Spatial and temporal variation in mycorrhizal associations in a rare North American orchid
By
Pablo Antonio Tovar, B.S.
A Thesis In
PLANT AND SOIL SCIENCE
Submitted to the Graduate Faculty of Texas Tech University in Partial Fulfillment of the Requirements for the Degree of
MASTER OF SCIENCES
Dr. Jyotsna Sharma Chair of Committee
Dr. Richard Strauss
Dr. Stephen Maas
Dr. Mark Sheridan Dean of the Graduate School
December, 2015
© 2015 Pablo Antonio Tovar
Texas Tech University, Pablo Antonio Tovar, December 2015
ACKNOWLEDGEMENTS
First, I would like to express my sincere gratitude to my advisor Dr. Jyotsna Sharma for her guidance, support, motivation and contribution throughout the development of my master’s degree. In addition, I would like to acknowledge the other members of my advisory committee,
Dr. Richard Strauss and Dr. Stephen Maas for the great assistance and support, and patience for helping me out with the data analyses that were carried out for this research. Moreover, I am thankful to Dr. Madhav Pandey, a former Senior Research Associate in Dr. Sharma's lab, for providing aid with statistical analyses.
I wish to extend my appreciations to the U.S Fish and Wildlife Service for providing funding to Dr. Sharma for this investigation.
To my lab mates, I want to thank you all for your support revising my proposal, multiple times, and for helping me practice my thesis oral presentation. I hope we can work together again in the close future. To my parents, thanks for all the long distance support through this journey.
Being away from home is never easy, and I will keep following your constant advice always.
Finally, I am thankful to my girlfriend Lila for all the encouragement and affection over this process.
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TABLE OF CONTENTS
ACKNOWLEDGEMENTS ...... ii
LIST OF TABLES ...... vii
LIST OF FIGURES ...... xii
I. INTRODUCTION AND BACKGROUND ...... 1
Introduction ...... 1
Orchid diversity ...... 1
Plant habit ...... 4
Platanthera praeclara Sheviak and Bowles ...... 6
Morphology of Platanthera praeclara ...... 7
Habitat of Platanthera praeclara ...... 8
Reproduction in Platanthera praeclara ...... 10
Mycorrhizal associations ...... 12
Threats to Platanthera praeclara ...... 15
Why study this taxon? ...... 17
Objectives of the study...... 20
Significance of the study ...... 20
Literature Cited ...... 22
II. MYCORRHIZAL ASSOCIATIONS OF PLATANTHERA PRAECLARA
ACROSS ITS NATURAL DISTRIBUTION ...... 35
Abstract ...... 35
Introduction ...... 37
Materials and methods ...... 41
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Collection sites ...... 41
Sample collection ...... 43
Fungal identification from root sections ...... 44
PCR and sequencing ...... 45
Cloning ...... 47
OTU clustering ...... 48
Mean pairwise sequence distance (pi, π) ...... 48
Fisher’s exact test ...... 49
Two-way hierarchical ordination ...... 49
Multiple response permutation procedure ...... 50
Diversity curves ...... 50
Nonmetric multidimensional scaling ...... 51
Phylogenetic analyses ...... 51
Results ...... 53
Operational taxonomic unit (OTU) diversity ...... 53
Mean pairwise sequence distance (π) ...... 55
Fisher’s exact test ...... 55
Two-way hierarchical ordination ...... 56
Multiple response permutation procedure ...... 56
Diversity curves ...... 57
Nonmetric multidimensional scaling ...... 57
Phylogenetic analyses ...... 58
Discussion ...... 60
Literature Cited ...... 68
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III. MYCORRHIZAL ASSOCIATIONS OF PLATANTHERA PRAECLARA IN
RESPONSE TO PRESCRIBED FIRE AND HAYING ...... 114
Abstract ...... 114
Introduction ...... 116
Materials and methods ...... 118
Collection sites ...... 118
Sample collection ...... 119
Environmental data ...... 120
Fungal identification from root sections ...... 121
PCR and sequencing ...... 122
Cloning ...... 123
OTU clustering ...... 124
Mean pairwise sequence distance (pi, π) ...... 125
Fisher’s exact test ...... 125
Two-way hierarchical ordination ...... 126
Nonmetric multidimensional scaling ...... 126
Results ...... 127
Operational taxonomic Units diversity ...... 127
Mean pairwise sequence distance (pi, π) ...... 128
Fisher’s exact test ...... 129
Two-way hierarchical ordination ...... 130
Nonmetric multidimensional scaling ...... 130
Discussion ...... 131
Literature Cited ...... 135 v
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IV. CONCLUSIONS...... 163
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LIST OF TABLES
2.1. Number of plants sampled and number of samples that yielded useable sequences from roots of Platanthera praeclara in 2013 and 2014. Plant roots
were sampled for mycorrhizal diversity studies across 11 sites representing the natural distribution of the species………………………………………… 77
2.2. Soil chemical characteristics of 11 sites from where Platanthera praeclara roots were sampled in 2013 for mycorrhizal analyses. OM, organic matter; ENR, estimated nitrogen release; P1, phosphorus 1 (readily available to plants); P2, phosphorus 2 (readily available to plants plus part of the active reserve in the soil); K, potassium; Mg, magnesium; Ca, calcium; Na, sodium and CEC, cation exchange capability...... 78
2.3. Soil chemical characteristics of 11 sites from where Platanthera praeclara roots were sampled in 2014 for mycorrhizal analyses. OM, organic matter; ENR, estimated nitrogen release; P1, phosphorus 1; P2, phosphorus 2; K, potassium; Mg, magnesium; Ca, calcium; Na, sodium; H, hydrogen and CEC, cation exchange capability.…………………………………………….. 80
2.4. Primer pairs utilized, oligo sequences, and annealing temperatures to amplify the fungal ITS region of the nuclear ribosomal DNA via polymerase chain reaction from the DNA extracted from root fragments of Platanthera praeclara……………………………………………………………………... 82
2.5. Number of root sections (i.e. sequences) representing each of the 19 operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal
fungi at 11 sites in the year 2013. Values inside the parenthesis denote the number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was observed……………………………. 83
2.6. Number of root sections (i.e. sequences) representing each of the 19 operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal fungi at 11 sites in the year 2014. Values inside the parenthesis denote the
number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was observed……………………………. 84
2.7. Number of root sections (i.e. sequences) representing each of the 39 operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal fungi at 11 sites in the combined data set (years 2013 and 2014). Values
inside the parenthesis denote the number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was observed……………………...... 85
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2.8. Mean pairwise sequence distances and standard errors (SE) for all observed fungal ITS sequences from roots of Platanthera praeclara sampled across 11 sites in years 2013 and 2014, and for the combined data set (years 2013 and 2014), separated by fungal family. The values presented within the parenthesis are the numbers of individual ITS sequences (n) composing each group………………………………………………………………………….. 86
2.9. Mean pairwise fungal ITS sequence distances (pi-distance) estimated based on combined transitions and transversions substitution model, within fungal families identified in roots of Platanthera praeclara across 11 sites in North America. Mean pairwise sequence distances from publically available fungal ITS sequences obtained from orchids other than Platanthera praeclara and Nervilia nipponica were calculated by Pandey et al., 2013.………………….. 87
2.10. Statistical significance from the Fisher’s exact test comparing the fungal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) at each site for 2013 below the diagonal. Pairwise geographic distances (kilometers) between population pairs are presented above the diagonal. MB, Manitoba; PC, Pembina Control; PH, Pembina Hay; PF, Pembina Fall fire; PS, Pembina Spring fire; NDG, North Dakota Graze; NDVK, North Dakota Viking; BU, Bluemound Uphill; BD, Bluemound Downhill; NE, Nebraska and IA, Iowa…………….. 88
2.11. Statistical significance from the Fisher’s exact test comparing the fungal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) at each site for 2014 below the diagonal. Pairwise geographic distances (kilometers) between population pairs are presented above the diagonal. MB, Manitoba; PC, Pembina Control; PH, Pembina Hay; PF, Pembina Fall fire; PS, Pembina Spring fire; NDG, North Dakota Graze; NDVK, North Dakota Viking; BU, Bluemound Uphill; BD, Bluemound Downhill; NE, Nebraska and IA, Iowa.……………. 89
2.12. Statistical significance from the Fisher’s exact test comparing the fungal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) at each site using the combined data set (years 2013 and 2014) below the diagonal. Pairwise geographic distances
(kilometers) between population pairs are presented above the diagonal. MB, Manitoba; PC, Pembina Control; PH, Pembina Hay; PF, Pembina Fall fire; PS, Pembina Spring fire; NDG, North Dakota Graze; NDVK, North Dakota Viking; BU, Bluemound Uphill; BD, Bluemound Downhill; NE, Nebraska and IA, Iowa………………………………………………………………….. 90
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2.13. Multiple response permutation procedure (MRPP) showing the significance of the variation of the mycorrhizal fungal communities by using abundance data observed at 11 sites of Platanthera praeclara by grouping the sites according to the type of land management (prescribed fire, haying or grazing, or no management). The MRPP was performed with the combined data set (years 2013 and 2014) as well as years 2013 and 2014 separately.…. 91
3.1. Soil chemical characteristics in each prairie management treatment (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) from where Platanthera praeclara roots were sampled in 2013 for mycorrhizal analyses. OM, organic matter; ENR, estimated nitrogen release; P1, phosphorus 1 (readily available to plants); P2, phosphorus 2 (readily available to plants plus part of the active reserve in the soil); K, potassium;
Mg, magnesium; Ca, calcium; Na, sodium; CEC, cation exchange capability; and NO3, nitrate. Please see next page for the remaining table.……………... 139
3.2. Soil chemical characteristics in each prairie management treatment (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) from
where Platanthera praeclara roots were sampled in 2014 for mycorrhizal analyses. Mg, magnesium; Ca, calcium; Na, sodium; CEC, cation exchange capability; NO3, nitrate; S, Sulphur; Zn, zinc; Mn, manganese; Fe, iron; Cu, copper and B, boron and soil textural components (sand, silt and clay).…….. 141
3.3. Number of plants sampled and number of samples that yielded useable sequences from roots of Platanthera praeclara in 2013 and 2014. Plant roots were sampled for mycorrhizal diversity studies across four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) at Pembina Trail Preserve in Minnesota, USA……….. 143
3.4. Number of root sections (i.e. sequences) representing each of the four operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal fungi in each of the four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) in 2013. Values inside the parenthesis denote the number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was observed………………………………………………………………………. 144
3.5. Number of root sections (i.e. sequences) representing each of the 20 operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal fungi in each of the four prairie management treatment plots (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) in 2014. Values inside the parenthesis denote the number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was 145 observed………………………...... ix
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3.6. Number of root sections (i.e. sequences) representing each of the 20 operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal fungi in each of the four prairie management treatment plots (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) using the combined data set (years 2013 and 2014). Values inside the parenthesis denote the number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was observed…………… 146
3.7. Mean pairwise sequence distances and standard errors (SE) for all observed fungal ITS sequences from roots of Platanthera praeclara sampled from four experimental prairie management treatments in years 2013 and 2014, and for the combined data set (years 2013 and 2014), separated by fungal family Ceratobasidiaceae and Tulasnellaceae. The values presented within the parenthesis are the numbers of individual ITS sequences (n) composing each group…….……………………………………………………………… 147
3.8. Mean pairwise sequence distances and standard errors (SE) for all observed fungal ITS sequences from roots of Platanthera praeclara in each of four prairie treatment plots (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) for years 2013 and 2014 separated by land management treatment. The values presented within the parenthesis are the numbers of individual ITS sequences (n) composing each group………………………… 148
3.9. Statistical significance from the Fisher’s exact test comparing the fungal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) observed in each of the four prairie management treatment plots (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) for 2013……………………………………….. 149
3.10. Statistical significance from the Fisher’s exact test comparing the mycorrhizal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) observed in each of the four prairie
management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) for 2014………………………………………………. 150
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3.11. Statistical significance from the Fisher’s exact test comparing the fungal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) observed in each of the four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) for the combined data set (years 2013 and 2014)…….. 151
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LIST OF FIGURES
1.1. An inflorescence of Platanthera praeclara. Photo by Jyotsna Sharma.... 32
1.2. A partial map of the US and Canada showing the natural distribution of Platanthera praeclara. The states where the plant is currently known to be present (orange), the counties where the populations occur (gray) and the counties where roots were collected for the studies (yellow) are highlighted.……………………………………………………………... 33
1.3. A transverse section of a root of Platanthera praeclara showing colonization by fungal pelotons (examples indicated by arrows). The scale-bar represents 200 microns………………………………………. 34
2.1. A partial map of the US and Canada showing the natural distribution of Platanthera praeclara. The states where the plant is currently known to be present (orange), the counties where the populations occur (gray) and the counties where roots were collected for the studies (yellow) are highlighted.……………………………………………………………… 92
2.2. A map of the nuclear fungal ribosomal internal transcribed spacer (ITS) region. The two ITS regions are between the SSU18S and LSU28S ribosomal RNA genes and are separated by the 5.8S rRNA gene. Image taken from http://www.gatc-biotech.com...... 93
2.3. A Platanthera praeclara plant excavated for sampling while keeping the root system intact. Photo on the left was taken by Nancy Sather…… 94
2.4. A whole seedling of Platanthera praeclara along with severed root and tuber...... 95
2.5. Root fragments collected from plants of Platanthera praeclara………. 96
2.6. A transversal section of a root of Platanthera praeclara showing colonization by fungal pelotons in cortical cells (examples indicated by arrows). The scale-bar represents 400 microns………………………… 97
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2.7. Pie charts showing the distribution of the fungal families Ceratobasidiaceae and Tulasnellaceae observed in roots of Platanthera praeclara at 11 sites in the year 2013. Numeric values represent the individual ITS sequences belonging to each fungal family. of in the year 2013. Numbers in the pie charts represent the amount of ITS sequences from each fungal family in a site……………………………. 98
2.8. Pie charts showing the distribution of the fungal families Ceratobasidiaceae and Tulasnellaceae observed in roots of Platanthera
praeclara at 11 sites in the year 2013. Numeric values represent the individual ITS sequences belonging to each fungal family.…………… 99
2.9. One to four operational taxonomic units in individual plants of Platanthera praeclara (a) and number of individual plants that hosting one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) (b) across 11 sites in 2013.……………………………………………… 100
2.10. One to four operational taxonomic units in individual plants of Platanthera praeclara (a) and number of individual plants that hosted one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) (b) across 11 sites in 2014.………… …………………………………... 101
2.11. One to four operational taxonomic units in individual plants of Platanthera praeclara (a) and number of individual plants that hosted one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) across 11 sites (b) for the combined data set (years 2013 and 2014)…… 102
2.12. Two-way hierarchical cluster tree and matrix coding based on abundance of 19 fungal operational taxonomic units (OTUs) observed at 11 sites where roots of Platanthera praeclara were sampled in the year 2013……………………………………………………………….. 103
2.13. Two-way hierarchical cluster tree and matrix coding based on abundance of 19 fungal operational taxonomic units (OTUs) observed at 11 sites where roots of Platanthera praeclara were sampled in the year 2014……………………………………………………………….. 104
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2.14. Two-way hierarchical cluster tree and matrix coding based on abundance of 39 fungal operational taxonomic units (OTUs) observed at 11 sites where roots of Platanthera praeclara were sampled in the combined data set (years 2013 and 2014)…………………………….... 105
2.15. Sample based observed (Mao Tau) and rarefaction (Chao 1) cumulative fungal operational taxonomic unit (OTU) abundance curves for Platanthera praeclara. Fungal communities within the roots of plants were determined by sampling plants from 11 sites that host the orchid species.…………………………………………………………………... 106
2.16. Nonmetric multidimensional scaling (NMS) ordination of the mycorrhizal fungal communities within the roots of Platanthera praeclara sampled from 11 sites with correlations to soil variables [Na, Mn, Zn, Ca, OM (organic matter), CEC (cation exchange capability), soluble salts and pH] for the year 2013……………………………….... 107
2.17. Nonmetric multidimensional scaling (NMS) ordination of the fungal communities of Platanthera praeclara sampled from 11 sites with correlations to soil variables (S, B, H, clay and silt) for the year 2014... 108
2.18. Partial Maximum likelihood tree of the fungal family Ceratobasidiaceae constructed with operational taxonomic unit (OTU) sequences observed in Platanthera praeclara roots collected from 11 sites across its natural habitat. Sequences of known orchid mycorrhizal fungi were added to the alignments to place our sequences in a broader context. Scale bar represents estimated number of DNA substitutions per site. Only branch support values over fifty percent are displayed (maximum likelihood / posterior probability). Please see the next page for the remaining tree…. 109
2.19a. Maximum likelihood tree of the fungal family Tulasnellaceae constructed with operational taxonomic unit (OTU) sequences observed in Platanthera praeclara roots collected from eleven populations across its natural habitat. Sequences of known orchid mycorrhizal fungi were added to the alignments to place our sequences in a broader context. Scale bar represents estimated number of DNA substitutions per site. Only branch support values over fifty percent are displayed (maximum likelihood / posterior probability)……………………………………….. 111
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2.19b. Maximum likelihood tree of the fungal family Tulasnellaceae constructed with operational taxonomic unit (OTU) sequences observed in Platanthera praeclara roots collected from eleven populations across its natural habitat. Sequences of known orchid mycorrhizal fungi were added to the alignments to place our sequences in a broader context. Scale bar represents estimated number of DNA substitutions per site. Only branch support values over fifty percent are displayed (maximum likelihood / posterior probability)……………………………………….. 112
2.19c. Maximum likelihood tree of the fungal family Tulasnellaceae constructed with operational taxonomic unit (OTU) sequences observed in Platanthera praeclara roots collected from eleven populations across its natural habitat. Sequences of known orchid mycorrhizal fungi were added to the alignments to place our sequences in a broader context. Scale bar represents estimated number of DNA substitutions per site. Only branch support values over fifty percent are displayed (maximum likelihood / posterior probability)……………………………………….. 113
3.1. Satellite image of the arrangement of the four prairie management treatment plots (PC, no management; PS, spring burn; PF, fall burn, and PH, annual haying) where mycorrhizal fungi of Platanthera praeclara were sampled in years 2013 and 2014. Experimental plots were located at Pembina Trail Preserve, Polk County, Minnesota, USA. Image taken from Google Earth.……………………………………………………… 152
3.2. Soil chemical characteristics in each prairie management treatment (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) from where Platanthera praeclara roots were sampled in 2014 for mycorrhizal analyses. Mg, magnesium; Ca, calcium; Na, sodium; CEC, cation exchange capability; NO3, nitrate; S, Sulphur; Zn, zinc; Mn, manganese; Fe, iron; Cu, copper and B, boron and soil textural components (sand, silt and clay)………………………………………... 153
3.3. Pie charts showing the distribution of the fungal families Ceratobasidiaceae and Tulasnellaceae observed in roots of Platanthera praeclara sampled from four experimental prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) in the year 2014. Numeric values represent the number of individual ITS sequences belonging to each fungal family……………... 154
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3.4. One to three operational taxonomic units in individual plants of Platanthera praeclara (a), and number of individual plants hosting one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) (b), across four experimental prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) in 2013…. 155
3.5. One to four operational taxonomic units in individual plants of Platanthera praeclara (a), and number of individual plants hosting one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) (b), across four experimental prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) in 2014…. 156
3.6. One to four operational taxonomic units in individual plants of Platanthera praeclara (a), and number of individual plants hosting one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) (b), across four experimental prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment). Data represent samples collected from each experimental treatment over two years (2013 and 2014).………………………………………………….. 157
3.7. Two-way hierarchical cluster tree and matrix coding based on abundance of four fungal operational taxonomic units (OTUs) within the roots of Platanthera praeclara representing four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment). Data represent samples collected from each experimental treatment in 2013…………………………………… 158
3.8. Two-way hierarchical cluster tree and matrix coding based on abundance of 15 fungal operational taxonomic units (OTUs) within the roots of Platanthera praeclara representing four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment). Data represent samples collected from each experimental treatment in 2014……………………………………………………….. 159
3.9. Two-way hierarchical cluster tree and matrix coding based on abundance of 20 fungal operational taxonomic units (OTUs) within the roots of Platanthera praeclara representing four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment). Data represent samples collected from each experimental treatment over two years (2013 and 2014).…………………………….. 160 xvi
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3.10. Nonmetric multidimensional scaling (NMS) ordination of the fungal communities of Platanthera praeclara in 4 land management treatment plots (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) with correlations to soil variables (clay, silt, sand, S, P1, Na, B and pH) for the year 2013……………………………………………. 161
3.11. Nonmetric multidimensional scaling (NMS) ordination of the fungal communities of Platanthera praeclara representing four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying, and PC, no treatment) with correlations to soil physicochemical variables (clay, silt, sand, S, P1, OM and pH) and soil environmental variables (electrical conductivity in June, EC J; electrical conductivity in May, EC M, and volumetric water content in June, VWC J) for the year 2014..…………………………………………………………...... 162
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CHAPTER I
INTRODUCTION AND BACKGROUND
Introduction
Orchid diversity
The Orchidaceae is one of the largest and arguably the most diverse families of angiosperms. Estimates suggest a range between 20,000 and 35,000 species in 5 subfamilies and approximately 870 genera, occurring in all vegetated continents and some Antarctic islands (Dressler, 1993; Mabberley, 1997; Chase et al., 2003). The highest abundance and distribution occurs towards the tropics, following hotspots of species richness and endemism (Myers et al., 2000; Swarts & Dixon, 2009). Relatively little is known about the origin and biogeography of the Orchidaceae. Ramirez et al.
