foliage. Shoot and root N values are averages of the appropriate seedlings from the following treatments: W-W = Treatments 5, 8; N-W = Treatment 5; W-N = Treatment 3, 4, 6, 7; N-N = Treatments 3, 4). Horizontal lines above bars indicate N inputs for that treatment, assuming no N was absorbed from refreshed wells. Percent values above bars represent percent of N inputs. See Appendix A, Table A.2 for N contents of less available sources, such as agar and charcoal-infused filter paper.

Results from the severing control microcosms provided strong evidence that the reductions in nitrogen in the seedlings were triggered by the fungus. These microcosms had hyphal connections between the seedlings and wells severed immediately before

NH4 or glycine additions to the wells. The shoots and roots of the severing control seedlings (Figure 2.6) had similar nitrogen concentrations to the early harvest seedlings

(Figure 2.2a; two-factor ANOVA comparing severing and foliar treatment from

Treatments 1, 2, 9, and 10, shoots: foliage effect F1,20 = 0.2241 p = 0.6, severing effect

F1,20 = 0.0073 p = 0.9, foliage x severing F1,20 = 0.1998 p = 0.7, roots: foliage effect F1,16

= 0.6834 p = 0.4, severing effect F1,16 = 0.0208 p = 0.9, foliage x severing F1,16 = 0.0738 p = 0.8). In summary, seedlings with severed hyphae showed no signs of nitrogen loss in response to nitrogen addition to the wells. Instead, they had approximately three times the shoot nitrogen concentrations as seedlings that were treated identically except for the severing of hyphae (two-factor ANOVA comparing severing and foliar treatments from Treatments 3, 4, 9, and 10; severing effect: F1,40 = 171.60, p < 0.0001).

Figure 2.6 Nitrogen concentrations (% N; mean ± SD) of root and shoots of P. contorta seedlings after 70 d of growth followed by 3 d of foliage treatment (H2O or 4.7 mM NH4) and finally 2 d of hyphal well addition (9.46 mM N from glycine or NH4Cl). Red lines indicate manual severing of hyphae at the time of N addition to the wells. Treatment numbers are in the top left corner of the boxes. One-factor ANOVA; shoots: F1,7 = 3.8664 p = 0.09; roots: F1,6 = 0.1191, p = 0.7). See Appendix A, Table A.1 for replicate numbers and Table A.2 for nitrogen inputs.

2.3.2 Nitrogen concentration in naturally-occurring field seedlings

The foliar nitrogen concentrations observed in the early harvest seedlings

(Treatments 1,2, Figure 2.2a) were considerably above those expected in foliage of mature lodgepole pine trees in British Columbia forests (Brockley, 2001). Because these experimental seedlings were considerably younger (10 weeks old) than trees used to generate nutritional guidelines, I sampled naturally regenerating, 16 to 20-week- old, P. contorta germinants from two recently logged sites in order to determine typical tissue nitrogen concentrations for seedlings of this age. Nitrogen concentrations in foliage of the naturally-regenerating lodgepole pine seedlings covered a similar range to that seen in the experimental seedlings (Figure 2.7). The highest foliar nitrogen observed in the field seedlings was similar to that of the early harvest seedlings

(approximately 4%), whereas the lowest was just slightly higher than those of seedlings that had experienced rapid nitrogen loss (approximately 1.2%). Nitrogen concentrations in roots of field seedlings tended to be slightly lower than shoots, with 75% of the roots between 1.4 and 1.7%. Two seedlings had higher nitrogen concentrations, but these were still slightly lower than the values observed in experimental seedlings.

In most studies of ectomycorrhizal seedlings, colonization by the ectomycorrhizal fungus is associated with increased accumulation of nitrogen compared to non- mycorrhizal plants. This is especially true when the source nitrogen is in complex organic forms, because most tree seedlings cannot access these without ectomycorrhizal fungi (Abuzinadah et al., 1986). As a result, ectomycorrhizal fungi are viewed as an essential conduit of nitrogen from soil to their plant partners (Smith &

Read, 2010). It is important to note, however, that ectomycorrhizal fungi can vary in the proportion of absorbed nitrogen they allocate to their plant partner based on fungal species (Colpaert et al., 1996), and this retention of nitrogen by ectomycorrhizal fungi may be proportional to increasing nitrogen limitation (Wallander, 1995; Wallander et al.,

1999; Hobbie & Colpaert, 2003). In nitrogen-limited forests, increasing allocation of plant carbon to the ectomycorrhizal fungus can also result in decreased allocation of nitrogen to the tree (Hasselquist et al., 2016; Högberg et al., 2017). To our knowledge,

the results described in the current study are the first to clearly demonstrate a loss of plant nitrogen associated with colonization by an ectomycorrhizal fungus. What is especially remarkable is the amount and rate of translocation of nitrogen out of seedlings: a 70% reduction in plant nitrogen concentration over 48 h. There is some evidence, however, that nitrogen-rich conifer germinants can lose nitrogen under field conditions. Zackrisson et al. (1997) measured the nitrogen status of Pinus sylvestris L. germinants over their first growing season and found that seedlings experienced a 35% nitrogen loss compared to original seed reserves, except in the treatment where ectomycorrhizal hyphae were severed. Seedlings with severed hyphae retained all their seed nitrogen. This indicates that a similar phenomenon might occur under natural field conditions.

The design of our microcosms, where the fungus grew among cellulose fibers on filter paper, meant that I could not extract hyphae to compare hyphal nitrogen concentrations among treatments. Nevertheless, I expect that nitrogen was translocated into the fungus in our study, based on the retention of nitrogen by seedlings with severed hyphae. Higher concentrations of nitrogen in colonized roots of seedlings where foliage was exposed to ammonium than seedlings where foliage was treated with water also suggested that nitrogen was moving from shoots in the direction of the fungus in the roots. Based on the nitrogen budget, I speculate that hyphae had depleted the nutrients placed in the well at the beginning of the experiment and then became nutrient limited (Tibbett & Sanders, 2002). The addition of nitrogen to the wells on Day

73 would then have stimulated the metabolism of the hyphae (Perez-Moreno & Read,

2000; Hasselquist et al., 2016). Ectomycorrhizal fungi experiencing starvation are

known to increase expression of nitrogen transporters (Elleström et al., 2015). If that happened at the root-fungal interface in the microcosms, it could have triggered movement of nitrogen from plant to fungus. The function of nitrogen transporters at the root-fungal interface is not completely understood, but it is possible that some could mediate nitrogen transfers in both directions (Garcia et al., 2016a).

The pine seedlings in our study, like pine germinants growing in the field, had very high tissue nitrogen concentrations, likely derived primarily from the seed

(Abuzinadah & Read, 1986). Under such conditions, transfer of nitrogen to a fungal symbiont would not be expected to inhibit seedling growth, especially if the fungus began to translocate nitrogen to its plant partner shortly thereafter. Indeed, supply of nitrogen to their fungal symbiont could act as a form of priming by the seedling, similar to that documented for carbon, where soil microorganisms and fungi are unable to mineralize soil organic matter optimally unless an initial supply of labile carbon is provided via litterfall, root exudates or plant symbioses (Hamer & Marschner, 2005a).

Priming enables microbes to increase their production of excreted enzymes, giving the microbe a significant resource exploitation advantage (Hamer & Marschner, 2005b;

Chen et al., 2014). A nitrogen-priming mechanism would be especially beneficial for ectomycorrhizal symbionts of lodgepole pine seedlings, which naturally germinate after severe wildfires that have killed trees in the existing stand and removed the nitrogen- rich forest floor. Under such conditions most ectomycorrhizal fungi colonize pine germinants from spores (Glassman et al., 2016) and, therefore, would not have access to the carbon and nutrients from a hyphal network associated with larger trees. Because spores have such small reserves to fuel hyphal growth, a young seedling is a potential

source of nitrogen. Importantly, seedlings that lost nitrogen in response to fungal triggers had foliar nitrogen concentration just slightly within the deficient range

(Brockley, 2001), but still well within the range for survival.

