AN ABSTRACT OF THE DISSERTATION OF
Wendy S. Phillips for the degree of Doctor of Philosophy in Environmental Sciences presented on September 6, 2012
Title: Drivers of Arbuscular Mycorrhizal Fungal Community Composition in Roots:
Hosts, Neighbors, and Environment
Abstract approved:
____________________________________________________________________
Virginia M.Weis Eric W. Seabloom
The vast majority of terrestrial plant species live in symbiosis with arbuscular mycorrhizal fungi (AMF). AMF and plants live in complex networks, with roots of individual plants hosting multiple AMF, and single AMF colonizing multiple plants concurrently. Through the exchange of resources, the two partners of this symbiosis can have great effects on each other, effects which can ripple through both communities. What determines the patterns of associations between the partners is still largely unknown. In this dissertation, I examine a variety of factors, and in particular host identity, that could drive the community composition of AMF in roots.
I began by surveying the diversity of AMF in roots of 12 plant species at a remnant bunchgrass prairie in Oregon, U.S.A. (Chapter 2). To do that, I first designed new primers for use in polymerase chain reaction (PCR) to specifically amplify DNA from all Glomeromycota species. Using those primers, I found 36 distinct AMF phylogenetic groups, or operational taxonomic units (OTUs) in the roots from the

prairie. The proportion of OTUs in the basal order Archaeosporales was greater than in many other environmental surveys. I also conducted an in silico analysis to predict how effectively previously published primers would detect the whole diversity of
OTUs I detected.
I then assayed AMF community composition in the roots of 50 plants from nine plant species (Chapter 3). To do that, I designed primers specific to 18 of the
OTUs detected in the initial field survey and used them to test for the presence of each
OTU in the roots individual plants. I used that data to test if AMF community composition in individual roots correlated with host identity, spatial distribution, or soil characteristics. I found host identity was associated with both the richness and the structure of root AMF communities, while spatial distribution and soil characteristics were not.
Finally, I performed an experimental test of the effect of host identity and community context on AMF community assembly (Chapter 4). I grew plants from four native perennial plant species, including two common and two federally endangered plants, either individually or in a community of four plants (with one plant of each species). I analyzed the AMF community composition in the roots of all plants after 12 weeks of growth with exposure to a uniform mix of field soil as inoculum. I found that host species identity affected root AMF richness and community composition, and community context affected AMF richness. Only one of the endangered species was highly colonized by AMF, and I did not detect unique AMF communities associated with it.

This dissertation provides information on the diversity of AMF at a remnant bunchgrass prairie, an ecosystem which has been the subject of very few studies of
AMF. Although a complex mix of factors interact to determine AMF community composition in roots, this work provides strong evidence that host identity plays a major role in that process.

©Copyright by Wendy S. Phillips
September 6, 2012
All Rights Reserved

Drivers of Arbuscular Mycorrhizal Fungal Community Composition in Roots: Hosts,
Neighbors, and Environment by
Wendy S. Phillips
A DISSERTATION
Submitted to
Oregon State University in partial fulfillment of the requirements for the degree of
Doctor of Philosophy
Presented September 6, 2012
Commencement June 2013

Doctor of Philosophy dissertation of Wendy S. Phillips presented on
September 6, 2012.
APPROVED:
_____________________________________________________________________
Co-Major Professor, representing Environmental Sciences
_____________________________________________________________________
Co-Major Professor, representing Environmental Sciences
_____________________________________________________________________
Director of the Environmental Sciences Graduate Program
_____________________________________________________________________
Dean of the Graduate School
I understand that my dissertation will become part of the permanent collection of
Oregon State University libraries. My signature below authorizes release of my dissertation to any reader upon request.
__________________________________________________
Wendy S. Phillips, Author

ACKNOWLEDGEMENTS
I cannot say enough to thank Virginia Weis, my Ph.D. co-advisor and longtime mentor before that. Thank you for letting me do my dissertation research in your lab, even before you graciously took on the role of co-advisor. I am so lucky to have been under your wing. I am brimming with gratitude for the help, guidance, and resources provided by Eric Seabloom, my other co-advisor. Thank you, Eric, for pushing me to take a wider perspective of my research (and also sometimes to narrow it). I thank my other committee members, Bruce McCune and Joseph Spatafora, for being there to answer my questions and for the general mentoring they provided. I thank my current graduate council representative, Russell Ingham, and former grad reps Yevgeniy Kovchegov and Eric Skyllingstad for their service. I would also like to thank former committee members Deborah Clark, Jane Smith, Elizabeth Borer and
Mark Wilson. I very much appreciate the guidance all of you gave me, particularly during the early phases of my program.
Thank you to current and former Weis lab members from my time as a Ph.D. student, Angela Poole, Camille Paxton, Christine Schnitzler, and Sheila Kitchen, for so kindly sharing your lab space with me and my roots. Thank you to the Borer-
Seabloom crew that overlapped with me and welcomed me into the world of plants
(and aphids): Angela Brandt, Kelly Farrell, Lydia O’Halloran, Phoebe Zarnetske, Sean
Moore, and Shawn Gerrity. I especially appreciate Kelly Farrell, my office mate of a few years, for her supportive conversations about all sorts of matters.

I could not have done my Ph.D. program without the financial support provided by a National Science Foundation Graduate Research Fellowship and research grants from Prairie Biotic Research, Inc. and The Portland Garden Club
Pamplin Fund. I would like to thank Andrew Blaustein, Director of the Environmental
Sciences Graduate Program, for admitting me to the program and providing financial assistance along the way.
I am blessed by the support of wonderful friends and family. I thank my sister and lifetime friend, Sharon, for being there for me through thick and thin. Brad Upton and Elizabeth Sokolowski, you two are life savers! Thank you for your friendship through these years and in particular for your help both with editing my dissertation and hanging out with my kids in the last few weeks while I worked to get this done on time. I am part of an amazing women’s group. You know who you are. Words cannot express how much I have learned from you all or how much I appreciate the wisdom and guidance of our group leader. Thank you all for your caring presence.
I have two utterly priceless treasures in my life, Abby and Jessica. Thank you for sharing your grade school years with my grad school years and for helping me to keep my feet on the ground.

CONTRIBUTION OF AUTHORS
Deborah Clark contributed to the experimental design of Chapter 3.

TABLE OF CONTENTS
Page
Chapter 1. Introduction. ......................................................................................... 1
Organization of Introduction .............................................................................. 1
Origins of mycorrhizal symbiosis ...................................................................... 1
Biology and ecology of arbuscular mycorrhizas ................................................ 2
Functional differences in AMF .......................................................................... 4
Taxonomic diversity of AMF and molecular tools for identification ................ 6
The pairing of AMF and plants ........................................................................ 11
Mechanisms of partner selection ...................................................................... 14
Broader impacts ............................................................................................... 15
Summary .......................................................................................................... 16
References ........................................................................................................ 18
Chapter 2. Newly designed primers detect a diverse community of arbuscular mycorrhizal fungi at a remnant bunchgrass prairie ...................................... 28
Abstract ............................................................................................................ 29
Introduction ...................................................................................................... 29
Methods ............................................................................................................ 33
Results .............................................................................................................. 39
Discussion ........................................................................................................ 45
Conclusions ...................................................................................................... 50
Acknowledgements .......................................................................................... 51
References ........................................................................................................ 52

TABLE OF CONTENTS (Continued)
Page
Chapter 3. Host species identity but not spatial distribution influences arbuscular mycorrhizal community in plant roots ......................................................... 66
Abstract ............................................................................................................ 67
Introduction ...................................................................................................... 67
Methods ............................................................................................................ 70
Results .............................................................................................................. 76
Discussion ........................................................................................................ 79
Conclusions ...................................................................................................... 87
Acknowledgements .......................................................................................... 88
References ........................................................................................................ 89
Chapter 4. Host species identity and community context influences arbuscular mycorrhizal fungal community in plant roots ............................................ 102
Abstract .......................................................................................................... 103
Introduction .................................................................................................... 103
Materials and Methods ................................................................................... 107
Results ............................................................................................................ 114
Discussion ...................................................................................................... 116
Conclusions .................................................................................................... 122
Acknowledgements ........................................................................................ 123
References ...................................................................................................... 124
Chapter 5. Conclusions ....................................................................................... 137
Summary of findings ...................................................................................... 137

TABLE OF CONTENTS (Continued)
Page
Comparison of results from Chapters 3 and 4 ................................................ 139
Future directions............................................................................................. 142
References ...................................................................................................... 146
Bibliography ........................................................................................................ 149

LIST OF FIGURES
Figure Page
1.1 Illustration of characteristic structures of arbuscular mycorrhizal fungi ..... 25
1.2 A representative phylogenetic tree showing the relationships of the orders and genera within the Glomeromycota to each other ................. 26
1.3 Representation of the complex network of arbuscular mycorrhizas
(AM) ...................................................................................................... 27
2.1 Primer binding sites of the newly designed primers. ................................... 56
2.2. A consensus phylogenetic tree showing the relationship of the
Kingston Prairie operational taxonomic units to other
Glomeromycota sequences .................................................................... 60
2.3. Comparison of published primers targeting large subunit (LSU) rDNA to Glomeromycota sequences found at Kingston Prairie Preserve ........ 62
2.4. Number of Kingston Prairie sequences in each order that are predicted to be amplified by published primers ..................................................... 65
3.1. Phylogenetic tree showing the Kingston Prairie operational taxonomic units (OTUs) used in the community analysis. ...................................... 95
3.2. The mean number (± S.D.) of AMF operational taxonomic units detected per host individual within the plant species studied . .............. 97
3.3. Two-way dendrogram of AMF OTU occurrences in individual plants ....... 98
3.4. The total number of OTUs detected within plants of different species, separated by AMF family .................................................................... 100
3.5. The network of interactions between plants (left) and arbuscular mycorrhizal fungi operational taxonomic units (right) ........................ 101
4.1. Typical root systems at the end of the experiment: a) Achillea millefolium , b) Prunella vulgaris , c) Lomatium bradshawii , and d)
Erigeron decumbens ............................................................................ 130
4.2. Rarefaction of sequencing effort showing the expected number of
OTUs detected with the sample sizes on the x-axis ............................. 131

LIST OF FIGURES (Continued)
Figure Page
4.3. A consensus phylogenetic tree showing the Kingston Prairie operational taxonomic units (OTUs) used in the community analysis ................................................................................................. 132
4.4. Richness of AMF communities (mean ±1 S.E.) in roots of 3 different host species ( Achillea millefolium , Erigeron decumbens , Prunella vulgaris ) grown individually or in community with other species ...... 134
4.5. Total AMF richness in pots as measured by mean number (±S.E.) of
OTUs detected within a pot for Prunella vulgaris grown individually, all plants ( Prunella vulgaris , Achillea millefolium and
Erigeron decumbens ) grown individually, and plants grown in community ........................................................................................... 135
4.6. Two-way dendrogram of AMF OTU occurrences in individual plants ..... 136

LIST OF TABLES
Table Page
2.1. PCR and cloning success rates ..................................................................... 55
3.1. Primers used in the PCR of individual operational taxonomic units
(OTUs) in plant root DNA samples. ...................................................... 93
3.2. Paired comparison values from multi response permutation procedure
(MRPP) of root AMF community composition between plant species with p-values < 0.05.. ................................................................ 94
4.1. Primers used in the PCR of individual operational taxonomic units
(OTUs) in plant root DNA samples. .................................................... 128
4.2. Significantly different AMF communities between host groups based on MRPP pairwise comparisons of AMF community dissimilarities ....................................................................................... 129
5.1. The proportion of individual occurrences from different
Glomeromycota orders found in A. millefolium and P. vulgaris plants from the field (Chapter 2) and the proportion of
Glomeromycota orders found overall in field plants (Chapter 2) and in experimental plants (Chapter 4) ................................................ 148

For Abby and Jessica

Drivers of Arbuscular Mycorrhizal Fungal Community Composition in Roots: Hosts,
Neighbors, and Environment
Chapter 1: Introduction
Organization of Introduction
In this dissertation, I investigate different aspects of the arbuscular mycorrhizal symbiosis. I begin this chapter by giving background information on the symbiosis, including the origin of the symbiosis, the biology and ecology of arbuscular mycorrhizas, and functional differences in arbuscular mycorrhizal fungi (AMF). I then explore the current knowledge regarding the diversity of AMF and the use of molecular tools as a part of understanding that diversity, which are topics of Chapter
2. In the last sections, I discuss patterns in the pairings of host and symbiont, the subject of Chapters 3 and 4, and include some background on biological mechanisms that might be involved in that process.
Origins of mycorrhizal symbiosis
The mycorrhizal symbiosis formed between terrestrial plants and mycorrhizal fungi is as old as terrestrial plants themselves. Of the land plants, 80% of plant species and 92% of plant families are mycorrhizal. In the largest of the plant groups, the
Angiosperms, the percentages are even higher: 85% of species and 94% of families.
Some of those plants species, about 72%, are obligately mycorrhizal, that is partnering in the symbiosis is essential to their survival, while about 12-13% are facultatively mycorrhizal (Wang & Qiu 2006). The ancestral type of mycorrhizal association in land plants is with the arbuscular mycorrhizal fungi (AMF) (Wang & Qiu 2006).

There are fossils of arbuscules in the protostele of the 400 million year old Devonian
2 protracheophyte Aglaophyton major (Remy et al. 1994) and 460 million year old fossil evidence of AM fungal spores and hyphae from the mid-Ordovician, a time before terrestrial plants had evolved (Redecker et al. 2000). Some suggest that association with AM fungi was necessary for plants to colonize land, a hypothesis which is supported by an abundance of AM fungal associations in non-vascular plants (Wang
& Qiu 2006, Winther & Friedman 2008).
Biology and ecology of arbuscular mycorrhizas
The arbuscular mycorrhizal fungi derive their name from highly branched hyphal structures known as arbuscules formed intracellularly in the cortex of plant roots, although only about 80% of AMF species actually form arbuscules (Fig. 1.1).
The foundation of the symbiosis is considered to be the exchange of nutrients between plant and fungus, presumed to occur primarily at the arbuscular-plant cell interface.
Plants provide the fungi with fixed carbon, with up to 20% of a plant’s fixed carbon going to the fungal partner (Jakobsen & Rosendahl 1990). The fungi are obligate symbionts as they have no other known mechanism of obtaining carbon. In addition to forming arbuscules, the fungi have other hyphae running through the root cortex, and most AMF form structures in roots called vesicles that store lipids. The fungi also form extensive hyphal networks in the soil, and because the hyphae can extend farther and are much smaller than plant root hairs, they can access smaller soil pore spaces and thus provide increased access to nutrients and water (Smith & Read 2008).

Assisting in their provision of nutrients is their unique biochemistry which allows them to mobilize otherwise immobile elements. The primary nutrient provided by the
3 fungi is phosphorus, which is relatively immobile in many soil types, but the fungi also transfer other nutrients such as N, Zn, Mg, and Ca (Bucher 2007, Sikes et al.
2010).
AMF affect plants in more ways than just supplying nutrients. The presence of
AMF impacts the movement of water through plants and can provide protection from the stress incurred by moderate drought conditions (Augé 2001, Ruiz-Sánchez et al.
2010). AMF can affect rates of leaf herbivory (Gange & West 1994) and the growth responses of plants affected by herbivory (Bennett & Bever 2007). Colonization of roots by AMF results in declines in root pathogen growth (Azcón-Aguilar & Barea
1996, Borowicz 2001, Cordier et al. 1998). Under saline soil conditions, plant growth can be improved by AMF colonization (Al-Karaki 2000, Evelin et al. 2009, Ruiz-
Lozano et al. 1996). AMF can increase growth rates of plants grown in soils with heavy metals by immobilizing the metals in particular areas of root tissues
(Hildebrandt et al. 1999, Kaldorf et al. 1999).
Given that AMF affect plants in so many ways, it is not surprising that AMF can have significant impacts on plant communities. AMF can impact plant seedling success (Kytöviita et al. 2003, van der Heijden 2004), alter the outcome of plant competition (Scheublin et al. 2007), and ultimately increase plant community biodiversity and productivity (van der Heijden et al. 1998b). AMF also influence ecosystem processes through various pathways (Rillig 2004). Some of these functions

are directly related to their effects on plants: as mentioned above, AMF impact both
4 the productivity and diversity of plant communities (van der Heijden et al. 1998a). The transfer of fixed carbon from plants to AMF results in a substantial sink of carbon to the soil (Olsson & Johnson 2005), making AMF key in nutrient cycling. The presence of extensive AMF hyphal networks in soils influences soil aggregation through biochemical, biophysical and biological processes (Rillig & Mummey 2006). One ecosystem level consequence of increased soil aggregation by AMF is a reduction in nutrient loss from soil (van der Heijden 2010). Also, AMF can make important contributions to soil organic matter upon their decay (Wilson et al. 2009).
Functional differences in AMF
While the aggregate effects of AMF are important, the group consists of many different species that have a variety of functional differences. For example, AMF species differ in the rate and extent of colonization of roots and soil (Hart & Reader
2002a, Hart & Reader 2005), number of spores produced per root weight under various soil nutrient conditions (Douds & Schenck 1990), and how much of their lipids are allocated to storage (van Aarle & Olsson 2003). They differ in nutritional characteristics such as their mechanisms of phosphate metabolism (Boddington &
Dodd 1999) and their growth and nutrient uptake responses to localized nutrient patches (Cavagnaro et al. 2005). They differ in the rate of nutrient transfer such as the phosphorus inflow per unit mycorrhizal root length (Jakobsen et al. 1992) and the amount of phosphorus transferred per unit area of interface (Dickson et al. 1999).

The functional differences in AMF species are manifested in their impacts on their hosts. For example, AMF species differ in their effect on phosphorus
5 concentrations in leaf tissue (van der Heijden 2004) and in how they affect plant growth responses under different soil phosphorous conditions (Graham & Abbott
2000). The identity of AMF partners influences the degree of effect of AMF seen on plants grown under some of the previously mentioned conditions such as with herbivory (Bennett & Bever 2007), with pathogens such as nematodes (Zhang et al.
2008), and in soils containing heavy metals (Tonin et al. 2001) or salt (Ruiz-Lozano et al. 1996). AMF community composition also influences the outcome of competitive interactions between plant species (Wagg et al. 2011).
In fact, plant responses to different species of AMF vary with the particular combination of the plant and AMF species in the partnership. In a classic study,
Klironomos (2003) examined the effect of one AMF species ( Glomus etunicatum ) on
64 different plant species and also the effect of different combinations of AMF and plant species (with all combinations of 10 AMF species and 10 plant species). The relative change in total biomass in mycorrhizal plants compared to non-mycorrhizal controls in the two experiments ranged from -46% to +48% and from -49% to +46%, respectively. Although the symbiosis between plants has historically been classified as a mutualism, these results indicate that the relationship may not always provide a net benefit to the plant. Given that the effects of different AMF on plants varies from positive to negative, some prefer to think of the symbiosis as a continuum that ranges from parasitism to mutualism (Johnson et al. 1997). Others question whether

describing the relationship as sometimes parasitic is appropriate when other unmeasured benefits may be conferred to hosts even if whole-plant measures such as
6 total biomass are negatively impacted by some AMF (Smith & Smith 2012).
Taxonomic diversity of AMF and molecular tools for identification
Despite the ecological importance of AMF and the recognition that AMF taxa have relevant functional differences, we lack basic knowledge of the diversity of
AMF. First, as soil-dwelling, microscopic organisms, they live hidden from the human eye, so that discovery of species diversity requires more sampling effort than most above-ground macroscopic organisms. Second, delineation of species in the AMF poses unique challenges. Historically, species were delineated based on the morphological species concept, focusing primarily on spore morphology. By those standards, there are approximately 230 named, described AMF species. But the use of spore morphology is dogged with a variety of problems, including ontogenetic differences in spore characteristics and dimorphism in the spores of some species
(Morton & Redecker 2001, Rosendahl 2008). Because no sexual reproduction has ever been observed in the AMF, the biological species concept cannot be employed for these organisms. As with most other taxonomy, the development of molecular tools, particularly DNA sequencing, has renovated phylogenetic classifications in the AMF.
The use of DNA sequences for building phylogenies of AMF has the same challenges as for other groups of organisms and some unique challenges.