(2007) suggested that the subfamilies had diverged around 65 mya between the late
Cretaceous and early Paleocene when global temperatures were rising (Zachos et al.,
2008). By using molecular dating techniques on orchid fossils of the two major subfamilies Orchidoideae and Epidendroideae, Gustafsson et al. (2010) suggested that major diversification within these two subfamilies occurred during a cooler period by the end of the Eocene and into the Oligocene.
A number of hypotheses owing to pollination specialization, specialized mycorrhizal interactions, and niche partitioning attempt to explain the diversity within the
Orchidaceae.
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Pollinator specialization hypothesis is one of them (Cozzolino & Widmer, 2005).
The Orchidaceae is marked by a degree of pollinator specialization that probably has not been attained by any other family of angiosperms (Ibish et al., 1996). Certain pollination strategies are argued to involve remarkably specialized relationships between plants and pollinators. Some orchids produce highly specific suites of olfactory and visual stimuli that attract specific pollinator species. For example, in orchids that utilize pseudo- copulation strategies, a male bee or wasp is attracted to these stimuli, visually mistaking the orchid flower for a female conspecific and pollinates through repeated mistaken copulations (Dafni & Bernhardt, 1990). It is also reported that recently derived sub- families show a decrease in pollinator specialization (Tremblay, 1992). Additionally,
Gravendeel et al. (2004) did not record evidence for orchid speciation being driven by pollinator specialization. Instead, the study reports that Epidendroideae does not present a different level of pollinator specialization from other subfamilies. This is especially notable because Epidendroideae represents the most species rich subfamily in
Orchidaceae.
Orchids exemplify diversity in specialized mycorrhizal interactions (van der Pijil &
Dodson, 1966; Dressler, 1981, Rasmussen, 1995; Taylor et al., 2002; Otero & Flannagan,
2006; Barret et al., 2010). Examples for this are the common and widely distributed orchid species like Neottia ovata that associate with a broad spectrum of mycorrhizal fungi from families including Tulasnellaceae, Ceratobasidiaceae, Thelephoraceae, and
Sebacinaceae (Jacquemyn et al. 2015). Also, some species with restricted distributions, such as Piperia yadonii, host a large variation in mycorrhizal communities among
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2002) to restricted distribution like Hexalectris colemanii (Kennedy et al., 2011)
Considering that orchid seeds are rudimentary and lack endosperm, they lack stored energy resources for germination and establishment (Smith and Read, 2008). Thus, they rely upon mycorrhizal symbiosis to provide nutrients and carbohydrates to germinating seeds. Orchid mycorrhizae differ from other types of endomycorrhizae like arbuscular or ericoid mycorrhizae in that hyphae actually penetrate root cell membranes and form hyphal coils (Smith and Read, 2008). Other endomycorrhizae invaginate the cell membrane, but do not penetrate the cell membrane (Peterson et al., 2004). In addition, studies have shown a high level of orchid specificity towards mycorrhizal fungi
(Taylor and Bruns, 1997; Otero et al., 2002; Otero et al., 2004), where most of the fungal associates belong to three families in the basidiomycetes: Ceratobasidiaceae,
Tulasnellaceae and Sebacinaceae (Dearnaley et al., 2012), among others (Cortinariaceae,
Atheliaceae, Auriculariaceae, Cantharellaceae, Telephoraceae). Irregular distributions of orchid mycorrhizal fungi, combined with a diversity of mycorrhizal strategies ranging from broad to highly specific could explain the high diversity of orchid species (Otero and Flannagan, 2006; McCormick and Jacquemyn, 2014).
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Environmental niche partitioning or microhabitat specialization has been demonstrated in tropical epiphytic environments (Gentry and Dodson, 1987; Wolf and
Flamenco, 2003). In tropical forests, is not uncommon to find orchids restricted to different layers across the forest stratification. Biotic and abiotic factors change across the gradient, affecting species distribution (Diez & Pulliam, 2007). Still, this hypothesis has yet to be extensively researched in orchids.
Plant habit
Approximately 70% of the Orchidaceae are photosynthetic epiphytes (Zotz, 2013) that are primarily distributed in the tropical and subtropical regions of the planet.
Epiphytic orchids typically grow numerous roots that increase adhesion to the host
(Gravendeel et al., 2004). Tropical epiphytic species may experience periodic drought due to seasonal rains. Adaptations to an epiphyte habit include modifications in the root system. Roots of epiphytic orchids may be photosynthetic and aid in uptake and storage of water and nutrients (Gravendeel et al., 2004). Succulence, crassulacean acid metabolism (CAM), and sequential production of individual shoots operating as independent physiological units are among the adaptations for nutrient and water uptake in epiphytic orchid taxa (Gravendeel et al., 2004).
The remaining 30% of the orchid taxa occupy terrestrial habitats distributed across all continents. Like epiphytic orchids, terrestrial orchids have specific adaptations according to the environmental pressures they have to face within a habitat. Orchids may exhibit plant organs that are modified for mycorrhizal associations. A protocorm, for 4
Texas Tech University, Pablo Antonio Tovar, December 2015 example, is a tuber-shaped body with rhizoids that make up the entire seedling for a period of time before producing a shoot. Terrestrial orchids also may produce thickened roots, tubers, corms, and / or rhizomes that can serve as belowground storage structures
(Rasmussen, 1995). As stated by Rasmussen (1995), terrestrial orchids are dependent on mycorrhizal association for germination and plant development, especially in the protocorm and seedling stages.
When a seed from terrestrial orchids germinates, it first develops into a protocorm, a structure that bears primordial leaves but no roots. Eventually, the mycorrhizal stage develops when the apical meristem elongates and the first root initials develop
(Rasmussen, 1995). The protocorm meristem produces rhizomes that may vary in shape according to the species (Rasmussen, 1995). This rhizome is a modified subterranean stem that sends out roots and shoots from its nodes. Rhizomes start with mycotrophic functions, but eventually this function is transferred to the roots and the mycotrophic tissue in the rhizome decreases. Storage of water, starch, proteins and other nutrients is another function of the rhizome in orchids (Jang et al., 2006).
Terrestrial orchids typically have thick, fleshy, brittle unbranched roots; root systems are generally simple and small in relation to the plant size. Apart from mycotrophy, root functions include absorption of water and nutrients, storage, and anchorage (Rasmussen, 1995). Several terrestrial orchids produce root modifications that are called tubers for nutrient storage as their principal function. Tubers vary considerably in habit and shape, and usually they are colonized by fungi only in peripheral extensions and in superficial tissue (Rasmussen, 1995). The old tuber senesces during the growing 5
Texas Tech University, Pablo Antonio Tovar, December 2015 season as the new tuber begins to grow (Medina et al., 2009). During the dormant season
(winter, or dry season), the aboveground portions of the plant senesce except for the stem tuber and rhizome that grow a new shoot in the subsequent growing season, producing stems and leaves.
Platanthera praeclara Sheviak and Bowles
Platanthera praeclara (Orchidaceae) (Figure 1.1) is placed in the subfamily
Orchidoideae, tribe Orchideae and subtribe Orchidinae. Plants in this subtribe have a terrestrial habit. Orchidinae are usually non-saprophytic orchids with root-stem tuberoids.
Their roots do not have velamen (usually present in epiphytic orchids) and they have a slender stem (Dressler, 1993). The genus Platanthera includes approximately 200 species that are distributed across temperate and tropical North America, North Africa, Central
America and Eurasia (U.S. Fish and Wildlife Service, 2009). In North America, 24
Platanthera species occur from northern Mexico to the southern province of Manitoba in
Canada (USFWS, 2009). Platanthera is one of the most species-rich genera of temperate orchids in the northern hemisphere (Hapeman and Inoue, 1997), having most of its diversity in North America (Sheviak & Catling, 2002) and in East Asia (Inoue, 1983).
Species of the genus Platanthera occur in a wide variety of habitats and soils, usually acidic, ranging from swamps and meadows to fairly dry woodlands (Correll, 1978;
Williams and Williams 1983; Rasmussen, 1995).
Platanthera praeclara, previously included in the P. leucophaea taxonomic concept, was described by Sheviak and Bowles in 1986 when they encountered plants on 6
Texas Tech University, Pablo Antonio Tovar, December 2015 the Sheyenne National Grassland in North Dakota that exhibited distinct features. The species is currently listed as 'Threatened' by the U.S. Fish and Wildlife Service under the
Endangered Species Act of 1973 in the United States and as Endangered in Canada
(USFWS, 2009; Environment Canada, 2006). Natural populations of the species are known to occur in 6 mid-western states in the United States (Kansas, Missouri, Iowa,
Nebraska, Minnesota, and North Dakota) and in the province of Manitoba, Canada
(USFWS, 2009) (Figure 1.2).
Platanthera praeclara may have been historically distributed through a large portion of the western Central Lowlands and eastern Great Plains of the central United
States, and through the Interior Plains in south-central Canada (Brownell, 1984). The species had historically occurred in South Dakota and Oklahoma, but conversion of prairies to agriculture in the latter half of the nineteenth century caused extinction of the species in these states (Bowles and Duxbury, 1986; USFWS, 2009).
Morphology of Platanthera praeclara
Platanthera praeclara, as described by Sheviak and Bowles (1986), is an erect, stout, herbaceous perennial. It grows from 30 to 85 cm tall. Typically, the plant has multiple coarse, fleshy roots. Each plant usually has a single, glabrous, unbranched stem bearing two to five oblong-elliptic to lanceolate, keeled, glabrous leaves. These are usually 7 to 15 cm long and 1 to 4 cm wide. The inflorescence is 5 to 22 cm long and 4 to
7 cm in diameter and is a raceme of 5 to 25 flowers, with lanceolate, acuminate bracts 1.5
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Texas Tech University, Pablo Antonio Tovar, December 2015 to 4 cm long and 0.4 to 0.7 cm wide. Flowers are white or creamy white and have a perianth which is directed forward and forms a hood over the column. The dorsal sepal is ovate to suborbiculate, concave, 9 to 13 mm long and 5 to 8 mm wide. The lateral sepals are obliquely- obovate, asymmetrical, 7 to 14 mm long and 5 to 10 mm wide. The lateral petals are cuneate to flabelliform, rounded to truncate, 9 to 16 mm long, and 6 to 13 mm wide with lacerate distal margins. The labellum (modified petal) is deeply 3-lobed, usually 2 to 3.2 cm long, 2 to 4 cm wide, fringed, and bears a slender, arcuate, clavate, 4 to 5.5 cm long, sometimes as short as 2 cm long spur. The ellipsoid capsule is 2 to 2.5 cm long and 4 to 6 mm in diameter and releases seeds upon dehiscence.
Habitat of Platanthera praeclara
The western prairie fringed orchid is a perennial herbaceous species occurring in the North American tallgrass prairie. Tallgrass prairie plant communities are historically fire and grazing adapted and are usually dominated by Andropogon gerardii, A. scoparius, Sorghastrum nutans, Deschampsia caespitosa and Panicum virgatum, all members of the Poaceae (USFWS, 1996; Ghimire et al., 2011). Tallgrass prairie remains in the eastern fringe of the Red River Valley in Manitoba in Canada and in Minnesota in the USA. In eastern North Dakota, it survives in the Sheyenne National Grassland. From
South Dakota, through Minnesota and Iowa tallgrass prairie can be found in the Coteau des Prairies. Through east-central Kansas it can be found in the Flint Hills, and in the south, in can be found in northern Oklahoma. Tallgrass prairie extends further into the east through Illinois, Indiana and a portion of south western Michigan (Jones and
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Cushman, 2004). Platanthera praeclara plants generally occur in unplowed, calcareous prairies and sedge meadows (USFWS, 1996). The largest known population of P. praeclara occurs in southeastern Manitoba, Canada, with 23,530 flowering plants documented in the year 2003 (Environment Canada, 2006).
Platanthera praeclara in Manitoba occur in wet prairies or meadow vegetation developed on Alfisols (poorly drained grey wooded soils) with a thin sandy mantle overlying stony calcareous reworked till (Collicutt, 1992; USFWS, 1996). In North
Dakota, Platanthera praeclara is usually observed in the sedge meadows in the Glacial
Sheyenne Delta in tallgrass prairies (USFWS, 1996). Calcium rich wet prairie soils
(calciaquolls); wet prairie soils with minimum horizon development (haploquolls) and cool prairie soils with minimum horizon development (haploboralls) are the typical soils that support orchid growth on this area (United States Department of Agriculture, 1999;
Wolken, 1995). In Minnesota, populations in the Pembina Trail prairie complex are associated with poorly drained to moderately well–drained soils with gentle slopes
(USFWS, 1996). Populations in Iowa, southwestern Minnesota, northeastern Nebraska, northeastern Kansas and northwestern Missouri occur in wet-mesic to mesic tallgrass prairie (Freeman and Brooks, 1989). Udolls or Udic Ustolls (humid to intermittently dry mollisols, or prairie soils) constitute soils in these areas, on gentle to moderate slopes
(USDA, 1999). Plants in north-central Nebraska typically grow in tall grass prairies or sedge meadows in swales among the Nebraska Sand Hills in poorly developed sandy soils of warm climates (USDA, 1999). Platanthera praeclara growing in eastern
Nebraska occurs in wet mesic prairies and sedge meadows along the floodplain of the
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Platte River (Nagel and Kolstad, 1987). In eastern Kansas P. praeclara is observed in mesic to wet-mesic upland prairies on unglaciated, level to hilly, Pennsylvanian-age sediments covered with a thin, discontinuous mantle of loess residuum (USFWS, 1996).
Platanthera praeclara generally occurs in the wetter facies of such prairies or in associated sedge meadows (USFWS, 1996). Perennial taxa from the family Cyperaceae, with Carex spp. and Eleocharis spp. dominate sedge meadows. In addition, annual and perennial grasses and forbs (like Poa pratensis, Carex lanuginose and Juncus balticus) also occur in this community type (USFWS, 1996). Changes in the plant community might alter orchid presence (Wolken, 2001). Swales in sedge meadows supporting P. praeclara were compared with swales lacking orchids and it has been suggested that plant canopy cover of Juncus balticus, Stachys palustris and Carex spp. was higher in swales supporting the orchids (Wolken, 2001).
Reproduction in Platanthera praeclara
Platanthera praeclara has evolved an obligate outcrossing pollination system, i.e. pollen is delivered to a flower from a different individual (Sheviak and Bowles, 1986).
The flowers lack nectar guides, bear long nectariferous spurs and are fragrant at night; these are features that typify hawkmoth-pollinated plants (Phillips, 2003). In an effort to reach the nectar contained within the flower’s spur, the hawkmoth unintentionally brushes up against the pollinia, and they generally attach to its eyes (USFWS, 1996). The hawkmoth then transfer the pollinia to another P. praeclara flower (Phillips, 2003). Fox et al. (2013) documented six different moth species (including two native species) of 10
Texas Tech University, Pablo Antonio Tovar, December 2015 hawkmoth as pollinaria carrying pollinators of P. praeclara over nine years of observations at the Sheyenne National Grasslands in North Dakota, USA. These include:
Lintneria eremitus, Hyles lineata, Sphinx drupiferarum, Eumorpha achemon, Hyles euphorbiae and Hyles gallii, two of which (S. drupiferarum and H. gallii) had already been reported as pollinators of P. praeclara in Manitoba, Canada (Westwood and
Borkowsky, 2004; Borkowsky and Westwood, 2009).
Plant species with more diverse pollinators are typically at a lower extinction risk than those that rely on a few specialized pollinators (Bond, 1994; Marten-Rodriguez et al., 2010). Abundance of the pollinator species also plays a major role (Tremblay et al.,
2005). Unfortunately, only one (H. euphorbiae) of the known pollinators of P. praeclara is considered abundant. The other pollinator species are not abundant, and two of them,
E. achemon and S. drupiferarum, now appear to be less common (Fox et al., 2013). In addition, it is important to note that the hawkmoths do not depend on the orchid as they can feed from a number of non-orchid nectar sources to support their populations, whereas the orchid depends on the hawkmoths for pollination (USFWS, 1996).
As mentioned above, P. praeclara depends on outcrossing pollination for reproduction (Sheviak and Bowles, 1986). However, low pollination rates and seed production have been observed in most populations (Phillips, 2003). Borkowsky and
Westwood (2009) assessed pollination in the Manitoba Tall Grass Prairie Preserve in
Canada by recording number of removed and remaining pollinaria and number of seed capsules, among other observations, from flowers of P. praeclara. The study reported a
11
Texas Tech University, Pablo Antonio Tovar, December 2015 significant increase in seed capsule production in P. praeclara plants that had ultraviolet lights placed by them. However, the study describes that only 7% of flowers produced seed capsules, a smaller percentage of what has been reported in P. praeclara from
Minnesota and North Dakota in other similar studies (Pleasants, 1993; Pleasants and
Moe, 1993).
Mycorrhizal associations
Mutualistic mycorrhizal symbiosis involves reciprocal transfer of nutrients between a plant and its fungal partner (Kiers et al., 2011). Plants may reward fungal hyphae that are able to allocate greater phosphorus (P) resources. Consequently, fungi respond to this cooperation by supplying increased phosphorus (P) when receiving more carbon (C)
(Kiers et al., 2011).
Members of the Orchidaceae have evolved their own form of mycorrhiza. All orchid seeds are believed to be mycoheterotrophic in nature, or fully dependent on fungi for germination and early protocorm development before plants gain photosynthetic capability (Smith and Read, 2008). Coils of fungal hyphae, called pelotons (Figure 1.4), are digested by the host plant to acquire nitrogen (N), carbon (C), and other nutrients
(Smith, 1967). It is important to add that the fungi involved in the symbiosis do not appear to depend on the host plant to survive (Rasmussen, 1995; Bunch et al., 2013).
Orchids across the planet have been reported to be most commonly associated with members of the fungi previously referred to as ‘rhizoctonia’, which were recorded in older literature as the anamorphic states of heterobasidiomycetes (Currah and Zelmer,
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1992; Sen et al., 1999; Sharma et al., 2003a; Taylor et al., 2003; Zettler et al., 2003;
Zettler and Piskin, 2011). 'Rhizoctonias' were divided into two genera, Ceratorhiza and
Epulorhiza (Roberts, 1999), but this assemblage contains three different Agaromycetes taxa including Sebacinales, Ceratobasidiaceae and Tulasnellaceae. However, as explained by Dearnaley et al. (2012), this classification is not entirely correct since none of these taxa is adequately described by the definition (De Candolle, 1815) of the now obsolete asexual genus Rhizoctonia.
Orchid mycorrhizal fungi are widely diverse outside of their ecological niche inside orchid roots. The fungal family Ceratobasidiaceae includes economically important pathogenic and parasitic species such as Ceratobasidium noxium, which causes black rot of coffee, and Ceratobasidium cereale that causes sharp eyespot of cereals (Roberts,
1999). Many species are saprophytic, occurring in the soil and plant detritus (Smith and
Read 2008). Tulasnellaceae, along with the Ceratobasidiaceae have been detected as taxa in clades forming ectomycorrhizae (Bidartondo et al., 2003; Yagame et al., 2012). Fungi from the Order Sebacinales have been observed as mycorrhizae forming fungi in the
Orchidaceae and Ericaceae, as well as occurring as common endophytes in the roots of many plant species (Selosse et al., 2009; Weiβ et al., 2011). In orchids, molecular taxonomic identification of fungal associates have revealed that Ceratobasidiaceae,
Sebacinaceae and Tulasnellaceae generally associate with orchids worldwide (Dearnaley et al., 2012), with Tulasnellaceae observed as the most frequently fungal family in temperate and tropical regions (Rasmussen, 1995; Yuan et al., 2010).
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Despite the wide diversity and distribution of orchid mycorrhizal forming fungi, different orchid taxa often restrict the associations to specific fungi. The specificity in the association between fungi and orchids can be defined as the phylogenetic breadth in the diversity of the associates (Shefferson et al., 2010). Narrow fungal specificity is more noticeable usually in holomycoheterotrophic (i.e., fully dependend on fungi) orchid species that typically associate with fungi from one fungal family (Selosse et al., 2002;
Kennedy et al., 2011). Conversely, low specificity has been observed in orchids associating with two or three fungal families at the same time (Bonnardeaux et al., 2007;
Pandey et al., 2013).