In conclusion, I report here on an observation of rapid and substantial loss of nitrogen from lodgepole pine germinants, apparently mediated by ectomycorrhizal fungi that had experienced a rapid change in their abiotic environment. Further research is required to confirm that this phenomenon occurs in the field, determine which environmental changes may trigger this response, and discover the physiological mechanism behind this phenomenon.

3 The Saprotrophic Potential of a Mycorrhizal Fungus in

Symbiosis

3.1 Background

Generally, the key defining characteristic of ectomycorrhizal symbioses is the allocation of photosynthetic carbon from the plant to the fungus, and the transfer of mineral nutrients from the fungus to its plant partner. By contrast, saprotrophic fungi acquire nutrients and carbon from the decomposition of soil organic matter (SOC;

Abuzinadah et al., 1986; Abuzinadah & Read, 1986; Brundrett, 2004; Bidartondo, 2005).

Nevertheless, over the last decade it has become clear that ectomycorrhizal fungi excrete a diverse set of extracellular enzymes (Pritsch & Garbaye, 2011; Rineau et al.,

2012; Hacquard et al., 2013; Bödeker et al., 2014). These enzymes have high target specificity (Luis et al., 2005; Shah et al., 2013), and have functionality and efficiency that have been equated to the decomposition capabilities of saprotrophic fungi (Rineau et al., 2012; Phillips et al., 2014; Zhao et al., 2013; Bödeker et al., 2014). Hyphae of ectomycorrhizal fungi even appear to competitively interfere with those of saprotrophic fungi in patches of soil organic matter (Bödeker et al., 2016; Fernandez & Kennedy,

2016). It is therefore of interest to determine whether or not ectomycorrhizal fungi take up and metabolize SOC (de Vries & Caruso, 2016), i.e., behave saprotrophically. Given the global coverage and ecological impact of ectomycorrhizae, if even 1% of mycorrhizal carbon is derived from soil organic matter, this would equate to millions of tons of soil carbon being released globally as respiration or incorporated into fungal tissues. Examining the potential for saprotrophy by ectomycorrhizal fungi is important

because the conventional understanding of these fungi considers all fungal carbon to come from the plant host (Cullings & Courty, 2009).

There is some evidence for saprotrophy among ectomycorrhizal fungi (Durall et al., 1994b; Hobbie et al., 2002; Courty et al., 2007; Cullings et al., 2008; Hobbie et al.,

2013; Hobbie et al., 2014). For example, Durall et al. (1994b) formed mycorrhizas between Pseudotsuga menziesii and four different fungal species and discovered that, when grown in media containing 14C-labelled hemicellulose, cellulose, pine needles, or humic polymers, the labelled carbon could be detected as respiration and was fixed in the seedling tissues for most of the substrates and species tested. As that experiment was conducted in sterile vessels, it is likely that either the fungus or plant mediated this release of carbon from the labelled substrate and metabolized the carbon so that it could be detected as respiration, and found in the plant tissues. Another source of soil carbon known to be taken up by ectomycorrhizal fungi is amino acids (e.g., Abuzinadah et al., 1986; Näsholm et al., 2000; Teramoto et al., 2016), where the amino-acid carbon likely becomes incorporated into fungal cell walls or proteins together with nitrogen

(Lipson & Näsholm, 2001; Baldrian, 2009; Jones et al., 2009).

Proposed reasons for saprotrophic behaviour by ectomycorrhizal fungi are: (i) as an alternative carbon source when supplies of photosynthate from the host plant are low, (ii) as a by-product of mining the soil for organic nutrients, or (iii) when allocation of plant photosynthate to mycorrhizal roots is high and the fungus can expend more resources on enzymes (Talbot et al., 2008; Baldrian, 2009; Lindahl & Tunlid 2015;

Hupperts et al., 2017). A fungal carbon exporter at the plant-fungal interface has not yet been characterized, yet we know that ectomycorrhizal hyphae can facilitate carbon

transfer from tree-to-tree (Simard et al., 1997; Song et al., 2015; Klein et al., 2016;

Pickles et al., 2017). This indicates that they can transport carbon toward the plant- fungal interface, not just away from it (Song et al., 2015; Teramoto et al., 2016; Pickles et al., 2017). Song et al. (2015) hypothesized that source-sink gradients direct this transfer.

It is difficult to study saprotrophy in the field; therefore, I decided to use a simple study system: a single plant host, a single species of ectomycorrhizal fungus, and a single SOC source, in a controlled and sterile environment. I chose to use Pinus contorta and Suillus tomentosus ectomycorrhizae because I had previous experience with these species. These organisms were grown symbiotically in silica sand-filled microcosms that also contained fungal and plant cell wall material, enriched in 13C or at natural abundance levels, in mesh bags accessible to hyphae but not roots. Evidence that S. tomentosus was acting saprotrophically while in symbiosis with P. contorta would

13 be indicated by detecting respired CO2 that is enriched with C in the mycorrhizal microcosms that received enriched SOC.

3.2 Methods

The experiment comprised a completely randomized design with six treatments

(Table 3.1). Microcosms contained the fungus plus seedlings, fungus alone, or no organisms. All three organism treatments were subjected to two labelling treatments:

13C-enriched SOC or SOC at natural abundance. Fifty microcosms were originally established.

Table 3.1 Experimental treatments and replicates. Original Survived to Not Treatment SOC 2 Organisms Replicates Harvest 1 Contaminated

M1 Enriched Mycorrhizal 17 11 3 M2 Natural Abundance Mycorrhizal 17 13 1 F1 Enriched Fungus-Only 5 5 0 F2 Natural Abundance Fungus-Only 5 5 0 E1 Enriched No Organisms 3 3 3 E2 Natural Abundance No Organisms 3 3 3

1 = Harvest was 6 months after seedling germination (Pinus contorta), and 5 months post-inoculation (Suillus tomentosus). 2 = Enriched soil organic carbon (SOC) was at 4.1 at% 13C and natural abundance SOC was 1.1 at% 13C.

3.2.1 SOC creation

In this experiment, 13C-labelled and unlabelled fungal cell wall, as well as unlabelled plant cell wall material were used as the sole source of organic carbon in microcosms. To produce enough labelled and unlabelled fungal cell wall for this experiment, 200 L of ½ Modified Melin-Norkrans (MMN) medium (Marx & Bryan, 1975), with D-glucose as the only carbon source, were distributed into Erlenmeyer flasks ranging in volume from 750 mL to 8 L; additionally, 1 L of ½ MMN, with 13C D-glucose

(99%+, LC Scientific Ltd. Concord, ON) as the only carbon source, was distributed equally into ten 750 mL flasks. The volume of medium placed in the flasks was only a quarter of the flask’s designed volume to increase the surface area in contact with the air. The flasks were then inoculated with a Geomyces sp., which had been in culture in our lab. Geomyces is a widely-distributed genus, commonly found in northern temperate soils. The flasks were then shaken at low speed for two months to allow the fungus to grow and use all of the carbon in the medium. I separated the fungal cell wall

components from cytoplasmic and aqueous fractions using a procedure from Kerley and

Read (1997; Appendix B).

The extracted fungal cell wall material represented 40% of the carbon required for the experiment. The remainder of the SOC was obtained from woody and herbaceous litter collected from a ponderosa pine forest adjacent to UBC Okanagan campus. This material was dried and ground to 20 mesh using a Mini Wiley Mill

(Thomas Scientific, Swedesboro, NJ) and subjected to the same extraction protocol

(Kerley & Read, 1997).