7
AMF hyphae are aseptate and coenocytic, that is they are multinucleate, with perhaps hundreds of haploid nuclei in a single connected cytoplasm. AMF spores, too, generally contain hundreds of nuclei. Molecular studies have found an atypically high rate of intra-individual variation in spores, and there is some controversy over whether all the genetic diversity is contained within genetically similar nuclei, or homokaryons, or whether the diversity is spread over divergent nuclei, or heterokaryons (Hijri & Sanders 2005, Kuhn et al. 2001, Pawlowska & Taylor 2004).
Also, although there is no direct observational evidence for sexual reproduction in the
AMF, levels of genetic diversity are greater than would be expected in completely asexual organisms (Vandenkoornhuyse et al. 2001). At least some of this enhanced genetic diversity can be explained by genetic exchange that occurs when hyphae from genetically different individuals join via anastomosis (Croll et al. 2009). Rates of anastomosis vary by AMF genus and species and by whether hyphae are in symbiotic versus asymbiotic phases (Purin & Morton 2011). Whatever the cause, intraspecific genetic diversity varies between species (Rodriguez et al. 2005), which complicates the determination of appropriate DNA identify cut-off values for molecular delineation of species.
Despite this complicated genetic structure in AMF, use of DNA sequences has provided great insights into the taxonomic relationships within the AMF. AMF are a monophyletic group of fungi given phylum status: the Glomeromycota (Schüßler &
Walker 2001). The precise location of AMF on a common fungal phylogenetic tree is as yet unclear, with molecular studies based on nuclear ribosomal RNA gene (rDNA)

sequences supporting the Glomeromycota as a sister group to the Dikaryon, which
8 includes the Ascomycetes and Basidiomycetes, but studies of mitochondrially encoded proteins concluding it is not a sister group to but basal to the Dikaryon (Lee & Young
2009). All known species within the Glomeromycota are classified as arbuscular mycorrhizal forming fungi except one unique species, Geosiphon pyriformis , which lives symbiotically with the cyanobacteria in the genus Nostoc . The Glomeromycota consists of four orders: Glomerales, Diversisporales, Archaeosporales, and
Paraglomerales (Fig. 1.2), with the first two of those orders containing roughly 100 described species, whereas the latter contain just 11 and 3 described species, respectively (A. Schüßler’s
Glomeromycota phylogeny, http://www.lrz.de/~schuessler/amphylo/; accessed 19 June 2012). A greatly overhauled taxonomy of the Glomeromycota, with many newly renamed genera, has recently been proposed (Schüßler & Walker 2010).
The use of molecular tools has not only helped with taxonomic delineation, it also has added to the toolbox for surveying for AMF. Before molecular methods of studying AMF were developed, AMF communities were surveyed by examining spores – either directly from the soil of interest or by taking environmental samples as inoculum for pot cultures from which the resulting spore community was examined.
The use of spores has the advantage of requiring less specialized equipment than most molecular work, but identifying spores to the species level is a time consuming skill to develop, and, as previously stated, spore morphology as a basis of species identification is complicated by ontogenetic differences in spore characteristics and

dimorphism in the spores of some species (Morton & Redecker 2001, Rosendahl
2008). Another disadvantage of using spores to evaluate community composition is
9 that as a dormant structure, spores may not quantitatively or qualitatively reflect the distribution of AMF species active in nearby roots or in the rhizosphere (Hempel et al.
2007). Because sporulation occurs in different AMF species at very different rates,
AMF with active hyphal networks but low rates of sporulation would be underrepresented in spore based community analysis.
By sampling for DNA in soil, one can survey for presence of both hyphal and spore structures. Sampling for DNA in roots is the only tool currently available for identifying below the genus level the AMF associating with roots in mixed plant and
AMF communities. Through phylogenetic analysis of sequences from environmental samples, individual sequences can be assigned to groups and their relationships to sequences from morphologically described species can be determined. Groupings of sequences meant to roughly represent species level delineations have been called
“phylotypes” (Vandenkoornhuyse et al. 2002), “operational taxonomic units” =
“OTUs” (Pivato et al. 2007), and “virtual taxa” (Öpik et al. 2010). However, even using molecular tools, the choice of sampling method impacts the results (Sýkorová et al. 2007b). Although it is also an imperfect system, the use of DNA sequences and phylogenetic tools, especially in environmental studies where many undescribed AMF species may be present, expands the available resources for addressing many questions.

10
As data from more environmental surveys for AMF become available, we are gaining ground on understanding diversity levels and distribution patterns. One clearly emerging pattern is that most new studies of environmental samples find sequences not matching any of those in sequence databases at what would likely be species level similarity. Because of this, many have proposed the true diversity of AMF species to be much higher than that determined from morphologically described species. Ö pik et al. (2010) investigated the worldwide distribution of AMF by analyzing 282 “virtual taxa” based on SSU rDNA phylogenetic analysis using sequences compiled from 105 studies. They found one, five, 17, 26, 62 and 168 virtual taxa had been detected on six, five, four, three, two and one continents, respectively. Of those found on just one continent, 95 were found at just one location. They also found that individual virtual taxa that were detected in a wide range of plants were more likely to be geographically widespread. In contrast, other studies show that often times AMF communities in a particular ecosystem have locally dominant AMF species that do not show widespread patterns of dominance (Dumbrell et al. 2010b, Rosendahl 2008). It is noted that these are emergent patterns only, as the field is far from having exhaustively sampled AMF worldwide. In Chapter 2, I contribute to our understanding of AMF diversity in natural environments by surveying the AMF in the roots of 12 plant species at a remnant bunchgrass prairie in Oregon.
One complicating factor in studies using molecular methods to identify AMF is that to date, different studies have used different parts of the rDNA, either the SSU,
ITS or LSU. One great disadvantage to this is the difficulty it creates in comparing

11 studies that have identified AMF types by different regions of the rDNA, particularly because so many of the sequences detected in environmental studies do not match described, sequenced species. As an example, the Ö pik et al. (2010) study referred to above used SSU sequences to analyze worldwide distribution patterns, yet the many environmental surveys that have used the LSU region to type AMF could not be included in that study because the SSU and LSU sequences do not overlap. Another problem with many molecular-based environmental surveys to date is that primers used to amplify Glomeromycotan sequences often failed to amplify all orders in the phylum. In more recent years, improved primers have been designed (Krüger et al.
2009, Lee et al. 2008). In Chapter 2, I describe primers I designed to amplify
Glomeromycota rDNA and analyze the likelihood that previously designed primers would capture the full diversity of AMF that I detected in my field survey.
The pairing of AMF and plants
In Chapters 3 and 4, I investigate factors that potentially influence the pattern of associations between plants and AMF, including host identity, geographic distance, soil types, and community context. What governs the pairing of the partners between plant and AMF species, including to what degree random vs. non-random processes contribute, is still an open question. Non-random patterns in the pairings between
AMF and plants could result directly from interactions between the organisms but also from environmentally and biologically based factors that cause co-variation in their distribution. Correlations between plant and AMF species in characteristics such as

12 abiotic habitat preferences (Dumbrell et al. 2010a), life histories (Sýkorová et al.
2007b), or local abundances (Dumbrell et al. 2010b) could yield non-random patterns in partner associations even without selection mechanisms exerted by either partner.
Yet experimental studies with controlled conditions suggest interactions between partners do factor into AMF community composition in roots (Helgason et al. 2002,
Scheublin et al. 2004).
Because there are variations in functional characteristics of AMF and differences in the effect of AMF on plant species, selection pressure could exist for plants to associate with those AMF that most benefited them, either in current time or in evolutionary time. Likewise, even though AMF are obligate biotrophs and the worst plant partner is better than none, AMF would be expected to gain increased fitness from associating with plants from which they derive the most benefit. Although less is known about variations in plant contribution to AMF fitness because of the difficulties in measuring that parameter, at least one aspect of fungal fitness, sporulation rate, has been shown to vary with host identity (Bever et al. 1996).
Various terms have been used to describe pairings regulated in some way by one or both partners, with “specificity” (Smith & Read 1997),“selectivity” (Helgason et al. 2002, Scheublin et al. 2004), and “partner preference” (Vandenkoornhuyse et al.
2002) being the most common. Here, I use the definition provided by Taylor et al.
(2002) for specificity: it is “the phylogenetic breadth of the mycorrhizal associations of that particular plant or fungus.” Put another way, specificity represents a continuous axis and the positions of organisms on that axis represent their relative specificity.

13
Both the terms selectivity and partner preference are often used in ways that infer that one or the other partner is selecting or having the preference, even when the particular research being discussed provides no evidence for which, if any, partner is acting as the selector. Another confusing aspect is the use of the term “host preference” to sometimes mean selection by plant of AMF partner, e.g. Aguacil et al. (2011),
Sýkorová et al. (2007a) and Vandenkoornhuyse et al. (2002), and sometimes to mean selection by AMF of plant partners, e.g. Johnson et al. (2005). Here, I use the terms specificity or selectivity when referring to non-random pairings, acknowledging that when using the latter it may be without direct evidence that one or both partners is actively selecting.
For some time, it was considered that there was little to no specificity within the symbiosis (Smith & Read 1997). This idea was based on the very large ratio of mycorrhizal plant species to described AMF species, likely on the order of 1,000:1.
Clearly with a ratio like that, prevalent one-to-one species specificity would be impossible. In the last dozen years, numerous studies have shown that associations between partners are not entirely random, including both observational (Helgason et al. 2002, Vandenkoornhuyse et al. 2002) and experimental studies (Eom et al. 2000,
Helgason et al. 2002). Except for some unique cases of specialization (Bidartondo et al. 2002), the evidence for selectivity is ‘fuzzy’ in many studies, and some researchers suggest that plants vary in their degree of selectivity (Scheublin et al. 2004). I investigate partner specificity as a cause of non-random AMF community structure in roots in Chapters 3 and 4.

Mechanisms of partner selection
Partner selection could occur by various mechanisms. One possibility is that
14 plants or AMF have a-priori partner sets with which they will or will not associate. In that scenario, one would expect interplay of recognition molecules and receptors between organisms that would either promote or inhibit the establishment of symbiosis with each other. Much work has been done to identify the so-called “Myc factors” secreted by AMF that promote root branching and AM formation, and only very recently have fungal lipochitooligosaccharides been identified as possible components of these factors (Maillet et al. 2011). In one fungus studied, a mixture of four lipochitooligosaccharides was found (Maillet et al. 2011), but it is as yet undetermined if other AMF species produce the same mix or unique mixes of lipochitooligosaccharides. It is also unknown what receptors for these molecules plants have and how they would work in recognizing AMFs (Bonfante & Requena
2011). Therefore, it is as yet unclear whether different AMF might have differences in signaling molecules that could identify them to their potential hosts.
Another possibility is that partner selection occurs after the fungus colonizes the host. In that case, it may be that one or both partners “choose” whether or not to maintain the relationship depending on the amount of benefit they are receiving. Kiers
& van der Heijden (2006) propose and review evidence that the host might have mechanisms for controlling aspects of symbiont growth at the cellular, organismic or whole root level. Very recent studies show that “plants can detect, discriminate, and reward the best fungal partners with more carbohydrates” (Kiers et al. 2011). The

plant, therefore, may influence its AMF community by differential rewards to its different partners. Fungi that are in concurrent relationships with other plants could
15 exit from hosts of less benefit. Such processes could lead to non-random pairings between plant and fungal species if there was consistency in benefits within species.
Plants and AMF usually live in complex communities with each other (Fig.
1.3). As such, even if hosts exert control over their root AMF communities, interactions with neighboring plants may have significant impacts on those root communities as well (Hawkes et al. 2006, Mummey & Rillig 2006). For the research presented in Chapter 4, I grew plants both individually and in community and compared resulting root AMF communities to determine if host influences seen outside of a plant community remain when a plant grows in community.
Broader impacts
There are many reasons why it would be valuable to understand if AMF and plants form partnerships randomly or non-randomly. The AM symbiosis affects many aspects of plant fitness, as previously described, and the degree of effect often varies with the particular combination of AMF and plant species. Incorporating an understanding of the patterns in plant-AMF associations with knowledge of the effects of the symbiosis on various plant measures could improve our understanding of the role of the symbiosis in shaping plant communities. At a more theoretical level, understanding partner pairing patterns could provide insight into the evolutionary mechanisms that have allowed for the long term maintenance of this symbiosis as a

mutualism. Because nutrients can flow between plants through their mycorrhizal connections (Bidartondo et al. 2002, Mikkelsen et al. 2008), if some plant and AMF
16 combinations are more or less likely, this would similarly affect the chance that nutrients may be transported between them.
Lastly, if AMF are to be used as a tool in plant conservation, as has been proposed by some, e.g. Bothe et al. (2010), knowledge of processes governing plant-
AMF combinations could help make their use more effective. To that end, I chose to incorporate two federally listed endangered plants into the experiment of Chapter 4 to see if they formed unique AMF communities. If plants that are conservation targets benefit from associations with particular AMF, then providing those AMF could help in the introduction and maintenance of plant populations. I take the first step in that by determining what AMF these endangered plants associate with and whether those associations are unique.
Summary
In this dissertation, I explore the arbuscular mycorrhizal symbiosis through studies on members of the plant and fungal communities at a remnant bunchgrass prairie, Kingston Prairie Preserve, in the Willamette Valley, Oregon, U.S.A. For my first study (Chapter 2), I conducted a field survey to identify and describe the diversity of AMF occurring in the roots of plants at the prairie. As part of that survey, I designed new primers for selective amplification of the Glomeromycota large subunit
(LSU) ribosomal gene. Primers that had been published and used previous to the start

17 of my study had problems with not amplifying all Glomeromycota while also amplifying non-Glomeromycota fungi. Since that time, a variety of new primers have been designed and used by other researchers, and I use the sequences from my study to assess what portion of the diversity of the Kingston Prairie sequences likely would be amplified by different primers. In the second study (Chapter 3), I examine the community composition of AMF in roots of individual plants to determine if it correlates with host identity, geographic distance, or soil characteristics. In the third study (Chapter 4), I test the effect of host identity and community context on AMF root community composition through a growth chamber experiment with plants and
AMF from Kingston Prairie Preserve. I include in that study two endangered plants. I conclude in Chapter 5 with a discussion tying the results of Chapters 2-4 together and presenting thoughts on future experiments that could help elucidate the role of partner choice in the arbuscular mycorrhizal symbiosis.

18
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24 van der Heijden M.G.A., Klironomos J.N., Ursic M., Moutoglis P., Streitwolf-Engel R., Boller
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Extraradical hypha
Spore
Epidermis
Hypodermis
Cortex
Intraradical hypha
Arbuscule
Vesicle
Figure 1.1.
Illustration of characteristic structures of arbuscular mycorrhizal fungi. The design of the illustration is adapted from http://mycorrhizas.info/vam.html, accessed 15
August 2012.

26
Funneliformis
Glomus
Rhizophagus
Sclerocystis
Claroideoglomus
Acaulospora
Redeckera
Diversispora
Scutellospora
Racocetra
Gigaspora
Scutellospora
Pacispora
Ambispora
Geosiphon
Archaeospora
Paraglomus
Figure 1.2.
A representative phylogenetic tree showing the relationships of the orders and genera within the Glomeromycota to each other. A few genera are not shown.
Scutellospora appears twice to show its current paraphyletic status. The relationships shown are based on those established in Krüger et al. (2012).

27
Figure 1.3. Representation of the complex network of arbuscular mycorrhizas (AM).
Single AM fungi can colonize multiple plants, and individual plants can host multiple
AM fungi. Different AM fungal taxa are represented as different colors.

Chapter 2. Newly designed primers detect a diverse community of arbuscular mycorrhizal fungi at a remnant bunchgrass prairie
Wendy S. Phillips
To be submitted to:
Mycorrhiza
Berlin/Heidelberg, Germany
28

Abstract
Despite the importance of arbuscular mycorrhizal fungi (AMF) to ecosystem
29 functioning, our knowledge of AMF taxonomic diversity is incomplete. While environmental surveys for AMF have been conducted in many ecosystem types, there remain un-sampled or under-sampled ecosystems which may hold undiscovered AMF diversity. To best detect all AMF taxa in molecular environmental surveys and improve our understanding of AMF taxonomic diversity, well-designed primers must be used. I designed new primers that target nuclear rDNA sequences for the detection of all AMF taxa and used those primers to survey the AMF in plant roots from a remnant bunchgrass prairie. I then performed in silico tests to predict the ability of previously published AMF primers to amplify all the sequences I had detected. With a single sampling time point, a total of 36 AMF operational taxonomic units (OTUs) were detected from the prairie, with a higher proportion from basal Glomeromycota than usual in environmental samplings. The predicted ability of previously published primers to detect all 36 OTUs was highly variable, with predicted detection rates ranging from 64% to 100%. Analyses with the presented sequence data provide guidance for selection of primers to use in future studies. Use of primers that target all
AMF will provide improved ability to detect complete AMF diversity, thereby yielding improved data for use in ecological analysis.
Introduction

Arbuscular mycorrhizal fungi (AMF), which are thought to have coevolved with land plants and now live symbiotically with 70-80% of angiosperms (Brundrett
30
2009, Wang & Qiu 2006), perform a variety of ecosystem functions. The AMF-plant symbiosis plays an important role in plant nutrient acquisition, and the transfer of fixed carbon from plants to AMF results in a substantial sink of carbon to the soil
(Olsson & Johnson 2005), making AMF key in nutrient cycling. AMF impact both the productivity and diversity of plant communities (van der Heijden et al. 1998a). AMF also affect plant herbivory (Klironomos et al. 2004) and plant susceptibility to root pathogens (Wehner et al. 2010). In addition to impacting plants, extraradical AMF hyphae reduce nutrient loss from soil, increase soil aggregation via excretion of glomulin (van der Heijden 2010), and add to soil organic matter upon their decay
(Wilson et al. 2009).
Despite the ecological importance of this group of fungi, there are large gaps in our knowledge of AMF taxonomic diversity. Over 400 million years of evolution within the phylum Glomeromycota (Schüßler & Walker 2001), to which all AMF belong, has produced AMF currently classified into four orders: Glomerales,
Diversisporales, Archaeosporales, and Paraglomerales (Schüßler & Walker 2010), with approximately 100, 100, 11 and 3 morphologically described species in each order, respectively (A. Schüßler’s Glomeromycota phylogeny, http://www.amfphylogeny.com; accessed 7 December 2010). Because species description of AMF is largely based on spore morphology, described AMF species are limited to those from which spores have been collected. Molecular studies, which can detect the fungi even

31 if they are not sporulating, indicate that unidentified AMF likely outnumber described species (Öpik et al. 2010).
The gaps in our knowledge of AMF taxonomic diversity transmit to gaps in our understanding of the ecological roles of AMF. Functional diversity, including differences in the extent of pathogen protection and rates of hyphal growth in root tissue versus soil, has been shown to vary between AMF taxonomic groups (Hart &
Reader 2002b, Powell et al. 2009, Wehner et al. 2010). To better understand the functional role of AMF communities in ecosystems, we must have better knowledge of the taxonomic and functional diversity of those AMF communities. Additionally, knowledge of AMF diversity is integral to understanding specificity between plant and
AMF partners. Historically both plants and AMF were considered to be generalists in terms of partner pairing, particularly because so few AMF species had been described while so many plant species exist. Research in the last decade has provided evidence for some degree of partner specificity (Helgason et al. 2002, Vandenkoornhuyse et al.
2003). With improved knowledge of the AMF taxa available as partners, we can better investigate patterns of specificity in the symbiosis.
Only molecular tools allow identification of AMF taxa to the species level within roots. Before the advent of molecular techniques, environmental AMF were identified via microscopic examination of spore morphology, a technique that is still employed. A limitation of using spores to identify AMF community constituents of plant roots is that soil spore populations may have little relation to the AMF actively colonizing root tissue (Hempel et al. 2007). Yet molecular tools have their own

limitations. When using PCR as a molecular detection technique, the quality of the
32 results depends on how well the primers detect all members of the subject group while excluding non-group-members. Although the study of AMF diversity has been much enhanced by the use of molecular tools, many of the historically used primers are imperfectly discriminatory, either not detecting all taxa in the Glomeromycota or additionally detecting non-arbuscular mycorrhizal (AM) fungi (Krüger et al. 2009, Lee et al. 2008). To address this problem of imperfect discrimination of AMF by PCR primers, as others have recently done (Krüger et al. 2009, Lee et al. 2008), I developed primers to amplify all Glomeromycota taxa while discriminating between the
Glomeromycota and all other taxa.
Much of the sequencing for phylogenetic analysis and nearly all molecular environmental sampling of AMF to date has targeted genomic DNA encoding ribosomal RNA (rDNA): the small subunit (SSU); the internal transcribed spacer
(ITS) region; and/or the large subunit (LSU) regions. Given that the majority of sequence information for AMF in sequence databases is for some section of rDNA, it remains the most informative region for designing all-inclusive AMF primers and for identifying ‘unknown’ sequences obtained in environmental sampling. I examined existing databases of the entire SSU, ITS and LSU regions of rDNA to find regions of shared derived sequence identity within the Glomeromycota that diverged with sequences from other fungal phyla.
I report here on the primers I developed. I then used the primers in an environmental sampling of a remnant bunchgrass prairie. I report on the phylogenetic

33 diversity detected at the prairie and how it compares with that found in other studies. I also examine whether other published primers would likely detect the same diversity of AMF as found in this study.
Methods
Site description
Grasslands are one of the most critically endangered biomes globally
(Hoekstra et al. 2005) and the Willamette Valley prairies of Oregon are no exception, having been reduced to 0.1% of their historical range (Noss et al. 1995). In June 2007, plant roots were collected from a 61-hectare remnant prairie owned by the Nature
Conservancy, Kingston Prairie Preserve (approximately 44°46’N, 122°44’W; elevation 225 m). This prairie includes both wet meadow and dry upland vegetation, is dominated by bunchgrasses such as Festuca idahoensis Elmer ssp. roemeri (Pavlick)
S. Aiken and Poa secunda J. Presl, and is home to two federally listed endangered plant species: Lomatium bradshawii (Rose ex Mathias) Mathias and Constance and
Erigeron decumbens Nutt. The climate is Mediterranean, with cool, wet winters and warm, dry summers and an average annual precipitation of 133 cm. The soil is generally shallow clay loam to loam with numerous cobblestones overlaying basalt bedrock. Although soil was not sampled at the time of root collection, the range of values for three soil samples taken in June 2009 were: nitrate N = 5-16 ppm, P = 4-9 ppm, soil pH = 5.5-6.1, organic matter = 3.5-4.6%.