Mycorrhizal associations in Platanthera praeclara have been studied previously to some extent. However, like majority of the other orchids, these critical associations have not been investigated range-wide or over time. In 1995, Zelmer and Currah first isolated and characterized eight isolates of Ceratorhiza pernacatena from the family
Ceratobasidiaceae by sampling roots from seven plants of P. praeclara from a single population in Manitoba, Canada. From the same orchid population, one isolate belonging to the Tulasnellaceae and fifteen Ceratobasidiaceae isolates were subsequently observed in 11 plants of P. praeclara (Zelmer et al., 1996). Later, Sharma et al. (2003a) isolated, characterized and identified fungi from 21 plants of P. praeclara in its native habitat in
Minnesota (Clay county, Norman county, Polk county and Mower county) and Missouri
(Harrison county). In this study, 78 isolates belonged to the fungal family
Ceratobasidiaceae (e.g. Genbank accession number DQ088771) and nine belonged to the fungal family Tulasnellaceae (e.g. Genbank accession number DQ068772). Results from
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Texas Tech University, Pablo Antonio Tovar, December 2015 these studies in Manitoba, Minnesota, and Missouri indicated that the orchid may prefer members of the Ceratobasidiaceae when forming mycorrhizal associations. However, a large part of the range was not represented in these studies, which limits the interpretation of range-wide patterns of mycorrhizal diversity and specificity in the species.
Additionally, temporal variation in mycorrhizal associations (phenological stages and / or yearly variation) was not captured in these studies (Zelmer and Currah, 1995; Zelmer et al. 2006; Sharma et al. 2003a). However, studies to test in vitro symbiotic germination of
P. praeclara seeds indicated that germination is more successful when cold-moist stratified seeds are cultured with fungal isolates derived from naturally occurring protocorms of P. praeclara (Sharma et al., 2003b) suggesting that there might be a successional patterns in fungal associations through the various phenological stages of plant growth.
Threats to Platanthera praeclara
The tallgrass prairie ecosystem is one of the most threatened ecosystems in North
America if not on Earth (Jones and Cushman, 2004). Approximately 1% of the original tallgrass prairie remains. Some of the reasons for this loss include the nutrient rich soils formed by prairie ecosystems that are also highly desirable for agricultural use.
Conversion to crop fields physically removes the native prairie ecosystem in addition to causing hydrological changes in any remaining adjacent prairies. Platanthera praeclara prefers late-successional prairies that do not undergo rapid changes, which renders them exceptionally sensitive to habitat destruction and degradation (USFWS, 1996; Goedeke et al., 2008). Anthropogenic factors can induce habitat alteration and degradation while
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Texas Tech University, Pablo Antonio Tovar, December 2015 inappropriate land management results in ecological changes that disadvantage P. praeclara populations. For example, non-application or misapplication of prescribed fire, grazing or mowing could have a detrimental impact on orchid survival, growth and reproduction (USFWS, 1996). Habitat deterioration and loss could lead to a decline in soil microbiota, local hydrology, and changes in plant and animal community compositions. As a result, exotic or invasive plants that P. praeclara could not out- compete find it easier to establish, displacing the orchid (Kirby et al., 2003). Agricultural production, especially row-cropping has damaged large tracts of mature, late-succession prairie habitat that is considered ideal for the orchids to persist (USFWS, 1996). If, instead, land is under grazing usage, cattle can also damage individual plants by grazing and / or trampling them. While natural low-density grazing may have been viable under historical circumstances when the plant was widely distributed, high-density commercial grazing can be a considerable threat especially for populations expressing low numbers of individuals (Goedeke et al., 2008). Moreover, remnant prairie and grazing lands are decreasing under pressure of urban development (Goedeke et al., 2008).
While the biology and ecology of P. praeclara are somewhat understood, concern increases about the persistence of the species. As populations decline, irregular flowering, dormancy periods, the need for appropriate symbiont and the need for particular pollinators might reduce the likelihood of population survival. Further, with smaller and isolated populations, genetic diversity is reduced and a decline in reproduction is likely to occur (Sharma et al., 2002).
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Other miscellaneous threats to the native Tallgrass Prairie communities and specifically to P. praeclara include damage or destruction by herbicides used against invasive species, roadway maintenance activities, introduction of new species, and / or plant collection by hobbyists (Goedeke et al., 2008).
Conservation status and measures
Platanthera praeclara is currently listed in the USA as 'Threatened' by the U.S.
Fish and Wildlife Service under the Endangered Species Act of 1973 in the United States and as Endangered in Canada (USFWS, 2009; Environment Canada, 2006). The focus of the recovery efforts has been on “maintaining the habitat of known populations on native prairie and providing the highest level of protection appropriate for all populations”
(USFWS, 2009). These goals will be accomplished by warranting that a minimum proportion of P. praeclara occur on lands that are being converted to non-grassland, and by ensuring that appropriate management is administrated where P. praeclara populations are established. However, many orchid populations are in private owned lands, making it challenging to manage plants properly (Goedeke et al., 2008).
Why study this taxon?
The biology and ecology of P. praeclara has been investigated in a little more detail than many other North American taxa native to the prairies due to its relatively wide distribution, its threatened status, and mostly because it is a large-flowered and a charismatic plant, which makes it a more desirable target than diminutive species that might be equally unique and rare (USFWS, 2009). Further, the species is associated with
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Texas Tech University, Pablo Antonio Tovar, December 2015 a highly unique, endangered, and iconic ecosystem that is vanishing quickly. Despite the body of knowledge available for changes in habitat (Collicut, 1993; Fauske and Rider,
1996; Alexander, 2001; Self and Anthonisen, 2005; USFWS, 2009), reproduction and pollinators (Cuthrell, 1994; Westwood and Borkowsky, 2004; Jordan et al., 2006;
Borkowsky and Westwood, 2009; Fox et al., 2013), germination (Sharma et al., 2003a;
Sharma et al., 2003b), and assessment of mycorrhizae from parts of its range (Zelmer and
Currah, 1995; Zelmer et al., 1996; Sharma et al., 2003a), there is an overall decline in populations, mostly because of habitat loss and / or degradation.
Some of the challenges for conservation of this species include a lack of understanding of its microhabitat requirements and how those conditions shift over space and time. Sieg and King (1995) highlighted changes in soil moisture in the landscape across different land management treatments, showing that soils subjected to long season grazing tend to have lower percentages of soil moisture. Hydrological changes can also influence the availability and community composition of mycorrhizae (Kivlin et al.,
2011). Little or no data are available for range-wide temporal and spatial variation in mycorrhizal associations and / or microhabitat conditions that support the growth of P. praeclara. Considering the critical involvement of fungi in germination, and a life-long association with adult plants, knowledge of distribution, diversity, and shifts in mycorrhizal associations is a necessary component toward understanding the ecological constraints on the persistence of the individuals and populations of any orchid taxon.
Changes in the soil microflora can have implications on the composition and function of the prairie communities and / or on the distribution, abundance, and
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Texas Tech University, Pablo Antonio Tovar, December 2015 persistence of individual species (Eom et al., 1999). Obligate fungal partners of plants are implicated in influencing vegetative community composition via positive or negative feedback mechanisms (Bertness & Callaway, 1994). These effects are often compounded when the plant-fungus relationships are highly exclusive, as can be the case for many orchid species and their mycorrhizal fungi. Shifts in fungal communities in tall grass prairies have been linked to changes in soil variables due to different types of land management (Murty et al., 2002). As plant community composition is immediately affected by land management, microbial communities are ultimately affected, too (Bossio et al., 1998; Johnson et al., 2003; Wu et al., 2015). Studies have described that shifts in the structure of fungal communities are associated with changes in soil properties like texture (Girvan et al., 2003), soil pH (Fierer and Jackson, 2006) and soil nitrogen and phosphorus availability (Frey et al., 2004; Lauber et al., 2008). Although these variables change depending on the land use, it is not clear if and how they affect biogeographical patterns in soil microbial communities (Lauber et al., 2008) at macro or micro spatial scales.
An assessment of both biotic (fungal associates) and abiotic (physical and chemical soil characteristics) components of the microhabitat of P. praeclara is thus warranted to assist with identifying its ecological niches and for optimizing conservation decisions for the taxon. This study was undertaken to include the objectives listed below:
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Texas Tech University, Pablo Antonio Tovar, December 2015
Objectives of the study
1) To assess the variation in the distribution and diversity (species richness and
abundance) of mycorrhizal fungal communities within the roots of Platanthera
praeclara in response to space, time, and edaphic characteristics.
It was expected that P. praeclara will exhibit high specificity towards its fungal associates across space, time, and edaphic characteristics despite its wide distribution, given the rarity of the orchid and the previously reported literature on mycorrhizal associations within the taxon.
2) To assess the effect of four experimental land management treatments (Spring burn
every four years, Fall burn every four years, Annual fall haying, and no treatment) on
the temporal variation in the diversity of mycorrhizal fungal communities within the
roots of Platanthera praeclara.
The diversity of fungi associated with P. praeclara was expected to be similar across prairie management treatments and over the two years because of the expectation of overall high mycorrhizal specificity in the species.
Significance of the study
The data from this study will quantify whether and how the distribution and diversity of mycorrhizal fungi associated with Platanthera praeclara shifts spatially and
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Texas Tech University, Pablo Antonio Tovar, December 2015 temporally. Majority of orchid mycorrhizal assessments are conducted and reported by assuming that relatively few plant roots sampled from few locations at any one time are reliable measures of diversity and specificity in orchid-fungal associations. Range-wide, multi-year investigations by using robust statistical approaches are rare, and only a few other orchid species have been studied in sufficient detail to develop generalized theories about temporal or spatial stasis in orchid-mycorrhizal associations. While spatial shifts are more easily expected, mycorrhizal fungi of orchids are generally assumed to be static over temporal scales. An assessment of the variability in orchid-fungal associations in response to edaphic characteristics are less common still.
The studies described here were designed to reveal both the spatial and temporal variation in orchid mycorrhizal symbiosis by using an emblematic and rare North
American orchid taxon native to an equally threatened ecosystem that is the North
American tallgrass prairie. Along with explaining the dynamics of the orchid mycorrhizal relationships in response to space, time, and soil edaphic variables, the studies described herein will assist conservation biologists with management decisions. Knowledge of the identity and diversity of fungi associated with the orchid under specific management treatments and how they change over time may be utilized to assist in habitat preservation and management programs.
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Borkowsky, C., & Westwood, A.R. (2009). Seed capsule production in the endangered western prairie fringed orchid (Platanthera praeclara) in relation to sphinx moth (Lepidoptera: Sphingidae) activity. J. Lepid. Soc, 63, 110-117.
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Brownell, V.R. (1984). Status report on the prairie white-fringed orchid Platanthera leucophaea: a rare species in Canada. Unpublished report to the Canadian government.
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Figure 1.1. An inflorescence of Platanthera praeclara. Photo by Jyotsna Sharma
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Figure 1.2. A partial map of the US and Canada showing the natural distribution of Platanthera praeclara. The states where the plant is currently known to be present (orange), the counties where the populations occur (gray) and the counties where roots were collected for the studies (yellow) are highlighted.
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Figure 1.3. A transverse section of a root of Platanthera praeclara showing colonization by fungal pelotons (examples indicated by arrows). The scale-bar represents 200 microns.
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CHAPTER II
MYCORRHIZAL ASSOCIATIONS OF PLATANTHERA PRAECLARA ACROSS ITS NATURAL DISTRIBUTION
Abstract
Platanthera praeclara is a rare endemic orchid native to parts of the North
American tallgrass prairie. As every other orchid species, P. praeclara survival relies on mycorrhizal associations at early developmental stages. Yet, the associations remain through the rest of the orchid’s life cycle. This study assessed the mycorrhizal diversity from several sites to describe mycorrhizal communities where P. praeclara occurs naturally across its natural distribution. Moreover, mycorrhizal assessments were conducted in two consecutive years to observe specificity and variation through time.
Types of land management that are usually applied to the sites where these orchids naturally occur were taken into account, in addition to edaphic characteristics, in an attempt to explain variation in the orchid mycorrhizal communities that were observed.
Mycorrhizal associations of Platanthera praeclara were studied in plants at 11 sites across its natural distribution in two consecutive years, 2013 and 2014. Across the two years, 39 Operational Taxonomic Units (OTUs) in two fungal families, Ceratobasidiaceae and Tulasnellaceae, were observed in roots of P. praeclara. Ceratobasidiaceae was the dominant fungal family at majority of the sampled sites, and it was represented by 77% of the analyzed nuclear ribosomal internal transcribed spacer sequences. Short pairwise sequence distances (π = 0.039 ± 0.03, N=238) between the sequences of
Ceratobasidiaceae and phylogenetic clustering of the OTUs suggested that P. praeclara 35
Texas Tech University, Pablo Antonio Tovar, December 2015 forms specific associations toward fungi from Ceratobasidiaceae. Fungi from the fungal family Tulasnellaceae (π = 0.135 ± 0.06, N=69) represented a broader phylogenetic breadth, with the orchid being less specific in its associations with the OTUs from this family. Despite the observed specificity, fungal communities in roots tended to vary
(P<0.05) among a large majority of the sites. Changes were observed in mycorrhizal fungal community composition from one year to the next. Clustering of sites with similar land management treatments was not detected. Nonmetric multidimensional scaling analysis did not segregate sites by land management; however, fungal communities appeared to be more similar in northern sites when compared to the fungal communities associated with the species in southern sites. Correlations between the observed fungal communities and edaphic characteristics were not evident.
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Introduction
Mycorrhizal fungi in the plant family Orchidaceae are characterized by intracellular coils of hyphae termed pelotons (Smith and Read, 2008). A large majority of the known orchid mycorrhizae are formed with fungi from the fungal phylum Basidiomycota
(Rasmussen, 1995; Taylor et al., 2002; Taylor and McCormick, 2008; McCormick and
Jacquemyn, 2014). Orchid mycobionts may be identified by isolating the fungi from the roots of the orchids and culturing them (Hadley and Williamson, 1972; Currah et al.,
1987; Currah et al., 1990; Zettler et al., 2001; Sharma et al., 2003b). However, identifying cultured isolates of fungi that associate specifically with orchids presents a number of challenges. Complex fruit bodies are absent or rarely seen, vegetative hyphal morphologies are mostly homogenous within genera (Taylor and McCormick, 2008), and many characters overlap between species, or vary because of the environment or changes during development stages within individuals (Andersen, 1990). Additionally, fungal isolation success may vary depending on the season, capacity of fungi to respond to culture conditions, or disturbance of plant roots previous to isolation (Ramsay et al.,
1986).
A more comprehensive method for characterizing the species richness and abundances of mycorrhizal fungi from plant roots is by amplifying specific DNA barcode regions by performing a polymerase chain reaction (Gardes and Bruns, 1993). Often, fungal DNA needs to be specifically amplified from mix of plant and fungal DNA, which requires targeting barcode regions that are specific to the organism of interest. 37
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In fungi, the nuclear ribosomal internal transcribed spacer (nrITS) region (between
700 and 800 nucleotides long) contains two variable non-coding regions (ITS1 and ITS2) that are nested within DNA repeats between the highly conserved small subunit (18S),
5.8S, and the large subunit (28S) rRNA genes (Figure 2.2). The internal transcribed spacer is the most widely sequenced target region for molecular identification of fungi because it can be amplified with ‘universal primers’ that are complementary to sequences within the rRNA genes (White et al., 1990). It has a multi-copy nature that makes this region relatively easy to amplify from small, diluted or degraded DNA samples (Gardes and Bruns, 1993). Further, the high variability of the nrITS region among closely related species makes it suitable to differentiate species or operational taxonomic units (OTUs)
(Gardes et al., 1991; Gardes and Bruns, 1996; Anderson and Cairney, 2004; Martin and
Rygiewicz, 2005; Taylor and McCormick, 2008; Seifert K.A., 2009; Schoch et al., 2012;
Oja et al., 2015).
Polymerase chain reaction amplification followed by Sanger sequencing of the nrITS has improved identification and characterization of different types of mycorrhizae
(Gardes et al., 1991; Gardes and Bruns, 1993; Redecker, 2000). In 2008, Taylor and
McCormick published primers that were developed specifically to amplify the nrITS region of fungi known to commonly associate with orchids. While these primers have helped to clarify distributions of orchid mycorrhizae in natural environments, their exclusive use was not implied. Capturing the complete profiles of communities of peloton forming fungi in orchid roots can in fact require a multi-primer approach or use of cloning techniques. With the advent of multiplex sequencing tools such as Next
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Generation Sequencing (NGS), identification of fungi in roots is technically possible to generate large amounts of sequences from individual samples, however separating the multitudes of endophytes from peloton-forming fungi can heavily skew the interpretations with respect to specificity if generic fungal nrITS barcoding is performed.
Platanthera praeclara is an herbaceous, perennial terrestrial orchid endemic to the
North American tallgrass prairie. It is listed as Threatened by the U.S. Fish and Wildlife
Service under the Endangered Species Act of 1973 in the United States and as
Endangered in Canada (USFWS, 2009; Environment Canada, 2006). Natural populations of P. praeclara are known to occur in six mid-western states in the United States
(Kansas, Missouri, Iowa, Nebraska, Minnesota, and North Dakota) and in the province of
Manitoba, Canada (USFWS, 1996) (Figure 2.1). According to the U.S Fish and Wildlife
Service (USFWS, 2009), there were 29,140 individuals of P. praeclara in 2008 in the
United States. Other plants that compose vegetation communities in the North American tallgrass prairies are species historically adapted to fire and grazing like Andropogon gerardii, A. scoparius, Sorghastrum nutans, Deschampsia caespitosa and Panicum virgatum, all members of the Poaceae (USFWS, 1996; Ghimire et al., 2011). Still, there is a vast presence of exotic or invasive plants that individuals of P. praeclara cannot out- compete [e.g., Poa pratensis, Bromus inermis, Trifolium campestre, Festuca arundinaceae and Sorghum halapense (Cully et al., 2003)] and displace the orchid (Kirby et al., 2003; Biederman et al., 2014). As an obligate outcrossed pollinated species, P. praeclara relies on hawkmoth species for reproduction (Fox et al., 2013). However, low
39
Texas Tech University, Pablo Antonio Tovar, December 2015 pollination rates and seed production have been observed in most populations (Phillips,
2003).
Mycorrhizal associations in Platanthera praeclara have been studied previously to some extent. Zelmer and Currah (1995) first isolated and characterized eight isolates of
Ceratorhiza pernacatena from the family Ceratobasidiaceae by sampling roots from seven plants of P. praeclara from a single population in Manitoba, Canada. From the same orchid population, one isolate belonging to the Tulasnellaceae and fifteen
Ceratobasidiaceae isolates were subsequently observed in 11 plants of P. praeclara
(Zelmer et al., 1996). Sharma et al. (2003a) assessed mycorrhizal fungi from P. praeclara from 21 plants in its native habitat in Minnesota (Clay county, Norman county,
Polk county and Mower county) and Missouri (Harrison county). In this study, 78 isolates belonged to the fungal family Ceratobasidiaceae (e.g. Genbank accession number
DQ088771) and nine belonged to the fungal family Tulasnellaceae (e.g. Genbank accession number DQ068772). However, like majority of the other orchids, these critical associations have not been investigated range-wide or over time.
While the ecology and biology of P. praeclara has been investigated in more detail than other North American orchids because of its threatened status and because of its association with a unique and endangered ecosystem that is disappearing rapidly, there is still a lack of understanding of its microhabitat requirements and how those conditions shift over space and time. Because different prairie management treatments are utilized in areas where populations of P. praeclara occur, soil variables known to affect dynamics of soil fungal or mycorrhizal communities likely change in response (Girvan et al., 2003;
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Texas Tech University, Pablo Antonio Tovar, December 2015
Fierer and Jackson, 2006; Frey et al., 2004; Lauber et al., 2008). Although the edaphic characteristics can vary depending on the ecosystem management, it is not clear if and how they affect biogeographical patterns in soil microbial or mycorrhizal communities
(Lauber et al., 2008) at macro or micro spatial scales.
The objective of the study reported herein was to assess the variation in the distribution and diversity (species richness and abundance) of mycorrhizal fungal communities within the roots of Platanthera praeclara in response to space, time, and edaphic characteristics. Given the rarity of the orchid and the previously reported literature on mycorrhizal associations within the taxon, high specificity towards its mycobionts was expected in the species across time, space, and edaphic characteristics despite its wide distribution.
Materials and methods Collection sites
Root tissues of Platanthera praeclara were sampled from 11 extant sites across four states in the US and one province in Manitoba, Canada. The locations in the United
States included Dinesen Prairie in Shelby County, Iowa; Valentine National Wildlife
Refuge in Cherry County, Nebraska; two different sites separated by a slope at
Bluemound State Park in Rock County, Minnesota, four long-term treatment sites with different management including prescribed fire in spring every 4 years (PS), prescribed fire in fall every 4 years (PF), annual haying in fall (PH), and no management (PC) at
Pembina Trail Preserve) in Polk County, Minnesota; and two sites within the Sheyenne 41
Texas Tech University, Pablo Antonio Tovar, December 2015
National Grasslands in Ransom County, North Dakota. In Canada, plants were sampled at the Manitoba Tallgrass Preserve in the Province of Manitoba. The largest distance between any two sites (Manitoba and Iowa) was 838 km whereas the shortest distance between any two sites (Blumound uphill and Bluemound downhill) was 3 km.