3.2.2 Microcosm construction

To create the microcosms for this experiment, 59-mL flat-bottom culture tubes

(#9850-25XX, Corning Inc., Corning, NY) were filled with 20 g of silica sand, which had been heated in a muffle furnace to 360 °C for 2 h to remove all carbon (Craft et al.,

1991). I added 0.44 g of fungal and plant cell wall material to each microcosm to achieve 1% SOC (0.2 g) content in the microcosms (assuming 45% carbon content of the fungal and plant cell wall material; Swift et al., 1979; Bauer et al., 2002). This is similar in magnitude to the 1.5-2% SOC found in mineral soils in the Okanagan Valley

(Zebarth et al., 1999; Hannam et al., 2016; Munro, 2017). The SOC for each microcosm was divided into two 1.5 cm x 3 cm bags (85 mg of fungal material and 135 mg of plant material per bag) made of 25 µm nylon mesh (Plastok Associates Ltd., Wirral, UK), sealed with a heat sealer. The two bags in the 13C treatments each contained 78.4 mg of natural abundance fungal material, 6.6 mg of 99+% 13C fungal material, and 135 mg of plant material. This resulted in the enriched bags containing SOC at 4.1 at% 13C and the natural abundance bags containing SOC at 1.1 at% 13C. The bags were placed

vertically in the sand on opposite sides of the microcosm cylinder and at different depths

(Figure 3.1). The sand-filled tubes and the SOC bags were autoclaved separately and assembled in a biological safety cabinet. A subset of both enriched and natural- abundance microcosms were set aside to (a) receive only Suillus tomentosus (i.e., no seedling), to examine how much carbon the fungus could mobilize when free living; or

(b) have neither fungus nor plant, to determine if any carbon was released from the

SOC in gaseous form.

To facilitate air sampling from the microcosms, the culture tubes were capped with a two-position polypropylene closure (26795-25, 25 mm, Corning Inc.) attached to two 16-gauge, 3 cm needles (BD Biosciences, San Jose, CA). The closures were left in the open position, but wrapped with Parafilm™ (Bemis NA, Neenah, WI). To ensure sterility of the air entering the microcosms, the needle tips were combined with 1 mL aerosol-filter pipette tips (16466-008, VWR International, Radnor, PA). The interfaces between the needle tips and the pipette tips, and the needle tips and microcosm caps was sealed with contact sealing adhesive (‘Amazing Goop: Plumbing’, Eclectic Products

Inc., Eugene, OR). The needles, tube closures, culture tubes, and pipette tips were sterilized in an autoclave and the microcosms were assembled in a biosafety cabinet.

Figure 3.1 Diagram of microcosm used in the labelling experiment.

3.2.2 Preparation and incorporation of organisms

Suillus tomentosus Singer was grown on solid 1/10 MMN medium (Marx &

Bryan, 1975) for one month. Then 1-cm3 pieces of medium, each containing a distal portion of a colony, were placed on strips of porous cellophane (GE80-1129-38, GE

Healthcare, Little Chalfont, UK), located on the upper surface of solid 1/10 MMN medium. Fungi were allowed to grow for one month, after which they were used in the microcosms.

Pinus contorta Dougl. var. latifolia Engelm. seeds (Seedlot 30556, British

Columbia Ministry of Forests, Lands and Natural Resource Operations, Tree Seed

Center, Surrey, British Columbia, Canada, [email protected]) were surface sterilized in aerated 30% hydrogen peroxide for 15 min and then shaken at low speed in sterile deionized water overnight. Three seeds were then directly placed in each microcosm, and kept at a constant 22°C, with a 16/8 h (light/dark) photoperiod of 370 μmol m-2 s-1 photosynthetically active radiation in a plant growth chamber (GC-

20, Biochambers Inc., Winnipeg, MB). All microcosms had the lower half of the culture tubes covered with opaque black plastic throughout the experiment to shade the fungus growing within the substrate.

Seedlings were inoculated twice, one and two months after seeding. The inoculum consisted of S. tomentosus hyphae that had been peeled off of the cellophane. During both inoculation events, excess seedlings and empty seed coats were removed from the microcosms. During the first inoculation event, extra germinated seedlings were transferred into microcosms where none of the seeds had germinated.

Seedling transfer happened only between microcosms with identical SOC treatments.

As a result of poor seedling germination, the number of replicates differed slightly between the two main treatments (Table 3.1).

3.2.3 Growth of seedlings

The microcosms were visually monitored for the presence of new hyphae and moisture status of the substrate. Nutrient solution was added (0.5 mL) to the microcosms when the seeds were first put in the microcosms and again, one and two months later, when the S. tomentosus inoculum was added. The nutrient solution was

specially formulated for the experiment (Appendix C). First, a nitrogen concentration of

20 mg N L-1 was chosen as an optimal nitrogen concentration to maintain growth of

Pinus seedlings, when administered repeatedly (Hartley et al., 1999). Ratios of macro and micro nutrients were then set using Ingestad and Kähr’s (1985) recommendations for Pinus, but with the P concentration halved to promote mycorrhization (Jones et al.,

1990). The microcosms received only deionized water for the remaining three months of the experiment. Sufficient deionized water was added to each microcosm to moisten the sand substrate while only saturating the lower 1 cm of sand in the culture tube. During the first four months, several microcosms became visibly contaminated and were discarded.

3.2.4 Analysis of respired CO2

After four months of seedling growth (i.e., three months post-inoculation), the at%

13 C and ppm of the CO2 in the microcosms was measured with a Picarro cavity ring- down spectrometer (G2131-i, Picarro Inc., Santa Clara, CA). The CO2 generated through respiration of hyphae and seedlings will be referred to henceforth as ‘total respiration’. To ensure that all ambient air in the Picarro had been flushed with sample air and the sample readings had stabilized, 3 min of sampling time (20 mL of sample) was required. The mean values recorded for each sample were taken after the readings had stabilized for at least 1 min. For each sample, 25 mL of air was extracted from the microcosms with a syringe and transferred to a 0.5 L Tedlar® bag with a Kynar® valve. A

® Tedlar bag filled with CO2-free air was attached to microcosm so that neutral air pressure was maintained in the microcosms during sampling. To ensure the sample was within the recommended CO2 ppm range of 500 ± 200 ppm CO2 (P. Millard and A.

Midwood, pers. comm.), 3 mL of air were first extracted from the bag and analyzed using a static sampling EGM-4 gas analyzer (PP Systems, Amesbury, MA). Any samples that exceeded 700 ppm CO2 were diluted with CO2-free air until they were in the recommended range. For any samples lower than 300 ppm CO2, a 15 mL sample was taken 24 h later from the same microcosm and used to augment the CO2 in the appropriate Tedlar® bag. In all cases where this was done, the extra sample contained enough CO2 to make the original sample usable. Between all sample transfers, sampling equipment (syringes and Tedlar® bags), as well as the external components of

® the Kynar valves, were flushed twice with CO2-free air, verified to be 0 - 3 ppm CO2 with the EGM-4. The respiration analysis was conducted again, one month after the first analysis (5 months after seedling germination, 4 months post-inoculation), to test for any changes.

3.2.5 Harvest

Seven days after the second CO2 analysis had been conducted, total respiration readings were again obtained for the microcosms. Seedlings were removed from the microcosms and immediately frozen at -80°C. A small sample of the sand was aseptically removed from each microcosm and spread onto a plate of potato dextrose medium (BD Difco™ PDA, Fisher Scientific Co., Ottawa, ON) to test for any contamination of the substrate by bacteria or by fungi other than S. tomentosus. The microcosms were then resealed, flushed with CO2-free air, and placed back in the growth cabinet, in the dark. After 24 h, air was sampled and analyzed as described in

13 section 3.2.4 to determine the at% C and ppm of CO2 respired by S. tomentosus

alone, hereafter referred to as ‘fungal respiration’. All microcosms were then sealed and stored at -20°C, for any further analyses.

Three weeks later the Petri plates were visually inspected for the presence of contaminants. The roots from the microcosms were examined with a compound microscope to verify the presence of ectomycorrhizae. DNA extraction, amplification and Sanger sequencing were subsequently conducted by an Undergraduate Research

Award student on fungal colonies cultured from these microcosms using the approach of Kranabetter et al. (2012).

3.2.6 Statistical analyses

13 To test whether CO2 was significantly enriched in C in labelled microcosms compared to natural abundance values from the unlabelled microcosms, both the total respiration readings at each sampling time, and the fungal respiration of the two mycorrhizal treatments were compared using one-tailed t-tests (Treatments M1, M2). To test whether there was a change in the relative enrichment of total respiration over the three sampling times, a repeated measures analysis with a linear mixed model approach was also used: specifically, the data were analyzed with a restricted maximum likelihood (REML) ANOVA, with treatment, time, and treatment x time as fixed effects.