34
Root collection
Plants were sampled by first assigning random GPS points within the prairie to each of 12 host species: Achillea millefolium L., Brodiaea coronaria (Salisb.) Engl.,
Prunella vulgaris L., Zigadenus venenosus S. Watson, Eriophyllum lanatum (Pursh)
Forbes, Festuca idahoensis Elmer ssp. roemeri (Pavlick) S. Aiken, Poa secunda J.
Presl, Potentilla gracilis Douglas ex Hook, Delphinium nuttallii A. Gray, Perideridia gairdneri (Hook. & Arn.) Mathias, Leucanthemum vulgare Lam., and Sidalcea campestris Greene. At the given GPS point, the closest plant or cluster of plants of that species was identified. If the closest plant was part of a cluster, a flag was thrown blindly in the direction of the cluster, and the closest plant to the flag was sampled.
The actual GPS location of the sampled plant was recorded. The rhizosphere to approximately 15-20 cm depth and the stem material of an entire small plant or a portion of larger plants were collected and stored in a plastic cup at room temperature until processing. Root systems were washed thoroughly and only root segments that could be traced back to the stem were collected. If the collected root system appeared to have less than approximately 300 mg of root tissue, the entire root system was collected. If the root system collected was larger than that a subset of fine root material was collected haphazardly with an attempt to sample representative and various sections of the root system. Within 3 days of collection, all root tissue was washed, sampled, and frozen.
DNA extractions

35
DNA was extracted using the DNAeasy Plant Mini-kit (Qiagen, Valencia, CA,
U.S.A.). Root tissue ranging from 14 to 216 mg (mean 71 mg) wet weight was used in the extraction. Root tissue and three 3 mm glass balls in a 2 ml cryovial freshly pulled from liquid nitrogen was subjected to grinding for 30 s using a Mini-Beadbeater-1
(Biospec, Bartlesville, OK, U.S.A.) set to 4200 rpm. Following a brief centrifugation,
Buffer AP1 was added and the tubes were placed in the Beadbeater for another 10 s.
Further extraction steps followed the manufacturer’s protocol. Final DNA concentrations in the extracts were measured using a NanoDrop ND-1000 spectrophotometer (Thermo Scientific, Wilmington, DE, U.S.A.). DNA was diluted to create a 0.5 ng/µl solution, 5 µl of which was used in a 20 µl PCR reaction. Unusual absorption spectra and failure of PCR indicated PCR inhibiting contaminants occurred in some samples. For those samples, DNA was further diluted until a positive PCR reaction was obtained.
Primer design
New primers were designed from existing sequences of fungal ribosomal RNA genes in regions where there was 100% conservation of nucleotides for
Glomeromycota sequences but non-conservation with non-Glomeromycota fungal sequences at the 3’ end of the primer sequence. Focus was given to the 3’ end of potential primer sequences as primers with mismatches at the 3’ end have much lower amplification efficiency (Lee et al. 2008, Sarkar et al. 1990). After surveying alignments of the entire small subunit (SSU) and large subunit (LSU) rDNA

36 sequences of multiple Glomeromycota and non-Glomeromycota fungi, only one site in the SSU and two sites in the LSU that matched those criteria were found for the sequences available at that time. To maintain consistency with the description of primer locations by Krüger et al. (2009), Rhizophagus irregulare DAOM197198 sequences are used for referencing primer location sites. I designed one forward primer in the SSU rRNA gene (AM-SSU-F: 5’-CGCGCTACACTGATGAAGT-3’ at positions 1435-1453 in accession AY635831) and two reverse primers in the LSU
(AM-LSU-Rx: 5’-GTTTGAGAATAGGTTAAGGC-3’ and AM-LSU-Ri: 5’-
GTTAAGGCTGTTTCAACCCT-3’ at positions 1193-1212 and 1181-1200, respectively, in accession DQ273790; Fig 2.1). As the reverse primers are overlapping, both the AM-SSU-F – AM-LSU-Rx and AM-SSU-F – AM-LSU-Ri primer sets yield a similarly sized product of approximately 2 kb.
Since designing the primers, new sequences have become available, and I conducted a reanalysis of the degree to which the primer sequences are identical to known sequences. A comprehensive phylogenetic analysis of the Glomeromycota was recently conducted by Krüger et al. (2012), and I compared my new primers to their sequence alignments. Those alignments contained ~110 species covering the AM-
SSU-F primer region, whereas sequences for only seven species covered the AM-
LSU-Rx and AM-LSU-Ri primer region.
Amplification and cloning of rDNA sequences

AMF rDNA was amplified from roots of individual plants in two PCR steps.
37
The 20 µl PCR reactions contained 5 µl template DNA, 1X PCR buffer, 2 mM MgCl
2
,
0.2 mM dNTPS, 2.5 µM each primer, and 1 unit of standard Taq DNA Polymerase.
An initial reaction with AM-SSU-F and AM-LSU-Rx primers was performed with cycling parameters: 2 m initial denaturing at 94°C; 30 cycles of 94°C for 30 s, 58°C for 30 s, 72°C for 2 m; and a final extension at 72°C for 10 m. That was followed by a
20 µl reaction using 1 µl of a 1/50 dilution of the first reaction as a template and primers AM-SSU-F and the semi-nested AM-LSU-Ri. PCR parameters were the same for the semi-nested reaction with the exceptions that only 26 cycles were performed and an annealing temperature of 59.5°C was used. DNA from individual plants was amplified separately, and then the AM-SSU-F – AM-LSU-Ri products from individuals of the same host plant species were pooled (Renker et al. 2006) before cloning them into the TOPO-TA vector (Life Technologies, Grand Island, NY,
U.S.A.). To pool PCR products for cloning, DNA quantities in the PCR bands were visually estimated in 1.5% agarose gels and then approximately equal amounts of
DNA from each PCR reaction were included in the pooled cloning reaction. The reactions from four plant hosts ( Potentilla gracilis , Perideridia gairdneri ,
Leucanthemum vulgare and Sidalcea campestris ) with limited root collections or few successful PCRs were pooled for a single mixed species cloning reaction.
To screen clones for sequence variation, AM-SSU-F and AM-LSU-Ri primers were used to PCR amplify inserts. PCR products of the appropriate size were digested with the restriction enzyme AluI and run on a 2% agarose gel to identify distinct

38 restriction fragment length polymorphism (RFLP) patterns. Because the 2 kb fragment included the ITS region, which is highly polymorphic in the Glomeromycota, RFLP banding patterns were complex and many different patterns were often observed within what was later determined to be a single OTU (see Results). From that screening process, two clones for RFLP types occurring more than once or the single clone of distinct RFLP types were chosen for sequencing. When the same RFLP pattern occurred in more than one cloning reaction, i.e. in more than one host, the host from which it was sequenced was chosen haphazardly. To minimize the costs associated with sequencing the whole 2 kb fragment, I sequenced only the D2 region of the LSU, for which there is abundant sequence data in public databases to use in phylogenetic analysis and which allows species level delineations (da Silva et al.
2006). For this, a universal reverse primer was designed for sequencing (AM-Seq, 5’-
GGCATAGTTCACCATCTTTC-3’ at LSU accession DQ273790 nucleotides 967-
986) that sits approximately 200 bases in from AM-LSU-Ri. Sequencing reactions were performed by the Nevada Genomics Center (Reno, Nevada, U.S.A.).
Sequence and phylogenetic analysis
Sequencing in the LSU D2 region yielded approximately 600 bases of high quality sequence. Sequences were checked for chimeras using Bellerophon (Huber et al. 2004) and for unique sequences using MOTHUR (Schloss et al. 2009). Combining an initial alignment and phylogenetic tree constructed using SATe (Liu et al. 2009) and analysis using MOTHUR (Schloss et al. 2009), operational taxonomic units

(OTUs) were defined with a 3% sequence identity cutoff using the average neighbor
39 clustering method. At least one sequence from each OTU was compared against those in public databases using a BLASTN search (http://www.ncbi.nlm.nih.gov).
I used one representative sequence for each Kingston Prairie OTU
(approximately 600 bp) for phylogenetic analysis to determine their relationship to database sequences. A selection of the consensus SSU-ITS-LSU sequences
(approximately 3200 bp) of described species from Figure 1 of Krüger et al . (2012) were used to create a robust backbone for the phylogeny. Also included was one of the closest BLASTN matches for sequences with <97% identity to a Krüger et al . (2012) sequence. Alignments were done using the MAFFT online server (MAFFT version 6; http://mafft.cbrc.jp/alignment/server/; Katoh et al. 2002) using the Q-INS-i strategy of iterative refinement. Following alignment, the ITS regions, which are highly variable in the Glomeromycota, were removed before subsequent phylogenetic analysis.
Phylogenetic analysis was performed using the PHYLIP program suite v3.69
(Felsenstein 1993). A consensus maximum likelihood tree was constructed from100 bootstrap replicates.
Results
PCR of plant root DNA
For eight of the twelve plant species, every root system yielded a PCR product within the expected amplicon size range (Table 2.1). Of the four remaining plant species, Potentilla gracilis had the fewest successful PCR reactions – with

amplification from only two out of five root systems, while the other three hosts had
40 successful amplifications in most of the roots systems used. Because root systems were not stained to ascertain mycorrhizal colonization, it is unclear if amplification failed in these samples due to lack of mycorrhizal colonization or to experimental error such as poor DNA template quality or poor primer binding.
In the first step of screening clones, the entire insert of roughly 2 kb was amplified via PCR. Varying numbers of colonies were screened for each cloning reaction (Table 2.1), generally with more screening when a cloning reaction had a lower rate of colonies containing full-length inserts. The portion of clones with inserts of the expected size varied between plant species, ranging from 37% to 100% (Table
2.1).
In total, 229 clones were sequenced. Of those, two were found to be chimeras and were discarded. Three were found to be non-AM fungi most likely in the
Mucoromycotina based on BLASTN search results. Fourteen sequences were identical to another sequence, and only non-identical sequences are reported in the subsequent analysis. All non-identical sequences have been deposited in GenBank (accession numbers JF717423-JF717633).
Comparison of new primers with Genbank sequences
Because many new sequences have been submitted to public databases since I designed these primers, I conducted a new comparison of the primers to currently available database sequences. With few exceptions the primers match Glomeromycota

sequences perfectly while mismatching at the more critical 3’ end with other fungal
41 sequences (Fig 2.1). A notable exception for the AM-SSU-F primer is a single base pair mismatch five base pairs in from the 3’ end in the four
Claroideoglomus species analyzed. Two Redeckera species had mismatches in the 5’ half of the primer site.
Polymorphisms in Acaulospora lacunosa and Funneliformis sp. WUM3 resulted in some sequences for those species having perfect identity while some had single base pair mismatches. The non-AM fungi analyzed, with the exception of Tilletiaria anomala and Taphrina wiesneri
, have either a mismatch in one of the three 3’ terminal nucleotides of or more than two mismatches throughout the AM-SSU-F primer site.
Analysis of basal plant sequences (representative sequences from hornworts and one member of each superfamily in the mosses and liverworts) indicates AM-SSU-F mismatches all analyzed plant sequences at the second to last 3’ nucleotide (Fig. 2.1c).
Public sequence databases contain LSU sequences of only seven described
AMF species that extend to the region in which I designed the two reverse primers.
For those that do, all match perfectly with both reverse primer sequences, with the exception of the recently described Diversispora celata (Gamper et al. 2009).
Although the Diversispora sequence matches the external AM-LSU-Rx primer perfectly, it shares with most of the other non-AM fungi analyzed a 3’ terminal mismatch to the nested primer AM-LSU-Ri. Two of the 11 non-AM fungi analyzed,
Spiromyces aspiralis and Mortierella verticillata , have no mismatches with the external reverse primer AM-LSU-Rx but do have a mismatch with the nested AM-
LSU-Ri primer at its final 3’ base. In contrast, Endogone pisiformis has a 3’ terminal

42 base mismatch with the external AM-LSU-Rx primer but only a single internal mismatch with the nested AM-LSU-Ri primer. Analysis of basal plant sequences
(representative sequences from at least one member of each superfamily in the mosses and liverworts) indicates both LSU reverse primers mismatch the plant sequences at their final 3’ terminal nucleotides (Fig. 2.1d).
Phylogenetic analysis
As part of assigning the sequences to OTUs, an initial phylogenetic analysis with all sequences from Kingston Prairie was constructed using SATe (Liu et al.
2009). OTU analysis was done with MOTHUR (Schloss et al. 2009) at identity levels
0.02, 0.03, 0.04, and 0.05. At the 0.02 level, many of the resulting OTUs were paraphyletic based on the SATe tree, suggestive that it is too fine an identity threshold for meaningful taxonomic differentiation. At identity level 0.03 the topography of the
SATe tree was consistent with the results of OTU analysis done, such that all sequences in an OTU were in well supported monophyletic groups, with the exception of one paraphyletic grouping: OTU 2. At the 0.04 identity level OTUs 9 and 18 collapse to a single OTU, and at the .05 identity level they join with OTU 10 in a single OTU. All other OTUs remain the same at the 0.03, 0.04 or 0.05 identity level, suggesting that 0.03 is an appropriate level for OTU definition with this data set. In total, 36 OTUs at the 0.03 identity level were found at Kingston Prairie Preserve.
Some OTUs had only a single sequence while OTU 2, the largest group, contained 48 sequences (Fig. 2.2).

analysis (Fig. 2.2), was used for BLASTN searches. Based on results of BLASTN
43
A representative sequence for each OTU, the same one used for phylogenetic searches, 31 of the OTUs shared ≥97% identity with a database sequence (for a sequence length of at least 200 nucleotides). Of those, seven shared ≥97% identity with published sequences of described AMF: three in the Glomerales, four in the
Diversisporales, and none in the Archaeosporales. Five OTUs did not match any database sequence at ≥97% identity, including three OTUs in the Glomerales (KP13,
15, 29), one in the Diversisporales (KP22), and one in the Archaeosporales (KP33)
(Fig. 2.2).
Bootstrap analysis yielded strong support for the Glomerales and
Diversisporales as monophyletic clades, consistent with recent phylogenies (Krüger et al. 2012). Although there was only modest support for the Archaeosporales as a monophyletic group, the placement of several described Archaeosporales species throughout the cluster provides support for that group as the Archaeosporales clade.
Of the 36 OTUs found at Kingston Prairie Preserve, 23 OTUs were detected in
Glomerales; eight in the Diversisporales: two in Gigasporaceae and six in
Acaulosporaceae; and five in the Archaeosporales (Fig. 2.2). There was a large monophyletic clade in the Glomerales containing twelve KP OTUs with sequence matches only to environmental sequences.
Analysis of published primers

The sequences obtained from Kingston Prairie were used in a theoretical analysis of previously published primers (located within the LSU rDNA region sequenced in this study) to determine if other primers were likely to detect all AMF sequences detected here (Fig. 2.3 and 2.4). The more recently designed LSUmBr
44 nested reverse primer set by Krüger et al. (2009) would likely detect all Kingston
Prairie sequences, although there is a mismatching nucleotide to all Archaeosporales sequences in the middle of the primer set. The LSU-Glom1 primer (Renker et al.
2003) is predicted to detect all but one Kingston Prairie sequence, although there are nucleotide mismatches in internal positions with five of the Kingston Prairie sequences. The only forward primer analyzed here, primer FLR3 (Gollotte et al.
2004), spans the same sequence as the reverse primer LSU-Glom1 but is one nucleotide longer on both sides. The FLR3 primer is predicted to detect just 64% of the Kingston Prairie sequences, failing to detect mostly sequences in the Glomerales because of a mismatch at its 3’ terminus. FLR2 (Trouvelot et al. 1999), FLR4
(Gollotte et al. 2004) and 28G2 (da Silva et al. 2006) are all designed in the same region and overlap with each other. Among those three, FLR4 is predicted to detect the most Kingston Prairie sequences but has critical mismatches with four of the
Archaeosporales sequences. Both FLR2 and 28G2 are predicted to detect approximately 70-75% of the Kingston Prairie sequences, with FLR2 failing to detect mostly sequences in the Glomerales and 28G2 in the Diversisporales and
Archaeosporales.

45
Discussion
Field survey results
From a single sampling event at Kingston Prairie Preserve, 23 OTUs were detected in the Glomerales, five OTUs in the Archaeosporales, and eight OTUs in the
Diversisporales: six in the Acaulosporaceae and two in the Gigasporaceae. No
Paraglomerales taxa were detected. A higher proportion of diversity in the basal AMF lineages was found at Kingston Prairie than in global averages. Öpik et al.
(2010) used sequences from 102 publications to characterize global distribution patterns of AMF.
Whereas 14% of OTUs detected at Kingston Prairie were in the basal
Archaeosporales, the percent Öpik et al.
(2010) recorded in that order was 4.6% of the
AMF diversity in temperate biomes and 3.9% of the total global AMF diversity. It should be noted that many of the studies used in Öpik et al .
(2010) used older primers that have been recognized to not detect all AMF taxa, making it unclear how accurately a compilation of those sequences truly describes environmental diversity.
Therefore, it is possible that either the AMF community at Kingston Prairie Preserve hosts a unique distribution of taxa from the various AMF orders or that the difference between this study and global averages is a result of historical bias in primer specificity within the Glomeromycota.
In terms of total site richness, 36 OTUs were detected at Kingston Prairie, much greater than the mean of 12.5 (±5.0 S.D.) OTUs detected per site for 12 sites included in a review of the AMF diversity in grassland ecosystems (Öpik et al. 2006).
The finding of both greater richness and basal AMF diversity at this prairie could

reflect an actual greater diversity, or it could reflect the incomplete capturing of the
46 diversity in other studies due to methodological considerations. In the studies reviewed by (Öpik et al. 2006), an average of only 2.7 (±1.7 S.D.) plant species were sampled per site as compared to the 12 host species sampled here. Also, most of those studies used primers now known to not detect Archaeosporales and Paraglomerales taxa.
Based on BLASTN searches, seven OTUs (19% of the total) from Kingston
Prairie had ≥97% identity to published sequences from AMF of described species; four of those were in the Acaulosporaceae and three were in the Glomerales (Fig. 2.2).
Five OTUs (14%) – three in the Glomerales, one in the Archaeosporales and one in the Diversisporales – did not have highly matching sequences (>97% identity) in sequence databases. It should be noted that most described AMF species do not have
LSU rDNA sequences publically available, which limits the ability to ascertain the relationship between environmental OTUs and described species. None of the
Archaeosporales OTUs match described species at >97% sequence identity.
Analysis and performance of newly designed primers
In silico comparisons of the new primers to database sequences of described
AMF species confirms the primers match well with AM fungi while excluding non-
AM fungi (Fig 2.1). There are many more available sequences of described AMF covering the region of primer AM-SSU-F than the AM-LSU-Rx and AM-LSU-Ri primers. There are a few exceptions to perfect identity with the sequence of primer

AM-SSU-F. The most notable is a single base pair mismatch in all Claroideoglomus
47 speces. That mismatch is 5 bp in from the 3’ end of the primer, so it is likely that the primer will still anneal to and amplify those sequences. Indeed one OTU (KP 30) which shares 98% identity with Cl. claroideum BEG14 in the analyzed LSU region was detected. The few mismatches found in other sequences are even further towards the 5’ end of the primer, thus even less likely to affect annealing and amplification.
There is less theoretical evidence for how well the reverse primers will detect
AMF diversity because there are few LSU sequences of described species available in public databases that extend to the region of those primers. Sequences of the basal
Glomeromycota species Geosiphon pyriformis and Paraglomus occultum match these new primers, although no Paraglomerales were detected in the Kingston Prairie samples. Since designing these primers, the sequence of Diversispora celata has been made available (Gamper et al. 2009). The external reverse primer AM-LSU-Rx matches that sequence, but the nested AM-LSU-Ri primer mismatches the
Diversispora sequence at its 3’ terminal nucleotide, likely resulting in amplification with primer AM-LSU-Rx but not with primer AM-LSU-Ri. A single base pair abbreviation of AM-LSU-Ri to remove the non-identical base with Diversispora would result in a sequence lacking a 3’ distinction from non-AM fungi. A new primer abbreviated at 3’ end by four bases but sharing more sequence with AM-LSU-Rx
(such as 5’-GAGAATAGGTTAAGGCTGTTTCAA-3’) might capture the
Diversispora with the other Glomeromycota while still eliminating amplification of most non-AM fungi. Although no sequences clustering within the family

48
Diversisporaceae were detected, eight OTUs were detected in the order
Diversisporales. Unfortunately, theoretical analysis is limited by the dearth of database sequences extending to the portion of the LSU against which the reverse primers were designed, but empirical results from this field survey suggest that these new primers detect a wide range of AMF diversity.
Based on comparisons with database sequences, it appears that the AM-SSU-F and AM-LSU-Rx/AM-LSU-Ri primers will discriminate against non-AM fungi.
Although each primer analyzed separately matches a few non-AM fungi well enough to likely result in amplification, the primers were designed as a semi-nested set so that when used in combination they would discriminate against and obstruct amplification of non-AM fungi. While very few (three out of 229) sequenced clones were non-AM fungi and none were from plants, clones were prescreened for proper insert size before sequencing. Some plant species had 100% of clones of the expected length, while other plants had a majority of clones of unexpected length (Table 1), generally less than 1 kb rather than the predicted 2 kb. Therefore, caution is advised in the use of these primers alone for community analysis applications that don’t have a clone screening step included (e.g. T-RFLPs).
Analysis of published primers
Using the sequence information from the Kingston Prairie survey to predict the ability of other published primers to detect the entire diversity of AMF found at
Kingston Prairie shows a wide range in likely detection. As previously noted (e.g.