All locations and sites from where plants were sampled for this study represented some form of prairie management practice including no management, prescribed fire, grazing, or mowing, etc. While the historic information on frequency and intensity of these practices was not always precise or complete, information on recent applications during at least the past 5 years was obtained and recorded. The site in Manitoba has typically received prescribed burns every 5 to 7 years, unless there are severe droughts preventing such events or wildfires occur undesirably. For example, an unprescribed fire occurred at the study site in MB in 2012. In ND, high intensity grazing is applied to the
AA unit from May to November, however variable sections are fenced off each year to protect the individuals of P. praeclara. The Viking unit is not subjected to any kind of land management, and for that reason it was categorized as a control/no management treatment although a wildfire occurred in 2012 at this site. The two sites separated by a slope in Rock County, MN are subjected to prescribed burn periodically. In NE, prescribed grazing with light pressure is administrated during the growing season once every 5 to 6 days with rest periods of 2 to 3 months. Annual late summer haying is also administrated at this site while individually marking and protecting the reproductive plants of P. praeclara so that the capsules can mature and release seeds naturally.
Prescribed burns are administrated once every three years at Dinesen Prairie in IA.
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Texas Tech University, Pablo Antonio Tovar, December 2015
Sample collection
In 2013 and 2014, roots from individual plants were sampled between mid-June and early July. By this time in the growing season, differentiating between seedlings, juveniles or vegetative (individuals that will not produce an inflorescence in the growing season), and reproductive individuals was possible. Roots from two seedlings, two vegetative and two reproductive plants were sampled at each of the 11 sites. Roots from one or two extra plants were sampled with permission. Seventy-three and 66 plants were sampled across the 11 sites during 2013 and 2014, respectively (Table 2.1), which is a remarkably significant sampling effort considering the status of the species. Whole plants were carefully excavated to examine the complete root system (Figure 2.3). To keep plants alive after sampling, just a few roots were extracted from the plant, and the sampled plant was replanted. Sampled plants were watered thoroughly immediately after replanting to maximize reestablishment. Because seedlings have very small root systems, the whole seedling was collected and all roots were processed for fungal identification
(Figure 2.4). Roots were stored at 4°C until they were shipped overnight to the laboratory at Texas Tech University for further processing.
In each of the two years, one aggregate soil sample was collected at each site where orchids were sampled by adding together at least three subsamples. Approximately 500g of soil were extracted from sites where the plants were present. The soil samples were stored at room temperature until processed further. Physicochemical analyses were carried out by A&L Plains Labs, Inc., Lubbock, Texas. Organic matter (OM), pH, macro 43
Texas Tech University, Pablo Antonio Tovar, December 2015
nutrients (K, Mg, Ca, Na), cation exchange capacity (CEC), NO3-N, other nutrients (S,
Zn, Mn, Fe, Cu, B) and textural components (% sand, silt and clay) were measured in each sample.
Fungal identification from root sections
Roots were rinsed under tap water to wash them free of soil and debris before photo-documentation (Figure 2.5). Roots from each individual plant were subsequently placed in individual 50 mL plastic tubes. Inside a laminar flow hood and under sterile conditions, the surface sterilization was carried out by following these steps: (1) roots were first rinsed in a 70% EtOH (ethanol) solution by shaking thoroughly for 50 seconds;
(2) roots were then rinsed with a 3% NaOCl (sodium hypochlorite) solution by shaking thoroughly for 30 seconds; (3) a 70% ethanol rinse for 50 seconds was carried out; (4) roots were finally rinsed with ultrapure, sterile water several times to remove residues of
EtOH and NaOCl. Surface sterilization treatment was modified as needed based on the thickness and the number of the roots (Pandey et al., 2013).
The epidermis of each root was gently shaved off using a sterile scalpel to remove additional microbes that might have survived the surface sterilization treatment. Peloton inspection was conducted by cutting transversal thin slices of root tissue and examining under a compound microscope (Figure 2.6). If pelotons were observed, 2.5 to 3 cm long segments adjacent to the inspected slice were collected, sliced thinly and stored
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Texas Tech University, Pablo Antonio Tovar, December 2015 individually in 2 mL plastic tubes at -80°C until DNA was extracted (Pandey et al.,
2013).
DNA extraction was conducted using Qiagen DNeasy Plant Mini Kit (Qiagen,
Valencia, CA, USA) by following the manufacturer's protocol with a slight modification.
All samples were incubated in a 3.3% solution of polyvinylpyrrolidone in AP1 lysis buffer at 65°C for 2 hours while mixing once every 30 minutes before the RNase digestion step. Subsequently, 4 μL of RNase (supplied with the kit) was added to each sample and samples were incubated at 65°C for 15 minutes, mixing once every 4 minutes. Total DNA was eluted in 50 μL of elution buffer and stored at -80°C until further processing.
PCR and sequencing
Polymerase chain reaction was performed using total DNA extracted from individual root segments. Each reaction was prepared to a final volume of 25 μL by using
Promega GoTaq Flexi DNA Polymerase reagent kit (Promega, Madison, Wisconsin,
USA). Concentrations for each reagent were: 5x Green GoTaq Flexi Buffer, 10 mM/μL dNTPs (100 mM of each dNTP), 25 mM of MgCl2, 10 μM/μL of each primer, 10 μg/μL of BSA and 5 u/μL GoTaq DNA polymerase (Pandey et al., 2013). PCR consisted of 35 cycles in an epGradients Master Cycler (Eppendorf, Hamburg, Germany) and included a
2 minute initial denaturation at 94°C before thermocycling with a 45 seconds denaturation at 94°C followed by a 45-second annealing at different temperatures
45
Texas Tech University, Pablo Antonio Tovar, December 2015 depending on the primer pair (ITS1-OF and ITS4-OF; ITS4-TUL and ITS1) used (Table
2.4 in Chapter II) and 72°C elongation for 1 minute. Finally, the last cycle was followed by an extension at 72°C for 5 minutes.
Two primer pairs ITS1-OF/ITS4-OF and ITS1/ITS4-TUL (Taylor & McCormick,
2008) were used. A total of 498 root segments were assayed for molecular analysis. Each
DNA sample was first subjected to PCR using the ITS-OF primer pair at 58°C annealing temperature. Samples that did not amplify were then run with the same primer pair at
52°C annealing temperature. If amplification was still not obtained, ITS1/ITS4-TUL was used with 54°C annealing temperature on the samples that did not amplify previously. A
2% agarose gel electrophoresis was run to verify amplification. Samples showing a single band of the expected size range (600 – 800 bp) were cleaned using DNA Clean and
Concentrator 5 kit (Zymo Research, Irvine, CA, USA). DNA quantity and quality were measured using a NanoDrop 2000c spectrophotometer (Thermo Scientific, Wilmington,
DE, USA). Samples showing multiple bands were processed using GenElute Gel
Extraction Kit (Sigma-Aldrich, Poole, United Kingdom) by isolating the band closer to the expected size range. Sanger sequencing of all cleaned PCR products was carried out by the DNA Analysis Facility on Science Hill at Yale University. Samples were sequenced in one direction using the reverse primer (ITS4-OF or ITS4-TUL) (Pandey et al., 2013).
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Texas Tech University, Pablo Antonio Tovar, December 2015
Cloning
Polymerase chain reaction products from samples with sequencing difficulties were purified either using PureLink PCR Purification kit (Invitrogen Life Technologies,
Carlsbad, California) or GenElute Gel Extraction Kit (Sigma-Aldrich, Poole, United
Kingdom). Once a clean PCR product was obtained, it was used to carry out a cloning reaction and cloning transformation using the TOPO TA Cloning kit for Sequencing along with One Shot TOP10 and DH5α-T1 competent cells (Invitrogen Life technologies,
Carlsbad, California), following the manufacturer protocol.
To analyze the transformants, 3-5 colonies were selected and cultured overnight in
1 mL of LB (Luria-Bertani) (ThermoFisher Scientific, Waltham, MA, USA) medium containing 50 μg/mL of ampicillin. Plasmid DNA was isolated using the PureLink Quick
Plasmid Miniprep Kit, (Invitrogen Life Technologies, Carlsbad, CA) following the protocol included in the kit.
Polymerase chain reaction was performed on the extracted plasmid DNA following the amplification protocol described earlier using ITS1-OF/ITS4-OF primer pair.
Polymerase chain reaction products were purified using GenElute Gel Extraction Kit
(Sigma-Aldrich, Poole, United Kingdom) and DNA concentrations were measured using a NanoDrop 2000c spectrophotometer (Thermo Scientific, Wilmington, DE, USA).
Sanger sequencing of all cleaned PCR products was carried out as described above.
Data analyses 47
Texas Tech University, Pablo Antonio Tovar, December 2015
DNA Analysis Facility on Science Hill at Yale University performed a basic trimming on the sequences. However, all sequences were double-checked and a more thorough trim was implemented using Geneious 4.8.5. Trimmed sequences were then identified through BLAST searches using the Megablast option for finding highly similar
DNA sequences.
OTU clustering
Sequences were grouped into operational taxonomic units (OTUs) at 97% sequence similarity criterion using the OTU pipeline from the University of Alaska at Fairbanks
Life Science Informatics Portal (http://www.borealfungi.uaf.edu; Taylor & Houston
2011). The longest and highest quality representative sequence from each OTU was used for further analyses. Operational taxonomic Units were obtained separately for sequences obtained from root fragments collected in year 2013 and 2014. Additionally, for general diversity estimation, sequences were also clustered into OTUs by combining the data sets for both years.
Mean pairwise sequence distance (pi, π)
To estimate sequence divergence within the identified fungal families, separate multiple alignments of sequences were constructed for years 2013 and 2014 data sets, and for the combined data set, using MAFFT version 7 with L-INS-I model (Katoh and
Standley, 2013). Mean pairwise sequence distance (pi, π; Nei and Kumar, 2000) among
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Texas Tech University, Pablo Antonio Tovar, December 2015 all individual sequences within a fungal family was then estimated using MEGA 6
(Tamura et al., 2013). Mean pairwise sequence distances obtained using the combined 2- year data set from P. praeclara mycorrhizal fungi were compared with the previously reported pi distances among mycorrhizal fungi from other orchids (Pandey et al., 2013).
Fisher’s exact test
The statistical significance of the variation in fungal diversity between sites was tested using the Fisher’s exact test based on abundances of individual OTUs across the sampled sites per year of collection. The test was performed in R (R Development Core
Team 2010) using all the OTUs detected per sampled population for 2013 and 2014 separately and also for the combined data set.
Two-way hierarchical ordination
Two-way hierarchical ordination analysis (Bray and Curtis., 1957) using OTU abundances was performed to test the relationships between the mycorrhizal fungal communities of the sites based on dissimilarities. PC-ORD (McCune et al., 2002) was used to perform the two-way hierarchical ordination using the Sorensen (Bray-Curtis) distance measure, and setting the group linkage criterion as ‘average’. This test was conducted for each sampling year separately and also for the combined two-year data set.
The resulting dendrogram clustered sites according to similarity in mycorrhizal fungal communities held by the sampled plants across the sampled sites, and served as an aid to
49
Texas Tech University, Pablo Antonio Tovar, December 2015 observe whether sites sharing land management types (prescribed burn, haying or grazing, no management) clustered together.
Multiple response permutation procedure
We divided all sampled sites into groups by land management and by latitude to test whether fungal community composition abundance varies among the groups. Sites including MB, PF, PS, BU, BD and IA represented prescribed fire, NDG, NE, and PH represented hayed or grazed prairies, and hayed or grazed prairies, and no active management / control was represented by NDVK and PC. When grouped by latitude,
MB, NDG, NDVK, PC, PH, PF and PS were grouped as northern sites while BD, BU, IA and NE were grouped as southern sites. A multiple response permutation procedure
(MRPP) using PC-ORD (McCune et al., 2002) was carried out by using these groups for data sets from year 2013 and 2014 separately as well as for both data sets combined.
Diversity curves
Cumulative OTU diversity curves across populations were estimated using the sample-based Mao Tao and Chao 1 methods to examine whether the abundance of OTUs increases with increasing sample size using EstimateS 9.0 (Colwell, 2005). This analysis was conducted by using out OTU abundance data obtained from the dataset that represented both sampling years together.
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Texas Tech University, Pablo Antonio Tovar, December 2015
Nonmetric multidimensional scaling
Nonmetric multidimensional scaling (NMDS) in PC-ORD (McCune et al., 2002) was used to ordinate OTU abundance in roots of P. praeclara and to visualize the correlation between edaphic characteristics [OM, organic matter; ENR, estimated nitrogen release; P1, phosphorus 1 (readily available to plants); P2, phosphorus (readily available to plants plus part of the active reserve in the soil) 2; K, potassium; Mg, magnesium; Ca, calcium; Na, sodium; CEC, cation exchange capability; NO3, nitrate; S,
Sulphur; Zn, zinc; Mn, manganese; Fe, iron; Cu, copper and B, boron) and OTUs observed at each sampled site separately for each sampling year. The abundance-based version of the Sorensen index (Bray-Curtis) was used to calculate the distances.
Dimensionality of the ordination was determined to choose the lowest dimensionality that captured most of the variation. Data sets from 2013 and 2014 were both best described by
3-dimensional solutions with instabilities below 0.00001.
Phylogenetic analyses
To examine the phylogenetic relationships among the fungi associated with
Platanthera praeclara and other known orchid fungi, known orchid mycorrhizal sequences were added to the data used to generate the trees to place the fungi identified in this study in the context of previously known orchid mycorrhizae. Separate multiple alignments were constructed using MAFFT version 7 with L-INS-I model (Katoh and
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Texas Tech University, Pablo Antonio Tovar, December 2015
Standley, 2013) for each of the fungal families, that is, Ceratobasidiaceae and
Tulasnellaceae (Taylor and McCormick, 2008; Pandey et al., 2013) by using the sequences generated in this study and the respective reference sequences. Reference sequences were selected from previously published studies on orchid mycorrhizae. The resulting alignments were improved manually in AliView Version 1.17.1 (Larsson,
2014).
Phylogenetic relationships within each fungal family were estimated by determining the best-fit substitution model in ModelTest (Posada and Crandall, 1998) to construct maximum-likelihood trees using MEGA 6 (Tamura et al., 2013). Support values were estimated via 10000 bootstrap replicates in the same program. Because of the accelerated diversification of Tulasnella nuclear ribosomal ITS regions, a family level alignment using only the conserved 5.8S region extracted from the full length ITS sequences was first constructed (Shefferson et al., 2007; Taylor & McCormick, 2008) to build a maximum-likelihood tree in MEGA 6. The resulting tree was used to identify subclades within Tulasnellaceae containing the Platanthera praeclara observed OTUs, and full-length ITS alignments from these subclades were then constructed to generate the trees presented herein. Sistotrema sp. (Genbank accession number EU218893) was used for rooting Ceratobasidiaceae tree. Tulasnellaceae tree was midpoint rooted and subclade trees were rooted based on relationships shown in the broader 5.8S tree.
Additionally, posterior probability trees were built in MrBayes version 3.2.5 (Ronquist and Huelsenbeck, 2003) using the substitution models T92+G for the Ceratobasidiaceae tree, and K2+G for the Tulasnellaceae trees. The priors of the substitution models were
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Texas Tech University, Pablo Antonio Tovar, December 2015 set to default values. More than one million generations were carried out for analysis and continued until the standard deviation value of <0.01 was obtained. Cold chains were sampled after 100 generations and cross diagnosis of two runs was carried out after 1000 generations with the 25% burn in fraction and 0.2 temperature. All phylogenetic trees were visualized and edited in FigTree software (http://tree.bio.ed.ac.uk/).
Results
Operational taxonomic unit (OTU) diversity
Of the 498 root fragments from 139 sampled plants across 2013 and 2014, 306 root fragments from 109 plants produced workable DNA sequences after processing sequentially with several primer pairs and subject to cloning as needed (Table 2.1).
Ceratobasidiaceae and Tulasnellaceae were the two fungal families associated with
Platanthera praeclara across all 11 sites from where plants were sampled (Figures 2.7 and 2.8).
2013
Nineteen OTUs representing both mycorrhizal fungal families were identified from samples collected in the year 2013 (Table 2.5). Tulasnellaceae was represented by 10
OTUs, and Ceratobasidiaceae by 9 OTUs. Across all plants and sites, infection of a single root system by 2 to 4 fungal OTUs was observed in 46% of all sampled plants (Figure
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Texas Tech University, Pablo Antonio Tovar, December 2015
2.9a). The remaining 54% of plants contained one OTU each. A majority of the plants
(48 of 54 plants) possessed fungi belonging to a single family, while only six plants had fungi belonging to both families (Figure 2.9b).
2014
When additional plants were sampled in 2014, Ceratobasidiaceae and
Tulasnellaceae were represented by 14 and 5 OTUs, respectively (Table 2.6). Across all plants and sites, infection of a single root system by 2 to 4 fungal OTUs was observed in
27% of all sampled plants (Figure 2.10a). The remaining 73% of plants had only one
OTU each. A majority of the plants (51 of 55 plants) possessed fungi belonging to a single family, while only four plants had fungi belonging to the two families (Figure
2.10b).
2013 and 2014 combined
Thirty-nine OTUs representing both fungal families were identified when data from
2013 and 2014 were pooled. The Fungal family Ceratobasidiaceae was represented by 27
OTUs and Tulasnellaceae was represented by 12 OTUs (Table 2.7). Across all sampled plants and sites, infection of a single root system by 2 to 4 fungal OTUs was observed in
50% of all sampled plants (Figure 2.11a). The remaining 50% of sampled individuals yielded only one OTU. A majority of the plants (100 of 109 plants) possessed fungi 54
Texas Tech University, Pablo Antonio Tovar, December 2015 belonging to a single family, while roots of nine plants yielded fungi belonging to both families (Figure 2.11b).
Mean pairwise sequence distance (π)
Mean pairwise distance of sequences from Tulasnellaceae was consistently larger than for Ceratobasidiaceae regardless of the year of sampling, and in the combined data set (Table 2.8). By combining the data sets for both years (2013 and 2014), the mean distance observed among Ceratobasidiaceae was 0.039 ± 0.003 and the mean distance for
Tulasnellaceae was 0.135 ± 0.006. Additionally, mycorrhizal fungal associates of P. praeclara from the family Ceratobasidiaceae had the lowest mean pairwise sequence distance among orchid species with similar wide distribution except for Serapias vomeraceae. Mean pairwise distances among the Tulasnellaceae documented in the roots of P. praeclara were larger than most of the orchids with similar distribution patterns
(Table 2.9).
Fisher’s exact test
The family Ceratobasidiaceae was represented in roots of P. praeclara from all populations sampled in the year 2013 and 2014. The fungal family Tulasnellaceae was absent in roots sampled from four of the sampled sites (PC, PH, NDVK and NE) in 2013
(Figure 2.7). Significant differences were observed in 84% of the sites when the
55
Texas Tech University, Pablo Antonio Tovar, December 2015 mycorrhizal fungal community abundances were compared to each other by using the
Fisher’s exact test (Table 2.10). In the year 2014, Tulasnellaceae was not documented in the roots sampled from three sites (PH, NDVK and BD) (Figure 2.8). The Fisher’s exact test indicated that for 2014, 66% of the sites had significant differences in the composition of their mycorrhizal fungal abundance (Table 2.11). In the combined data set representing both years, the Fisher’s exact test including all OTUs per population revealed that P. praeclara roots host similar fungal communities in 13% of the sampled sites (Table 2.12).
Two-way hierarchical ordination
The site-wise two-way hierarchical ordination based on the OTU abundances indicated that sites did not cluster according to the type of land management (prescribed burn, haying/grazing, and no management) (Figure 2.12 and 2.13). When the two-way hierarchical ordination analysis was conducted for the combined data set (years 2013 and
2014), no clear clustering of sites based on management or latitude was detected (Figure
2.14).
Multiple response permutation procedure
The multiple response permutation procedure did not explain the variation in the mycorrhizal fungal communities to be related to type of land management (P > 0.05 in
2013, 2014 and combined data sets; Table 2.13). The variation was neither explained by
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Texas Tech University, Pablo Antonio Tovar, December 2015 geographical positioning of the sites in 2013 or 2014. However, variation in the mycorrhizal fungal communities was observed in the combined data set when separating most northern and most southern sites (Table 2.13).
Diversity curves
Sample-based cumulative OTU diversity curves built from the combined data set
(2013 and 2014) showed that estimates of Chao 1 (55 OTUs) are higher than the estimates of Mao Tau (39 OTUs), indicating that OTU diversity was not saturated with the current sampling and that more OTUs would be detected with additional sampling
(Figure 2.15).
Nonmetric multidimensional scaling
Nonmetric multidimensional scaling (NMS) ordination analysis of OTU abundances in orchid roots from samples collected in 2013 showed that distinction in orchid-associated fungi was correlated to Mn and Zn in NE; ENR and Ca in PC; OM,
CEC, S, and Salts in PH; and Na and pH in BD (Figure 2.16). For the 2014 data set, distinction in orchid-associated fungi was correlated to Silt, Clay and H in BU; S in
NDVK; pH and P2 in PF; and B in BD (Figure 2.17).