Replicate, nested within treatment, was set as a random effect to account for multiple sources of variation. When the ANOVA detected significant treatment effects (α = 0.05), differences among means were determined using Tukey’s Honest Significant Difference

Tests.

The two treatments that did not contain any organisms were compared using a one-tailed t-test (Treatments E1, E2) to examine whether the labelled SOC was

13 releasing elevated levels of CO2 in the absence of organisms. The mycorrhizal treatment with enriched SOC was compared to the no-organism treatment that received enriched SOC with a one-tailed t-test (Treatments M1, E1), to determine if the presence of mycorrhizal S. tomentosus in the 13C-enriched-microcosms increased the isotopic signature of CO2 found in the microcosms. All data showed normal distribution (Shapiro-

Wilk tests). All analyses were conducted using JMP (α = 0.05; V10, SAS Institute Inc.,

Cary, USA) and all results are shown with standard deviation unless otherwise noted.

3.2.7 Isotope mixing model

An isotope mixing model can be used to determine what proportion of CO2-C in a sample originated from two different sources (Fry, 2006; Millard et al., 2010; Walder et al., 2012), as long as the isotopic carbon signature of the two sources (referred to as the

‘end members’) is known. In my experiment, the two potential sources of carbon for the fungus were plant carbon and SOC. Equation (1) uses the 13C at% of the end members, the enriched SOC (‘SOC’) and the natural abundance fungal respiration (‘NTLabun’), in combination with the proportional contribution of each source, where 푓푆푂퐶 and 푓푁푇퐿푎푏푢푛 respectively represent this proportion, to determine the at% 13C of the sample.

푎푡%푠푎푚푝푙푒 = 푎푡%푆푂퐶 ∗ 푓푆푂퐶 + 푎푡%푁푇퐿푎푏푢푛 ∗ 푓푁푇퐿푎푏푢푛 (1)

Since the 13C at% abundance of the sample, but not the proportional contributions, are known, the equation can be rearranged to solve for 푓푆푂퐶 and 푓푁푇퐿푎푏푢푛.

Given that 푓푆푂퐶 + 푓푁푇퐿푎푏푢푛 = 1, 푓푁푇퐿푎푏푢푛 can be substituted with 1 − 푓푆푂퐶, the equation can then be rearranged to (2), which can be solved to determine the relative contributions each source made to the 13C at% of the sample (Fry, 2006).

푓푆푂퐶 = (푎푡%푠푎푚푝푙푒 − 푎푡%푁푇퐿푎푏푢푛)/(푎푡%푆푂퐶 − 푎푡%푁푇퐿푎푏푢푛) (2)

I used the fungal respiration readings from the mycorrhizal treatment that received natural abundance SOC (Treatment M2) as the ‘plant-carbon’ end member

(NTLabun) because they represent the at% of fungal respiration when all components of the microcosm were at natural abundance. In an experiment where artificial enrichment is not used, it is essential that the source values are as accurate as possible because differences in 13C isotopic signature can be orders of magnitude smaller than in this case. In those experiments, minute differences in the isotopic signatures of the unlabelled SOC and plant photosynthate in the mycorrhizal, natural abundance SOC, treatment (Treatment M2) would be important. However, in my study, the difference in at% between the two end members: enriched SOC and natural abundance plant carbon, was substantial enough that minor isotopic discrimination processes could be ignored. Therefore, even if the S. tomentosus in the mycorrhizal, natural abundance

SOC, treatment was behaving saprotrophically and metabolizing the unenriched SOC, any effects on the isotopic ratio of the respired carbon would be too small to affect the end member value used here.

3.3 Results

3.3.1 Contamination

Many PDA plates inoculated with sand from the microcosms showed obvious signs of bacterial or non-Suillus-like fungal contamination; however, other plates had hyphal growth and colour characteristics that were typical for S. tomentosus. The two fungus-only treatments (F1, F2; Table 3.1) were lost entirely due to contamination, and only one microcosm from the mycorrhizal, natural abundance SOC, treatment

(Treatment M2) was uncontaminated. Consequently, data from these treatments were eliminated from any statistical tests. Nevertheless, three replicates each of the two no- organism treatments showed no signs of contamination. Three of the mycorrhizal, 13C- labelled SOC, microcosms contained only S. tomentosus (Anton Hsu, unpublished), and, hence, could be used to evaluate saprotrophic behaviour.

3.3.2 Total respiration

Figure 3.2 13C atom percent values of total respiration readings. Treatment M1 and M2 comprised of one Pinus contorta in symbiosis with Suillus tomentosus with 1% substrate SOC content at: M1) 4.1 at% 13C or M2) 1.1 at% 13C. Some microcosms contained no organisms but still had the SOC addition (E1, 4.1 at% 13C; or E2, 1.1 at% 13C). Readings were taken 111 (t1), 142 (t2), and 148 d (t3) post-germination. S. tomentosus inoculation occurred 30 d after germination. n = 3 for all except Treatment M2 (n = 1). Letters indicate significant differences among treatments (Tukey’s HSD following REML ANOVA, Treatment M2 excluded). Bars indicate mean ± SEM. Star indicates a significant within-treatment difference (p < 0.05).

Suillus tomentosus appeared to be able to access and assimilate carbon from the SOC even when growing in symbiosis. Natural abundance values for total respiration samples, based on all of the mycorrhizal, natural abundance SOC, treatment microcosms, including the contaminated ones, were: Day 111 = 1.09 (0.004), Day 142 =

13 13 1.09 (0.006), Day 148 = 1.09 (0.006) C-CO2 at% (Treatment M2, n = 12). The at% C of carbon dioxide sampled from microcosms containing enriched SOC and P. contorta ectomycorrhizal with S. tomentosus (Treatment M1) was elevated relative to these natural abundance values (Figure 3.2; day 111: t-ratio2 = 6.63, p-value = 0.010; day

142: t-ratio2 = 4.23, p-value = 0.026; day 148: t-ratio2 = 5.12, p-value = 0.018). CO2 from microcosms with ectomycorrhizal seedlings and labelled SOC (Treatment M1) was also significantly more enriched than microcosms lacking organisms, with either labelled or unlabelled SOC treatments (Treatments E1, E2; Tukey’s test following repeated measures REML ANOVA; treatment effect: F2,6 = 18.46, p = 0.003). Interestingly, enrichment also increased with time (F2,12 = 4.07 p = 0.04), more so where mycorrhizal seedlings were present than in microcosms where no organisms were present (time x treatment: F2,12 = 3.16, p = 0.05).

3.3.3 Fungal respiration

Respiration samples were taken after the seedlings were removed and the microcosms underwent a 24 h dark incubation in order to isolate CO2 generated by the fungi from CO2 generated by the seedlings. Natural abundance of CO2 from fungal

13 respiration only was 1.09 (0.001) C-CO2 at% (Treatment M2, n = 13) using all of the mycorrhizal, natural abundance SOC, treatment microcosms, including the contaminated ones. The results were similar to the total respiration readings, with 13C-

CO2 at% higher in mycorrhizal microcosms with labelled SOC (Figure 3.3; one-tailed t- test from natural abundance, t-ratio14 = 10.53, p-value = < 0.0001). This is consistent with metabolism of the labelled SOC by S. tomentosus. It is interesting to note that the empty microcosms containing labelled SOC (E1) also showed significant CO2 enrichment over the same microcosms without labelled SOC (E2), indicating that some labelled carbon may have been volatilized or otherwise released as CO2 (one-tailed t-

13 test, t-ratio4 = 4.19, p-value = 0.007). It is important to note that the CO2 enrichment of the no-organism microcosms that received enriched SOC was still significantly lower than fungal respiration in the mycorrhizal microcosms that received enriched SOC (one- tailed t-test, t-ratio4 = 3.14, p-value = 0.017), meaning that the labelled CO2 release observed in the no-organism microcosms that received enriched SOC was not the only factor contributing to the observed CO2 enrichment. Additionally, the uncontaminated, mycorrhizal, enriched SOC, microcosm that had the highest 13C at% value observed during the total respiration analysis (Figure 3.2) also had the highest 13C at% fungal respiration (Figure 3.3), corroborating the two analyses.