Gamper et. al 2009), the FLR3 and FLR4 primers show poor binding site matching
49 with various AMF groups, biasing detection towards a subset of AMF. Analysis with
Kingston Prairie sequences indicates that FLR3 mismatches with many Glomerales sequences and both FLR3 and FLR4 mismatch with some (three and four respectively)
Archaesporales sequences. The primer FLR2 was designed as a universal fungal primer for use in an initial PCR to create a fungal rDNA enriched product that could be used as template in a subsequent nested reaction with AMF specific primers
(Trouvelot et al. 1999). However, in silico analysis indicates that 25% of the Kingston
Prairie AMF sequences, mostly in the order Glomerales, would not be amplified when using FLR2. The primer 28G2 (da Silva et al. 2006) is predicted to miss only three out of 36 Kingston Prairie sequences, two of those in the Archaeosporales. In a field survey of plant roots using 28G1/28G2 primers, Li et al. (2010a) reported less than 4% of sequences were non-AMF or chimeras, indicating that the 28G2 primer likely discriminates against non-AM fungi well as 28G1 is identical to some non-AM fungi
(da Silva et al. 2006).The same field survey did not detect any sequences in the
Archaesporales, yielding a lack of positive evidence that 28G2 does detect that AMF group (Li et al. 2010a). One of the best predicted detectors of Kingston Prairie sequences, likely amplifying all but one sequence, is LSU-Glom1, although it also detects some non-AM fungal taxa (Renker et al. 2003). The other best predicted detector of Kingston Prairie sequences, likely amplifying all sequences, is the primer set LSUmBr (Krüger et al. 2009). Although Krüger et al. (2009) did not include
Archaeosporaceae sequences in their LSU alignments for primer design (but did

include Geosiphon pyriformis ), they did detect two Archaeosporales sequences from
50 spores and environmental samples. Comparing their LSUmBr primers to the Kingston
Prairie sequences shows that within the Archaeosporales there is a G in the middle of the sequences where no G is included in the LSUmBr pimer set (Fig. 2.3). Because that mismatch is in the middle of the primer, their LSUmBr primer set still is likely to detect those Archaeosporales sequences. Nonetheless, adding another primer matching most Archaeosporales sequences reported here, e.g. 5’-
AACACTCGCACGTATGTTAGA-3, to the LSUmBr primer mix may improve detection of the Archaeosporales.
Conclusions
It is clear that AMF serve a variety of functions in ecological communities, yet we have much to learn about how their functional diversity relates with their taxonomic diversity. Without a firm understanding of AMF taxonomic diversity, our understanding of their functional diversity will remain incomplete. Many molecular sampling studies done to date consistently indicate that actual global AMF diversity is much greater than the current morphologically described species (Öpik et al. 2006,
Öpik et al. 2010). However, many environmental studies have used primers that are known to miss at least some Glomeromycota taxa. Future use of primers designed to better target all AMF will allow for improved ability to detect total AMF diversity, thereby providing much improved data for use in ecological analysis.

51
Acknowledgements
Funding for this work was provided by a National Science Foundation
Fellowship and a grant from Prairie Biotic Research, Inc. I thank Christine Hawkes for helping with the SATe phylogenetic analysis. I thank Deborah Clark for helpful conversations during the planning stages of this project. I thank Eric Seabloom for his writing guidance and Virginia Weis and Joseph Spatafora for their comments on the manuscript. I thank the Nature Conservancy of Oregon for allowing access to
Kingston Prairie Preserve for this research.

52
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223-241.
Powell J.R., Parrent J.L., Hart M.M., Klironomos J.N., Rillig M.C., Maherali H. 2009.
Phylogenetic trait conservatism and the evolution of functional trade-offs in arbuscular mycorrhizal fungi. Proceedings of the Royal Society B-Biological Sciences
276: 4237-4245.
Renker C., Heinrichs J., Kaldorf M., Buscot F. 2003. Combining nested PCR and restriction digest of the internal transcribed spacer region to characterize arbuscular mycorrhizal fungi on roots from the field. Mycorrhiza 13: 191-198.
Sarkar G., Cassady J., Bottema C.D.K., Sommer S.S. 1990. Characterization of polymerase chain-reaction amplification of specific alleles. Analytical Biochemistry 186: 64-68.

54
Schloss P.D., et al. 2009. Introducing mothur: Open-Source, Platform-Independent,
Community-Supported Software for Describing and Comparing Microbial
Communities. Applied and Environmental Microbiology 75: 7537-7541.
Schüßler A., Walker C. 2010. The Glomeromycota: a species list with new families and new genera. The Royal Botanic Garden Edinburgh, The Royal Botanic Garden Kew,
Botanische Staatssammlung Munich, and Oregon State University.
Schüßler A., Schwarzott, D., Walker C. 2001. A new fungal phylum, the Glomeromycota : phylogeny and evolution. Mycological Research 105: 1413–1421.
Trouvelot S., van Tuinen D., Hijri M., Gianinazzi-Pearson V. 1999. Visualization of ribosomal DNA loci in spore interphasic nuclei of glomalean fungi by fluorescence in situ hyhridization. Mycorrhiza 8: 203-206.
van der Heijden M.G.A. 2010. Mycorrhizal fungi reduce nutrient loss from model grassland ecosystems. Ecology 91: 1163-1171.
van der Heijden M.G.A., Boller T., Wiemken A., Sanders I.R. 1998a. Different arbuscular mycorrhizal fungal species are potential determinants of plant community structure.
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461.

55
Table 2.1. PCR and cloning success rates. The number of root systems of individual plants initially used in PCR reactions, the number of those that produced an amplification product of the predicted size (2 kb) used in cloning reactions, the number of colonies screened for each cloning reaction and the number and portion of those that had inserts in the predicted size range is shown.
Festuca roemerii
Achillea millefolium
Zigadenus venenosus
Brodiaea coronaria
Delphinium nuttallii
Eriophyllum lanatum
Poa secunda
Prunella vulgaris
Perideridia gairdneri
6
7
6
3
Leucanthemum.
vulgare 2
Sidalcea campestris 3
Potentilla gracilis
Totals
5
62
6
6
6
6
6
6
6
6
4
6
6
7
6
2
2
2
2
55
94
152
120
96
144
70
147
89
56
120
85
55
59
64
1015 706
0.95
0.37
1.00
0.89
0.38
0.84
0.44
96 91 0.95
96* 87* 0.91*
0.70
*Cloning reaction of the pooled species mixture containing P. garidneri , L. vulgare , S. campestris , and P. gracilis .

56
Figure 2.1
.
Primer binding sites of the newly designed primers. Comparisons to fungal sequences are shown in: (a) forward primer AM-SSU-F and forward strands of database SSU rDNA sequences and (b) reverse primers AM-LSU-Rx and AM-LSU-Ri and the reverse complement sequence of database LSU rDNA sequences. For each, the sequence set at the top is AM fungi and those following the gap between sequences are non-AM fungi. Where polymorphism was found within a species at a primer binding site, a representative of each sequence variant for that species is shown. Comparisons to database sequences of basal plants are shown in (c) for AM-
SSU-F and in (d) for AM-LSU-Rx and AM-LSU-Ri. Nucleotides that are identical to those of the primer sequence are given with “.” and variant bases from the primer sequence are given.

(a)
AM-SSU-F 5’- CGCGCTACACTGATGAAGT - 3’
Rh. irregulare DAOM179198
Fu. caledonium BEG20 Y17635
Fu. mosseae UT101 AY635833*
...................
...................
...................
Fu. species WUM3 AJ301864
Fu. species WUM3 AJ301865
Cl. claroideum BEG14 AJ301851
Cl. luteum SA101-1 AJ276089
Gl. versiforme BEG47 X86687
Ac. laevis AU211-3 AJ250847
Ac. longula W3302/Att698-3 AJ306439
...................
............G......
..............A....
..............A....
...................
...................
...................
Ac. rugosa WV935 Z14005
Ac. cavernata BEG33 AJ306442*
Ac. spinosa WV860 Z14004
Ac. sp . W3424/Att729-0 AJ306440
Ac. colombiana WV877 Z14006
Ac. lacunosa BEG78 HE610426
Ac. lacunosa BEG78 HE610427
Ac. lacunosa BEG78 FR719957
Di. spurca W3239 AJ276077
Di. sp . W2423/Att382-16 AJ301863
Gi. sp . W2992 AJ276090*
Gi. rosea DAOM194757
Re. fulva AM418545*
Re. pulvinata AM418550*
...................
...................
...................
...................
...................
...................
............T......
..............A....
...................
...................
...................
...................
..T................
T......T...........
Sc. cerradensis MAFF520056 AB041345
Am. fennica W3847/Att200-21 AM268194*
Am. leptoticha MAFF520055 AB047304*
Ar. trappei NB112 AJ243420
Ge. pyriformis AM183923
Sc. heterogama FL225 AY635832*
Pac. scintillans
Ar. schenckii CL401 AM743189*
Pa. brasilianum
W3793 AJ619940
WV219 AJ012112
Pa. occultum clone IA702-3 AJ276081
Capronia pilosella DQ823106
Taphrina wiesneri AY548293
Penicillium chrysogenum M55628
Neurospora crassa X04971
Peziza badia L37539
Saccharomyces cerevisiae J01353
Platygloea disciformis DQ234563
Tilletiaria anomala AY803752
Endocronartium harknessii M94339
Russula compacta U59093
Parasitella parasitica AF157149
Endogone pisiformis DQ322628
Mortierella verticillata AF157145
Spiromyces aspiralis AF007543
Piptocephalis corymbifera NG_017192
...................
...................
...................
...................
...................
...................
...................
...................
...................
...................
.............CAG...
.............C.G...
.............CAGG.C
.............CAC..C
.............CAG..C
.............C.G..C
.............CCG...
.............C.G...
.............CC...C
.............CAG..C
.................A.
.............C....T
..............C...T
..............C...T
.............C.....
Figure 2.1
57

(b)
AM-LSU-Rx 5’-GTTTGAGAATAGGTTAAGGC-3’
AM-LSU-Ri 5’-GTTAAGGCTGTTTCAACCCT-3’
3’
Rh. irregulare DAOM179198 DQ273790 ................................
Rh. intraradices 4695rac-11G2 DQ273828 ................................
Fu. mosseae UT101 DQ273793 ................................
Sc. heterogama FL225 DQ273792 ................................
Di. celata AM713417* ...............................C
Pa. occultum IA702 DQ273827 ................................
Ge. pyriformis AM183920 ................................
Capronia pilosella DQ823099 ...................T...........C
Taphrina wiesneri AY548292 ...................T...........C
Penicillium sp . KH00312 GU017561 ...................T...........C
Platygloea disciformis AY629314 ...................A....G..T...C
Tilletiaria anomala AY745715 ...................A....A..T...C
Endocronartium harknessii AY700193 ...................A.A..A..T...C
Endogone pisiformis DQ273811 ...................A.......T....
Mortierella verticillata DQ273794 ...........................G...C
Spiromyces aspiralis DQ273801 ...........................G...C
Piptocephalis corymbifera AY546690 ...................GCA..C.GC...C
Figure 2.1 (Continued)

(c)
AM-SSU-F1 5’-CGCGCTACACTGATGAAGT-3’
Anthoceros agrestis X80984 .................T.
Folioceros fuciformis DQ845699 ..T..............T.
Sphaerosporoceros adscendens DQ845703 .................T.
Phaeoceros laevis U18491 .................T.
Sphagnum fuscum GQ375070 .................T.
Pallavicinia lyellii AY688723 .................T.
Leptopteris hymenophylloides AY612727 .................T.
Lepidogyna hodgsoniae DQ629371 .................T.
Haplomitrium blumei AY688711
Blasia pusilla DQ629388
Hymenophyton flabellatum DQ629391
Marsupella emarginata DQ629377
.................T.
.................T.
.................T.
.................T.
Notothylas breutelii
Thuidium recognitum
Andreaea wilsonii
DQ629408
DQ629403
Takakia ceratophylla DQ629407
AY330416
.................T.
...............G.T.
.................T.
.................T.
Andreaeobryum macrosporum AJ275005 .................T.
Lycopodium clavatum DQ629409 .................T.
Equisetum arvense DQ629411 .................T.
Polytrichastrum alpinum GU569642
Pogonatum urnigerum GU569632
Trichocoleopsis saccatula EF460691
Selaginella apoda AF313575
Burmannia hexaptera EU420994
Burmannia capitata DQ786066
.................T.
.................T.
.................T.
.................C.
...............T.C.
...............T.C.
(d)
AM-LSU-Rx 5’- GTTTGAGAATAGGT-TAAGGC-3’
AM-LSU-Ri 5’-GT-TAAGGCTGTTTCAACCCT-3’
Coleochaete scutata AB491669 ..............-CG...A.......T...C
Ambuchanania leucobryoides GQ375083 ..............-CG...G....A..C...C
Leucobryum sp. GQ375084
Targionia hypophylla GQ910743
..............-CG...G....A..C...C
..............GCG...G.......C...C
Athalamia hyalina GQ910714 ..............-CG...G...W...C...C
Adiantum capillus-veneris EU161365 ..............-CG..AG.......C...C
Neohodgsonia mirabilis DQ265789 ..............-CG...G.......C...C
Temnoma pilosum DQ026569
Ptilidium ciliare AY608247
..............-CG...G.......C...C
..............-CG...G.......C...C
Petalophyllum ralfsii AY877377 ..............-CG...G.......C...C
Hypopterygium tamarisci EF680816 ..............-CG...G....A..C...C
Andreaeobryum macrosporum AY330426 ..............-CG...G....A..C...C
Andreaea nivalis AJ271023 ..............-CG...G....A..C...C
Takakia lepidozioides AJ271020 ..............-CG...G....A..C...C
Treubia lacunosa DQ026571
Allisonia cockaynii DQ026523
..............-CG...G.......C...C
..............-CG...G.......C...C
Figure 2.1 (Continued)
59

60
Figure 2.2. A consensus phylogenetic tree showing the relationship of the Kingston
Prairie operational taxonomic units to other Glomeromycota sequences. A consensus maximum likelihood tree was constructed based on ribosomal RNA gene sequences with Paraglomus occultum used as the outgroup. Branch support values from 100 bootstrap replicates are shown. The Kingston Prairie sequences are labeled with ‘KP,’ their assigned OTU number, the accession number for the sequence used, and in parentheses is the number of unique sequences obtained for that OTU. (Note, because clones were screened by RFLP before selection for sequencing, the number of unique sequences does not represent the relative abundance of OTUs.) Black diamonds indicate Kingston Prairie sequences with ≥97% identity to a described AMF species sequence; black triangles indicate Kingston Prairie sequences with ≥97% identity to environmental (labeled ‘ENV’), unpublished (labeled ‘UnPub’), or AMF cultures of non-described species; and grey squares indicate sequences with <97% identity to a database sequence. Sequences from Genbank begin with their accession number.
Consensus sequences from Krüger et al. (2012) Figure 1 begin with “C ##.”

77
98
69
90
97
98
99
89
97
99
100
73
97
89
38
100
65
100
100
98
92
95
98
70
99
81
92
61
77
100
84
99
44
74
30
77
54
95
100
95
100
60
100
100
77
72
89
61
100
100
49
100
100
32
71
100
100
100
49
100
100
41
94
99
100
97
100
97
47
100
98
100
42
67
100
100
93
77
100
42
50
91
100
93
100
43
39
63
100
93
100
KP01 JF717469 (8)
HQ895792 ENV
EU380045 ENV
KP02 JF717517 (48)
KP03 JF717464 (7)
HE775302 ENV
KP04 JF717432 (13)
AM040432 ENV
EU379997 ENV
KP05 JF717434 (4)
JF439189 UnPub
KP06 JF717607 (2)
AB369768 ENV
KP07 JF717488 (10)
EU380048 ENV
KP08 JF717527 (3)
KP09 JF717614 (5)
AB369760 ENV
KP18 JF717500 (4)
KP10 JF717452 (9)
AB369759 ENV
KP29 JF717462 (1)
FR750095 Rhizophagus sp. MUCL 43208
AB369739 ENV
KP12 JF717487 (29)
KP13 JF717583 (3)
KP14 JF717511 (1)
FR772327 ENV
C 33 Sclerocystis sinuosa MD126
KP15 JF717578 (3)
HE775302 ENV
KP16 JF717570 (4)
AJ854593 ENV
KP17 JF717495 (18)
AJ854592 ENV
KP19 JF717596 (6)
KP21 JF717538 (3)
C 29 Rhizophagus irregularis DAOM197198
KP20 JF717471 (5)
C 30 Rhizophagus irregularis AFTOL-ID845
C 31 Rhizophagus proliferus MUCL41827
C 32 Rhizophagus intraradices FL208
C 24 Funneliformis mosseae BEG12
C 25 Funneliformis coronatus BEG28
C 26 Funneliformis sp. WUM3
C 27 Funneliformis caledonius BEG20
C 22 Glomus macrocarpum
KP11 JF717475 (3)
FR750203 Glomus sp. Att565-7
C 20 Glomus sp. W3347/Att565-7
C 34 Claroideoglomus sp. W3349/Att565-11
KP30 JF717489 (1)
C 35 Claroideoglomus luteum SA101
KP24 JF717621 (1)
KP25 JF717587 (6)
C 05 Acaulospora brasiliensis
C 04 Acaulospora laevis AU211
FR750172 Acaulospora entreriana
KP26 JF717598 (1)
FR750156 Acaulospora spinosa
C03 Acaulospora spinosa W3574
FR692351 Acaulospora scrobiculata
AM040293 Acaulospora longula
KP27 JF717591 (2)
FM876789 Acaulospora cavernata
C 01 Acaulospora cavernata BEG33
C 02 Acaulospora sieverdingii WUM18
KP28 JF717567 (3)
KP36 JF717558 (1)
C 06 Acaulospora lacunosa BEG78
C 11 Diversispora epigaea BEG47
C 07 Diversispora spurca
C 09 Diversispora celata BEG231
KP22 JF717509 (1)
EU252109 ENV
KP23 JF717626 (1)
C 18 Scutellospora spinosissima W3009/Att664-1
C 15 Gigaspora rosea DAOM194757
C 12 Scutellospora heterogama BEG35
C 19 Pacispora scintillans W4545
KP31 JF717581 (1)
KP32 JF717553 (1)
FR750037 Archaeospora trappei
C 38 Archaeospora schenckii
KP35 JF717625 (2)
HQ128660 ENV
AY639362 ENV
KP33 JF717594 (1)
FN547527 Ambispora appendicula
C 36 Ambispora fennica
FJ461888 UnPub
KP34 JF717576 (1)
C 37 Geosiphon pyriformis GEO1
C 39 Paraglomus occultum IA702
Figure 2.2
61

62
Figure 2.3. Comparison of published primers targeting large subunit (LSU) rDNA to
Glomeromycota sequences found at Kingston Prairie Preserve. The most common sequence type for each Kingston Prairie OTU is given. A degenerate nucleotide indicates equal frequency of bases at that site within the OTU. The reference sequence and position numbers are from Rhizophagus irregulare MUCL43194 accession number DQ273790. A ‘.’ is shown where the Kingston Prairie sequence is identical with the reference sequence, except where the nucleotide differs with a published primer. Where a nucleotide in a Kingston Prairie sequence differs from a listed primer, the differing nucleotide is given in bold. Italicized reference or primer sequence names indicate that the complementary sequence is shown. Primers shown are FLR3 and
FLR4 (Gollotte et al. 2004), LSU-Glom1 (Renker et al. 2003), FLR2 (Trouvelot et al.
1999), 28G2 (da Silva et al. 2006), and LSUmBr1-5 (Krüger et al. 2009).