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Texas Tech University, Pablo Antonio Tovar, December 2015
Phylogenetic analyses
Maximum likelihood trees and posterior probability Bayesian trees showed similar relationships among the OTUs detected in this study and reference sequences. The maximum likelihood tree built with the OTUs from the fungal family Ceratobasidiaceae observed in P. praeclara along with reference nrITS sequences from published research showed four major clades (Figure 2.18). Ninety percent of the OTUs clustered together in one clade, along with one representative Ceratobasidium sp. sequence obtained from previously sampled roots of P. praeclara (GenBank accession DQ068771, J. Sharma unpublished data). The sister clade, without any OTUs from this study, was represented by mycorrhizal fungi belonging to the genus Ceratobasidium reported from roots of epiphytic neotropical orchids Ionopsis utricularoides (Otero et al., 2007; Borges et al.,
2013) and Tolumnia variegate (Otero et al., 2002). The same clade was also represented by a Ceratobasidium sp. obtained from the roots of tropical orchids in Singapore (Ma et al., 2003). Two of the OTUs from this study were separated from these two clades. One
OTU (OTU 24) was located in the third clade and was closely related to Ceratobasidium sp. observed in decayed roots of conifer seedlings in Lithuania (Menkis et al., 2006) and in Picea mariana forest soils in Alaska in the United States (Taylor et al., 2014). In the fourth clade, OTU 23 was closely related to Thanatephorus sp. observed in roots of
Vanilla aphylla in Cuba (Porras-Alfaro and Bayman, 2007).
The first sub-tree of the Tulasnellaceae (Figure 2.19a) contained five of the 12
OTUs observed for this mycorrhizal fungal family in P. praeclara. Operational
Taxonomic Units 1, 10 and 11 were not closely related to any previously accessioned 58
Texas Tech University, Pablo Antonio Tovar, December 2015 sequences. However, the sister clade of the clade described before contained OTU 3, which is closely related to a reported Tulasnella sp. from P. praeclara roots (GenBank accession DQ068773, J. Sharma unpublished data) and to a Tulasnella calospora obtained from roots of the orchid Paphiopedilum charlesworthii in Thailand
(Nontachaiyapoom et al., 2010). A third clade with OTU 12 was related to Tulasnella calospora obtained from roots of Acianthus exsertus, an orchid from the Andean cloud forest (Suarez et al., 2006) and is also related to Tulasnella sp. observed in the New
Zealand orchid Nematocera iridescens (Watkins, 2012). The second sub-tree of the
Tulasnellaceae (Figure 2.19b) also included five OTUs. Two of the five OTUs (OTU4 and OTU6) were closely related to Tulasnella bifrons, which have been related to mycorrhizal fungi from the North American native orchid Goodyera pubescens
(McCormik et al., 2004) and to Tulasnella sp. obtained from roots of Dactylorhiza sp.
(Jacquemyn et al., 2012). Tulasnella sp. associated with the orchid Tipularia discolor
(McCormik et al., 2004) were closely related to OTU2 and OTU 8. Operational
Taxonomic Unit 7 was closely related to Tulasnella irregularis observed in the roots of an orchid from Thailand (Nontachaiyapoom et al., 2010). In the third sub tree (Figure
2.19c), OTU 5 and OTU 9 were observed closely related to an uncultured Tulasnellaceae isolate from the roots of the orchid Cypripedium parviflorum (Shefferson et al., 2007) from Illinois in the United States.
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Discussion
Studies quantifying range-wide mycorrhizal communities of grassland orchids are severely limited, and are rarer still, or non-existent, within North America. Moreover, majority of the studies that report mycorrhizal association in orchids include samples collected only once from a few sites, thus restricting the interpretations with respect to the species-wide patterns of specificity over space and time (Rasmussen, 1995;
McCormick et al., 2006; Smith and Read, 2008; Dearnaley et al., 2012; Ercole et al.,
2015). We report both spatial and temporal variation patterns in the fungal communities within the roots of a rare terrestrial orchid native to North American tallgrass prairie.
Roots of Platanthera praeclara associated with two of the most common fungal families, Ceratobasidiaceae and Tulasnellaceae, that orchids across the world typically associate with. The fungal family Ceratobasidiaceae was represented at all of the 11 sampled sites in each of the two sampling years. Within the roots of the orchid, fungi from this family were more abundant than those of the fungal family Tulasnellaceae; however, an exception to this pattern was noted at the site BU in Minnesota across both sampling years. These patterns suggest that the orchid is specific in its choice of mycorrhizal fungi, and prefers the members of Ceratobasidiaceae (Figures 2.7 and 2.8).
Majority of the Ceratobasidiaceae OTUs identified in this study clustered with a previously known isolate from P. praeclara (Sharma et al. 2003) suggesting that the orchid has retained the associations with this narrow group of fungi for over a decade.
Rarity in orchids has been related to their specific associations with mycorrhizal fungi (Selosse et al., 2004; Shefferson et al., 2008; McCormik and Jacquemyn, 2014). In 60
Texas Tech University, Pablo Antonio Tovar, December 2015 addition to orchid rarity, geographic range of a species should be considered when assessing specificity in the association between orchids and mycorrhizal fungi (Otero et al., 2007). Data obtained over two years from >100 plants yielded 39 fungal OTUs across
11 sampled sites. Although 39 OTUs might appear to be a large number of different taxonomic units to have relationships with, it is important to note that specificity is defined by the phylogenetic breadth of the symbionts (Molina et al., 1992; Taylor et al.,
2002; McCormick et al., 2004). While Ceratobasidiaceae was observed to be the dominant fungal family across the natural range of the species when data from 2013 and
2014 were combined (represented by 238 nrITS sequences and 27 OTUs), their sequence distance (π = 0.039 ± 0.003) is lower than that of Tulasnellaceae (π = 0.135 ± 0.006). Its sequence distance is also lower than the distances from the same fungal family in other widely distributed orchids (Table 2.9) except for Serapias vomaracea. Interestingly, when comparing pi-distances from sequences of the fungal family Ceratobasidiaceae observed in P. praeclara with Ceratobasidiaceae known from orchids with limited distributions, only Hexalectris grandiflora, a nonphotosynthetic orchid, exhibited a smaller sequence distance. Pairwise sequence distances were supported by the constructed phylogenetic trees, displaying a narrow phylogenetic breadth in the OTUs observed in the family Ceratobasidiaceae, and a broader phylogenetic breadth in the
OTUs of the Tulasnellaceae. Among the Ceratobasidiaceae, 25 of 27 OTUs clustered together in a separate clade. Operational taxonomic units 3 and 12 were the most closely related to a previously published nrITS sequence of Ceratobasidiaceae from P. praeclara
(Genbank accession code DQ068771) (Figure 2.18). This particular genotype was
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Texas Tech University, Pablo Antonio Tovar, December 2015 detected in 2003, suggesting that the same isolate can remain associated with the species for over a decade. In previous studies (Sharma et al. 2003b), this isolate induced the highest germination in vitro when seeds of P. praeclara were cultured symbiotically.
Tulasnellaceae phylograms on the other hand are an example on a wider phylogenetic breadth, as the OTUs are distributed through the constructed sub trees (Figures 2.19a, b and c). Differences between the fungal communities associated with P. praeclara were noted when analyzing the data from each year separately. In the year 2013,
Ceratobasidiaceae was composed of 9 OTUs (from 122 nrITS sequences) versus 10
OTUs (from 48 nrITS sequences) in Tulasnellaceae. Despite having more OTUs in
Tulasnellaceae, sequence distances were smaller for Ceratobasidiaceae (π = 0.017 ±
0.003) in comparison to Tulasnellaceae (π = 0.109 ± 0.006). In the year 2014, the mycorrhizal fungal community of P. praeclara shifted to 14 OTUs in Ceratobasidiaceae
(116 nrITS sequences) and 5 OTUs in Tulasnellaceae (21 nrITS sequences). Still, sequence distances remained smaller for Ceratobasidiaceae (Table 2.8). For P. praeclara, the number of OTUs observed coupled with the sequence distances provides strong quantitative evidence that this orchid has high specificity compared to other orchids, majority of which have been studied with a much smaller sample size. Still, it is important to notice that rarefaction curves (Mao Tao and Chao 1 in Figure 2.15) showed that higher number of OTUs might have been detected if more plants were investigated although our results suggest that the increased diversity will most likely be attributed to variants within a narrow clade of Ceratobasidiaceae.
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Operational Taxonomic Unit data for the years 2013 (Table 2.5) and 2014 (table
2.6), and for the combined data set (Table 2.7) showed the community composition in terms of OTU for each sampled site. Although there are a few OTUs representing more than 5 sites in each data set (OTUs c1, c4 and c8 in 2013; OTUs c1 and c11 in 2014;
OTUs c1, c12 and c20 in the combined data set), most of the less common OTUs were observed to be restricted to one or two different sites. Moreover, composition of the mycorrhizal fungal communities at each sampled site varied from one year to the other.
Nine OTUs from the fugal family Ceratobasidiaceae were observed in 2013. The next year, 14 OTUs were detected for the same fungal family. For Tulasnellaceae, 10 OTUs were identified in 2013, and 5 were observed for 2014. Shifts in the community of fungal mycorrhizae are not unusual. Seasonal changes might alter the communities through time
(Ercole et al., 2015). Research in other, non-orchid mycorrhizal associations have shown temporal dynamics of fungal assemblages (Twieg et al., 2007; Dumbrell et al., 2011;
Bennett et al., 2013) to change within a year (Cowden and Peterson, 2013). Others have described no change in the community across seasons (Richard et al., 2011). For Orchid mycorrhizae, McCormick et al., (2006) observed that individuals of the orchid Goodyera pubescens maintained associations with single fungal genotypes for periods longer than four years. The reasons that explain shifts in fungal mycorrhizal communities are not clearly elucidated yet. Using high throughput sequencing, studies have shown stable communities of arbuscular mycorrhizae fungi in soil through time (Davison et al., 2012), as well as shifts in ECM fungi in soils of Quercus plants (Jumpponen et al., 2010).
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Multiple fungal taxa were detected in root systems of individual plants of P. praeclara. Several studies have reported this situation in other photosynthetic orchids:
Platanthera spp. (Zelmer et al., 1996); Tipularia discolor (McCormick et al. 2004);
Cephalantera damasonium, Epipactis atrorubens and Platanthera chlorantha
(Bidartondo et al., 2004); Aphyllorchis spp. (Roy et al., 2009); Orchis spp. (Jacquemyn et al., 2010; Lievens et al., 2010); Piperia yadonii (Sharma et al., 2007; Pandey et al., 2013)
Nervilia nipponica (Nomura et al., 2013); Neottia ovata (Jacquemyn et al., 2015).
However, photosynthetic orchid Goodyera pubescens (McCormick et al., 2006) roots were observed to be colonized by a single fungal taxon. Individual root systems associating with several fungal taxa are usually documented in non-orchid species that typically display low fungal specificity (Lodge & Wentworth, 1990; Perotto et al., 1994 and 1996; Monreal et al., 1999). Mycorrhizae from nonphotosynthetic orchids, which usually have high specificity towards mycorrhizal fungi, are colonized only by a single fungal taxon. Examples of nonphotosynthetic orchids presenting these pattern include
Corallorhiza maculata (Taylor and Bruns, 1999), Hexalectris revoluta (Taylor et al.,
2003) and Neottia nidus-avis (Selosse et al., 2002).
Variation in the dominance of orchid mycorrhizal fungi across sites was observed in P. praeclara. The Fisher’s test for the combined data set (Table 2.12), as well as for the data sets for years 2013 and 2014 independently (Table 2.10 and 2.11), indicated significant differences between the mycorrhizal fungal communities in most of the sampled sites. Since communities are different from one year to the other, similarities or differences between populations vary from one year to the next. The two-way
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Texas Tech University, Pablo Antonio Tovar, December 2015 hierarchical clustering analyses based on Bray-Curtis dissimilarities elucidated the relationships between sampled sites based on their mycorrhizal fungal communities by exhibiting different relationships in both years. Sites with similar mycorrhizal fungal communities were not subjected to the same type of land management in any of the data sets (2013, Figure 2.12; 2014, Figure 2.13; combined data set, Figure 2.14). The multiple response permutation procedure with all data sets ((P > 0.05) (Table 2.13) further confirmed that the variation of the mycorrhizal fungal communities across sites is not influenced by the overall prairie management practices. Also, nonmetric multidimensional scaling ordination analysis did not segregate sites according to their type of land management in either of the yearly data sets (Figures 2.16 and 2.17).
However, studies have shown how changes in habitat management can have significant effects on microbial communities (Carlile et al., 2001; Steenwerth et al., 2002; Johnson et al., 2003). Research has also shown that changes in the structure of fungal communities can be associated with alterations in soil properties like soil nitrogen availability (Frey et al., 2004), soil texture (Girvan et al., 2003), or soil pH (Fierer and Jackson, 2006).
Moreover, soil fungi might be sensitive to vegetation shifts that come along with different types of land management; this may be especially true for mycorrhizal fungi that form associations with particular plants (Heinemeyer et al., 2004). Lauber et al. 2008 observed significant differences in soil texture, carbon, nitrogen, and phosphorus measured under different land management practices. In the same study, phosphorus concentrations were correlated with fungal communities in that similar concentrations of the element fostered similar fungal communities, but no correlation was observed between land management
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Texas Tech University, Pablo Antonio Tovar, December 2015 and fungal communities in roots of P. praeclara. This suggests that phosphorus might be an important factor that regulates the biogeographical patterns in fungal communities in both soil and plant roots. Rousk et al. (2010) observed that pH and fungal community composition can be related, but the relationships are often weak in part because of the wide pH tolerance that fungal species usually have (Wheeler et al., 1991; Nevarez et al.,
2009). However, few soil variables with strong vectors appeared to correlate with the mycorrhizal fungal communities in orchid roots observed at different sites. In 2013, pH and sodium, exhibited a degree of correlation with the fungal community within orchid roots at PC (Pembina Control plot). Additional strong vectors like Mn and P1 did not appear to correlate with any of the remaining sites. In 2014, clay and H appeared to correlate with the mycorrhizal community within orchid roots at BU. Sites PH and
NDVK seemed to be correlated with S, and site BD may have a correlation with B.
Although the strength of the observed correlations can be inferred from the length of the vector, correlation analysis remain to be performed between these variables and the observed mycorrhizal fungal communities observed at each site to further infer the relationship between edaphic variables and fungal communities within the orchid roots.
Sharma et al. (2003a) isolated and identified mycorrhizae from roots of 21 P. praeclara plants sampled form four sites in Minnesota and one site in Missouri. They reported a higher proportion of the fungal family Ceratobasidiaceae (78 isolates) and a lower proportion for Tulasnellaceae (9 isolates). The current study analyzed 109 plants from 11 sites spanning a much wider geographic area to represent the entire natural range of the species and confirmed that the fungal family Ceratobasidiaceae was dominant and
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Texas Tech University, Pablo Antonio Tovar, December 2015 more abundant throughout the entire range. Our expectation with respect to the mycorrhizal specificity exhibited by the species was largely corroborated given the narrow phylogenetic breadth observed in the nrITS sequences of the fungal family
Ceratobasidiaceae. However, while the associations are specific, mycorrhizal communities of P. praeclara were not static, as shifts over time (from year 2013 to 2014) and space (from site to site) were evident.
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Table 2.1. Number of plants sampled and number of samples that yielded useable sequences from roots of Platanthera praeclara in 2013 and 2014. Plant roots were sampled for mycorrhizal diversity studies across 11 sites representing the natural distribution of the species.
Samples collected Samples yielding useable sequences
2013 2014 Total 2013 2014 Total
Number of plants 73 66 139 54 55 109
Number of root fragments 267 231 498 168 138 306
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Table 2.2 Soil chemical characteristics of 11 sites from where Platanthera praeclara roots were sampled in 2013 for mycorrhizal analyses. OM, organic matter; ENR, estimated nitrogen release; P1, phosphorus 1 (readily available to plants); P2, phosphorus 2 (readily available to plants plus part of the active reserve in the soil); K, potassium; Mg, magnesium; Ca, calcium; Na, sodium and CEC, cation exchange capability. Please see next page for the remaining Table.
Soil characteristics
OM ENR P1 P2 K Mg Ca Na CEC pH Site (%) (g/ha) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (meq/100g) PC 2.5 89668 27 39 8.2 167 607 3155 53 21 PH 3.1 103118 29 46 8.5 150 852 2582 54 20 PF 1.6 70613 40 45 8.3 137 516 1578 52 13 PS 2.7 94151 33 39 7.8 168 673 1605 57 14 NDG 0.9 53800 29 40 7.2 156 117 749 33 5 NDVK 1.2 60526 30 37 7.0 124 176 923 34 6 BU 3.1 101997 28 31 6.8 314 503 1923 45 15 BD 2.9 99755 28 41 7.8 262 552 4198 42 26 NE 3.1 104239 27 44 7.1 85 86 2145 43 12 IA 1.7 72855 36 39 6.7 413 320 1701 36 13
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Table 2.2 Continued Additional soil chemical characteristics at 11 sites from where Platanthera praeclara roots were sampled in 2013 for mycorrhizal analyses. NO3, nitrate; S, Sulphur; Zn, zinc; Mn, manganese; Fe, iron; Cu, copper; B, boron; Soluble salts and soil textural components (sand, silt and clay).
Soil characteristics
NO3 S Zn Mn Fe Cu B Soluble salts Sand Silt Clay Site (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (mmhos/cm) (%) (%) (%)
PC 4 9.6 1.2 11.9 60 1.0 0.9 0.4 74 10 16 PH 5 7.8 1.6 10.4 53 1.2 0.7 0.3 72 10 18 PF 3 6.9 0.8 6.4 44 0.8 0.9 0.3 74 12 14 PS 4 6.8 1.5 13.0 89 0.7 0.9 0.4 66 20 14 NDG 6 6.7 0.7 9.4 35 0.5 1.1 0.2 84 6 10 NDVK 6 7.7 1.4 14.6 105 0.5 0.8 0.1 82 6 12 BU 4 7.1 2.5 28.9 241 1.2 1.3 0.1 38 42 20 BD 3 7.7 1.5 17.2 52 1.6 1.0 0.2 32 40 28 NE 3 10.7 2.1 30.1 166 0.7 1.2 0.3 72 14 14 IA 2 6.2 2.1 41.9 76 1.3 0.9 0.2 28 48 24
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Table 2.3 Soil chemical characteristics of 11 sites from where Platanthera praeclara roots were sampled in 2014 for mycorrhizal analyses. OM, organic matter; ENR, estimated nitrogen release; P1, phosphorus 1; P2, phosphorus 2; K, potassium; Mg, magnesium; Ca, calcium; Na, sodium; H, hydrogen and CEC, cation exchange capability. Please see next page for the remaining Table.
Soil characteristics
OM ENR P1 P2 K Mg Ca Na H CEC pH Site (%) (g/ha) (uL/L) (uL/L) (uL/L) (uL/L ) (uL/L) (uL/L) (uL/L) (meq/100g)
MB 3.4 110964 7.9 28 7.7 130 574 4497 50 0.0 28 PC 3.8 118810 14.3 38 6.9 141 665 2702 54 0.3 20 PH 4.0 123294 14.3 75 7.4 151 913 4407 72 0.0 30 PF 4.0 124411 15.3 36 7.0 161 698 2653 57 0.0 20 PS 4.4 133381 17.1 25 6.8 170 732 2300 65 0.6 19 NDG 1.5 67251 10.9 21 6.2 94 127 1152 36 1.0 8.0 NDVK 1.2 59405 21.5 33 6.2 260 192 841 31 0.9 7.0 BU 3.3 107602 14.4 15 5.8 326 632 3281 36 5.3 28 BD 3.2 105360 11.2 26 6.6 270 641 4720 40 1.9 32 NE 3.3 108723 10.0 14 5.8 112 84 2366 47 3.0 16 IA 3.3 106481 30.5 41 6.1 553 447 2950 37 3.3 23
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Table 2.3 Continued Additional soil chemical characteristics of 11 sites from where Platanthera praeclara roots were sampled in 2014 for mycorrhizal analyses. NO3, nitrate; S, Sulphur; Zn, zinc; Mn, manganese; Fe, iron; Cu, copper and B, boron; Soluble salts and soil textural components (sand, silt and clay).
Soil characteristics
NO3 S Zn Mn Fe Cu B Soluble salts Sand Silt Clay Site (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (mmhos/cm) (%) (%) (%)
MB 3 4.7 0.9 18 118 1.2 0.3 0.2 72 12 16 PC 1 4.1 1.3 30 144 1.8 0.8 0.3 74 14 12 PH 1 10.6 1.6 36 126 2.4 0.5 0.4 72 12 16 PF 1 11.0 1.5 19 93 1.8 0.7 0.3 74 12 14 PS 1 3.7 1.4 30 128 1.4 0.7 0.4 76 12 12 NDG 3 5.8 0.7 14 60 0.6 0.5 0.1 82 4 14 NDVK 4 5.4 1.6 5 32 0.3 0.4 0.1 82 8 10 BU 3 4.4 3.4 39 153 1.5 0.5 0.2 38 38 24 BD 3 3.3 1.7 33 70 2.0 0.5 0.1 36 38 26 NE 2 3.1 0.6 11 173 0.6 0.5 0.1 76 10 14 IA 4 3.9 3.4 43 78 1.5 0.4 0.1 36 40 24
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Table 2.4 Primer pairs utilized, oligo sequences, and annealing temperature to amplify the fungal ITS region of the nuclear ribosomal DNA via polymerase chain reaction from the DNA extracted from root fragments of Platanthera praeclara.