Figure 3.3 13C atom percent values of fungal respiration, following a 24 h dark incubation after seedling removal. Treatment M1 and M2 shown with all replicates included (n = 11 and 13; black = not contaminated; grey = contaminated) and without contaminated replicates (red). Microcosms consisted of one Pinus contorta in symbiosis with Suillus tomentosus with 1% substrate SOC content at: M1) 4.1 at% 13C or M2) 1.1 at% 13C. Some microcosms contained no organisms but still had the SOC addition (E1, 4.1 at% 13C; or E2, 1.1 at% 13C). Readings were taken 149 d post-germination. S. tomentosus inoculation occurred 30 d after germination. Uncontaminated n = 3 for all except Treatment M2 (n = 1). Bars indicate mean ± SEM.

An isotope mixing model was used to determine the proportion of fungal respiration that was derived from the labelled SOC. The end member for the SOC was set at 4.1 at%, and the mean fungal respiration from the mycorrhizal microcosms that received natural abundance SOC (1.09 at%) was used as the end member for plant- carbon. For the sample value in Equation 2 the mean fungal respiration of the mycorrhizal microcosms that received enriched SOC (2.52 at%) was used, but this number was altered because the no-organism microcosms that only received enriched

13 SOC had significantly enriched CO2. The cause of this enrichment is unknown, but the mean of this treatment (E1) was enriched by 0.37 at% over natural abundance; thus this enrichment (0.37 at%) was used as a correcting factor for the sample value in the isotope mixing model (2.52 at%). This resulted in the sample value being 2.15 at% 13C.

Using these values, the isotope mixing model estimated that 35% of fungal respiration was derived from the labelled SOC tissue. Using the standard deviation from the fungal respiration of the mycorrhizal microcosms that received enriched SOC, the isotope mixing model calculated a minimum value of 17% and a maximum value of 54% of carbon respired by S. tomentosus came from the labelled SOC.

3.4 Discussion

In this experiment, CO2 generated by Suillus tomentosus in symbiosis with P. contorta and grown in microcosms with 13C-labelled SOC was enriched in 13C relative to labelled microcosms lacking organisms. To the best of my knowledge, no other microorganisms were present in these microcosms; therefore, the S. tomentosus associated with the pine roots is the most likely source of the enriched CO2. Given that hyphal respiration is commonly associated with recently acquired carbon (Clemmensen

et al., 2013) it appears that S. tomentosus was readily accessing and metabolizing the

SOC placed in the microcosms. This experiment supports my hypothesis that an ectomycorrhizal fungus can behave saprotrophically while in symbiosis. These results are consistent with a similar experiment by Durall et al. (1994b) where they noted enriched fungal respiration when 14C-labelled hemicellulose, cellulose, humic polymers, and conifer needles were provided to cultures of several different species of ectomycorrhizal fungi. Also, Dighton et al. (1987) previously noticed that mycorrhizal P. contorta in sterile microcosms caused significant decomposition of cotton, relative to non-mycorrhizal pine. More recently, while examining protease activity in 13 different

Suillus species growing on bovine serum albumin (BSA), Rineau et al. (2016) found that increasing glucose additions inhibited protease activity. As high concentrations of glucose, or other carbon catabolites, repress carbon metabolism pathways, Rineau et al. (2016) surmised that this would only occur if the fungi were using BSA as a carbon source.

An advance made by this study was the quantification of the relative contributions of SOC and plant photosynthetic C to fungal respiration. The isotope mixing model indicated that approximately 17-54% of the carbon respired by S. tomentosus came from the SOC. A previous study with a detection limit of 5%, did not detect any use of SOC by ectomycorrhizal fungi associated with Quercus alba in the field (Treseder et al., 2006). A value of 35% is surprisingly high. If ectomycorrhizal fungi metabolize SOC to this extent in the field then it should have been detected in earlier studies. The fungus may have been behaving more saprotrophically in this lab study because the plant was providing small amounts of carbon. This could have been due to

i) the lower irradiance of our growth cabinet (less than ¼ natural sunlight), ii) the early stages of mycorrhizal formation, iii) the young age of the seedlings, iv) CO2 depletion of the microcosm air due to limited gas diffusion through the Parafilm and aerosol-filter pipettes, or v) the high humidity in the sealed microcosms (can cause a reduction in photosynthesis due to stomatal closure under high humidity). In addition, S. tomentosus had no competition from obligate saprotrophic fungi or bacteria (Bödeker et al., 2016). A potential source of artificial inflation is that the empty microcosms that contained the enriched SOC (Treatment E1) had significantly enriched respiration compared to the mycorrhizal microcosms that received natural abundance SOC (Treatment M2; 1.46 ±

0.15 at% compared to 1.09 ± 0.00 at%, respectively); the SOC may have released

13 some carbon as CO2. Despite this artificial source of CO2, respiration in the mycorrhizal microcosms that received enriched SOC (2.52 ± 0.56 at%) was still significantly higher than respiration in the no-organism microcosms that contained the enriched SOC by 1.05 at% 13C, so I am confident that a saprotrophic effect was observed.

With fungi having a high turnover rate and constituting around 40 (Högberg &

Högberg, 2002) to 60% (Fernandez & Koide, 2012) of soil microbial biomass, chitin, and other fungal cell wall necromass, forms a large carbon and nitrogen pool in the soil; chitin alone can represent up to 3% w/w of ectomycorrhizal soils (Kerley & Read, 1997).

The saprotrophy observed in this experiment should not be surprising as the fungal- specific chitinase activity of several ectomycorrhizal species has been characterized in situ with high activity (Buée et al., 2007), and the saprotrophic behaviour of ectomycorrhizal fungi has been demonstrated with several other ‘recalcitrant’ carbon-

containing structural compounds, such as hemicellulose, cellulose, and humic polymers

(pure culture; Durall et al., 1994b). Nevertheless, a debate around ectomycorrhizal saprotrophy still persists in the literature despite all of the theoretical work and modelling

(Talbot et al., 2008; Baldrian, 2009; Lindahl & Tunlid 2015; Hupperts et al., 2017), inferential evidence for ectomycorrhizal saprotrophy (Courty et al., 2007; Cullings et al.,

2008; Rineau et al., 2016), saprotrophic enzyme evidence (Pritsch & Garbaye, 2011;

Rineau et al., 2012; Hacquard et al., 2013; Zhao et al., 2013; Bödeker et al., 2016;

Fernandez & Kennedy, 2016), and direct evidence of ectomycorrhizal saprotrophy

(Dighton et al., 1987; Durall et al., 1994b).

In this experiment, only one ectomycorrhizal fungus was studied. However, S. tomentosus was a relevant choice because it colonizes lodgepole pine in stands of different ages, from two-month-old seedlings to 90-year-old mature trees (Bradbury,

1998; Rineau et al., 2016). Moreover, S. tomentosus forms a major component of the ectomycorrhizal fungal communities across this range of ages (Wurzburger et al.,

2001). It was identified on 25% of P. contorta trees in a mature forest in Oregon

(Ashkannejhad & Horton, 2006), and comprised 20% of the sporocarps in an undisturbed lodgepole pine forest in Utah (Kropp & Albee, 1996). S. tomentosus has also remained abundant even after thinning (Kropp & Albee, 1996). During primary succession of lodgepole pine on coastal sand dunes in Oregon, S. tomentosus was present on 30% of the successional seedlings, with 5% of fresh deer droppings found on the site containing viable S. tomentosus spores (Ashkannejhad & Horton, 2006).

Because of its abundance, any saprotrophic activity by S. tomentosus has the potential to influence carbon cycling in lodgepole pine stands in western North America.