414 435
SSU Ref TTGAAAGGGAAACGATTGAAGC
FLR3 TTGAAAGGGAAACGATTGAAGT
KingPr01 .....................T
KingPr02 .....................T
KingPr03 .....................T
KingPr04 .....................T
KingPr05 .....................T
KingPr06 .....................T
KingPr07 .....................T
KingPr08 .....................T
KingPr09 .....................
C
KingPr10 .....................
C
KingPr11 .....................T
KingPr12 .....................T
KingPr13 .....................T
KingPr14 .....................T
KingPr15 .....................
C
KingPr16 .....................
C
KingPr17 .....................
C
KingPr18 .....................
C
KingPr19 .....................
C
KingPr20 .....................
C
KingPr21 .....................
C
KingPr29 .....................
C
KingPr22 .....................T
KingPr23 .....................T
KingPr24 .....................T
KingPr25 .....................T
KingPr26 .....................T
KingPr27 .................
A ...T
KingPr28 ..
A ..................T
KingPr36 .....................T
KingPr30 .....................T
KingPr31 ...................
G .
C
KingPr32 .........
G .........
G .
C
KingPr33 .................
A ...T
KingPr34 .....................
C
KingPr35 ....
T ................T
Figure 2.3
434 415
LSU Ref CTTCAATCGTTTCCCTTTCA
LSU-Glom1 CTTCAATCGTTTCCCTTTCA
....................
....................
....................
....................
....................
....................
....................
....................
...................
.
....................
....................
....................
....................
....................
....................
....................
....................
....................
....................
....................
....................
..................
T .
....................
....................
....................
....................
....................
...
T ................
....................
....................
....................
.
C ..................
.
C .........
C ........
...
T ................
....................
................
A ...
63

802 767
LSU Ref GTCGTTTAAAGCCATTACGTCAACATCCTTAACGAA
FLR2 GTCGTTTAAAGCCATTACGTC
28G2 CCATTACGTCAACATCCTTAACG
FLR4 ACGTCAACATCCTTAACGAA
KingPr01 ...................C................
KingPr02 ...................C....G...........
KingPr03 ...................C....G...........
KingPr04 ...................C................
KingPr05 ...................C................
KingPr06 ...................C....G...........
KingPr07 .......................G...........
KingPr08 ....................................
KingPr09 ....................................
KingPr10 ....................................
KingPr11 ...................a............T...
KingPr12 ...........T........................
KingPr13 ...........T........................
KingPr14 ....................................
KingPr15 ....................................
KingPr16 ....................................
KingPr17 ....................................
KingPr18 ....................................
KingPr19 ....................................
KingPr20 ....................................
KingPr21 ....................................
KingPr29 .............G.....C................
KingPr22 ......................G.G...........
KingPr23 ......................G.G...........
KingPr24 ......................G........G....
KingPr25 ......................G........G....
KingPr26 ......................G........G....
KingPr27 ......................G........G....
KingPr28 ......................G........G....
KingPr36 ......................G........G....
KingPr30 ....................T...............
KingPr31 .........GA......T....G.......GG.A..
KingPr32 .........G.T..........G.......GG.A..
KingPr33 ......................G........T....
KingPr34 ......................G............G
KingPr35 ...............................G...G
Figure 2.3 (Continued)
64
849 828
LSU Ref TAACACTCGCATATATGTTAGA
LSUmBr1 DAACACTCGCATATATGTTAGA
LSUmBr2 AACACTCGCACACATGTTAGA
LSUmBr3 AACACTCGCATACATGTTAGA
LSUmBr4 AAACACTCGCACATATGTTAGA
LSUmBr5 AACACTCGCATATATGCTAGA
G.....................
G.....................
A.....................
GG....................
G.....................
G.....................
G.....................
G.....................
G.....................
G.....................
A.....................
G.....................
G.....................
G.....................
A.....................
A.....................
......................
G.....................
......................
......................
......................
G.....................
.............C........
.............C........
A..........C.C........
A..........C..........
A..........C.C........
A..........C.C........
A..........C.C........
A..........C.C........
A................C....
A..........CG.........
A..........CG.........
A...........G.........
A..........CG.........
A..........CG.........

65
Figure 2.4. Number of Kingston Prairie sequences in each Glomeromycota order that are predicted to be amplified by published primers. Amplification is predicted to be unsuccessful if any of the three 3’ nucleotides mismatch, based on Lee et al. (2008), or if three or more nucleotides within the primer sequence mismatch.

Chapter 3: Host species identity but not spatial distribution influences arbuscular mycorrhizal community in plant roots
Wendy S. Phillips
Deborah L. Clark
Eric W. Seabloom
To be submitted to:
66
Ecology
Ithaca, New York, U.S.A.

67
Abstract
Arbuscular mycorrhizal fungi (AMF) form symbioses with ~80% of plant species. They play a critical role in plant communities, affecting both plant productivity and richness. The structure of AMF-plant networks is complex, often with many different AMF colonizing the roots of a single plant and a single AMF colonizing multiple plants at the same time. In this study we investigated the effects of host identity, geographic distance and soil properties on root AMF communities. We also tested for nestedness in the plant-AMF network. To do this, we analyzed the composition of AMF communities in the roots of 50 plants from nine native plant species occurring at a remnant bunchgrass prairie in Oregon, U.S.A. We designed 18 primer pairs that specifically amplified sections of large subunit ribosomal RNA gene sequences from a subset of the operational taxonomic units found in a previous survey of the AMF occurring at the prairie. Host species identify affected both the richness and the community composition of AMF in roots. Neither geographic distance, soil type, nor distance from a creek correlated with root AMF community composition.
The AMF-plant interaction network was significantly nested.
Introduction
The symbiosis between land plants and arbuscular mycorrhizal fungi (AMF) is as old as land plants themselves (Redecker et al. 2000, Wang & Qiu 2006) and involves the largest number of plant species of any of the mycorrhizal symbioses
(Wang & Qiu 2006). Generally considered a mutualism, the fungi receive fixed carbon

68 in the form of sugars from their host, while plants receive benefits in the form of increased access to soil minerals and water (Smith & Read 2008) as well as reductions in herbivory and root pathogens (Azcón-Aguilar & Barea 1996, Bennett & Bever
2007). The critical role AMF play in plant communities is evidenced by the correlations of higher plant productivity and richness with increasing diversity of
AMF communities (van der Heijden et al. 1998b). AMF influence ecosystem processes through various other pathways (Rillig 2004), particularly impacting nutrient cycling (Olsson & Johnson 2005, van der Heijden 2010).
Despite the ecological importance of this symbiosis, little is known about the role plant species identity plays in structuring AMF community composition.
Historically the partners in the plant-AMF symbiosis were considered to have low specificity, particularly because of the small number of AMF species relative to the large number of plant species involved in the symbiosis (Smith & Read 1997).
Although most evidence still suggests a lack of tight specificity between hostsymbiont pairs, several studies indicate that host species identity does impact root
AMF community composition (Douds et al. 1998, Helgason et al. 2002,
Vandenkoornhuyse et al. 2003). Other studies indicate root AMF community composition is more highly correlated with the ecological grouping of host plants than the individual species’ identity is (Davison et al. 2011, Öpik et al. 2009). Still other studies have found no correlation between host species identity and AMF community composition (Öpik et al. 2008). Some have suggested that plant species vary in their

69 selectivity of AMF symbionts, with some plant species being highly selective and others being less discriminatory in their associations (Johnson et al. 2005).
One factor necessarily affecting the AMF with which individual plants partner is the distribution of AMF types in soil. As AMF have both extraradical (outside of roots, or in soil) and intraradical (within root) hyphae, soil characteristics would be expected to affect their distribution. Indeed, distributional correlations with factors such as soil sand vs. clay composition, nutrient and moisture content, and pH have been shown (Anderson et al. 1984, Egerton-Warburton & Allen 2000, Porter et al.
1987). Additionally, AMF distributions are impacted by dispersal limitation. AMF can disperse through spores, hyphal fragments in soil, or extensions of existing hyphal networks. For the first two, animals, water or wind may aid dispersal, although little research has been done to determine the distances such propagules can travel. These effects of soil characteristics and dispersal limitations on AMF distributions may contribute to spatial autocorrelation, such as that seen for soil spore densities of various AMF species (Carvalho et al. 2003, Klironomos et al. 1999). The spatial structuring of AMF soil communities due to such non-host related effects could overwhelm whatever role host selectivity plays in the assembly of AMF root communities.
In addition to examining the factors driving the assembly of individual root
AMF communities, we can also ask if there are larger patterns in the structure of the overall plant-AMF network of interactions. When specialist species of one of the symbiont pairs tend to interact only with generalists species of the paired symbiont,

70 this suggests non-random assembly processes and the interaction network is considered highly nested. Evidence for nestedness has been found in a diverse assortment of mutualistic networks, including plant-frugivore and plant-pollinator networks (Bascompte et al. 2003), anemonefish and sea anemones networks (Ollerton et al. 2007), and in reef cleaning symbioses (Guimarães et al. 2007). In recent work,
Montesinos-Navarro et al. (2012) found plant-arbuscular mycorrhizal networks also exhibit significant nestedness. We use our data to ask if similar nestedness patterns in the mycorrhizal community occur in a different type of vegetative community.
In this study we ask whether: i) host identity, ii) geographic distance between plants, and iii) environmental factors such as soil type and distance from creek bed are related to root AMF community composition; and iv) if the AMF-plant network shows nestedness. Many studies have used molecular methods to investigate AMF communities in plant roots in the last decade, but most have surveyed relatively few plants species within a field site (Davison et al. 2011). To our knowledge, very few studies have investigated AMF sequences from a variety of plants within naturally occurring plant communities in the United States, e.g. Appoloni et al. (2008), and no published studies have included spatial data with AMF root community analysis of individual plants of multiple species within a naturally occurring plant community.
Methods
Field site and plant sampling

71
We investigated the community of AMF in the roots of 50 plants naturally occurring at a remnant bunchgrass prairie in the Willamette Valley of Oregon:
Kingston Prairie Preserve, managed by the Nature Conservancy. This work is a continuation of the initial field survey of AMF occurring at the prairie that is presented in Chapter 2, and the prairie and the sampling methods are described there. Included in this study were two perennial grasses, Festuca idahoensis Elmer ssp. roemeri
(Pavlick) S. Aiken and Poa secunda J. Presl, and seven perennial forbs, Achillea millefolium L., Brodiaea coronaria (Salisb.) Engl., Delphinium nuttallii A. Gray,
Eriophyllum lanatum (Pursh) Forbes, Potentilla gracilis Douglas ex Hook, Prunella vulgaris L., and Zigadenus venenosus S. Watson. All plant species are natives of the
Willamette Valley of Oregon. For this study, we used four to seven individuals, previously determined as having AMF in their roots by positive PCR amplification with the AMF specific primers AM-SSU-F and AM-LSU-Rx (Chapter 2), from each species.
Molecular methods
In the initial field survey, we categorized AMF sequences of the large subunit of the ribosomal RNA gene (LSU rDNA) into operational taxonomic units (OTUs) based on phylogenetic relatedness. In this study, to assess the presence of a subset of those OTUs in individual plant samples, we designed primer sets to specifically amplify individual OTUs and used them in PCR. OTU specific primers were designed by aligning sequences of all the clones of a given OTU and the OTUs of its closest

phylogenetic neighbors. Primer locations were considered where there was sequence
72 conservation within the OTU but sequence divergence from other OTUs. To minimize amplification of non-target sequences, primer sites where chosen such that at least the
3’ terminal nucleotide of the primer differed from non-target sequences while maintaining 100% conservation within the OTU target sequences. The rest of the primer was generally conserved within all sequences of that OTU, but one or two discrepancies were allowed in nucleotides greater than 5 bases from the 3’ end of the primer where polymorphisms within the sequences of an OTU could not be avoided.
Where there were differences in nucleotides in the primer region within the sequences of an OTU, the most common nucleotide was used in the primer. Primer length ranged from 17 to 25 nucleotides, and primers were limited to locations where the G-C content of the sequence was between 32 and 71%, although most were between 40 and
60%.
We developed 18 primer sets that specifically amplified target groups. Fifteen of those primer pairs amplified single OTUs, and three primer pairs amplified larger monophyletic groups that included two OTUs (OTUs 1 and 2 and OTUs 9 and 10) or three OTUS (OTUs 15, 16 and 18), because it was only at that level that sequence conservation could be found within the group that differed from non-group sequences.
Including both single- and multi-OTU primer sets, 61% of the OTUs representing 79% of the sequences from the original field study were included in root AMF community assays (Fig. 3.1). Four of the five Archeaosporales OTUs and five of the eight
Diversisporales OTUs found in the original field survey (Chapter 2) were included in

the 15 single OTU primer sets. The OTUs not included from those two families all
73 were represented by a single sequence from the field survey. The primers used are given in Table 3.1.
Presence of individual OTUs in a plant sample was assessed by PCR using a two-step process. First, we PCR amplified a larger AMF rDNA fragment to use as template in a subsequent nested reaction. For that first PCR, root DNA templates were diluted and amplified as described in Chapter 2 using primers AM-SSU-F and AM-
LSU-Rx and 5 µl of DNA in a 20 µl PCR reaction that was cycled 25 times. The resulting PCR product was diluted 1:50 for use as template with individual OTU primer sets in reactions containing 0.25 µm primers and cycled 34 times. Prior to using the primer sets with actual samples, they were tested over a range of annealing temperatures on plasmid DNAs containing AM-SSU-F and AM-LSU-Rx inserts from the same OTU (positive control) and from several other OTUs most similar in sequence to that OTU (negative controls). An annealing temperature for each primer set was chosen that gave strong amplification in the positive controls but no visible product for the negative controls. Final annealing temperatures used are given in Table
3.1. Both positive and negative controls were included when amplifying root DNA samples with individual OTU primer sets, and results were only recorded when those controls were positive and negative, respectively, for products. Presence of a product was assessed by running 8 µl of the PCR reaction on a 1.5% agarose gel stained with ethidium bromide.

74
Statistical analyses
Host effects on AMF communities
To test whether detected numbers of AMF in roots differed between plant species, one-way ANOVA was used. Simple linear regression was used to determine if the number of OTUs detected correlated with mass of root tissue used for the DNA extraction.
To test the null hypothesis of no difference in AMF community composition between host species, we used multi-response permutation procedure (MRPP;
McCune & Grace 2002, Mielke 1984). This method compares dissimilarities of communities within and among groups using a Monte-Carlo approach and yields a chance-corrected within group agreement (A) and a p-value. For the MRPP, we used plant species identity as the grouping assignment and Bray-Curtis distances to measure between sample dissimilarities. We also performed a two-way agglomerative cluster analysis to visualize relationships among AMF OTUs and among sample units.
Indicator Species Analysis (ISA; Dufrene & Legendre 1997) is used most commonly to identify species that have high fidelity to and a high frequency of occurrence in particular ecological sites. Treating plant host species as an analog to ecological sites, we tested whether any AMF were good indicator species for plant hosts using ISA with a Monte Carlo test for significance with 1,000 randomizations.
Geographic distance and soil effects on AMF communities

To test if AMF community composition in roots was associated with geographic distance between plants in the field, we performed a Mantel test (Mantel
75
1967) comparing distance matrices of AMF community dissimilarity calculated using the Bray-Curtis method and the Euclidean distance between GPS coordinates of the samples. We ran a Monte Carlo randomization test (1,000 permutations) to test for significance of association between the two matrices.
To test if AMF community composition correlated with soil type or distance from the creek bed, MRPP was performed using those factors as grouping variables.
Based on soil maps obtained from the Web Soil Survey of the United States
Department of Agriculture Natural Resources Conservation Service (USDA-NRCS), soil was assigned to one of three types: Stayton (medial, mixed, mesic Lithic
Haploxerands) silt loam, Witzel (loamy-skeletal, mixed, superactive, mesic Lithic
Ultic Haploxerolls) very cobbly loam, and Witzel variant very cobbly silt loam. In addition, we delineated a riparian zone that bisects the study site based on satellite images of the prairie during a dry season showing still green vegetation extending approximately 5 m from the creek bed along most of its length. Samples were classified as riparian (within 5 m of a creek bed) or upland (greater than 5 m from a creek bed).
All above statistical tests were performed both on AMF community data sets containing the multi-OTU groups and data sets excluding those groups and excluding
AMF that occurred in only one sample. Results were qualitatively the same in all

76 cases. MRPP, ISA, the Mantel test, and cluster analysis were performed using PC-
ORD v6.07 (McCune & Mefford 2011).
Nestedness of the plant-AMF network
To test whether the AMF and plants form partnerships suggestive of reciprocal specialization, we analyzed the nestedness of the interaction network. A network is considered nested if species with few links interact with species with many links, i.e. specialist species interact with generalist partners rather than with a specialist partner
(Bascompte et al. 2003). We tested for nestedness by calculating nested overlap and decreasing fill (NODF) values (Almeida-Neto et al. 2008) using the program
Aninhado 3.0.3 (Guimarães & Guimarães 2006). Before calculating the nestedness metrics, we ordered rows and columns by interaction abundance. To test for significance, we then compared those values to 1,000 random values generated using the null model CE, or null model II (Bascompte et al. 2003), which fills cells using a probability of cell occupancy determined by averaging the probabilities of occupancy of its row and column.
Results
Host effects on AMF communities
Host identity was related to AMF communities both in terms of the richness of
OTUs per plant and AMF community composition in roots. Based on the testing of 18
OTUs, a mean of 5.7 (±2.6 S.D.) OTUs were detected per plant. The mean number of

77
AMF OTUs detected in individuals differed between plant species, with A. millefolium having the highest mean, 9.0 (±2.4 S.D.), and E. lanatum having the lowest, 3.8 (±1.5
S.D.) (ANOVA p=0.014; Fig. 3.2). The number of AMF detected per plant was not correlated with the mass of root tissue used for DNA extraction.
In addition to differing in richness, different host species had distinct AMF communities (MRPP, A=0.11, p<0.001, based on the smaller data set). MRPP of root
AMF community composition between plant species show 13 pairwise comparisons have a p-value <0.05 (Table 3.2). Although these p-values were uncorrected for multiple comparisons, based on the total of 36 pairwise comparisons calculated, a type
I error rate of 5% would yield less than two false positives. All species except P. gracilis were represented at least once in the significantly different pairwise comparisons. AMF community composition differed the most between F. idahoensis and P. vulgaris (A=0.21, p<0.001; Table 3.2). To determine if the difference in AMF community composition between those two host species was related to a difference in mean number of OTUs, we performed a two-way t-test that indicated no difference in the means (p=0.2). The AMF community composition in individual plants and the relatedness of those communities is shown in the two-way dendrogram presented in
Figure 3.3.
Despite the evidence for differences in AMF community composition between host species, no AMF OTU was uniquely associated with any host species. Indicator species analysis yielded only one AMF group highly correlated with a host species:

78
OTU 28 and P. vulgaris (ISA, p=0.002). OTU 28 occurs in five out of six P. vulgaris individuals but also occurs in three A. millefolium and two D. nuttallii (Fig. 3.3).
The distribution of Glomeromycota taxa among hosts
We observed a trend in the distribution of the different orders of the
Glomeromycota among host species. Five host species had associations with all three orders, three host species had associations with two of the orders, and one host species associated with only a single order (Fig. 3.4). Although diversity, in terms of the number of orders represented, varied between host species, the number of Glomerales
OTUs associated with host species remained relatively stable. When a lower total number of OTUs was detected in a host species, it was associations with the
Diversisporales and Archaeosporales taxa that were observed to be absent.
Geographic distance and soil effects on AMF communities
There was no correlation between AMF communities and geographic distance
(Mantel r=-0.01, p=0.4). There also was no relationship between AMF communities and soil type (MRPP, A=-0.001, p=0.49) or AMF communities and distance from creek (MRPP, A=0.012, p=0.11).
Nestedness of the plant-AMF network
The interaction network was found to be significantly nested, with a NODF value of 76.1, where a value of 100 indicates perfect nestedness (p<0.001; Fig. 3.5).

79
This indicates species with few links tend to interact with species with many links, i.e. specialist species interact with generalist partners rather than with a specialist partner.
Discussion
We found a relationship between host identity and both the richness and the community composition of the AMF colonizing roots. However, there was no relationship between geographic distance, soil type or distance from a creek and root
AMF community composition. AMF-plant interaction networks were significantly nested indicating few specialist-specialist pairings.
Host effects on AMF communities
Our results indicate that plant species identity impacts root AMF richness.
Other studies have shown such a difference in AMF hosting capacity between plant species (Alguacil et al. 2012, Öpik et al. 2008, Vandenkoornhuyse et al. 2003).
Differences in numbers of AMF in roots of different plant species could result from variations in the selection employed by plants. If some plants are more selective than others, as has been proposed by some (Scheublin et al. 2004), differences in AMF richness such as we observed would result. Yet mechanisms other than active plant selection could cause a reduced number of AMF in roots. For example, some plant species could be poorer hosts, providing a reduced resource supply and thus supporting fewer fungi than plants with greater resource supplies (Roumet et al. 2006,
Urcelay et al. 2011).