Annealing Paired Temperature Primer Sequence primer (°C)
ITS1 -OF Forward AACTCGGCCATTTAGAGGAAGT ITS4 -OF 58 or 52 (Mix these two primers) AACTTGGTCATTTAGAGGAAGT ITS4 -OF Forward GTTACTAGGGGAATCCTTGTT ITS1 -OF
ITS4-Tul Reverse CCGCCAGATTCACACATTGA ITS1 ITS1 Forward TCCGTAGGTGAACCTGCGG ITS4-Tul 54
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Table 2.5 Number of root sections (i.e. sequences) representing each of the 19 operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal fungi at 11 sites in the year 2013. Values inside the parenthesis denote the number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was observed.
Site
OTU MB PC PH PF PS NDG NDVK BU BD NE IA
Ceratobasidiaceae C1 2(2) 8(4) 6(3) 6(1) 7(3) 13(4) 8(4) 7(6)
C2 1(1) 1(1)
C3 1(1)
C4 1(1) 4(2) 1(1) 4(2) 5(2) 1(1) 1(1) 12(5) 9(3)
C5 1(1)
C6 2(1)
C7 1(1) 2(2)
C8 1(1) 3(1) 3(2) 1(1) 5(3) 1(1) 2(1)
C9 1(1)
Tulasnellaceae
T1 1(1)
T2 1(1)
T3 3(2) 13(3) 4(2)
T4 5(3) 5(2) 3(2) 1(1)
T5 4(2)
T6 2(1)
T7 2(2)
T8 1(1)
T9 1(1)
T10 1(1)
Total 10(6) 12(5) 9(4) 21(6) 12(5) 23(4) 5(2) 22(5) 16(4) 25(8) 13(5)
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Table 2.6 Number of root sections (i.e. sequences) representing each of the 19 operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal fungi at 11 sites in the year 2014. Values inside the parenthesis denote the number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was observed.
Site
OTU MB PC PH PF PS NDG NDVK BU BD NE IA
Ceratobasidiaceae C1 6(4) 2(2) 10(4) 14(2) 5(1) 15(6) 4(4) 5(4)
C2 2(1) 1(1)
C3 2(1)
C4 3(2)
C5 1(1)
C6 1(1)
C7 1(1)
C8 1(1) 1(1)
C9 2(1)
C10 2(1)
C11 7(4) 1(1) 3(3) 5(3) 1(1) 2(2) 4(2) 5(2)
C12 2(1)
C13 4(1)
C14 4(1)
Tulasnellaceae T1 2(1)
T2 1(1) 6(4) 1(1)
T3 2(1) 4(2) 1(1)
T4 1(1)
T5 3(1)
Total 16(8) 9(4) 18(4) 22(5) 17(7) 10(4) 22(6) 6(3) 4(4) 11(4) 6(3)
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Table 2.7 Number of root sections (i.e. sequences) representing each of the 39 operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal fungi at 11 sites in the combined data set (years 2013 and 2014). Values inside the parenthesis denote the number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was observed.
Site OTU MB PC PH PF PS NDG NDVK BU BD NE IA Ceratobasidiaceae C1 3(3) 8(3) 11(7) 15(3) 7(3) 17(5) 7(5) 12(8) 12(10) C2 3(2) C3 2(1) C4 2(1) C5 4(1) C6 1(1) 1(1) 8(2) 1(1) C7 1(1) 2(2) C8 2(1) 1(1) C9 4(4) C10 4(1) C11 1(1) 1(1) C12 2(2) 5(3) 6(5) 1(1) 5(4) 1(1) C13 1(1) C14 1(1) C15 1(1) C16 1(1) C17 1(1) C18 1(1) C19 1(1) C20 9(6) 2(2) 3(3) 7(3) 5(4) 5(3) 7(4) 1(1) 1(1) 16(7) 13(5) C21 1(1) C22 1(1) 1(1) C23 2(1) C24 2(1) C25 1(1) C26 1(1) C27 1(1) Tulasnellaceae T1 1(1) T2 6(5) 6(4) 5(2) 3(3) 1(1) T3 2(1) 2(1) 14(4) 4(2) 1(1) T4 5(1) T5 2(1) T6 1(1) T7 2(2) T8 2(1) 1(1) 1(1) T9 3(2) T10 1(1) T11 1(1) T12 1(1)
Total 27(13) 21(9) 25(8) 45(11) 29(11) 33(9) 23(8) 24(8) 21(8) 34(12) 19(8) 85
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Table 2.8 Mean pairwise sequence distances and standard errors (SE) for all observed fungal ITS sequences from roots of Platanthera praeclara sampled across 11 sites in years 2013 and 2014, and for the combined data set (years 2013 and 2014), separated by fungal family. The values presented within the parenthesis are the numbers of individual ITS sequences (n) composing each group.
Data set Fungal Family (n) Pi - Distance SE
2013 and 2014 Ceratobasidiaceae (238) 0.039 0.003 combined Tulasnellaceae (69) 0.135 0.006
Ceratobasidiaceae 2013 (122) 0.017 0.003 2013 Tulasnellaceae 2013 (48) 0.109 0.006
Ceratobasidiaceae 2014 (116) 0.072 0.005 2014 Tulasnellaceae 2014 (21) 0.073 0.006
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Table 2.9 Mean pairwise fungal ITS sequence distances (pi-distance) estimated based on combined transitions and transversions substitution model, within fungal families identified in roots of Platanthera praeclara across 11 sites in North America. Mean pairwise sequence distances from publically available fungal ITS sequences obtained from orchids other than Platanthera praeclara and Nervilia nipponica were calculated by Pandey et al., 2013.
Taxon Tulasnellaceae Ceratobasidiaceae Natural distribution n pi-distance n pi-distance
Platanthera praeclara 69 0.135 ± 0.006 238 0.039 ± 0.003 Wide Piperia yadonii 58 0.231 ± 0.026 71 0.077 ± 0.006 Restricted Anacampis laxiflora 12 0.186 ± 0.009 12 0.071 ± 0.006 Wide Ophrys fuciflora 12 0.225 ± 0.010 7 0.063 ± 0.006 Wide Orchis purpurea 9 0.089 ± 0.008 Wide Serapias vomeracea 27 0.097 ± 0.007 8 0.038 ± 0.004 Wide Chiloglottis aff. jeanesii 14 0.014 ± 0.003 Restricted Chiloglottis trapeziformis 12 0.006 ± 0.002 Wide Goodyera foliosa 7 0.062 ± 0.009 Wide Goodyera hachijoensis 5 0.103 ± 0.009 Moderate Goodyera tesselata 5 0.044 ± 0.006 Wide Cypripedium calceolus 12 0.005 ± 0.002 Moderate Cypripedium candidum 7 0.003 ± 0.002 Wide Cypripedium japonicum 18 0.033 ± 0.006 Wide Nervilia nipponica 9 0.157 ± 0.007 Wide Hexalectris grandiflora 6 0.023 ± 0.003 Moderate
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Table 2.10 Statistical significance (P values) from the Fisher’s exact test comparing the fungal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) at each site for 2013 below the diagonal. Pairwise geographic distances (kilometers) between population pairs are presented above the diagonal. MB, Manitoba; PC, Pembina Control; PH, Pembina Hay; PF, Pembina Fall fire; PS, Pembina Spring fire; NDG, North Dakota Graze; NDVK, North Dakota Viking; BU, Bluemound Uphill; BD, Bluemound Downhill; NE, Nebraska and IA, Iowa.
MB PC PH PF PS NDG NDVK BU BD NE IA
MB - 167 167 167 167 309 298 610 609 805 838
PC < 0.05 - 0 0 0 158 142 444 443 667 670
PH < 0.05 1.00 - 0 0 158 142 444 443 667 670
PF 0.111 < 0.05 < 0.05 - 0 158 142 444 443 667 670
PS 0.185 0.270 0.249 0.371 - 158 142 444 443 667 670
NDG < 0.05 0.922 0.671 < 0.05 0.308 - 22 319 319 509 553
NDVK < 0.05 < 0.05 < 0.05 0.050 < 0.05 < 0.05 - 321 321 527 554
BU < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 - 3 378 234
BD < 0.05 0.212 0.119 0.068 0.127 < 0.05 < 0.05 < 0.05 - 382 234
NE < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 0.649 < 0.05 < 0.05 - 443
IA < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 1.000 < 0.05 < 0.05 < 0.05 -
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Table 2.11 Statistical significance (P values) from the Fisher’s exact test comparing the fungal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) at each site for 2014 below the diagonal. Pairwise geographic distances (kilometers) between population pairs are presented above the diagonal. MB, Manitoba; PC, Pembina Control; PH, Pembina Hay; PF, Pembina Fall fire; PS, Pembina Spring fire; NDG, North Dakota Graze; NDVK, North Dakota Viking; BU, Bluemound Uphill; BD, Bluemound Downhill; NE, Nebraska and IA, Iowa.
MB PC PH PF PS NDG NDVK BU BD NE IA
MB - 167 167 167 167 309 298 610 609 805 838 PC < 0.05 - 0 0 0 158 142 444 443 667 670 PH 0.056 < 0.05 - 0 0 158 142 444 443 667 670 PF < 0.05 < 0.05 < 0.05 - 0 158 142 444 443 667 670 PS < 0.05 < 0.05 < 0.05 < 0.05 - 158 142 444 443 667 670 NDG 0.052 < 0.05 0.194 0.072 < 0.05 - 22 319 319 509 553 NDVK < 0.05 < 0.05 0.179 < 0.05 < 0.05 0.066 - 321 321 527 554 BU < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 - 3 378 234 BD 0.228 < 0.05 0.850 1.00 < 0.05 0.832 1.000 < 0.05 - 382 234 NE 0.648 < 0.05 0.340 < 0.05 < 0.05 0.517 0.102 < 0.05 0.384 - 443 IA 0.117 < 0.05 < 0.05 < 0.05 0.140 < 0.05 < 0.05 < 0.05 < 0.05 0.117 -
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Table 2.12 Statistical significance (P values) from the Fisher’s exact test comparing the fungal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) at each site using the combined data set (years 2013 and 2014) below the diagonal. Pairwise geographic distances (kilometers) between population pairs are presented above the diagonal. MB, Manitoba; PC, Pembina Control; PH, Pembina Hay; PF, Pembina Fall fire; PS, Pembina Spring fire; NDG, North Dakota Graze; NDVK, North Dakota Viking; BU, Bluemound Uphill; BD, Bluemound Downhill; NE, Nebraska and IA, Iowa.
MB PC PH PF PS NDG NDVK BU BD NE IA MB - 167 167 167 167 309 298 610 609 805 838 PC < 0.05 - 0 0 0 158 142 444 443 667 670 PH < 0.05 0.064 - 0 0 158 142 444 443 667 670 PF < 0.05 0.066 < 0.05 - 0 158 142 444 443 667 670 PS < 0.05 < 0.05 < 0.05 < 0.05 - 158 142 444 443 667 670 NDG < 0.05 0.114 0.381 0.05 < 0.05 - 22 319 319 509 553 NDVK < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 - 321 321 527 554 BU < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 - 3 378 234 BD < 0.05 < 0.05 < 0.05 0.06 < 0.05 0.281 < 0.05 < 0.05 - 382 234 NE < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 - 443 IA < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 < 0.05 -
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Table 2.13 Multiple response permutation procedure (MRPP) showing the significance of the variation of the mycorrhizal fungal communities by using abundance data observed at 11 sites of Platanthera praeclara by grouping the sites according to the type of land management (prescribed fire, haying or grazing, or no management). The MRPP was performed with the combined data set (years 2013 and 2014) as well as years 2013 and 2014 separately.
Combined data set (2013 and 2014) 2013 2014 p-value p-value p-value
Management 0.380 0.305 0.198 Latitude 0.034 0.070 0.192
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Figure 2.1 A partial map of the US and Canada showing the natural distribution of Platanthera praeclara. The states where the plant is currently known to be present (orange), the counties where the populations occur (gray) and the counties where roots were collected for the studies (yellow) are highlighted.
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Figure 2.2 A map of the nuclear fungal ribosomal internal transcribed spacer (ITS) region. The two ITS regions are between the SSU18S and LSU28S ribosomal RNA genes and are separated by the 5.8S rRNA gene. Image taken from http://www.gatc- biotech.com
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Figure 2.3 Plants of Platanthera praeclara excavated for sample collection while keeping the root system intact. Photo on the left was taken by Nancy Sather.
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Figure 2.4 A whole seedling of Platanthera praeclara along with severed root and tuber
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Figure 2.5 Root fragments collected from plants of Platanthera praeclara.
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Figure 2.6 A transversal section of a root of Platanthera praeclara showing colonization by fungal pelotons in cortical cells (examples indicated by arrows). The scale-bar represents 400 microns.
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MB PC PH 0 0
2
5 12 9
PF PS NDG 1
3 10 11 9 22
NDVK BU BD 0 2 6 10 5 20
NE IA 0 Ceratobasidiaceae 2
Tulasnellaceae 25 11
Figure 2.7 Pie charts showing the distribution of the fungal families Ceratobasidiaceae and Tulasnellaceae observed in roots of Platanthera praeclara at 11 sites in the year 2013. Numeric values represent the individual ITS sequences belonging to each fungal family.
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MB PC PH 0 3 3
6 13 18
PF PS NDG 1
3 2
19 16 8
NDVK BU BD 0 0
2
4 18 4
NE IA
Ceratobasidiaceae 1 1
Tulasnellaceae 10 5
Figure 2.8 Pie charts showing the distribution of the fungal families Ceratobasidiaceae and Tulasnellaceae observed in roots of Platanthera praeclara at 11 sites in the year 2013. Numeric values represent the individual ITS sequences belonging to each fungal family.
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(a) 35 30
25 20 15
# of plants of # 10 5 0 1 2 3 4 # of OTUs
(b) 60 50
40 30
# of plants of # 20 10 0 1 2 # of fungal families
Figure 2.9 One to four operational taxonomic units in individual plants of Platanthera praeclara (a), and number of individual plants hosting one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) (b), across 11 sites in 2013.
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(a) 50
40
30
20 # of plants of # 10
0 1 2 3 4 # of OTUs
(b) 60 50
40 30
# of plants of # 20 10 0 1 2 # of fungal families
Figure 2.10 One to four operational taxonomic units in individual plants of Platanthera praeclara (a), and number of individual plants that hosted one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) (b), across 11 sites in 2014.
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(a) 50
40
30
20 # of plants of # 10
0 1 2 3 4 # of OTUs
(b) 60 50
40 30
# of plants of # 20 10 0 1 2 # of fungal families
Figure 2.11 One to four operational taxonomic units in individual plants of Platanthera praeclara (a), and number of individual plants that hosted one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) (b), across 11 sites for the combined data set (years 2013 and 2014).
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Figure 2.12 Two-way hierarchical cluster tree and matrix coding based on abundance of 19 fungal operational taxonomic units (OTUs) observed at 11 sites where roots of Platanthera praeclara were sampled in the year 2013.
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Figure 2.13 Two-way hierarchical cluster tree and matrix coding based on abundance of 19 fungal operational taxonomic units (OTUs) observed at 11 sites where roots of Platanthera praeclara were sampled in the year 2014.
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Figure 2.14 Two-way hierarchical cluster tree and matrix coding based on abundance of 39 fungal operational taxonomic units (OTUs) observed at 11 sites where roots of Platanthera praeclara were sampled in years 2013 and 2014.
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60
50
40
30
Mao Tau # ofOTUs # Chao 1 20
10
0 27 55 82 109 137 164 192 219 246 274 301 # of sequences
Figure 2.15 Sample based observed (Mao Tau) and rarefaction (Chao 1) cumulative fungal operational taxonomic unit (OTU) abundance curves for Platanthera praeclara. Fungal communities within the roots of plants were determined by sampling plants from 11 sites that host the orchid species.
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Figure 2.16 Nonmetric multidimensional scaling (NMS) ordination of the mycorrhizal fungal communities within the roots of Platanthera praeclara sampled from 11 sites with correlations to soil variables [Na, Mn, Zn, Ca, OM (organic matter), CEC (cation exchange capability), soluble salts and pH] for the year 2013.
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Figure 2.17 Nonmetric multidimensional scaling (NMS) ordination of the fungal communities of Platanthera praeclara sampled from 11 sites with correlations to soil variables (S, B, H, clay and silt) for the year 2014.
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Figure 2.18 Partial Maximum likelihood tree of the fungal family Ceratobasidiaceae constructed with operational taxonomic unit (OTU) sequences observed in Platanthera praeclara roots collected from 11 sites across its natural habitat. Sequences of known orchid mycorrhizal fungi were added to the alignments to place our sequences in a broader context. Scale bar represents estimated number of DNA substitutions per site. Only branch support values over fifty percent are displayed (maximum likelihood / posterior probability). Please see the next page for the remaining tree. 109
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Figure 2.18. Continued Maximum likelihood tree of the fungal family Ceratobasidiaceae constructed with operational taxonomic unit (OTU) sequences observed in Platanthera praeclara roots collected from 11 sites across its natural habitat. Only branch support values over fifty percent are displayed (maximum likelihood / posterior probability).
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Figure 2.19a Maximum likelihood tree of the fungal family Tulasnellaceae constructed with operational taxonomic unit (OTU) sequences observed in Platanthera praeclara roots collected from eleven populations across its natural habitat. Sequences of known orchid mycorrhizal fungi were added to the alignments to place our sequences in a broader context. Scale bar represents estimated number of DNA substitutions per site. Only branch support values over fifty percent are displayed (maximum likelihood / posterior probability). 111
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Figure 2.19b Maximum likelihood tree of the fungal family Tulasnellaceae constructed with operational taxonomic unit (OTU) sequences observed in Platanthera praeclara roots collected from eleven populations across its natural habitat. Sequences of known orchid mycorrhizal fungi were added to the alignments to place our sequences in a broader context. Scale bar represents estimated number of DNA substitutions per site. Only branch support values over fifty percent are displayed (maximum likelihood / posterior probability).
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Figure 2.19c Maximum likelihood tree of the fungal family Tulasnellaceae constructed with operational taxonomic unit (OTU) sequences observed in Platanthera praeclara roots collected from eleven populations across its natural habitat. Sequences of known orchid mycorrhizal fungi were added to the alignments to place our sequences in a broader context. Scale bar represents estimated number of DNA substitutions per site. Only branch support values over fifty percent are displayed (maximum likelihood / posterior probability).
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CHAPTER III
MYCORRHIZAL ASSOCIATIONS OF PLATANTHERA PRAECLARA IN RESPONSE TO PRESCRIBED FIRE AND HAYING
Abstract
Grassland plant communities are affected by vegetation management practices including fire, grazing, haying, etc. Consequently, edaphic characteristics within the system can also change in response. Changes in soil can lead to shifts in functioning and composition of microbial communities, including mycorrhizal fungi, which are critical in germination and survival of orchid species. We tested the effect of prairie management treatments on mycorrhizal fungal communities within the roots of
Platanthera praeclara which is a rare terrestrial orchid native to the tallgrass prairies in North America. Root sampling, along with characterizing the edaphic variables, was conducted over two years at a long-term experimental site in Minnesota, USA.
Across the two years, Ceratobasidiaceae and Tulasnellaceae were observed as the fungal families composing mycorrhizal fungal communities in the roots of P. praeclara. Twenty operational taxonomic units (OTUs) were observed through both years of sampling. Fifteen OTUs were identified for Ceratobasidiaceae and five for
Tulasnellaceae. Variation in the mycorrhizal fungal communities was observed from
2013 (four OTUs) to 2014 (15 OTUs). Significant differences (P<0.05) were observed only between the composition of fungal communities of PC (no management) and PF (fall burn treatment) in 2013. In 2014, the composition of fungal communities between all four treatments was significantly different (P<0.05).
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Composition of mycorrhizal fungal communities did not appear to be associated with the type of prairie management treatment. However, temporal shifts appeared to be most noticeable under annual haying and both prescribed fire treatments by yielding an increase in the richness of OTUs. Further, fungal communities within orchid roots appeared to vary over time in response to edaphic variation across the four prairie management treatments. In 2013, sand and pH were correlated with the fungal community observed in treatment PC (no management); the fungal community in PH
(annual haying) was correlated with clay; silt and sodium (Na) were correlated to the fungal community observed in the treatment PS (spring fire); and the fungal community in PF (fall fire) was correlated with boron (B) and readily available phosphorus (P1). In 2014, the fungal community observed in treatment PC was correlated with volumetric water content in June (VWC J) and silt; sand, organic matter (OM) and P1 were correlated with the fungal community observed in treatment PS; clay and electrical conductivity in May (EC M) were correlated with the fungal community in PH; and sulfur was correlated with the fungal community observed in PF. We conclude that despite the relative specificity of the orchid toward its fungi (Ceratobasidiaceae π = 0.049 ± 0.04, N=97; Tulasnellaceae π = 0.203 ±
0.09, N=23), OTUs within the entire suite of its fungal associates change in response to time and management combinations.