Suillus tomentosus has some unique traits that may make saprotrophic behaviour more likely than in other ectomycorrhizal fungi. It is one of only a few ectomycorrhizal fungi capable of forming a tubercule (nodule-like clusters of ectomycorrhizal root tips; Paul et al., 2006) and is the only known tuberculate fungus that harbors nitrogen-fixing bacteria in its tubercules (Paul et al., 2007). Because S. tomentosus may have access to larger quantities of nitrogen than other ectomycorrhizal fungi, it may be able to produce an increased quantity or complexity of saprotrophic enzymes. Another factor to consider is that nitrogen fixation produces relatively large quantities of ammonium, which is toxic to fungi and must be rapidly converted into carbon-containing compounds (Scott, 2008); Suillus tomentosus may be more apt to behave saprotrophically because this fungus requires carbon to prevent ammonium toxicity. On a separate note, in the early successional scenarios this fungus appears to be well adapted to, plant hosts may not produce adequate quantities of photosynthate to sustain S. tomentosus, so the ability to supplement its carbon budget with SOC could benefit the fungus (Talbot et al., 2008).

Considerable numbers of replicates were lost in this experiment as a result of contamination. I suspect that the contamination was caused by the aerosol-filter pipette tips. During the course of the experiment the culture tube caps had to be removed and replaced several times for inoculation, and nutrient and water additions. During harvest,

I noticed that the positive air pressure that resulted from pushing the caps down onto the microcosms had enough force to push some of the filter disks partially out of the pipette cylinder, resulting in a slight opening in the sterile system. The saprotroph- contaminated mycorrhizal microcosms that received enriched SOC also readily used

13 13 the labelled C SOC (2.73 ± 1.11 at% CO2, n = 8). Nevertheless, it is clear from both the total respiration and fungal respiration data that Suillus tomentosus readily liberates, takes up and metabolizes carbon from recalcitrant fungal cell wall molecules (Kerley &

Read, 1997), which would primarily consist of chitin and β-glucans.

Because of the contamination, the replicate number was too low for a two-factor

ANOVA of the at% of fungal respiration using SOC type and organisms as factors.

Thus, a one-tailed t-test was used to compare the main treatment effect, respiration of microcosms with labelled SOC vs unlabelled SOC, to determine if the labelled carbon respiration was enriched relative to natural abundance respiration. To determine a

13 natural abundance respiration value for this analysis the mean CO2 respiration readings of all mycorrhizal microcosms that received natural abundance SOC, including

11 contaminated microcosms, was used. My reasoning for using the contaminated microcosms is that they contained microorganisms that respired carbon from natural abundance SOC in the same environment provided to S. tomentosus, and therefore could be combined with the one uncontaminated treatment to produce a reliable natural abundance value with appropriate error values. Any subtle differences in 13C- concentrations of CO2 respired by bacteria, ‘saprotrophic’ fungi and S. tomentosus utilizing a natural abundance SOC source would be multiple orders of magnitude lower

13 ‰ than CO2 being respired by S. tomentosus utilizing SOC enriched to 4.1 at% C.

There are limits to which the results of a laboratory study with three experimental replicates, one ectomycorrhizal species, 100-day-old Pinus contorta seedlings, and manufactured soil substrate can be extrapolated. Nevertheless, being able to show a significant saprotrophic effect, constituting approximately 35% (17-54%) of the recently

acquired carbon of an ectomycorrhizal fungus, growing symbiotically, challenges the current paradigm, where virtually the entire carbon budget of an ectomycorrhizal fungus is assumed to come from the plant partner.

4 Conclusion

4.1 Hypothesis summary and significance

4.1.1 Chapter 2

The hypotheses for Chapter 2, ‘Nitrogen Priming: Can an Ectomycorrhizal Fungus

Withdraw Nitrogen from its Plant Symbiont?’ were as follows:

H10 = Nitrogen transfer between symbionts in an ectomycorrhizal network containing

two pine seedlings associated with a single ectomycorrhizal mycelium is

unaffected by host nutrient status.

H1a = Nitrogen transfer between symbionts in an ectomycorrhizal network containing

two pine seedlings associated with a single ectomycorrhizal mycelium shows

preferential allocation from the fungus to the seedling that did not receive foliar

nitrogen.

H20 = Nitrogen transfer between symbionts in an ectomycorrhizal network containing

two pine seedlings associated with a single ectomycorrhizal mycelium is

unaffected by the form and amount of nitrogen available to the fungus.

H2a = Nitrogen transfer between symbionts in an ectomycorrhizal network containing

two pine seedlings associated with a single ectomycorrhizal mycelium shows

increased nitrogen transfer to the host when the fungus has access to inorganic

nitrogen.

Unfortunately, the results that were obtained in this study were not suited to addressing these hypotheses. I was able to detect that 15N labelled nitrogen, supplied

only to the fungal mycelium, had been taken up by the shoot and root tissues of the P. contorta germinants. This indicates that functional ectomycorrhizae had been established, but the enrichment of seedling tissues was very low, relative to the enrichment of the nutrients provided to the mycelium. Thus, with the information available to me, I was unable to disprove either H10 or H20. While going over the experimental data to try to figure out what had caused the unusually low volume of nitrogen transfer from the fungus to the host plant, I noticed that the seedlings in microcosms that received nitrogen additions to the mycelium had lost a significant proportion of their nitrogen over a very short period. To my knowledge, this is the first documented observance of this effect. My new hypothesis, based on the information reported in Chapter 2, is that ectomycorrhizal seedlings may possess a mechanism whereby they are able to allocate substantial proportions of their seed-nitrogen reserves to ectomycorrhizal fungal symbionts in situations where the ectomycorrhizal fungus would require nitrogen to, competitively and expediently, mobilize and take up newly accessed nutrients. The implication that there may be situations where ectomycorrhizal plant hosts allocate their nutrients to their mycorrhizal fungi contravenes over thirty years of ecophysiological research (Abuzinadah et al., 1986; Finlay et al., 1992;

Bending & Read, 1995; Högberg et al., 2007). My results do not invalidate any research involving nutrient transfer from an ectomycorrhizal fungus to their host plant. The fungus-to-plant direction of exchange has been documented extensively. However, my results indicate that there may be situations where transport happens in the opposite direction and that researchers in our field may have to revaluate the paradigm of

unidirectional nutrient transfers in ectomycorrhizae. Clearly the unusual transport detailed in this thesis deserves further research.

4.1.2 Chapter 3

The hypotheses for Chapter 3, ‘The Saprotrophic Potential of a Mycorrhizal Fungus in

Symbiosis’ were as follows:

H30 = No carbon is metabolized from labelled soil organic carbon by S. tomentosus

when growing symbiotically.

H3a = A detectable amount of carbon is metabolized from labelled soil organic carbon

by S. tomentosus when growing symbiotically.

Despite the considerable loss of replicates due to contamination, I was able to statistically reject H30, with my data supporting H3a. It is clear from my results that the ectomycorrhizal S. tomentosus had taken up and metabolized carbon from the labelled

SOC material while in symbiosis. Given the current debate in the field on the potential for ectomycorrhizal saprotrophy while in symbiosis, this research could be a critical methodological stepping stone for future researchers interested in examining this topic.

My experimental design was as simple as possible, with only two organisms and requiring a simple t-test to examine, yet remained as realistic as possible within these parameters, with relevant organisms, and SOC concentration and composition. It is my hope that future researchers can adapt this design to conduct future experiments. My results clearly challenge the paradigm of biotrophic, uni-directional carbon flux in ectomycorrhizae. I expect these results to encourage researchers in the ectomycorrhizal field to acknowledge that saprotrophy can occur, and may constitute a

significant portion of the fungal carbon budget. The next step is to begin to disseminate information about the saprotrophic role of ectomycorrhizal fungi to other academic fields, such as soil science, the ecological sciences, and climate modellers. This is important because over 25% of the global forest canopy (Chapin & Danell, 2001), constituting approximately 740 million trees (Crowther et al., 2015), allocates up to 20% of their net primary production to ectomycorrhizal fungi (Hobbie & Hobbie, 2008). If even

1% of the ectomycorrhizal fungal carbon budget is derived from SOC, this represents millions of tons of SOC that is respired globally. This potential carbon flux is not currently attributed (Cullings & Courty, 2009).