Looking beyond simple richness, we observed a relationship between host species identity and root AMF community composition (MRPP, A=0.11, p<0.001).
80
Although we found differences in mean numbers of AMF among host species, those differences alone do not account for the difference in community composition between species. For example, while F. idahoensis and P. vulgaris had the largest difference in
AMF community composition (Table 3.2), they did not differ in mean number of
AMF detected per plant (two-sided t-test, p=0.20). This observation of a relationship between host identity and AMF community composition is consistent with the results of many other studies (Helgason et al. 2002, Li et al. 2010b, Vandenkoornhuyse et al.
2003, Vandenkoornhuyse et al. 2002). While a few studies have found no relationship with host identity (Öpik et al. 2008) and others have found a larger relationship at the level of plant ecological group than at the species level (Davison et al. 2011, Öpik et al. 2009), the majority of published work indicates a connection between host identity and root AMF community composition.
Indicator species analysis also produces little evidence for specialist-specialist associations, with only one AMF having a significant indicator value for any plant:
KP28 and P. vulgaris . KP28 has up to 99% identity with a sequence in Genbank from a single spore of Acaulospora sp. WUM18 collected in Australia (Krüger et al. 2009) and also has >97% identity based on BLASTN searches with a few environmental samples from unpublished studies. With the limited data we have here, we cannot be certain if KP28 specializes on a limited host set. As well as occurring in five out of six
P. vulgaris , KP28 occurred in two other plant species in this study. That P. vulgaris

ranks first in mean number of OTUs per species and second in number of OTUs per
81 individual suggests it is on the generalist side of the generalist–specialist continuum, if such a spectrum exists for plants. The association of a specialist fungi with a generalist plant is consistent with the pattern of nestedness discussed later. If KP28 is indeed a fungus with relatively high specificity, our data suggest it is unique because if specialism in fungi was frequent, one would expect ISA to yield more significant associations. The lack of support our data shows for specialist-specialist associations is not suprising as to date little evidence exists for such mutual specialism in the AM symbiosis.
The distribution of Glomeromycota taxa among hosts
When we examine not just the mean number of AMF that individual plants host but the overall diversity of AMF phylotypes found within a plant species, we see variation in number of AMF OTUs found in different plant species that does not correspond directly with mean number of AMF in individual plants (Fig. 3.2 & 3.4).
As expected, the plant species with the greatest number of AMF OTUs per individual, such as P. vulgaris and A. millefolium , are also the species associating with the highest diversity of AMF, including AMF from all three orders surveyed. As number of AMF
OTUs found in a different host species declines, there is a tendency for AMF from the
Glomerales to be retained but those from the Diversisporales and Archaeosporales to be absent (Fig. 3.4). We note that the plant species with the lowest AMF richness, F. idahoensis and B. coronaria , were not the plant species with the lowest mean number

82 of AMF OTUs per individual, so low diversity of AMF orders is not necessarily associated with lower colonization levels.
Maherali and Klironomos (2007) proposed that competitive interactions involving AMF from different families, which have phylogenetically conserved traits associated with spatial niche requirements (Hart & Reader 2002a), would promote the assembly of a phylogenetically overdispersed community of AMF in roots (Cavender-
Bares et al. 2009). Indeed, they found evidence for such phylogenetic overdispersion in experiments using Plantago lanceolata as a host for manipulations with different starting AMF species pools. The hypothesis of phylogenetic overdispersion due to competitive interactions does not fit with our data indicating that as overall AMF species numbers in plant hosts declined, the remaining community tended towards
AMF from a single order, the Glomerales (Fig. 3.4). The common use of P. lanceolata in culturing AMF suggests it can host a broad range of AMF (or else it would be disfavored for use in culturing) and therefore can provide an environment in which multiple AMF orders can competitively interact to create phylogenetic overdispersion.
Our results suggest that phylogenetic dispersion characteristics of AMF communities in roots may vary with host species.
We found that one of the OTU groups (OTU 1-2) was present in every sample, consistent with previous findings that AMF communities tend to be dominated by a single taxon (Dumbrell et al. 2010b). A BLASTN search of GenBank against these sequences revealed sequences with ≥97% identity from environmental samples obtained within the United States (California and Michigan), the Czech Republic and

83
China. Despite the apparent global distribution of this OTU, it joins twelve other
Kingston Prairie OTUs in a monophyletic group without high sequence identity to any sequenced described AMF taxa (Fig. 3.1).
Geographic effects on AMF communities
The structure of the community of AMF in a plant’s rhizosphere necessarily affects the partnerships a plant can make. The distribution of AMF propagules will depend on the dispersal limitations of both spores and fungal hyphae. Therefore, spatial patterns in the distributions of AMF must play some role in individual plant root AMF community composition. Studies using geostatistical methods have found
AMF spores of individual species to exhibit spatial dependence in soil, ranging from
30 cm to 6.5 m (Carvalho et al. 2003, Klironomos et al. 1999), with some of that spatial distribution attributable to host location. Mummey & Rillig (2008) used molecular methods to assess composition of AMF DNA in soils and found AMF community composition to show positive spatial autocorrelation at distances of 40 cm or less, again with some of that spatial distribution attributable to host location. Other studies conducted at larger distance scales have found that distance between sites is correlated with site level AMF community composition, e.g. Lekberg et al. (2007) studied 10 sites within a 25 km range and found an effect of both soil type (clay vs. sand) and distance. Our sampling was between these scales, generally between 5 m and 250 m. We chose to sample plants from random locations to maximize analysis of host effect on AMF community composition rather than use a systematic sampling

strategy that would maximize geostatistical analysis and allow tests of spatial autocorrelation. Whereas we found a relationship between host species identity and
84 root AMF community composition, we did not find a linear relationship between geographic distance and root AMF community composition (Mantel, p=0.4).
Together, these results suggest that host identity has a greater effect on AMF community composition in roots than the spatial structuring of AMF in soil at these distances.
Soil effects on AMF communities
According to USDA-NRCS soil maps, the section of Kingston Prairie Preserve from which we sampled has a heterogeneous soil composition, with three different soil types, ranging from silt loam to very cobbly loam. The prairie is dissected by a seasonal creek that could function both in limiting dispersal (for example across the creek) and promoting dispersal (for example along the creek) as well as affecting soil moisture characteristics. We found no effect of soil type or proximity to the creek on
AMF community composition, although soil type and properties have been shown to affect AMF community in other studies (Anderson et al. 1984, Lekberg et al. 2007,
Porter et al. 1987).
Nestedness of the plant-AMF network
One approach to analyzing mutualistic networks is to assess the degree of nestedness of the interactions, i.e. whether specialist species of one of the symbiont

85 pairs tend to interact only with generalists species of the other symbiotic partners
(Bascompte et al. 2003). We found a significantly nesteded structure to the plant-AMF partnerships as also found by Montesinos-Navarro et al .
(2012), although we investigated a smaller data set. While such nested structure is not unexpected when analyzing a data set with a skewed distribution of species observations (Blüthgen et al.
2008), which the AMF OTUs showed, the NODF nestedness metric is more robust to such distributions than the original temparature metric for measuring nestedness
(Almeida-Neto et al. 2008). The finding of nestedness highlights a lack of evidence for specialist-specialist associations. Such specialist associations could be seen as connections between rare AMF, which more highly specialized AMF would be expected to be in a limited plant species sampling, and the host species associated with fewer AMF symbiont types.
Methodological considerations
We chose to assess AMF communities in roots via PCR of individual OTUs found in the pooled root AMF community, which is not a novel method (Gollotte et al.
2004) but is used infrequently. This approach has the benefit of being more affordable than next generation sequencing or cloning many individual amplicons and sequencing dozens of clones from each. It also may provide a more definitive assessment of the presence or absence of each OTU in a given sample than cloningand-sequencing based methods, which may fail to find, within the number of sequences performed, less abundant OTUs because of the higher probability of

86 sampling and sequencing bacterial colonies containing the more abundant OTU types.
Some newer environmental surveys of AMF use next-generation-sequencing approaches, a trend that is likely to continue, particularly if the price per sample continues to decline. One disadvantage in the application of that method is the tendency to pool samples, which impedes analysis at the individual plant level, because of the high per-sample cost.
Admittedly, the chosen method for surveying AMF communities has its limitations. First, we knowingly tested for only part of the OTUs found from an initial survey of roots due to the complexities of designing primers for all OTUs. Analyses of differences in community composition such as MRPP and Mantel tests remain informative even when a portion of the data reflecting actual community composition is missing. It is expected that numbers of OTUs per plant are higher in reality than what was detected from testing only a portion of the OTUs found at the prairie.
Therefore, it is possible that patterns in OTU numbers between host species could change with additional OTUs tested, and we interpret these results with that caution.
Additional OTUs tested also could lead to finding more AMF indicator species. The finding that Diversisporales and Archeosporales tended to be absent in host species with fewer AMF OTUs would be unlikely to change because most OTUs from those two orders found in the initial survey were included in this study. We found a higher mean number of OTUs per plant than in many studies that sequence multiple colonies from clonings of PCR amplifications from individual plants using conserved AMF primers (Alguacil et al. 2011, Öpik et al. 2008). Thus, the method used here of directly

surveying for each AMF type may allow for detection of more OTUs than if only a
87 limited number of clones are sequenced leading to the most abundant phylotypes having a higher likelihood of detection.
Second, using presence-absence of PCR products as a proxy for the presence of the actual OTUs is limited by the same issues that affect other PCR based methods, such as T-RFLPs, pyro-sequencing, and clone screening. In all PCR methods, various factors may cause a false negative, a most likely cause in this case being within-OTU polymorphisms at the primer binding sites. Using specific primer sets to survey for individual OTUs does not suffer the same challenges of PCR methods that amplify multiple OTUs with one set of primers targeting conserved sequence regions. With the latter method, there also are problems associated with primer annealing competition between OTUs differing in abundance in the template DNA leading to disproportionate amounts of OTUs in the products relative to starting abundances (von
Wintzingerode et al. 1997).
Third, as performed, this method of PCR is purely a qualitative rather than a quantitative measure of the individual components of the AMF community. We recognize that more sophisticated analyses could be performed with quantitative data, but the collection of such data (for example by using quantitative PCR) is a far more complex undertaking with particular challenges stemming from the general lack of information on the genetic structure of and gene copy numbers in different AMF taxa.
Conclusions

88
In conclusion, we found host identity effects on AMF richness and on the community composition of the AMF colonizing roots but found no effect of geographic distance, soil type or distance from a creek. The effect of host identity is subtle, with little evidence for AMF that specialize on particular plant species.
Understanding the factors that lead to host species’ effects on AMF communities will improve our understanding of the important role of these fungi in plant community dynamics and could facilitate the use of fungi in conservation efforts.
Acknowledgments
We are grateful to the Nature Conservancy for permitting us to sample plants from one of their prairies. We thank Dr. Virginia Weis for providing the lab space and equipment for conducting this research as well as for her helpful comments on the manuscript. We thank the Nevada Genomics Center at the University of Nevada, Reno
(who received funding from the National Center for Research Resources, NIH Grant
Number P20 RR-016464) for their help with the sequencing. Funding for this research was provided by a National Science Foundation Graduate Research Fellowship and a grant from Prairie Biotic Research, Inc. to W. Phillips.

89
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Table 3.1. Primers used in the PCR of individual operational taxonomic units (OTUs) in plant root DNA samples.
OTU Forward Primer (5'-3') Reverse Primer (5'-3')
Annealing temp (°C)
Size of product
(bp)
1_2 GCCTCGCTGTTTGCATG GTCGTTTAAAGCCATTACGC* 54.5 303
28
31
32
33
24
25
26
27
5
7
8
9_10
TGAAATTGTTGAAAGGGAAACGA**
GTGGGCGTATGAGATCTTCATT
CTTCGTCGCTTGGGATACC
GGCTTCGATCTCGTATCGG
11
12
GCCTTTGATTTTCGCATCATC
TTGGTGGAATGTAGCTTCGG
15_16_18 GAATGTAGCTTTGGTCTCGTATC
17 TACCTATGGGTAATCAGCCTTTT
CGAGGGAATGTGTCTCACCT
TCGTGCATTTTCTCGCTT
GTTTATGGTTAAACCGTGGATG***
CCCGTTTATGATTGTTCCTTTG***
TGTGTGGGTTCTCCCTCGTA
GACTGTCCAATGGGTGGT
TGATGGTTGTAAAATGGTGGTC
GAGTGTTATAGCCATCAGCCTA
TCTAACCTTTTGACCGTTGC
CTCGGTCCTAATCACCACATT
TCCGGTCATTTTATGGCA
AACCTTTTGACCGCCAACTA
CTTTCAGCACCAAAGTGACAAA
AAAGCACGCTTTCGATACCAA
CGTTCTARCCTTTTGAAAATCACT
CGAAGCTACATTCCTCCAGC
GTCAGCAAAGCTGTCGAAAGTA
TTTATGTTCACTCTAACTTCTAGGT
AACCATCAAGTTGATTCCCTATT
ACCATCTAGTTGATTCCCTACCAA
CCAGGTGACGAGCCACG
CATTCTATTCTGTTGAGGACACA
CTCTGCCCTATTGATTGAACG
TCCTTATCGAATTTATCGTATCTC
61
57
61
61
62.2
58
60
58
58
60
58
59.5
60
60
59
60
140
237
243
245
182
245
185
167
341
168
89
158
220
143
174
157
35 AAAATCAGCTCACAGGCTACTG CTCGTCGCGTCATCTAAAGT
* The reverse primer used for OTUs 1 and 2 was designed to also amplify OTUs 3, 4 and 5
** The forward primer for OTU 5 is a general primer that works for all OTUs
58 134
***The forward primer for OTUs 26 and 27 were designed to nucleotides upstream from the sequences that are currently available in GenBank .

94
Table 3.2. Paired comparison values from multi response permutation procedure
(MRPP) of root AMF community composition between plant species with p-values <
0.05. Although these p-values were uncorrected for multiple comparisons, based on the total of 36 pairwise comparisons calculated, a type I error rate of 5% would yield less than two false positives.
Plant 1 Plant 2
F. idahoensis P. vulgaris
A
0.21
p
0.0009
P. vulgaris E. lanatum
A. millefolium F. idahoensis
F. idahoensis Z. venenosus
F. idahoensis D. nuttallii
A. millefolium E. lanatum
P. vulgaris B. coronaria
F. idahoensis B. coronaria
A. millefolium D. nuttallii
F. idahoensis P. secunda
A. millefolium B. coronaria
A. millefolium P. vulgaris
A. millefolium Z. venenosus
0.12
0.13
0.13
0.14
0.12
0.17
0.16
0.11
0.12
0.13
0.08
0.08
0.0048
0.0087
0.0094
0.0140
0.0143
0.0152
0.0217
0.0229
0.0305
0.0397
0.0402
0.0434

95
Figure 3.1. Phylogenetic tree showing the Kingston Prairie operational taxonomic units (OTUs) used in the community analysis. Black circles indicate single OTUs tested by PCR, grey circles indicate OTUS tested as a group, and open circles show
OTUs found in the field survey but not tested by PCR. This consensus maximum likelihood tree was constructed based on ribosomal RNA gene sequences with
Paraglomus occultum used as the outgroup. Branch support values from 100 bootstrap replicates are shown. The Kingston Prairie sequences are labeled with ‘KP,’ their assigned OTU number and the accession number for the sequence used. Sequences from Genbank begin with their accession number. Consensus sequences from Krüger et al. (2012) Figure 1 begin with “C ##.”

98
70
98
Figure 3.1
99
77
98
69
90
97
89
98
99
97
99
100
97
73
89
100
38
65
100
92
100
98
95
81
92
77
100
84
99
44
61
30
95
54
74
77
95
100
100
60
100
100
77
72
89
61
100
100
49
100
100
32
71
100
100
100
49
100
100
41
94
99
100
97
100
97
47
100
42
100
42
67
100
100
93
77
100
50
91
100
93
100
43
39
63
100
93
100
KP01 JF717469
HQ895792 ENV
EU380045 ENV
KP02 JF717517
KP03 JF717464
HE775302 ENV
KP04 JF717432
AM040432 ENV
EU379997 ENV
KP05 JF717434
JF439189 UnPub
KP06 JF717607
AB369768 ENV
KP07 JF717488
EU380048 ENV
KP08 JF717527
KP09 JF717614
AB369760 ENV
KP18 JF717500
KP10 JF717452
AB369759 ENV
KP29 JF717462
FR750095 Rhizophagus sp. MUCL 43208
AB369739 ENV
KP12 JF717487
KP13 JF717583
KP14 JF717511
FR772327 ENV
C 33 Sclerocystis sinuosa MD126
KP15 JF717578
HE775302 ENV
KP16 JF717570 (4)
AJ854593 ENV
KP17 JF717495
AJ854592 ENV
KP19 JF717596
KP21 JF717538
C 29 Rhizophagus irregularis DAOM197198
KP20 JF717471
C 30 Rhizophagus irregularis AFTOL-ID845
C 31 Rhizophagus proliferus MUCL41827
C 32 Rhizophagus intraradices FL208
C 24 Funneliformis mosseae BEG12
C 25 Funneliformis coronatus BEG28
C 26 Funneliformis sp. WUM3
C 27 Funneliformis caledonius BEG20
C 22 Glomus macrocarpum
KP11 JF717475
FR750203 Glomus sp. Att565-7
C 20 Glomus sp. W3347/Att565-7
C 34 Claroideoglomus sp. W3349/Att565-11
KP30 JF717489
C 35 Claroideoglomus luteum SA101
KP24 JF717621
KP25 JF717587
C 05 Acaulospora brasiliensis
C 04 Acaulospora laevis AU211
FR750172 Acaulospora entreriana
KP26 JF717598
FR750156 Acaulospora spinosa
C03 Acaulospora spinosa W3574
FR692351 Acaulospora scrobiculata
AM040293 Acaulospora longula
KP27 JF717591
FM876789 Acaulospora cavernata
C 01 Acaulospora cavernata BEG33
C 02 Acaulospora sieverdingii WUM18
KP28 JF717567
KP36 JF717558
C 06 Acaulospora lacunosa BEG78
C 11 Diversispora epigaea BEG47
C 07 Diversispora spurca
C 09 Diversispora celata BEG231
KP22 JF717509)
EU252109 ENV
KP23 JF717626
C 18 Scutellospora spinosissima W3009/Att664-1
C 15 Gigaspora rosea DAOM194757
C 12 Scutellospora heterogama BEG35
C 19 Pacispora scintillans W4545
KP31 JF717581
KP32 JF717553
FR750037 Archaeospora trappei
C 38 Archaeospora schenckii
KP35 JF717625
HQ128660 ENV
AY639362 ENV
KP33 JF717594
FN547527 Ambispora appendicula
C 36 Ambispora fennica
FJ461888 UnPub
KP34 JF717576
C 37 Geosiphon pyriformis GEO1
C 39 Paraglomus occultum IA702
96

97
12
10
8
6
4
2
0
ACMI PRVU POGR BRGR FERO DEOR POSE ZIVE ERLA
Figure 3.2. The mean number (± S.D.) of AMF operational taxonomic units detected per host individual within the plant species studied. (ACMI= Achillea millefolium ,
BRGR= Brodiaea coronaria , DEOR= Delphinium nuttallii , ERLA= Eriophyllum lanatum , FERO= Festuca idahoensis , POGR= Potentilla gracilis , POSE= Poa secunda ,
PRVU= Prunella vulgaris , and ZIVE= Zigadenus venenosus )

Figure 3.3. Two-way dendrogram of AMF OTU occurrences in individual plants. A positive PCR for an OTU in a plant is represented by a filled in circle. Individual plants are in the column to the left, and AMF OTUs are listed at the top. Names of plants of the same species are the same color. ACMI= Achillea millefolium , BRGR=
Brodiaea coronaria , DEOR= Delphinium nuttallii , ERLA= Eriophyllum lanatum ,
FERO= Festuca idahoensis , POGR= Potentilla gracilis , POSE= Poa secunda ,
PRVU= Prunella vulgaris , and ZIVE= Zigadenus venenosus )
98

Matrix Coding
Presence Absence
0 25 50 75 100
POSE50
POSE37
ERLA34
ZIVE61
ZIVE14
ZIVE60
ERLA77
ACMI2
PRVU44
ERLA51
POGR71
FERO1
POSE46
DEOR35
DEOR24
DEOR12
DEOR8
PRVU5
BRGR78
ZIVE54
POSE41
ACMI62
ACMI70
PRVU56
PRVU55
PRVU16
PRVU53
DEOR18
ERLA7
ZIVE48
DEOR73
BRGR42
POGR17
POGR58
ACMI10
FERO21
ACMI59
ACMI40
POSE29
FERO27
FERO23
FERO13
POSE79
POGR38
FERO33
FERO30
ERLA3
ERLA11
BRGR45
BRGR15
Figure 3.3
99

20
15
10
5
0
Total PRVU ACMI POSE DEOR POGR ZIVE ERLA FERO BRGR
Figure 3.4. The total number of OTUs detected within plants of different species, separated by AMF family. The total column represents all OTUs surveyed for by individual PCRs. ACMI= Achillea millefolium , BRGR= Brodiaea coronaria ,
DEOR= Delphinium nuttallii , ERLA= Eriophyllum lanatum , FERO= Festuca idahoensis , POGR= Potentilla gracilis , POSE= Poa secunda , PRVU= Prunella vulgaris , and ZIVE= Zigadenus venenosus )
100

101
PRVU
KP01_02
KP9_10
ACMI
KP07
KP12
Glomerales
POSE
KP17
DEOR
POGR
ZIVE
ERLA
FERO
BRGR
KP08
KP05
KP11
KP15_16
KP25
KP24
KP27
KP28
KP26
KP35
KP32
KP31
KP33
Diversisporales
Archaeosporales
Figure 3.5. The network of interactions between plants (left) and arbuscular mycorrhizal fungi operational taxonomic units (right). ACMI= Achillea millefolium ,
BRGR= Brodiaea coronaria , DEOR= Delphinium nuttallii , ERLA= Eriophyllum lanatum , FERO= Festuca idahoensis , POGR= Potentilla gracilis , POSE= Poa secunda ,
PRVU= Prunella vulgaris , and ZIVE= Zigadenus venenosus )

Chapter 4: Host species identity and community context influences arbuscular mycorrhizal fungal community in plant roots
Wendy S. Phillips
Eric W. Seabloom
To be submitted to:
102
New Phytologist
Lancaster, U.K.

103
Abstract
Arbuscular mycorrhizal fungi (AMF) have important effects on plant fitness, and those effects vary depending on the specific combination of plant and AMF species. However, we have little understanding of the factors influencing the community of AMF with which plants associate. Here we examine the impacts of host identity and host community composition on AMF community composition. We grew two widespread AMF hosts, Achillea millefolium and Prunella vulgaris, and two endangered AMF hosts, Erigeron decumbens and Lomatium bradshawii , either individually or in a community with individuals of all four species. All plants were provided with an identical inoculum of AMF collected from a remnant prairie home to all four species. AMF community composition and diversity varied among hosts growing in identical soils inoculated with the same AMF species, confirming host identity impacts the identity of AMF with which plants associate. In addition, community context affected the richness of AMF in roots, causing a decline in the host that had the richest community when grown individually. Of the two endangered plants studied, only one was regularly colonized by AMF, but it did not form unique associations with any of the AMF observed, suggesting that it is not dependent on specialized AMF symbionts.
Introduction
Most terrestrial plant communities are interwoven with the hyphal networks formed by communities of underground arbuscular mycorrhizal fungi (AMF).