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Introduction
Anthropogenic factors continue to induce habitat destruction and degradation across the tallgrass prairies of the Great Plains in North America, which is one of most biodiverse ecosystems on earth. This ecosystem hosts many rare and endemic taxa including the federally threatened terrestrial orchid Platanthera praeclara (USFWS,
1996). It is known that management practices such as prescribed fire, grazing, haying, and/or fertilization can lead to changes in the ecosystem components above and below ground (Bentivenga et al., 1992; Dhilhon and Anderson, 1993; Bossio et al., 1998; Bossio et al., 2005). Habitat degradation and loss can also lead to loss of pollinators.
Additionally, local hydrology can change, and plant communities can shift over time. As the plant communities shift, soil microbial function and communities often shift concurrently. Soil respiration, metabolic profiles (Bendig et al., 2000) and enzyme activities (Bandick and Dick, 1999) are some of the measurements that have shown response to different types of land use because they rapidly respond to disturbances in the soil environment (Johnson et al., 2003). Land use changes may have significant and long- lasting effects on soil properties like carbon and nutrient content, soil texture and pH
(Murty et al., 2002). These effects are usually a consequence of changes in plant species composition. Changes in land use or management ultimately affect microbial communities (Bossio et al., 1998; Johnson et al., 2003; Wu et al., 2015). Studies have shown that shifts in the structure of fungal communities are associated with changes in soil properties such as texture (Girvan et al., 2003), soil pH (Fierer and Jackson, 2006) and soil nitrogen and phosphorus availability (Frey et al., 2004; Lauber et al., 2008).
Although these variables change depending on land use or management, it is not clear 116
Texas Tech University, Pablo Antonio Tovar, December 2015 how they affect biogeographical patterns in soil microbial communities (Lauber et al.,
2008).
Density of fungal propagules in soil is known to be affected by land management practices (Wu et al., 2007). Increased fungal hyphal biomass is usually observed in organically managed crop systems when compared to soil from conventionally managed systems (Sivapalan et al 1993; Shannon et al., 2002; Wu et al., 2007). Animal grazing, soil tillage and crop rotation had been observed to affect fungal densities in soil (Frey et al., 1999; Hedlund, 2002; Singh and Rai, 2004).
Mycorrhizal fungal communities may be susceptible to shifts in vegetation as they usually associate with specific plant types (Heinemeyer et al., 2004). For example, fungi from the phylum Basidiomycota that are usually involved in decomposing lignified plant detritus might be more affected by turning forest soils into agricultural soils (Bardgett and McAlister, 1999). Bentivenga and Hetrick (1992) examined arbuscular mycorrhizal
(AM) fungal species composition in tallgrass prairie subjected to annual fire, mowing and fertilization treatments, but no significant effects of these management types were observed on the AM fungal composition. On the other hand, Dhillion and Anderson
(1993) have shown that AM fungal communities in grasslands are influenced by application of fire, mowing, grazing and fertilization practices. Moreover, mycorrhizal root colonization has been reported to increase (Bentivenga and Hetrick, 1991), decrease
(Rashid et al., 1997) and be unaffected (Anderson and Menges, 1997) in response to prescribed fire. These conflicting results may suggest that fungal communities likely respond to a combination of factors including management and edaphic characters (Eom 117
Texas Tech University, Pablo Antonio Tovar, December 2015 et al., 1999). Also, the response time to allow detection of the shifts in communities may vary depending on the vegetation and / or fungal community.
Although an increasing number of mycorrhizal fungal communities are being described and characterized from roots of orchids, there is no evidence for how these communities are affected by vegetation management practices. We utilized a long-term
(over 20 years) experimental set-up at Pembina Trail Preserve in Polk County,
Minnesota, USA, where one of the largest populations of P. praeclara occurs to test the hypothesis that the diversity of fungi associated with P. praeclara will be similar across four management treatments (i.e., prescribed fire in spring every 4 years, prescribed fire in fall every 4 years, annual haying, and no active vegetation management) over the two sampling years because of the expectation of overall high mycorrhizal specificity in the species.
Materials and methods Collection sites
Root tissue from individual plants of P. praeclara were sampled at the Pembina
Trail Preserve in Polk County, Minnesota, USA. Four experimental treatment plots with different land management [prescribed fire in spring every 4 years (PS), prescribed fire in fall every 4 years (PF), annual haying (PH), and no vegetation management (control;
PC)] were sampled to allow a comparison of fungal associations under each prairie
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Texas Tech University, Pablo Antonio Tovar, December 2015 management treatment (Figure 3.1). The most recent fall burn was applied in November of 2011, whereas the last spring burn was applied in April, 2012.
Sample collection
In 2013 and 2014, roots from individual plants were sampled between mid-June and early July. By this time in the growing season, differentiating between seedlings, juveniles or vegetative (individuals that will not produce an inflorescence in the growing season), and reproductive individuals was possible. Roots from two seedlings, two vegetative and two reproductive plants were sampled within each prairie management treatment plot. Roots from one or two extra plants were sampled with permission.
Twenty-four and 20 plants were sampled across the 11 sites during 2013 and 2014, respectively (Table 3.3). Whole plants were carefully excavated to examine the complete root system (see Figure 2.3 in Chapter II). To keep plants alive after sampling, just a few roots were extracted from the plant, and the sampled plant was replanted. Sampled plants were watered thoroughly immediately after replanting to maximize reestablishment.
Because seedlings have very small root systems, the whole seedling was collected and all roots were processed for fungal identification (see Figure 2.4 in Chapter II). Roots were stored at 4°C until they were shipped overnight to the laboratory at Texas Tech
University for further processing.
In each of the two years, one aggregate soil sample was collected at each treatment where orchids were sampled by adding together at least three subsamples. Approximately
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500g of soil were extracted from sites where the plants were present. The soil was stored at room temperature until it was processed further. Physicochemical analyses were carried out by A&L Plains Labs, Inc., Lubbock, Texas. Organic matter (OM), pH, macro nutrients (K, Mg, Ca, Na), cation exchange capacity (CEC), NO3-N, other nutrients (S,
Zn, Mn, Fe, Cu, B) and textural components (% sand, silt and clay) were measured in each sample.
Environmental data
A cr800 data logger (Campbell Scientific, Salt Lake City, Utah) was installed with sensors recording continuous data (one reading per hour) on soil temperature, soil volumetric water content, and soil electrical conductivity. The data logger was set up at a location from where probes could simultaneously gather information in each of the four different treatment plots. Two sensors were utilized as replicates in each treatment plot.
Averages of the volumetric water content (VWC), electrical conductivity (EC) and soil temperature at orchid root depth were computed over the two replicate sensors for April,
May and June. We selected these months because the soil temperatures were below freezing prior to April and presumed to induce minimal or no mycorrhizal (plant or fungus) activity. Similarly, data beyond June were not used because root sampling occurred in the month of June (Table 3.1).
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Fungal identification from root sections
Roots were rinsed under tap water to wash them free of soil and debris before photo-documentation (see Figure 2.5 in Chapter II). Roots from each individual plant were subsequently placed in individual 50 mL plastic tubes. Inside a laminar flow hood and under sterile conditions, the surface sterilization was carried out by following these steps: (1) roots were first rinsed in a 70% EtOH (ethanol) solution by shaking thoroughly for 50 seconds; (2) roots were then rinsed with a 3% NaOCl (sodium hypochlorite) solution by shaking thoroughly for 30 seconds; (3) a 70% ethanol rinse for 50 seconds was carried out; (4) roots were finally rinsed with ultrapure, sterile water several times to remove residues of EtOH and NaOCl. Surface sterilization treatment was modified as needed based on the thickness and the number of the roots (Pandey et al., 2013).
The epidermis of each root was gently shaved off using a sterile scalpel to remove additional microbes that might have survived the surface sterilization treatment. Peloton inspection was conducted by cutting transversal thin slices of root tissue and examining under a compound microscope (see Figure 2.6 in Chapter II). If pelotons were observed,
2.5 to 3 cm long segments adjacent to the inspected slice were collected, sliced thinly and stored individually in 2 mL plastic tubes at -80°C until DNA was extracted (Pandey et al.,
2013).
DNA extraction was conducted using Qiagen DNeasy Plant Mini Kit (Qiagen,
Valencia, CA, USA) by following the manufacturer's protocol with a slight modification.
All samples were incubated in a 3.3% solution of polyvinylpyrrolidone in AP1 lysis buffer at 65°C for 2 hours while mixing once every 30 minutes before the RNase 121
Texas Tech University, Pablo Antonio Tovar, December 2015 digestion step. Subsequently, 4 μl of RNase (supplied with the kit) was added to each sample and samples were incubated at 65°C for 15 minutes, mixing once every 4 minutes. Total DNA was eluted in 50 μl of elution buffer and stored at -80°C until further processing.
PCR and sequencing
Polymerase chain reaction was performed using total DNA extracted from individual root segments. Each reaction was prepared to a final volume of 25 μL by using
Promega GoTaq Flexi DNA Polymerase reagent kit (Promega, Madison, Wisconsin,
USA). Concentrations for each reagent were: 5x Green GoTaq Flexi Buffer, 10 mM/μL dNTPs (100 mM of each dNTP), 25 mM of MgCl2, 10 μM/μL of each primer, 10 μg/μL of BSA and 5 u/μL GoTaq DNA polymerase (Pandey et al., 2013). PCR consisted of 35 cycles in an epGradients Master Cycler (Eppendorf, Hamburg, Germany) and included a
2 minute initial denaturation at 94°C before thermocycling with a 45 seconds denaturation at 94°C followed by a 45-second annealing at different temperatures depending on the primer pair (ITS1-OF and ITS4-OF; ITS4-TUL and ITS1) used (Table
2.4 in Chapter II) and 72°C elongation for 1 minute. Finally, the last cycle was followed by an extension at 72°C for 5 minutes.
Two primer pairs ITS1-OF/ITS4-OF and ITS1/ITS4-TUL (Taylor & McCormick,
2008) were used. A total of 498 root segments were assayed for molecular analysis. Each
DNA sample was first subjected to PCR using the ITS-OF primer pair at 58°C annealing
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Texas Tech University, Pablo Antonio Tovar, December 2015 temperature. Samples that did not amplify were then run with the same primer pair at
52°C annealing temperature. If amplification was still not obtained, ITS1/ITS4-TUL was used with 54°C annealing temperature on the samples that did not amplify previously. A
2% agarose gel electrophoresis was run to verify amplification. Samples showing a single band of the expected size range (600 – 800 bp) were cleaned using DNA Clean and
Concentrator 5 kit (Zymo Research, Irvine, CA, USA). DNA quantity and quality were measured using a NanoDrop 2000c spectrophotometer (Thermo Scientific, Wilmington,
DE, USA). Samples showing multiple bands were processed using GenElute Gel
Extraction Kit (Sigma-Aldrich, Poole, United Kingdom) by isolating the band closer to the expected size range. Sanger sequencing of all cleaned PCR products was carried out by the DNA Analysis Facility on Science Hill at Yale University. Samples were sequenced in one direction using the reverse primer (ITS4-OF or ITS4-TUL) (Pandey et al., 2013).
Cloning
Polymerase chain reaction products from samples with sequencing difficulties were purified either using PureLink PCR Purification kit (Invitrogen Life Technologies,
Carlsbad, California) or GenElute Gel Extraction Kit (Sigma-Aldrich, Poole, United
Kingdom). Once a clean PCR product was obtained, it was used to carry out a cloning reaction and cloning transformation using the TOPO TA Cloning kit for Sequencing along with One Shot TOP10 and DH5α-T1 competent cells (Invitrogen Life technologies,
Carlsbad, California), following the manufacturer protocol. 123
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To analyze the transformants, 3-5 colonies were selected and cultured overnight in
1 mL of LB (Luria-Bertani) (ThermoFisher Scientific, Waltham, MA, USA) medium containing 50 μg/mL of ampicillin. Plasmid DNA was isolated using the PureLink Quick
Plasmid Miniprep Kit, (Invitrogen Life Technologies, Carlsbad, CA) following the protocol included in the kit.
Polymerase chain reaction was performed on the extracted plasmid DNA following the amplification protocol described earlier using ITS1-OF/ITS4-OF primer pair.
Polymerase chain reaction products were purified using GenElute Gel Extraction Kit
(Sigma-Aldrich, Poole, United Kingdom) and DNA concentrations were measured using a NanoDrop 2000c spectrophotometer (Thermo Scientific, Wilmington, DE, USA).
Sanger sequencing of all cleaned PCR products was carried out as described above.
Data analyses
DNA Analysis Facility on Science Hill at Yale University performed a basic trimming on the sequences. However, the obtained sequences were double-checked and a more thorough trim was implemented using Geneious 4.8.5. Trimmed sequences were then identified through BLAST searches using the Megablast option for finding highly similar
DNA sequences.
OTU clustering
Sequences were grouped into operational taxonomic units (OTUs) at 97% sequence similarity criterion using the OTU pipeline from the University of Alaska at
Fairbanks Life Science Informatics Portal (http://www.borealfungi.uaf.edu; Taylor &
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Houston 2011). The longest and highest quality representative sequence from each OTU was used for further analyses. Operational taxonomic Units were obtained separately for sequences obtained from root fragments collected in year 2013 and 2014. Additionally, for general diversity estimation, sequences were also clustered into OTUs by combining the data sets for both years.
Mean pairwise sequence distance (pi, π)
To estimate sequence divergence within the identified fungal families, separate multiple alignments of sequences were constructed for years 2013 and 2014 data sets, and for the combined data set, using MAFFT version 7 with L-INS-I model (Katoh and
Standley, 2013). Mean pairwise sequence distance (pi, π; Nei and Kumar, 2000) among all individual sequences within a fungal family was then estimated using MEGA 6
(Tamura et al., 2013). Mean pairwise sequence distances obtained using the combined 2- year data set from P. praeclara mycorrhizal fungi were compared with the previously reported pi distances among mycorrhizal fungi from other orchids (Pandey et al., 2013).
Sequence divergence for the observed sequences groups by treatment
(independently of the fungal family they belonged to) were also computed.
Fisher’s exact test
The statistical significance of the variation in fungal diversity between treatments was tested using the Fisher’s exact test based on abundances of individual OTUs across 125
Texas Tech University, Pablo Antonio Tovar, December 2015 the sampled treatments per year of collection. The test was performed in R (R
Development Core Team 2010) using all the OTUs detected in each of the sampled treatment for 2013 and 2014 separately and also for the combined data set.
Two-way hierarchical ordination
Two-way hierarchical ordination analysis (Bray and Curtis., 1957) using OTU abundances was performed to test the relationships between the mycorrhizal fungal communities of the sites based on dissimilarities. PC-ORD (McCune et al., 2002) was used to perform the two-way hierarchical ordination using the Sorensen (Bray-Curtis) distance measure, and setting the group linkage criterion as ‘average’. This test was conducted for each sampling year separately and also for the combined 2-year data set.
The resulting dendrogram clustered sites according to similarity in mycorrhizal fungal communities held by the sampled plants across the four treatments, and served as an aid to observe whether sites sharing land management types (prescribed burn, haying or grazing, no management) clustered together.
Nonmetric multidimensional scaling
Nonmetric multidimensional scaling (NMDS) in PC-ORD (McCune et al., 2002) was used to ordinate OTU abundance in roots of P. praeclara and to visualize the correlation between edaphic characteristics [OM, organic matter; ENR, estimated nitrogen release; P1, phosphorus 1 (readily available to plants); P2, phosphorus (readily
126
Texas Tech University, Pablo Antonio Tovar, December 2015 available to plants plus part of the active reserve in the soil) 2; K, potassium; Mg, magnesium; Ca, calcium; Na, sodium; CEC, cation exchange capability; NO3, nitrate; S,
Sulphur; Zn, zinc; Mn, manganese; Fe, iron; Cu, copper and B, boron) and OTUs observed at each sampled site separately for each sampling year. The abundance-based version of the Sorensen index (Bray-Curtis) was used to calculate the distances.
Dimensionality of the ordination was determined to choose the lowest dimensionality that captured most of the variation. Data sets from 2013 and 2014 were both best described by
3-dimensional solutions with instabilities below 0.00001.
Results Operational taxonomic Units diversity
Out of 161 root fragments in 44 plants obtained from across all four treatments, 120 root fragments in 39 plants produced workable DNA sequences (Table 3.3).
Ceratobasidiaceae and Tulasnellaceae were identified as the mycorrhizal fungal families associated to Platanthera praeclara across the sampled land management treatments in both sampling years (Figures 3.2 and 3.3).
2013
Four OTUs representing both mycorrhizal fungal families were identified for samples collected in the year 2013 (Table 3.4). Tulasnellaceae was represented by 3
OTUs, and Ceratobasidiaceae by 1 OTU. Across all plants and treatments, infection of a single root system by 2 to 3 fungal OTUs was observed in 33% of all sampled plants 127
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(Figure 3.4a). The remaining 67% of plants contained only one OTU. A majority of the plants (18 of 19 plants) possessed fungi belonging to a single family, while only one plant had fungi belonging to the two families (Figure 3.4b).
2014
Ceratobasidiaceae and Tulasnellaceae were represented by 13 and 2 OTUs respectively (Table 3.5). Across all plants and land management types, infection of a single root system by 2 to 4 fungal OTUs was observed in 35% of all sampled plants. The remaining 65% of plants contained only one OTU. A majority of the plants (18 of 20 plants) possessed fungi belonging to a single family, while only two plants had fungi belonging to the two families (Figure 3.5a and 3.5b).
2013 and 2014 combined
Twenty OTUs representing both fungal families were identified when data sets for
2013 and 2014 where pooled (Table 3.6). Fungal family Ceratobasidiaceae was represented by 15 OTUs and Tulasnellaceae was represented by 5 OTUs. Across all plants and populations, infection of a single root system by 2 to 4 fungal OTUs was observed in 48% of all sampled plants. The remaining 52% of plants contained only one
OTU. A majority of the plants (36 of 39 plants) possessed fungi belonging to a single family, while only three plants had fungi belonging to the two families (Figure 3.6a and
3.6b).
Mean pairwise sequence distance (pi, π)
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Mean pairwise distance of sequences from Tulasnellaceae were always larger than those for Ceratobasidiaceae, regardless of the year of sampling or by combining the data sets for both years (Table 3.7). Pairwise sequence distance was also measured by treatment per year (Table 3.8). Both fire treatments (PS and PF) had a larger sequence distance in 2013 than the annual hay treatment or the control treatment (πPS = 0.201 ±
0.008 and πPF = 0.295 ± 0.01). This suggests a wider mycorrhizal fungal diversity in the roots of P. praeclara collected in the plots with prescribed fires. On the other hand, the treatments with a large sequence distance for the year 2014 were prescribed spring fire, prescribed fall fire and the control treatment.
Fisher’s exact test
The family Ceratobasidiaceae was represented in roots of P. praeclara from all populations sampled in the year 2013 and 2014. The fungal family Tulasnellaceae was not represented in two of the sampled treatments (PC, PH) in 2013. In the year 2014,
Tulasnellaceae was not observed in roots of one of the sampled treatments (PH). The
Fisher’s exact test revealed that mycorrhizal fungal communities were mostly similar in the year 2013 between treatments, except for the fungal communities in PF and PC (P <
0.05) (Table 3.9). Conversely, the communities observed in 2014 sampling are all significantly different (P < 0.05) (Table 3.10). Using the combined data set, the Fisher’s exact test for count data showed significant differences in all the treatments except between PH and PS (P > 0.05) (Table 3.11).
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Two-way hierarchical ordination
The treatment-wise two-way hierarchical ordination analysis based on the OTUs observed per year of sampling showed clustering between the treatments that were not significantly different according to the Fisher’s exact test for count data (Figures 3.7 and
3.8).
The two-way hierarchical ordination analysis was also run for the combined data set (Figure 3.9). Plots where prescribed fires were applied clustered together along with the annual haying treatment. This suggests that fungal communities were more similar between the fire treatments and the annual haying, and were more different from the control land management treatment.
Nonmetric multidimensional scaling
Nonmetric multidimensional scaling ordination analysis of fungal communities in samples collected in 2013 from the different land management treatments showed that distinction in orchid-associated fungi is most strongly related to sand and pH at PC (no management plot). Clay appeared to have a strong correlation with the mycorrhizal fungal community observed at PH (annual haying treatment). Silt and Na presented correlation with the fungal community observed at PF (fall burn treatment) (Figure 3.10).
In 2014, mycorrhizal fungal community observed in PH was correlated with clay and electrical conductivity measured in May. Mycorrhizal fungal community observed in PF appeared to have a strong correlation with S. Volumetric water content measured in June and silt presented correlation with the fungal community observed in PC. Sand and P presented correlation with the fungal community in PS. Other edaphic variables and 130
Texas Tech University, Pablo Antonio Tovar, December 2015 environmental data presented strong variables but no evident correlation with any of the sampled sites (Figure 3.11).