4.2 Strengths and limitations

Both of my studies concluded that ectomycorrhizal nutrients (C, N) can be transferred in directions that are not included in the prevailing paradigm. . A major challenge to publication of these studies is that supporting research with similar findings is either nonexistent (Chapter 2) or very scarce (Chapter 3; Dighton et al., 1987; Durall et al., 1994b). It remains to be seen whether the results I obtained are observable in the field or merely artifacts arising from laboratory conditions. Supporting research in field conditions would greatly strengthen the results and conclusions of these studies.

A major strength of the nitrogen experiment (Chapter 2) was the many treatments that were created for the experiment. While they were designed to serve a different purpose, they greatly increased my ability to interpret the results. More specifically, the early harvest treatments, originally designed to provide baseline 15N values, were invaluable in demonstrating the nitrogen concentration of the seedlings immediately prior to the well treatments, thereby helping to create a timeline for the

‘nitrogen priming’ effect. The severing controls were another fortuitous treatment group.

They were originally designed to test for any diffusion or spillage of 15N from the hyphal well to the plate media. However, these treatments formed a critical control because I was able to show that the severing of hyphae that had access to the fresh nitrogen source resulted in no nitrogen loss by the seedlings. This highlighted that it was hyphal access to the fresh nitrogen addition that caused the nitrogen loss from the seedling.

Another strength of the nitrogen experiment was the use of both inorganic and organic nitrogen forms as well treatments. These two nitrogen forms involve different stages of the soil nitrogen cycle and different uptake and mobilization strategies (Abuzinadah et al., 1986; Finlay et al., 1992; Bending & Read, 1995; Näsholm et al., 2000).

Consequently, they make the observed results more relevant to ectomycorrhizal forest soils (Chalot & Brun, 1998).

Unfortunately, because the nitrogen experiment was originally designed to examine differences in hyphal-15N allocation among the two seedlings, there were analyses that I could not perform on the microcosms. Due to the use of charcoal filter paper as a means of pressing the roots into the inoculum strip, I was unable to extract hyphal tissue for nitrogen concentration analysis, despite attempts using several methods. Additionally, using two seedlings in the microcosms was superfluous for the results that I present here. Another limitation of the experiment is that I had only two sampling times; if there had been several collection times over a longer time period, I could have gained more information on the extent, duration and lasting effects of the nitrogen loss from the seedlings.

For the carbon experiment, one of the biggest strengths of the experiment was the statistical results. A p-value of < 0.0001 (fungal respiration of M1 to natural abundance) is very robust; it is highly unlikely that the observed effect was an artifact or type 1 error (false positive). Another strength of the experiment was that it was planned to be as simple as possible. It is important to note, however, that the conclusions I have made relied on only three replicates. Additionally, I was unable to determine the cause

13 of the CO2 enrichment observed in the no-organism microcosms that received enriched SOC. It is possible that these microcosms were also contaminated as not all contaminants may have grown on the PDA plates used to verify contamination (L.

Nelson, pers. comm.). The extensive contamination that was detected also severely impacted the experiment, as it would have been very informative to examine the results from the microcosms that only contained S. tomentosus. This would have enabled me to compare the respiration values of the fungus while completely saprotrophic to those obtained while the fungus was mycorrhizal. I speculate that the fungus-only microcosms would have had a 13C respiration value higher than the value obtained for the mycorrhizal microcosms, because the free living fungus would not be receiving plant

13 photosynthate. If I had observed significantly higher CO2 at% for free-living than for symbiotic fungi in the presence of labelled SOC, it would have added strength to the experiment by indicating that the saprotrophy I observed may have functioned to supplement the plant-allocated photosynthetic carbon provided to the S. tomentosus. It would have also provided evidence that the symbiotic fungus was receiving carbon from the plant, in addition to the SOC.

Another treatment that would have been beneficial to my experiment would have been a non-mycorrhizal control that only contained P. contorta. This treatment would have enabled me to confirm that carbon was not released from the SOC by the roots. It is important to note that the 25 µm mesh constraining the SOC would have excluded roots, as fine root width is typically around 500 µm (Stögman et al., 2013), so the roots would have had limited contact with the SOC. Nevertheless, root exudates may have released carbon from the SOC. However, this is not supported by the results of non- mycorrhizal controls from previous research with hemicellulose, cellulose and needles

(Durall et al., 1994b), or cotton and chitin (Dighton et al., 1987), used as SOC substrates.

4.3 Potential applications and future research directions

4.3.1 Chapter 2 applications and future directions

There are a number of applications of my observation that nutrient-deprived ectomycorrhizal fungi may acquire nitrogen from germinant hosts when a new source of nutrients becomes available. This nutrient transfer from plant to fungus would be most beneficial in early succession situations, where the environment is devoid of established plants and ectomycorrhizal fungi. In this situation, pine germinants are primarily colonized from ectomycorrhizal spores (Glassman et al., 2016). Thus, the survival of both organisms depends on the ability of the fungus to competitively colonize and exploit nutrient patches. If the unusual nitrogen transfer I observed could serve to prime the fungus with nitrogen (Hamer & Marschner, 2005b; Chen et al., 2014), this investment of nutrients may confer a competitive advantage on the germinant and its ectomycorrhizal symbionts. A clear understanding of this mechanism has the potential

to increase our understanding of successional ecology and ecophysiology after fires, clear-cut logging or other stand-destroying disturbance in ectomycorrhizal forests.

Clearly more research needs to be done to elucidate the breadth and extent of this mechanism. Does it happen in field conditions? Is it observable with different ectomycorrhizal plant and fungus species? How much fungal nitrogen deprivation is required to cause the effect to occur? If this effect is confirmed in the field, and some of these questions are addressed, then the ecological effects of this mechanism could be utilized by policy makers and foresters. For example, it might greatly improve the survival percentage or stress tolerance of replanted seedlings if the seedlings were inoculated in the nursery with ectomycorrhizal fungi or spores that are known to use this nitrogen priming mechanism.

If I had the opportunity to further investigate nitrogen movement to the mycorrhizal fungus I would first attempt to observe the effect in field conditions. To accomplish this I would conduct a 2 x 2 x 3 factorial experiment. The factors would be i) presence/absence of mesh around the roots to exclude mycorrhizal colonization, ii) a logical nitrogen addition to the extramatrical mycelium (simulating the hyphae encountering a patch of nitrogen rich soil organic matter) or not, and iii) forest seral stage. To attempt to ensure that I would observe any nitrogen loss from the seedlings, if it occurs in field conditions, I would harvest seedlings from each treatment group every two days for the first several weeks, and then bi-weekly after that for the first growing season to observe the long-term nitrogen concentrations in the seedlings. The purpose of obtaining the long-term results would be to determine how long it takes seedlings to recover from nitrogen allocation to the fungi. If any seedling mortality occurred

throughout the course of this experiment then this would also be recorded and included in my data. If this field experiment provided support for my hypothesis I would transition from ecophysiological studies to metabolomic and genetic studies to attempt to determine how this effect functions at the molecular level.

4.3.2 Chapter 3 applications and future directions

It is surprising that the debate about the existence of ectomycorrhizal saprotrophy persists decades after this effect has been reported (Treseder et al., 2006;

Talbot et al., 2008; Baldrian, 2009; Cullings & Courty, 2009; Lindahl & Tunlid, 2015). My observation of ectomycorrhizal saprotrophy confirms a few other observations (Dighton et al., 1987; Durall et al., 1994b). A more concerted effort needs to be made by ectomycorrhizal researchers to conceptualize and understand the potential and capabilities for ectomycorrhizal saprotrophy while in symbiosis. Further research is certainly needed to examine this effect in a variety of field conditions with multiple plant and fungal species. Nevertheless, this effect is of tremendous relevance and importance because climate change is becoming a serious and critical issue for our society, and there is growing uncertainty of how and when climate change is going to affect humanity’s future (IPCC AR5, 2013). With the vast variability among climate models (Dolman et al., 2010, for a review) it is imperative that all sources, sinks and flux rates of carbon are examined and estimated to ensure that modellers are better able to predict global trends and trajectories, as well as provide the required foundational knowledge for proper employment of conservation, environmental, and habitat rehabilitation policies (Kranabetter et al., 2007; Talbot et al., 2008; Meyer et al., 2010;

Corrêa et al., 2011; Orwin et al., 2011; Bradford, 2014; Pritchard et al., 2014; Shantz et

al., 2015). Soil respiration is a major global carbon flux, which almost balances out all

CO2 fixation by terrestrial photosynthesis (Vargas & Allen, 2008), and is an order of magnitude greater than all anthropogenic CO2 emissions (Högberg & Read, 2006).