104
Through the exchange of resources, the two partners of this symbiosis can have great effects on each other. AMF affect various aspects of plant functioning, including growth rates (Klironomos 2003), tissue nutrient concentration (Hoeksema et al. 2010), and outcomes of competitive interactions (Scheublin et al. 2007). AMF effects are seen not just at the individual plant level, but also on community wide levels, with increasing AMF diversity resulting in increases in plant community diversity and productivity (van der Heijden et al. 1998b). Likewise, plants affect growth and sporulation rates of AMF (Bever et al. 1996), and plant identify affects AMF community composition (Eom et al. 2000).
In the plant-AMF relationship, partner affiliation often matters, with the direction and magnitude of a plant’s responses dependent on the identity of its AMF associate (Klironomos 2003). Thus, factors that determine AMF community composition have important implications for understanding plant community dynamics (van der Heijden et al. 1998a). A better understanding of the patterning of plant-AMF partnerships might also expand the tool set for plant conservation (Bothe et al. 2010). If plants, particularly ones that are restoration targets, are selective in their partnerships, supplying an appropriate mix of AMF could increase population survival rates (Zubek et al. 2009). Historically, plant-AMF pairings were considered to occur randomly with respect to partner identities (Smith & Read 1997), however more recent evidence, particularly from observational field studies, indicates that plant-AMF combinations are not random (Davison et al. 2011, Helgason et al. 2002,
Vandenkoornhuyse et al. 2003, Vandenkoornhuyse et al. 2002). In an analysis of the

105 root AMF communities in nine native plant species at a remnant bunchgrass prairie in
Oregon, we also found that distinct AMF communities form in different host species
(Chapter 3).
While informative, observational studies cannot distinguish the causal nature of the variation in AMF communities among host species. Specifically, observational studies cannot determine whether the non-random distribution of AMF among host species resulted directly from partner selectivity or indirectly from abiotic niche differentiation exhibited by both the plants and the fungi or other co-varying factors.
To control for the effect of possible co-varying environmental factors influencing species distributions, we conducted an experimental study with plants grown in the same abiotic environment and exposed to an identical pool of AMF propagules to see if AMF communities would assemble non-randomly by host species. Furthermore, we tested whether the formation of plant-AMF partnerships is mediated by host community interactions. Several studies have shown an effect of neighboring plants on
AMF community composition in soil and roots (Hausmann & Hawkes 2009,
Mummey et al. 2005, Stinson et al. 2006). Neighboring plants could indirectly affect
AMF communities in the roots of neighbors by altering the composition of AMF in the rhizosphere of it and its neighbor (Broz et al. 2007). Alternately, interactions between two plants could alter the root physiology of the plants, with subsequent effects on root colonization by AMF (Bais et al. 2003). To see if any host effects on
AMF community seen with plants grown individually would be maintained when neighbors are present, we also grew plants in communities with four species. Few

106 others have performed such experimental studies in order to isolate the effect of host identity on root AMF community composition (Pivato et al. 2007).
We used two common plant species that were included in the studies of
Chapters 2 and 3, Achillea millefolium and Prunella vulgaris , and added to this study two federally listed endangered plants, Erigeron decumbens and Lomatium bradshawii , that co-occur at the site of the original field survey. Because destructive sampling of naturally growing endangered plants could have substantial negative impacts on already threatened plant populations, an experimental study starting from seeds provides a less detrimental approach to investigating AMF communities in such plants. Determining the AMF with which endangered plants associate could benefit conservation of those plants, particularly if specialist relationships are formed between the two (Bothe et al. 2010). If these plants associate with unique AMF that increase their fitness, being sure the appropriate AMF taxa are available in the soil could improve plant establishment and persistence in existing or reintroduced populations.
In summary, in this research we focus on three questions: 1. Does host identity affect AMF community composition in roots?; 2. Does community context affect
AMF community composition in roots?; and 3. Do two endangered plant species associate with unique AMF? To address these questions, we grew plants both individually and in a community containing one plant of each of the four species. All plants were exposed to the same AMF inoculum collected from the Kingston Prairie
Preserve, where they all co-occur.

107
Methods
Seeds and soil
Seeds for the experiment were collected in the summer of 2008 from Kingston
Prairie Preserve (site described in Chapter 2) from two common native perennial forbs, Achillea millefolium L. and Prunella vulgaris L., and from two federally listed endangered (both status LE) native perennial forbs, Erigeron decumbens Nutt. and
Lomatium bradshawii (Rose ex Mathias) Mathias & Constance. Seeds were collected from at least 20 different plants (except for E. decumbens for which only about 10 seeding plants could be located) haphazardly chosen from throughout each plant species’ range at Kingston Prairie. Collected seeds within a species were mixed together for subsequent use. E. decumbens , P. vulgaris and L. bradshawii seeds were germinated on sterile coarse sand in petri dishes. Seedlings were initially transferred to
2.5 cm diameter/12.1 cm height/49 ml volume Cone-tainers (RLC3; Stuewe & Sons,
Tangent, OR, U.S.A.) filled with coarse sterile sand and grown until they were three to four weeks old, when they were transplanted to experimental pots. At the time seedlings were transplanted, seeds of A. millefolium were sown directly in the pots because preliminary germination trials indicated their seeds germinated and grew much more quickly than the other species. A. millefolium seedlings were thinned to a single individual in a pot within a week post-germination.
The potting mix consisted of field soil (collected from one spot immediately adjacent to Kingston Prairie and sieved through approximately 5mm mesh): sand: coconut fiber: worm castings: perlite (6:3:3:2:2, v:v). The potting mix components

were autoclaved pre-mixing two times over 2-3 days (except worm castings, which
108 were autoclaved 3 times) for at least 1.5 hours at 121°C per sterilization cycle. Poststerilization the potting mix had a pH of 5.7 with 16.3 ppm ammoniacal-N, 36.4 ppm nitrate-N, 1 ppm P and 80 ppm K, which was approximately 4x higher nitrate-N but
1/6 the P concentrations as in field soil. A field soil inoculum mix was created from a limited amount of soil, as per the collecting guidelines of the preserve steward, collected on 01 May 2009 from 20 locations within Kingston Prairie. In order to maximize the diversity of AMF from Kingston Prairie included in the soil inoculum, five soil samples were collected where L. bradshawii was abundant and five were collected where E. decumbens was abundant; the remaining 10 samples were taken from broadly distributed spots near plants of the more widely dispersed A. millefolium or P. vulgaris . The field soil was air dried so as to remove enough moisture for easy sieving, sieved through approximately 2 mm mesh, and thoroughly mixed before use as inoculum.
Plants were grown either individually in 7.3 cm width/22.9 cm height/960 ml volume pots (Anderson Band AB39, Stuewe & Sons Inc., Tangent, OR, U.S.A) or in a community of four with one individual from each species in 12.7 cm width/20.3 cm height/2830 ml volume pots (Anderson Band AB58). We used the larger size pot for community grown plants to minimize changes in overall root density, although the end soil volumes per plant still varied slightly, with 960 mls/plant when grown individually and 707 ml/plant when grown in community. Seedlings were randomly assigned to pots and planted on 18 or 19 May 2009. Pots were subjected to one of

109 three treatments (10 pots per treatment): sterile potting mix only, sterile potting mix with a microbial wash (as a control for effects of non-AMF soil microbes), or sterile mix with 50-51 g of field soil inoculum (containing AMF and other soil microbes).
For pots with field soil inoculum treatment, pots were filled to 8 cm from the rim with sterile potting mix, a layer of field inoculum was applied, and young plants were planted just above that in more sterile potting mix. The microbial wash was prepared by filtering a solution of 650 g soil inoculum in 1 L sterile water through a series of filters (850, 58 and 20 µm). On 20 May 2009, 20 ml of microbial wash was applied to treated pots. Plants were grown in a growth chamber under fluorescent grow lights with a 16 h photoperiod. Plants were watered every other day to field capacity. Once a week for ten weeks, as a measure of plant growth, the total number of leaves and the length of the longest leaf for each plant were recorded.
Plants were destructively harvested between 7 and 19 August 2009, after 11-12 weeks of growth in experimental pots. All root systems were thoroughly washed and those from community pots were carefully separated from each other in water baths. A subset of fine root tissue, checked for attachment to the appropriate stem, was collected and one portion was stored at -80°C for DNA extraction and another portion stored in 70% ethanol for later staining. Although we intended to collect all remaining root tissue for belowground biomass measures, the roots had so thoroughly integrated into the coconut fibers in the potting mix that separation of the two was untenable
(Fig. 4.1). Percent root length colonized was assessed by first staining roots with

110 trypan blue and then scoring them using a line intersect method under 100x magnification (Koide & Mooney 1987).
Molecular analysis of AMF communities in roots
DNA was extracted from roots of the plants exposed to field inoculum using a range of 13-284 mg tissue (median 91 mg) with the DNAeasy Plant Mini kit (Qiagen,
Valencia, CA, U.S.A. ) as described previously (Chapter 2). We conducted an initial survey to determine the collective diversity of AMF colonizing roots at the end of the experiment. First, we PCR amplified genomic DNA encoding large subunit ribosomal
RNA (LSU rDNA) from each plant using the PCR primers AM-SSU-F and
AM-LSU-Rx followed by a nested reaction with AM-SSU-F and AM-LSU-Ri using amplification conditions as described in Chapter 2. We pooled PCR products from samples of the same species and grown in the same community context so as to create approximately equimolar mixes of the products. We cloned those combined products into the pGEM-T Easy vector (Promega, Madison, Wisconsin, U.S.A.) Random colonies were picked (36 from A. millefolium grown individually, 36 from A. millefolium grown in community, 35 from E. decumbens grown individually, 41 from
E. decumbens grown in community, 53 from P. vulgaris grown individually, 53 from
P. vulgaris grown in community, and 16 from one plant of L. bradshawii ) and sequenced using the primer AM-Seq (Chapter 2). Sequencing was done by Functional
Biosciences, Inc. (Madison, Wisconsin, U.S.A.).

111
Sequences were checked for chimeras using Bellerophon (Huber et al. 2004).
Sequences were aligned using the MAFFT online server (MAFFT version 6; http://mafft.cbrc.jp/alignment/server/) with the FFT-NS-i strategy (Katoh et al. 2002), followed by additional editing by hand in MEGA4 (Tamura et al. 2007). Phylogenetic analysis was performed using the PHYLIP program suite v3.69 (Felsenstein 1993). A maximum likelihood consensus tree was constructed from100 bootstrap replicates.
Sequences were assigned to OTUs using the program MOTHUR based on ≥97% sequence identity using the average neighbor clustering method (Schloss et al. 2009), with the exception of OTUs 22, 32 and 37, where the sequence identity criteria was loosened to include sequences with ≥95% identity within a single monophyletic group.
To determine if sequence sampling was sufficient to uncover most AMF diversity, we performed rarefaction using Analytic Rarefaction 1.3 (Holland 2003). Representative sequences from each OTU were compared against those in public databases using a
BLASTN search (http://www.ncbi.nlm.nih.gov), and sequences from environmental samples were included in the presented phylogenetic tree when they were the closest matches and no sequences from described taxa with ≥97% identity were found.
Unique sequences detected in this study were deposited in GenBank under accession numbers JX848742-JX848972.
To assess AMF root community composition, we tested for PCR amplification of individual OTUs from the root DNA of individual plants. To do this we designed primers that would specifically amplify individual OTUs, as described in Chapter 3.
For those OTUs represented by more than a single sequence and for which at least two

separated sequence regions were found that were conserved within all sequences of
112 that OTU but that had nucleotide differences with sequences from all other OTUs and were amenable to primer placement, a pair of OTU specific primers were designed.
Newly designed primers for this study are given in Table 4.1. Altogether the presence of 10 OTUs was assessed. Presence of each OTU was assayed in each root system that had yielded a positive reaction with the general AMF primers. Five separate PCR runs were performed for each OTU. For three of those, templates were generated from three sets of AM-SSU-F – AM-LSU-Rx reactions using three separate dilutions of stock root DNA as template. For those reactions 5 µl of a 1/20 dilution of stock DNA was used in a 20 µl reaction and cycled 28 times (94°C for 30 s, 58°C for 30 s, 72°C for 120 s). Then for nested amplification of individual OTUs, 5 µl of 1/50 diluted
AM-SSU-F – AM-LSU-Rx PCR was used as template in a 20 µl reaction containing
0.25 µm OTU specific primers and cycled 35 times (94°C for 30 s, annealing see
Table 4.1 for 30 s, 72°C for 40 s). Two additional sets of reactions were performed directly from root DNA using 47 amplification cycles. An OTU was counted as present if it occurred in at least two of the five PCR reactions.
Statistical analyses
To test the null hypothesis of no difference in AMF community composition between host species, we used multi-response permutation procedure (MRPP;
McCune & Grace 2002, Mielke 1984) in PC-ORD v6.07 (McCune & Mefford 2011) with Bray-Curtis distances to calculate between sample dissimilarities. This method

compares dissimilarities of communities within and among groups using a Monte-
113
Carlo approach and yields an overall chance-corrected within group agreement (A) value and p-value as well as those values for all pairwise combinations. To test for an effect of host species on AMF community composition, we used plant species identity as the grouping assignment. To test for changes in AMF community based on community context, samples were classified into one of six groups based on both species and growing environment.
Indicator Species Analysis (ISA; Dufrene & Legendre 1997) is most commonly used to identify species that have high fidelity to and a high frequency of occurrence in particular ecological sites. Treating plant host species as analogs of ecological sites, we tested whether any AMF were good indicator species for plant hosts with ISA in PC-ORD v6.07 using a Monte Carlo test for significance with 1,000 randomizations.
Simple linear regression analyses were performed in Excel to test for correlations between % root length colonized and plant growth; number of OTUs and plant growth; number of OTUs and % root length colonized; and mass of root tissue used in the DNA extraction and number of OTUs detected. We used the number of leaves multiplied by the length of the longest leaf at week nine as a proxy for plant growth. Analysis of variance (ANOVA) was performed in Excel to test for differences in growth within species among treatments. We used R version 2.14 (R Development
Core Team 1990) to fit a general linear model for evaluating effects of host species identity and community context on AMF richness and also to evaluate the effects of

114 host identity and community context on overall AMF colonization using logistic regression.
Results
Plant growth
All plants grew over the course of the experiment, although at the end of the experiment L.
bradshawii had much less above-ground and below-ground biomass than the other species. Typical root systems at the end of the experiment are shown in
Fig. 4.1. There were no differences in plant growth within species between treatments
(ANOVA, p>0.05), and there were no correlations between plant growth and the number of OTUs detected (Linear regression, p>0.05) or between % root length colonized and plant growth (Linear regression, p>0.05).
Overall AMF diversity
Of the 248 clones sequenced, 241 yielded good quality sequences that were clearly in the Glomeromycota based on BLASTN alignments. Rarefaction of the sequences suggests the sample sizes were sufficient to capture most of the OTU diversity in five samples, where the rarefaction curve approached an asymptote (Fig.
4.2). The exception is the P. vulgaris individually grown set, the curve for which suggests some OTUs present in that sample remained undetected. We detected 13
OTUs, five in Glomerales, three in Diversisporales, and five in Archaeosporales (Fig.
4.3).

115
Host and context effects on AMF communities
Based on presence/absence of a PCR product with the general AMF primers, the endangered species, L.
bradshawii , had a lower overall colonization rate than the three other species with only two out of 20 plants showing colonization (Logistic regression, p<0.01). Positive amplification rates for the other plants were P. vulgaris
100%, A. millefolium 60%, and E. decumbens 63%. A variety of factors others than absence of AMF DNA in the template can cause amplification of AMF rDNA to fail.
We made numerous attempts at amplification with templates that did not immediately yield a PCR product of the appropriate size, particularly varying DNA dilutions, because we have found that root extractions often contain PCR inhibitors that lose their effect upon dilution. Because L.
bradshawii did not produce enough tissue for both DNA extraction and root staining, the only measure for its colonization available was PCR amplification.
Host species identity affected richness of the AMF root communities, with P. vulgaris having the richest AMF communities (Linear regression, p=0.03; Fig. 4.4).
Plant community context affected AMF richness (Linear regression, p=0.02; Fig. 4.4), with AMF richness in P. vulgaris roots lower in plants grown in community than in those grown individually. Cumulative AMF richness (AMF OTUs across all hosts in a pot) did not differ significantly between single species and four species pots (t-test, p=0.99). Furthermore there was no difference in total pot AMF richness between pots of P. vulgaris (the host with the highest AMF richness) grown individually and pots of plants grown in community (t-test, p=0.09; Fig. 4.5). Numbers of OTUs detected did

116 not correlate with the mass of root tissue used for DNA extractions (Linear regression, p=0.6).
Host species identity affected AMF community composition in roots (MRPP;
A= 0.03, p<0.05; Fig. 4.6). Pairwise comparisons of AMF communities indicate that the AMF community in roots of P. vulgaris grown with other plants differed from that in all other roots and also indicate that, although the P. vulgaris AMF community changed significantly with growth context (with or without other plants), the AMF communities of the other plant species did not (Table 4.2). Indicator species analysis showed three AMF OTUs, all in the Archaeosporales, characteristically associated with P. vulgaris : OTU 32 (p=0.02), OTU 39 (p=0.008), and OTU 40 (p=0.017). Of those, just OTU 39 was found exclusively in P. vulgaris (in seven individuals).
Discussion
We found strong differences in the richness and composition of AMF communities among four grassland species. In addition, we found that richness of
AMF communities in individual plant roots was reduced in mixed plant communities.
The correlation between host identity and root AMF community composition we detected in our field study persisted under experimental conditions. When we provided plant hosts with a uniform set of AMF propagules, AMF community assembly in roots varied between host species. In particular, we observed that plant species identity impacts both the richness and the structure of root AMF communities.

117
AMF inoculation and plant growth
We found no effect of AMF inoculation on plant growth. This result contrasts with results from meta-analyses in which the majority of studies recorded a positive growth response in plants exposed to AMF relative to non-exposed plants (Hoeksema et al. 2010, Lekberg & Koide 2005). Although it is a less common result, several studies included in those meta-analyses recorded no difference in growth between
AMF-treated and non-treated plants. Hoeksema et al. (2010) found that fertilization with nitrogen explained much of the growth differential, with larger plant responses to
AMF in plants not nitrogen fertilized. Although we did not fertilize the plants in our study, the potting mix we used had approximately 4x higher nitrate-nitrogen than the field soil of the prairie where these plants occur, possibly as a result of autoclaving the potting mix components. Another factor possibly contributing to the lack of increased growth response with AMF inoculation is the somewhat small quantity of field soil used as inoculum. Due to the limited inoculum supplied and the limited duration of the experiment, a sufficient enough extraradical hyphal network to effectively enhance nutrient supply to plants may not have been established. In natural settings, mycorrhizal colonization of seedlings is likely most often initiated from hyphae extending from pre-existing networks. Under those conditions, the hyphae already are linked to a carbon source and may provide seedlings with a higher nutrient supply to carbon demand ratio than do hyphae not connected to older plants (van der Heijden &
Horton 2009).

118
Host identity affects AMF community composition
When grown individually, P. vulgaris had a richer community of AMF than the other plants, indicating host identity can influence the number of AMF in plant roots. Other studies have found such differences between host species in the richness of their root AMF communities (Alguacil et al. 2012, Vandenkoornhuyse et al. 2003).
If hosts have mechanisms for regulating AMF colonization such differences in root
AMF richness could result. For example, plants could differ in their selectivity
(Scheublin et al. 2004), with more selective plants actively regulating their root community so as to host fewer AMF. Alternately, in the absence of host control, competitive interactions between fungi could act as the primary factor regulating final
AMF root community composition. Differences in the availability of root resources between host species (Roumet et al. 2006, Urcelay et al. 2011) could lead to host dependent effects on the competitive environments in roots that subsequently affect
AMF richness. Even without competitive interactions, assuming fungi would maintain associations with higher quality hosts, a higher diversity of AMF would be expected on hosts that provide more resources.
Notably, a series of experiments showing AMF taxa differentially affect a range of plant variables, such as overall biomass, leaf number, root length, nitrogen content and clonal growth rate, focused on P. vulgaris (Streitwolf-Engel & Boller
1997, van der Heijden 2004, van der Heijden et al. 2003). In contrast to our experiment, in all those studies P. vulgaris had large increases in growth when grown with at least one AMF taxon as compared to plants grown without any AMF,

consistent with its characterization as being highly mycorrhizal dependent (van der
119
Heijden 2002). That P. vulgaris often has greatly increased growth when colonized with AMF may relate to the high diversity and richness of AMF with which it associates. A plant that is highly dependent on AMF for its fitness would benefit from being a generalist, particularly in situations where AMF diversity is low. Future studies comparing the mycorrhizal responsiveness of plant species with the richness of
AMF community they usually carry could elucidate this possible correlation between mycorrhizal responsiveness and selectivity.
Not only did host species affect root AMF richness, it also affected AMF community structure, with the AMF community in roots of P. vulgaris grown individually differing from all other root communities (Table 4.2; Fig. 4.5).
Differences in AMF community could be due in large part to the higher richness of individually grown P. vulgaris . However, results of Indicator Species Analysis yield three AMF OTUs with relatively high fidelity to P. vulgaris that occur both in plants grown individually and in community. Interestingly, those three AMF OTUs are all in the basal Archaeosporales, an order for which less information is available due to the infrequency with which it is included in experimental work and a historic discrepancy against the group by primers commonly used in environmental surveys (Chapter 2,
Lee et al. 2008). The greater abundance of these Archaeosporales taxa particularly with P. vulgaris could be due to a lower selectivity of P. vulgaris or to unique root resources preferred or required by Archaeosporales species supplied by that host and not others.