Discussion
Across the two years of sampling, Platanthera praeclara associated with
Ceratobasidiaceae in each of the four land management treatments. The fungal family
Tulasnellaceae was not present in P. praeclara roots in PC (no management plot) in 2013 and was absent from PH (annual haying treatment) in both sampling years. Mean pairwise sequence distance separated by family (Table 3.7) suggested that due to the narrow distances within nrITS sequences in Ceratobasidiaceae, P. praeclara plants were specific towards its associations with this fungal family. Given the sequence distances within Tulasnellaceae, associations with the Tulasnellaceae were broader. Pi – distances were also calculated between sequences within each of the fungal families represented in each of the four management treatment (Table 3.8). In 2013, distances were much larger for both fire treatments (i.e., spring and fall burn) than distances from the haying and control treatments; the latter two were only composed by sequences belonging to the
Ceratobasidiaceae. In the year 2014, fungi from Tulasnellaceae were not detected in the annual haying treatment (PH), therefore presenting the lowest sequence distance between treatments for the year. These findings suggest that burn treatments might favor characteristics in the vegetation community or in the soil characteristics that allow for a greater microbial diversity within plant roots. Bleho et al. (2015) reported that P. praeclara plants might benefit from burn treatments applied once every 2 to 3 years. 131
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When comparing the fungal communities among treatments, the pairwise Fisher’s test revealed that in the year 2013, only PF presented a significantly different mycorrhizal fungal community in orchid roots as compared to PC, while the rest of the pairwise comparisons were not different (Table 3.9). Conversely, the four land management types presented significant differences in their mycorrhizal fungal communities in the year
2014 (Table 3.10) indicating temporal variation in mycorrhizal preferences of the orchid.
Although there is still high specificity according to the mean pairwise sequence distances, fungal community assemblies within the orchid roots from each treatment showed large variation. In 2013, four OTUs were observed across the four treatments. In the same year, the fungal family Tulasnellaceae presented three OTUs and Ceratobasidiaceae presented on. However, the single Ceratobasidiaceae OTU was observed in each treatment, whereas
Tulasnellaceae occurred only in the two burn treatments. In 2014, 15 OTUs (13 from
Ceratobasidiaceae) were observed across all treatments. These temporal differences suggest that fungal community assemblies in plant roots can shift drastically from one growing season to another, as has been previously recorded in other studies (Twieg et al.,
2007; Drumbell et al., 2011; Cowden and Peterson, 2013; Ercole et al., 2015).
The hierarchical clustering analysis suggested segregation of the mycorrhizal fungal communities by clustering together the communities observed in the prescribed fire treatments, and separating the fungal communities observed in the annual haying and control treatments when the total diversity from the combined data sets was analyzed
(Figure 3.9). However, the Fisher’s test for the combined data set suggested that mycorrhizal fungal community composition of PS and PH did not have significant
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Texas Tech University, Pablo Antonio Tovar, December 2015 differences. These inconclusive patterns implied that perhaps a stronger sampling effort adding more replicates of each treatment should be carried out to make the conclusions more robust.
Land management can affect soil moisture, temperature, carbon, nitrogen, and pH, etc., which are variables that are known to affect fungal communities in grasslands
(Bever et al., 1996). Annual spring fires in tallgrass prairie significantly increase soil temperature and alter soil moisture during the growing season (Knapp and Seastedt,
1986). However, fire treatments are applied to the experimental plots at Pembina Trail plots once every four years, and over one year had already passed by the time plants were sampled for this study in 2013. Regardless, management treatments can have long-lasting effects on edaphic characteristics (Post and Mann, 1990; Murty et al., 2002). A multi- year, long-term study of soil physicochemical profiles across multiple replicates of management treatments can likely improve our understanding of the effects management practices may have on the edaphic characteristics over space and time.
Components of soil texture, which have been associated with alterations in fungal communities previously (Girvan et al., 2003) appeared to be strongly correlated with the mycorrhizal fungal communities in both years 2013 and 2014 according to the nonmetric multidimensional scaling analysis. Phosphorus, pH, nitrates and other variables that have been associated to fungal communities are represented as strong vectors but given that only one replicate plot was sampled to represent each management treatment, strong conclusions cannot be drawn. With the use of a larger number of replicates, soil variables could be correlated to type of land managements although vegetation and soil 133
Texas Tech University, Pablo Antonio Tovar, December 2015 composition also varies in relation to other ecosystem features such as slope, water table, and proximity to edges, etc. Regardless, while the experimental prairie management treatments have been in place for over 20 years at Pembina Trail Preserve, we report the first comparison of the mycorrhizal fungal communities associated with P. praeclara under individual management treatments and over time.
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Ercole, E., Adamo, M., Rodda, M., Gebauer, G., Girlanda, M., & Perotto, S. (2015). Temporal variation in mycorrhizal diversity and carbon and nitrogen stable isotope abundance in the wintergreen meadow orchid Anacamptis morio.New Phytologist, 205(3), 1308-1319.
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Frey, S.D., Knorr, M., Parrent, J.L., & Simpson, R.T. (2004). Chronic nitrogen enrichment affects the structure and function of the soil microbial community in
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temperate hardwood and pine forests. Forest Ecology and Management,196(1), 159-171.
Girvan, M.S., Bullimore, J., Pretty, J.N., Osborn, A.M., & Ball, A.S. (2003). Soil type is the primary determinant of the composition of the total and active bacterial communities in arable soils. Applied and Environmental Microbiology,69(3), 1800-1809.
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Table 3.1 Soil chemical characteristics in each prairie management treatment (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) from where Platanthera praeclara roots were sampled in 2013 for mycorrhizal analyses. OM, organic matter; ENR, estimated nitrogen release; P1, phosphorus 1 (readily available to plants); P2, phosphorus 2 (readily available to plants plus part of the active reserve in the soil); K, potassium; Mg, magnesium; Ca, calcium; Na, sodium; CEC, cation exchange capability; and NO3, nitrate. Please see next page for the remaining table.
Soil characteristics
Prairie OM ENR P1 P2 pH K Mg Ca Na CEC NO3 management (%) (g/ha) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (meq/100g) (uL/L)
PC 2.5 89668 27 39 8.2 167 607 3155 53 21 4 PH 3.1 103118 29 46 8.5 150 852 2582 54 20 5 PF 1.6 70613 40 45 8.3 137 516 1578 52 13 3 PS 2.7 94151 33 39 7.8 168 673 1605 57 14 4
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Table 3.1 Continued Soil chemical characteristics in each prairie management treatment (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) from where Platanthera praeclara roots were sampled in 2013 for mycorrhizal analyses. S, Sulphur; Zn, zinc; Mn, manganese; Fe, iron; Cu, copper; B, boron and soil textural components (sand, silt and clay).
Soil characteristics
Prairie S Zn Mn Fe Cu B Soluble salts Sand Silt Clay management (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (mmhos/cm) (%) (%) (%)
PC 9.6 1.2 11.9 60.7 1.0 0.9 0.4 74.4 10 15.6 PH 7.8 1.6 10.4 53.5 1.2 0.7 0.3 72.4 10 17.6 PF 6.9 0.8 6.4 43.6 0.8 0.9 0.3 74.4 12 13.6 PS 6.8 1.5 13.0 88.8 0.7 0.9 0.4 66.4 20 13.6
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Table 3.2 Soil environmental data and chemical characteristics in each prairie management treatment (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) from where Platanthera praeclara roots were sampled in 2014 for mycorrhizal analyses. VMC, volumetric water content; EC, electrical conductivity; T, temperature, OM, organic matter; ENR, estimated nitrogen release; P1, phosphorus 1 (readily available to plants); P2, phosphorus 2 (readily available to plants plus part of the active reserve in the soil) and K, potassium.
Soil Characteristics
VWC VWC VWC EC EC EC T T T OM ENR P1 P2 K April May June April May June April May June pH Prairie (%) (g/ha) (uL/L) (uL/L) (uL/L) Management (m³/m³) (m³/m³) (m³/m³) (dS/m) (dS/m) (dS/m) (°C) (°C) (°C)
PC 0.49 0.50 0.51 0.23 0.35 0.50 4.0 10.3 17.3 3.8 118810 14 38 6.9 141 PH 0.43 0.48 0.49 0.22 0.39 0.60 1.7 8.8 18.0 4.0 123294 14 75 7.4 151 PF 0.51 0.50 0.50 0.28 0.39 0.54 1.8 10.1 17.4 4.0 124414 15 36 7.0 161 PS 0.44 0.48 0.49 0.20 0.36 0.51 2.6 11.0 17.9 4.4 133381 17 25 6.8 170
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Table 3.2 Soil chemical characteristics in each prairie management treatment (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) from where Platanthera praeclara roots were sampled in 2014 for mycorrhizal analyses. Mg, magnesium; Ca, calcium; Na, sodium; CEC, cation exchange capability; NO3, nitrate; S, Sulphur; Zn, zinc; Mn, manganese; Fe, iron; Cu, copper and B, boron and soil textural components (sand, silt and clay).
Soil characteristics
Prairie Mg Ca Na CEC NO3 S Zn Mn Fe Cu B Soluble salts Sand Silt Clay Management (uL/L) (uL/L) (uL/L) (meq/100g) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (uL/L) (mmhos/cm) (%) (%) (%)
PC 665 2702 54 20 1 4.1 1.3 30 144 1.8 0.8 0.3 74 14 12 PH 913 4407 72 30 1 11 1.6 36 125 2.4 0.5 0.4 72 12 16 PF 698 2653 57 20 1 11 1.5 19 93 1.8 0.7 0.3 74 12 14 PS 732 2300 65 19 1 3.7 1.4 30 128 1.4 0.7 0.4 76 12 12
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Table 3.3. Number of plants sampled and number of samples that yielded useable sequences from roots of Platanthera praeclara in 2013 and 2014. Plant roots were sampled for mycorrhizal diversity studies across four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) at Pembina Trail Preserve in Minnesota, USA.
Samples collected Samples yielding useable sequences
2013 2014 Total 2013 2014 Total
Number of plants 24 20 44 19 20 39 Number of root fragments 81 80 161 54 66 120
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Table 3.4 Number of root sections (i.e. sequences) representing each of the four operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal fungi in each of the four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) in 2013. Values inside the parenthesis denote the number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was observed.
Prairie management treatments OTU type PC PH PF PS Ceratobasidiaceae C1 12(5) 9(4) 11(3) 9(3)
Tulasnellaceae T1 6(2) 3(2) T2 2(1) T3 2(1)
Total 12(5) 9(4) 21(7) 12(4)
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Table 3.5 Number of root sections (i.e. sequences) representing each of the 20 operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal fungi in each of the four prairie management treatment plots (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) in 2014. Values inside the parenthesis denote the number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was observed.
Prairie management treatments OTU type PC PH PF PS Ceratobasidiaceae C1 3(2) 12(4) 13(2) C2 1(1) C3 3(1) C4 1(1) C5 1(1) C6 1(1) 4(3) C7 3(2) C8 2(1) C9 4(1) C10 4(1) C11 2(1) C12 1(1) C13 1(1)
Tulasnellaceae T1 6(4) 1(1) T2 3(1)
Total 9(5) 16(4) 24(5) 17(7)
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Table 3.6 Number of root sections (i.e. sequences) representing each of the 20 operational taxonomic units (OTUs) of Platanthera praeclara mycorrhizal fungi in each of the four prairie management treatment plots (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) using the combined data set (years 2013 and 2014). Values inside the parenthesis denote the number of sequences representing an OTU, while the values outside are the number of plants in which an OTU was observed.
Prairie management treatments OTU type PC PH PF PS Ceratobasidiaceae C1 9(3) 19(3) 20(8) 15(7) C2 4(3) 1(1) 1(1) C3 2(1) C4 2(1) C5 2(2) C6 1(1) C7 1(1) C8 1(1) C9 1(1) C10 3(2) C11 4(1) C12 4(1) C13 1(1) 4(2) C14 1(1) C15 1(1)
Tulasnellaceae T1 1(1) 3(3) T2 3(2) 6(2) 3(2) T3 2(1) T4 3(2) T5 2(1)
Total 29(11) 45(11) 25(8) 21(9)
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Table 3.7 Mean pairwise sequence distances and standard errors (SE) for all observed fungal ITS sequences from roots of Platanthera praeclara sampled from four experimental prairie management treatments in years 2013 and 2014, and for the combined data set (years 2013 and 2014), separated by fungal family Ceratobasidiaceae and Tulasnellaceae. The values presented within the parenthesis are the numbers of individual ITS sequences (n) composing each group.
Data set Fungal Family Pi - Distance SE Tulasnellaceae (23) 0.203 0.009 2013+2014 Ceratobasidiaceae (97) 0.049 0.004
Ceratobasidiaceae 2013 (41) 0.070 0.005 2013 Tulasnellaceae 2013 (13) 0.168 0.007 Ceratobasidiaceae 2014 (56) 0.009 0.002 2014 Tulasnellaceae 2014 (10) 0.257 0.010
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Table 3.8 Mean pairwise sequence distances and standard errors (SE) for all observed fungal ITS sequences from roots of Platanthera praeclara in each of four prairie management treatment plots (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) for years 2013 and 2014 separated by land management treatment. The values presented within the parenthesis are the numbers of individual ITS sequences (n) composing each group.
Data set Prairie management Pi - Distance SE PS (12) 0.201 0.008 PF (21) 0.295 0.010 2013 PH (9) 0.006 0.002 PC (12) 0.007 0.002
PS (17) 0.133 0.006 PF (24) 0.130 0.005 2014 PH (16) 0.038 0.004 PC (9) 0.259 0.010
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Table 3.9 Statistical significance (P values) from the Fisher’s exact test comparing the fungal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) observed in each of the four prairie management treatment plots (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) for 2013.
Prairie management treatments PC PH PF PS PC -
PH 1.00 -
PF < 0.05 0.111 -
PS 0.095 0.228 0.546 -
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Table 3.10 Statistical significance (P values) from the Fisher’s exact test comparing the mycorrhizal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) observed in each of the four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) for 2014.
Prairie management treatments
PC PH PF PS
PC -
PH < 0.05 -
PF < 0.05 < 0.05 -
PS < 0.05 < 0.05 < 0.05 -
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Table 3.11 Statistical significance (P values) from the Fisher’s exact test comparing the fungal communities of Platanthera praeclara based on the abundances of operational taxonomic units (OTUs) observed in each of the four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) for the combined data set (years 2013 and 2014).
Prairie management treatments PC PH PF PS
PC -
PH < 0.05 -
PF < 0.05 < 0.05 -
PS < 0.05 0.142 < 0.05 -
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Figure 3.1 Satellite image of the arrangement of the four prairie management treatment plots (PC, no management; PS, spring burn; PF, fall burn, and PH, annual haying) where mycorrhizal fungi of Platanthera praeclara were sampled in years 2013 and 2014. Experimental plots were located at Pembina Trail Preserve, Polk County, Minnesota, USA. Image taken from Google Earth.
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PC PH 0 0
12 9
PF PS
3
10
11
9
Ceratobasidiaceae
Tulasnellaceae
Figure 3.2 Pie charts showing the distribution of the fungal families Ceratobasidiaceae and Tulasnellaceae observed in roots of Platanthera praeclara sampled from four experimental prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) in the year 2013. Numeric values represent the individual ITS sequences belonging to each fungal family. See below please.
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PC PH 0
3
6 18
PF PS
3 1
19 16
Ceratobasidiaceae
Tulasnellaceae
Figure 3.3 Pie charts showing the distribution of the fungal families Ceratobasidiaceae and Tulasnellaceae observed in roots of Platanthera praeclara sampled from four experimental prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) in the year 2014. Numeric values represent the number of individual ITS sequences belonging to each fungal family.
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(a) 14 12
10 8 6
# of plants of # 4 2 0 1 2 3 # of OTUs
(b) 20 18 16
14 12 10 8 # of plants of # 6 4 2 0 1 2 # of fungal families
Figure 3.4 One to three operational taxonomic units in individual plants of Platanthera praeclara (a), and number of individual plants hosting one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) (b), across four experimental prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) in 2013.
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(a) 14 13 12
10 8 6 5
# of plants of # 4 2 1 1 0 1 2 3 4 # of OTUs
(b) 20
15
10 # of plants of # 5
0 1 2 # of fungal families
Figure 3.5 One to four operational taxonomic units in individual plants of Platanthera praeclara (a), and number of individual plants hosting one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) (b), across four experimental prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) in 2014.
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(a) 25
20
15
10 # of plants of # 5
0 1 2 3 4 # of OTUs
(b) 40 35
30
25 20
15 # of plants of # 10 5 0 1 2 # of fungal families
Figure 3.6 One to four operational taxonomic units in individual plants of Platanthera praeclara (a), and number of individual plants hosting one or both fungal families (i.e., Ceratobasidiaceae and Tulasnellaceae) (b), across four experimental prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment). Data represent samples collected from each experimental treatment over two years (2013 and 2014).
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Figure 3.7 Two-way hierarchical cluster tree and matrix coding based on abundance of four fungal operational taxonomic units (OTUs) within the roots of Platanthera praeclara representing four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment). Data represent samples collected from each experimental treatment in 2013.
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Figure 3.8 Two-way hierarchical cluster tree and matrix coding based on abundance of 15 fungal operational taxonomic units (OTUs) within the roots of Platanthera praeclara representing four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment). Data represent samples collected from each experimental treatment in 2014.
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Figure 3.9 Two-way hierarchical cluster tree and matrix coding based on abundance of 20 fungal operational taxonomic units (OTUs) within the roots of Platanthera praeclara representing four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment). Data represent samples collected from each experimental treatment over two years (2013 and 2014).
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Figure 3.10 Nonmetric multidimensional scaling (NMS) ordination of the fungal communities of Platanthera praeclara in 4 land management treatment plots (PS, spring burn; PF, fall burn; PH, annual haying and PC, no treatment) with correlations to soil variables (clay, silt, sand, S, P1, Na, B and pH) for the year 2013.
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Figure 3.11 Nonmetric multidimensional scaling (NMS) ordination of the fungal communities of Platanthera praeclara representing four prairie management treatments (PS, spring burn; PF, fall burn; PH, annual haying, and PC, no treatment) with correlations to soil physicochemical variables (clay, silt, sand, S, P1, OM and pH) and soil environmental variables (electrical conductivity in June, EC J; electrical conductivity in May, EC M, and volumetric water content in June, VWC J) for the year 2014.
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CHAPTER IV
CONCLUSIONS
The federally threatened orchid Platanthera praeclara was observed to form mycorrhizal associations with fungi from the families Ceratobasidiaceae and
Tulasnellaceae when sampled in two consecutive years (2013 and 2014) across eleven sites representing its natural distribution. Although ITS sequences from
Ceratobasidiaceae were detected more frequently, they also exhibited smaller pairwise sequence distances than ITS sequences from Tulasnellaceae. Short sequence distances suggest that P. praeclara makes more specific associations with very closely related fungi within Ceratobasidiaceae, while making associations with genetically diverse fungi from the Tulasnellaceae. In addition to the pairwise sequence distances, phylogenetic analyses consistently supported the observation that mycorrhizal fungal associates belonging to Ceratobasidiaceae have a rather narrow phylogenetic breadth. Moreover, fungi from Ceratobasidiaceae were present in roots of the sampled plants from each site, while fungi from Tulasnellaceae were absent or weakly represented at some of the sampled sites, suggesting that it is more important for P. praeclara to have more fungal associates belonging to Ceratobasidiaceae. Furthermore, shifts in the mycorrhizal fungal diversity in P. praeclara roots over time showed that although fungal community in the roots can be dynamic, the orchid restricted itself to associating with specific taxa from the fungal family Ceratobasidiaceae.
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Although the composition in the orchid mycorrhizal fungal communities within the roots was not linked to the type of land management, there were correlations with soil variables that have been reported to affect soil fungal community dynamics and biogeographical patterns. The overall mycorrhizal fungal diversity observed within the
Pembina Trail Preserve experimental prairie management treatments presented inconclusive patterns that could explain the observed mycorrhizal fungal communities in
P. praeclara in relation to the land management treatment that was administrated.
However, the two prescribed burn treatments hosted more diversity in terms of OTU richness, while orchid roots from the annual haying treatment lacked Tulasnellaceae symbionts at this site.
It is important to take into account the wide natural distribution of P. praeclara. It is usually expected that orchids with a wider distribution present more general associations towards its mycorrhizae. However, the orchid mycorrhizal fungal diversity observed in this study suggests that orchid taxa with wide distributions can also be specific towards their fungal associates. It is clear that few generalizations about orchid fungal specificity, diversity, or dynamics can be made for a plant family that holds between 20,000 and 35,000 species, and caution is warranted when extrapolating the findings to other orchid species with similar distributions or habits. At the same time, it is possible that terrestrial orchids native to grasslands exhibit similar patterns of mycorrhizal associations, and investigation of such patterns, along with an investigation of ecological drivers that influence the shifts in these associations, is warranted.
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