Therefore, an unattributed flux with potentially millions of tons of soil carbon being released to the atmosphere, such as ectomycorrhizal fungi behaving saprotrophically while in symbiosis, requires further research and attention, as well as consideration when constructing climate and carbon models.

If I had the opportunity to continue examining ectomycorrhizal saprotrophy while in symbiosis I would not expand upon this experiment excessively for my first follow up project. I would instead focus on resolving the methodological and design issues I encountered. I would potentially incorporate one or two more tree and ectomycorrhizal species, but my primary concerns would be resolving the contamination issues,

13 incorporating an air delivery system to remove any CO2 that may be in the headspace, and developing a less fiscally- and temporally-demanding method for producing labelled

SOC. If this first experiment provided further evidence to support my hypotheses I would proceed with ecophysiological field studies and then metabolomic and genetic studies, as with the nitrogen experiment.

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Appendices

Appendix A: Ch. 2 supplementary information

Table A.1 Replicates used for N% and total N analyses as well as mean final weight and 48h biomass increase for shoots and roots of each treatment.

%N Analyses Replicates (Foliar Application) Total N Analyses Replicates (Foliar Application) Biomass Calculations Shoot Root Shoot Root Biomass Biomass Foliar Harvest Shoot Shoot Root Root Shoot Shoot Root Root Weight Weight Increase6 Increase6 3 Treatment Number Treatment Hyphal Well Time (d) Hyphae (H2O) (N) (H2O) (N) (H2O) (N) (H2O) (N) n (mg ± SD) (mg ± SD) (% ± SD) (% ± SD)

1 Differential1 N/A 73 Intact 64 - 54 - 64 - 44 - 3 6.5 ± 3.7 1.7 ± 0.8 - - 2 5 2 H2O N/A 73 Intact 6 - 4 - 6 - 2 - 3 3.8 ± 1.7 0.6 ± 0.2 - - 1 3 Differential NH4 75 Intact 8 8 7 7 8 8 6 6 8 5.3 ± 2.1 1.8 ± 1.0 2 ± 40 54 ± 91 4 Differential1 Glycine 75 Intact 8 8 7 6 8 8 6 4 8 6.8 ± 1.9 1.5 ± 0.8 31 ± 38 34 ± 66 1 5 5 5 Differential H2O 75 Intact 4 4 3 3 4 4 2 2 4 6.8 ± 3.6 1.0 ± 0.5 32 ± 69 -11 ± 46 2 6 H2O NH4 75 Intact 8 - 7 - 8 - 6 - 4 6.2 ± 3.8 1.8 ± 1.1 20 ± 75 54 ± 92 2 7 H2O Glycine 75 Intact 8 - 6 - 8 - 4 - 4 6.3 ± 2.3 1.1 ± 0.3 22 ± 44 -9 ± 28 2 8 H2O H2O 75 Intact 8 - 5 - 8 - 3 - 4 4.8 ± 2.8 1.4 ± 0.9 -7 ± 55 23 ± 81 1 4 4 4 4 9 Differential NH4 75 Severed 4 - 4 - 4 - 4 - 4 7.7 ± 4.8 1.8 ± 1.2 49 ± 92 52 ± 102 10 Differential1 Glycine 75 Severed 54 - 44 - 54 - 44 - 4 5.1 ± 5.8 1.6 ± 1.5 -1 ± 114 42 ± 128 1 Microcosms where the foliage of one of the two P. contorta seedlings was treated with 4.7 mM-N NH4Cl and the other received deionized H2O. 2 Microcosms where the foliage of both seedlings received H2O. 3 + 15 + 15 At 73 d; 9.46 mM-N NH4Cl (98 at% N), 9.46 mM-N glycine (99 at% N), or deionized H2O were added to wells accessible by hyphae but not roots; N/A indicates treatments that were harvested before N additions were made to the hyphal wells. 4Analyses where differential foliar applications were merged due to low replicate counts. 5Analyses where sample merging for IR-MS resulted in insufficient replicate numbers; these were excluded from statistical analyses. 6Percent biomass increase calculated as increase from the mean of Day 73 treatments (1 and 2; shoots: 5.16 ± 3.1 mg, roots: 1.15 ± 0.82 mg).

Table A.2 Total N inputs into the microcosms. N Source N per Plate (µg ± SD) N per Seedling (µg ± SD)

Seed n/a 223 ± 48 Initial Well 38 19 Foliar N* n/a 53 Replenished Well† 38 19 MMN plate media 26 13 Agar 1057 516 Charcoal-infused 1083 ± 80 542 ± 40 paper

* An average of 9 µg N was absorbed from the N applied to foliage † An average of 8 µg N from 15N was found in seedlings with access to the second application of N to the hyphal wells.

Figure A.1 Shoot 15N at% values. Red line indicates upper 95% CI of natural abundance (0.3652). Natural abundance calculated as mean of differential foliar and foliar H2O treatments that received H2O well applications. Percent indicates how many replicates were above the 95% CI for natural abundance and n indicates total replicates.

Figure A.2 Root 15N at% values. Red line indicates upper 95% CI of natural abundance (0.3652). Natural abundance calculated as mean of differential foliar and foliar H2O treatments that received H2O well applications. Percent indicates how many replicates were above the 95% CI for natural abundance and n indicates total replicates.

100

Appendix B: SOC extraction protocol

From Kerley & Read, 1997:

1. Vacuum filtration with repeated DI-H2O washes, to wash out any remaining nutrient solution

2. Freeze tissues for 12 h, defrost at 2-3 °C for 12 h, resuspend in DI-H2O and vacuum filter, repeated five times, to fractionate cells and extract the aqueous cellular fraction

3. Grind tissues in liquid nitrogen with a mortar and pestle, to pulverize and homogenize the material

4. Resuspend in DI-H2O, centrifuge at 4000 rpm for 5 minutes, pour out supernatant and resuspend pellet with DI-H2O, repeated five times, to remove any soluble components released during the liquid nitrogen grinding

5. Sonicate for 3 minutes, to fracture cell walls completely

6. Resuspend in DI-H2O, centrifuge at 4000 rpm for 5 minutes, pour out supernatant and resuspend pellet with DI-H2O, repeated ten times, to isolate the solid cell wall fraction

7. Immerse in 70% ethanol for 16 h at 2-3 °C, to extract any cytoplasmic residue

8. Resuspend in DI-H2O, centrifuge at 4000 rpm for 5 minutes, pour out supernatant and resuspend pellet with DI-H2O, repeated ten times, to remove all ethanol

9. Store frozen until use

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Appendix C: Modified Ingestad’s pine medium

20mg/L N (Hartley et al., 1999) then ratios to match Ingestad and Kähr (1985) N:P ratio lowered from 100:14-100:7

Part A (Micro Concentrate)

ZnSO4*7H2O 0.0264 g CuSO4*5H2O 0.0236 g (NH4)6Mo7O24*4H2O 0.0181 g NaCl 0.00153 g

Water to 1 L

Part B (Macro Concentrate)

NH4Cl 7.64 g KCl 1.34 g MgSO4*7H2O 1.22 g CaCl2*2H2O 0.440 g K2SO4 0.0829 g Fe(III)Citrate*3H2O 0.0750 g MnSO4*H2O 0.0307 g H3BO3 0.0229 g

Water to 1 L

Part C (Final)

KH2PO4 0.00615 g Part A 1 mL Part B 10 mL 1 N (1 M) HCl 10 µL Agar 8 g

Water to 1 L

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