120
We know of two other studies investigating root AMF communities in the two common plants included in this experiment. A study comparing the root AMF communities in A. millefolium and Festuca pratensis collected from a field in Sweden found no host effect on AMF community composition (Santos et al. 2006). A different study comparing the root AMF communities in P. vulgaris and Antennaria dioica collected from the same field in Sweden found significantly different AMF communities between species and that the AMF community of P. vulgaris was richer
(Santos-González et al. 2007), consistent with our results. Here we detected a significant difference in AMF community between P. vulgaris and A. millefolium but not between the latter and E. decumbens .
Community context affects AMF community composition
The abundance of AMF taxa in P. vulgaris roots was affected by community context, with plants grown in community with other species hosting fewer AMF taxa than plants grown individually. Such a shift could result from biotic or abiotic changes caused by increased plant density or diversity in the pots, although larger pots were used for community grown plants to minimize density differences. For example, interactions between plants could cause changes to root physiology that then impact plant or AMF mechanisms controlling colonization by different AMF (Callaway et al.
2008, Stinson et al. 2006). Increased plant diversity also could lead to shifts in such abiotic conditions as soil nutrient composition, which could directly affect both plants and AMF and indirectly affect the interactions between the two (Zak et al. 2003).

121
Comparisons of overall AMF richness in pots between pots in which P. vulgaris was grown individually and in community showed a non-significant (p=0.09) decline in
AMF richness (Fig. 4.5). The decline in richness of AMF in roots of P.
vulgaris in community combined with similar AMF richness in pots of plant communities and pots of P. vulgaris grown alone suggests that some of the AMF colonizing P. vulgaris grown individually are migrating to other hosts when it is grown in community. We interpret this result loosely as this statistical result borders on significance and the size of this data set is relatively small.
AMF communities in endangered plants
We included two endangered plants in this experiment to address the question of whether these plants, which are in particular need of conservation, associate with unique AMF communities (Bothe et al. 2010). If so, such information could prove valuable to conservation efforts if any unique AMF were found and shown in future experiments to differentially promote growth and reproduction of those plants.
However, we did not find such unique associations in either of the plants, and indeed found very low rates of colonization at all in L. bradshawii . Roots of L. bradshawii were much smaller than in any of the other plants, leaving open the possibility that its reduced access to the layer of inoculum affected AMF colonization. Levels of colonization in E. decumbens were similar to that in the common A. millefolium . To our knowledge, very few studies have investigated AMF community composition in endangered plants (Bothe et al. 2010). In a study of the rare plant Pulsatilla patens ,

found in Estonia, Öpik et al.
(2003) did not find AMF that specifically or preferentially colonized it compared with the AMF community housed by a more
122 widespread sister species. Distinct AMF have been found in association with the endangered Ethiopian tree Juniperus procera (Wubet et al. 2006). Although no species preferentially associated with E. decumbens were found in this study, unique relationships between the plant and AMF remain a possibility. For example, colonization dynamics under these experimental conditions likely differed from what occurs in the field (discussed in Chapter 5) such that we may not have detected the same diversity of AMF one would find under natural conditions.
Conclusions
In conclusion, we demonstrated strong effects of host identity on root AMF community composition concordant with observational data. The richness of the AMF community varied between hosts, with a known highly mycorrhizal dependent plant hosting the richest AMF community. Plant community context affected the richness of
AMF in roots, with the richest community observed in P. vulgaris plants grown individually. Of the two endangered plants we studied, only one was regularly colonized by AMF, but it did not form unique associations with any of the AMF detected.
This study adds to the increasing evidence that host species identity affects root AMF communities (Alguacil et al. 2012a, Pivato et al. 2007, Vandenkoornhuyse et al. 2003, Vandenkoornhuyse et al. 2002). Understanding how host identity

123 interplays with other factors such as community context to shape AMF communities will move us towards understanding the complexity of the effects of the symbiosis on plant communities as a whole. In this time of rapidly changing environments and threats posed by quickly spreading invasive species, such an understanding could help protect native plant communities.
Acknowledgments
We are grateful to the Nature Conservancy for allowing us to collect seeds and soil from Kingston Prairie Preserve for this research. We thank Dr. Virginia Weis for providing the lab space and equipment for conducting this research. We acknowledge three undergraduate students who helped with parts of this work: Ellen Dowe,
Amanda Simonds and Charnelle Terawaki. Funding for this research was provided by a National Science Foundation Graduate Research Fellowship and grants from Prairie
Biotic Research, Inc. and the Portland Garden Club to W. Phillips.

124
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Table 4.1. Primers used in the PCR of individual operational taxonomic units (OTUs) in plant root DNA samples.
OTU Forward Primer (5'-3') Reverse Primer (5'-3')
Annealing temp (°C)
Size of product
(bp)
11 GCCTTTGATTTTCGCATCATC CTTTCAGCACCAAAGTGACAAA 60 220
35
36
37
39
22
29
32
34
40
TTTTTAATGTTCCGGGTTGGT
TTGGGATTGCAGCTCAAAAC
TGATGGTTGTAAAATGGTGGTC
CCAGTCGTAAAAAGTCCGTC
AAAATCAGCTCACAGGCTACTG
AAACGATTAAAGTCAGTCGTACTA
GAATCAACTTAGGGGAAACTC
CATCGGTTTCTGTTGCGTC
GTTGTAAAATGGTGGTCGGAAT
CCCGACATCGAATCAGAGG
GAAAGTACGAGGGAGAACCCA
CTCTGCCCTATTGATTGAACG
TCCTTAACGAGTTTACCGTGAC
CTCGTCGCGTCATCTAAAGT
TTCAGCACCATAGTGACATG
CAGCGTCCTTAACGAATTTAC
CAAAAGGCTCCCACCAAA
GCATCCCTGGCAAATTTATCA
60
60
63.8
63.8
63.8
60
60
62
62
502
210
185
205
134
284
268
332
209

129
Table 4.2. Significantly different AMF communities between host groups based on
MRPP pairwise comparisons of AMF community dissimilarities. “Single” indicates plants grown individually and “Community” indicates plants grown in community with other plants. Although these p-values were uncorrected for multiple comparisons, based on the total of 15 pairwise comparisons calculated, a type I error rate of 5% would yield less than one false positive.
Host 1
P. vulgaris Single
P. vulgaris Single
P. vulgaris Single
P. vulgaris Single
P. vulgaris Single vs. vs. vs. vs. vs.
Host 2
E. decumbens Single
P. vulgaris Community
A
0.15 p
0.0005
A. millefolium Community 0.14 0.0008
A. millefolium Single 0.13 0.0010
0.05 0.0292
E. decumbens Community 0.05 0.0299

a b
130 c d
Figure 4.1. Typical root systems at the end of the experiment: a) Achillea millefolium , b) Prunella vulgaris , c) Lomatium bradshawii , and d) Erigeron decumbens . The dark matter is coconut husks within which the roots intertwined.

131
12
10
8
6
4
2
Achillea S
Achillea C
Erigeron S
Erigeron C
Prunella S
Prunella C
0
0 10 20 30 40
50 60
Figure 4.2. Rarefaction of sequencing effort showing the expected number of OTUs detected with the sample sizes on the x-axis. S=plants grown individually, C=plants grown in community. Plant names are abbreviated with their genera.

132
Figure 4.3. A consensus phylogenetic tree showing the Kingston Prairie operational taxonomic units (OTUs) used in the community analysis. This consensus maximum likelihood tree is based on ribosomal RNA gene sequences with Paraglomus occultum used as the outgroup. Branch support values from 100 bootstrap replicates are shown.
Named AMF taxa followed by “C#” indicate consensus sequences from Krüger et al.
(2012). The tree contains one representative sequence for each OTU found in this study, labeled with “KingPr” and its OTU designation followed by the number of sequences from that OTU detected in the initial sequencing survey in parentheses.
Triangles and squares designate Kingston Prairie OTUs not included and included, respectively, in the community analysis. Environmental sequences are labeled with their Genbank accession number followed by “Env.”

98
99
50
64
36
57
86
99
60
88
100
100
100
100
100
100
100
54
100
100
65
100
96
78
100
96
96
63
100
100
100
64
100
70
67
100
50
KingPr02 (25)
KingPr03 (1)
AB561106 Env
KingPr09 (1)
AB369762 Env
Rhizophagus intraradices C32
Rhizophagus irregularis C29
Glomus sp. W3347/Att565-7 C20
KingPr11 (26)
KingPr41 (22)
Glomus macrocarpum C22
EU380157 Env
Funneliformis mosseae C23
Funneliformis caledonius C27
Acaulospora sieverdingii C02
KingPr29 (7)
Acaulospora cavernata C01
Acaulospora laevis C04
Diversispora celata C09
Diversispora spurca C07
Pacispora scintillans C19
KingPr22 (12)
Scutellospora spinosissima C18
Gigaspora rosea C15
Scutellospora heterogama C12
Racocetra castanea C17
KingPr37 (51)
Scutellospora nodosa C16
KingPr34 (17)
Ambispora fennica C36
Geosiphon pyriformis C37
Archaeospora trappei FR750037
KingPr40 (24)
KingPr32 (5)
Archaeospora schenckii C38
HQ128697 Env
KingPr39 (14)
AY639362 Env
KingPr35 (22)
Paraglomus occultum C39
Glomerales
Diversisporales
Archaeosporales

134
Figure 4.4. Richness of AMF communities (mean ±1 S.E.) in roots of 3 different host species ( Achillea millefolium , Erigeron decumbens , Prunella vulgaris ) grown individually or in community with other species .

135
3
2
5
4
1
0
7
6
PRVU indiv All Indiv Community
Figure 4.5: Total AMF richness in pots as measured by mean number (±S.E.) of OTUs detected within a pot for Prunella vulgaris grown individually, all plants ( Prunella vulgaris , Achillea millefolium and Erigeron decumbens ) grown individually, and plants grown in community.

136
Matrix Coding
Presence Absence
0 25 50 75 100
P114
P40
E19
P106
A113
A114
E110
P110
P113
E107
A111
P35
P37
P31
P36
P32
P112
P33
P34
P39
P108
P38
P105
P107
E108
A10
P109
E17
E106
E18
E13
A1
A9
E15
E14
E111
P111
A7
A108
A109
A106
E16
Figure 4.6. Two-way dendrogram of AMF OTU occurrences in individual plants.
(Circles filled=presence). Individual plants are on the vertical axis and AMF OTUs are on the horizontal axis. Plant sample names are color coded by plant species.
A= Achillea millefolium , E= Erigeron decumbens , P= Prunella vulgaris . Plants numbered with double digits were grown individually, and those with triple digit numbers were community grown samples.

137
Drivers of Arbuscular Mycorrhizal Fungal Community Composition in Roots: Hosts,
Neighbors, and Environment
Chapter 5. Conclusion
Summary of findings
Arbuscular mycorrhizal fungi (AMF) are integral contributors to plant communities, affecting both individual plant and community level processes.
Although generally considered a mutualism, the effect of the symbiosis on growth of individual plants depends upon the particular combination of partners, with some combinations leading to negative growth relative to non-mycorrhizal controls
(Klironomos 2003). An understanding of what drives AMF community composition in roots will provide insight into the formation of partner combinations that vary in their effects on plants. An important aspect of investigating that process is the discovery of the diversity of AMF available for symbiotic partnership.
In Chapter 2, I surveyed the AMF present in the roots of 12 plant species at a remnant bunchgrass prairie in the Willamette Valley of Oregon. No other published studies of AMF diversity have focused on the bunchgrass prairie ecosystem. For that study I also designed new primers to specifically amplify all Glomeromycota. I found
36 operational taxonomic units (OTUs), groups of sequences defined by ≥97% sequence identity, a relatively high diversity for a single site compared to studies of other grasslands. Additionally, I found a higher proportion of sequences in the basal order Archaeosporales than in world-wide averages (Öpik et al. 2010). These results could reflect a higher richness and unusual proportions of diversity from different

orders. It also could result from two methodological factors: I surveyed more plant
138 species than assessed in most studies, and I used newly designed primers.
In Chapter 3, I investigated three factors that could potentially influence AMF community assembly in roots: host identity, spatial distribution of plants, and soil characteristics. This involved assessing AMF community composition in the roots of
50 plants of nine species that were used in Chapter 2. I designed 18 OTU-specific primer pairs for a subset of the OTUs detected in Chapter 2 and used them to assay by
PCR for the presence of each OTU in the roots of each plant. AMF richness varied with host identity as did AMF community composition, while neither geographic distance between plants nor soil characteristics correlated with AMF community composition.
The research described in Chapter 4 involved an experimental study with plants grown in the same abiotic environment and exposed to an identical pool of
AMF propagules to determine if AMF communities would assemble non-randomly by host species. In addition, to examine the effect of plant community context on AMF community assembly, I grew plants either individually or in a community of four individuals, each from a different species. For that study, I used two common plants
( Achillea millefolium and Prunella vulgaris ) and two federally endangered species
( Erigeron decumbens and Lomatium bradshawii ). Integration of particular mycorrhizal fungi in the conservation of endangered species could be beneficial if endangered plant species associated with unique AMF species that also improved their fitness (Bothe et al. 2010). However, only one of the two endangered plant species, E.

decumbens , was regularly colonized by AMF in this experiment. I found no unique
139
AMF associated with that endangered plant. Similar to the results presented in Chapter
3, I found that host identity influenced both root AMF richness and community structure. Also, host community context affected the richness of the AMF community, with growth in a plant community resulting in lower richness for one species ( P. vulgaris ), the one with the highest richness when grown individually.
Comparison of results from Chapters 3 and 4
The two widespread plants used in Chapter 4 were also subjects of the AMF community analysis of field plants in Chapter 3. Interestingly, in the field study of
Chapter 3 A. millefolium hosted a richer root AMF community than did P. vulgaris
(Chapter 3: Fig. 3.2). Subsequent analysis of the field study data suggests differences in the overall proportion of AMF occurrences from the different Glomeromycota orders between the two species, with 70%, 20% and 9% of the AMF OTU occurrences in A. millefolium in the Glomerales, Diversisporales, and Archaeosporales, respectively, compared to 51%, 33% and 16% in P. vulgaris (Table 5.1). It is possible that experimental conditions and the manner in which the inoculum was prepared differentially affected the infectivity of propagules of the different orders of AMF. For example, if A. millefolium has a higher tendency to associate with AMF in the
Glomerales than P. vulgaris , as suggested by the field data, but the conditions of the experiment limited colonization by the Glomerales, one would see lower overall colonization in A. millefolium , such as was observed. Comparing the diversity of AMF

140 found in the field survey to that after experimental growth, the largest differences in proportional composition were with the Glomerales and Archaeosporales species, which accounted for 63% and 14%, respectively, of the diversity in the field but 38% and 38% of the diversity detected following the experiment (Table 5.1). The lower richness of Glomerales species following culturing supports the hypothesis that the lower diversity of AMF in cultured A. millefolium resulted from the combination of its propensity to associate with Glomerales and a decrease in the infectivity of
Glomerales taxa under the experimental conditions.
The advantage of growing the plants from seedlings in pots was the ability to control for variations in abiotic and biotic soil environment that would occur in the field. Particularly, by providing all pots with the same amount of thoroughly mixed field soil as a source of AMF, the AMF community available to plants for establishing associations was controlled between pots, which in a field situation is highly spatially variable (Klironomos et al. 1999, Mummey & Rillig 2008). A disadvantage to such experimental manipulation is that the conditions in a pot vary in many ways from the natural conditions usually encountered by seedlings. One important variation in this case is the type of inoculum for establishing symbiotic partnerships. New mycorrhizal associations can develop from spores; hyphal fragments that are or are not associated with root fragments (dead or alive); and hyphae extending from an active hyphal network. It is unknown what proportion of new associations is formed from each of these in a usual field setting, although new colonization is thought to be more commonly derived from hyphal networks or root fragments rather than spores, even

141 when many spores are present (Smith & Read 2008). Under the experimental conditions, there were no existing hyphal networks from which associations could be formed, but there would have been small root fragments and spores.
Experiments comparing the degree of infectivity of different sources of inoculum have shown variations between AMF taxa. Klironomos & Hart (2002) found that Glomus ( sensu lato ) and Acaulospora species colonized roots from spores, extraradical hyphae and infected roots, whereas Gigaspora and Scutullospora species colonized mostly from spores and not at all from extraradical hyphae. In other experiments, Scutellospora species were able to colonize from root fragments
(Braunberger et al. 1996, Tommerup & Abbott 1981). Given that published research indicates that Glomus species ( sensu lato ) colonize roots from a diversity of inoculum sources, it is surprising that the proportion of AMF in the Glomerales was much lower under the cultured conditions. However, in a study of agricultural soils, Glomerales taxa represented a lower amount of the total AMF diversity in tilled soils compared to untilled soils, suggesting they have higher colonization rates from intact hyphal networks (Schalamuk & Cabello 2010). I am unaware of research on infectivity rates of Archaeosporales propagules, but the proportional increase in their diversity detected in culture compared to the field suggests the experimental conditions had less of an effect on their infectivity than on the Glomerales.
Another factor possibly affecting colonization rates was the use of a relatively small amount of inoculum, 50 mg per pot, due to restrictions on the amount of soil I was permitted to collect from the remnant prairie. The inoculum was collected 18 days

142 prior to transplanting seedlings to pots. Other studies suggest that the longevity, including the retention of infectivity, of both spores and hyphal fragments in soil to be far longer than this, on the order of several months (Tommerup & Abbott 1981).
Although I hoped that starting with a small quantity of inoculum would limit initial competition between AMF in the smaller root system of young seedlings and thereby better test for direct host control of colonization, I recognize the possibility that high mortality rates of AMF propagules would more significantly impact smaller inoculum volumes.
Despite possible differences in inoculum potential for the various
Glomeromycota orders in the experiment described in Chapter 4, the evidence remains strong for an effect of host identity on root AMF community composition. All plants still had a uniform set of AMF with which to associate, even if it did not encompass the full diversity that would be found in a field situation. Indeed, I predict from the results of the field study that if there had been more colonization by Glomerales taxa, a larger difference in AMF community composition would have been observed due to a differential association of A. millefolium with Glomerales partners. Without having sampled E. decumbens from the field, it is difficult to predict how greater Glomerales infectivity in the experiment would have affected its resulting AMF community, leaving open the possibility that E. decumbens in the field setting associates with unique Glomerales species.
Future directions

143
More research is needed to make accurate estimations of the total number of
Glomeromycota species. Although there are approximately 230 described species, only about 120 have sequences in public databases, and many of those are only partial rDNA sequences (Krüger et al. 2012). Even though not all described species are maintained in culture collections, sequencing more of the rDNA of those available from culture would increase the data set against which to compare environmental sequences. The increased use of the more recently designed primers that capture more
Glomeromycota diversity with higher fidelity, such as the ones presented here or those by Lee et al. (2008) or Krüger et al. (2009), in environmental surveys will lead to more complete detection of AMF taxa. Given the current high rate of habitat loss and our impoverished knowledge of global Glomeromycota diversity, we may lose many AMF species without knowing they ever existed. Therefore, we would benefit from more environmental surveys of AMF, particularly in rare and endangered habitats before they are lost.
The research of this dissertation, in combination with the results of other published studies, provides strong support for an effect of host identity on AMF community composition in roots (Helgason et al. 2002, Vandenkoornhuyse et al.
2002). Given the strong support for the effect of host identity, it would be beneficial to focus future research on investigating the mechanisms of non-random association between partners. As briefly discussed in Chapter 1, there are various mechanisms that could contribute to non-random AMF community assembly in roots, including the involvement of recognition factors and the delivery of rewards to preferred partners.

144
The arbuscular mycorrhizal field would benefit from an ‘intradisciplinary’ approach, combining the research from studies of host selectivity, of differential effects of AMF on plants, and of differential rewarding by partners.
Of course, virtually all plants in natural settings live in communities with multiple species. Even as we determine the processes occurring in isolated plant-AMF interactions, evidence is compounding that interactions with neighboring plants also affect root AMF communities (Hausmann & Hawkes 2009, Mummey et al. 2005,
Stinson et al. 2006). Thus, we have the challenge of understanding how the effects of hosts and the effects of neighbors interact as determinants of final root AMF community composition.
In summary, the systems of interconnection and interaction between AMF and plants are quite complex. Individual plants associate with multiple AMF, some of which may, considered individually, provide a net benefit while others may be a net drain on the plant. The degree of benefits provided is likely to vary widely depending upon changing conditions related to factors such as nutrient levels, water and light availability, and the abundance of pathogens and herbivores. AMF associate with multiple plants, and a single AMF may provide a net benefit to one of its plant associates while yielding a net detriment to another. Additionally, nutrients may be transferred through AM fungal hyphae from one plant to another (Simard & Durall
2004). We have a long way to go to understand how the interplay of so many factors relates to the final community composition of AMF in plant roots and how to predict the impact of particular AMF combinations on individual plants, let alone plant

communities. As we pull together the knowledge gained from individual studies on
145 different aspects of the symbiosis, we get closer to that understanding.

146
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148
Table 5.1. The proportion of individual occurrences from different Glomeromycota orders found in A. millefolium and P. vulgaris plants from the field (Chapter 2) and the proportion of Glomeromycota orders found overall in field plants (Chapter 2) and in experimental plants (Chapter 4).
Order
Glomerales
Diversisporales
Archaeosporales
Field
Achillea
0.70
0.20
0.09
Field
Prunella
0.51
0.33
0.16
Field overall
0.64
0.22
0.14
Experimental overall
0.36
0.21
0.43

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