platanthera chapmanii: culture, population augmentation
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Platanthera chapmanii: culture, population augmentation, and mycorrhizal associations
By
Kirsten Poff, 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 SCIENCE
Approved
Dr. Jyotsna Sharma
Chair of Committee
Dr. Scott Longing
Dr. John Zak
Dr. Mark Sheridan
Dean of the Graduate School
August, 2016
© 2016, Kirsten Poff
Texas Tech University, Kirsten Poff, August 2016
ii
ACKNOWLEDGEMENTS
First I would like to thank my mentor and advisor, Dr. Jyotsna Sharma for all of
her help and support. She has challenged and encouraged me throughout my program and
the duration of this project. Thanks to her, I am light-years ahead of where I was two
years ago. Texas Parks and Wildlife is also gratefully acknowledged for funding portions
of this study.
I also wish to express my gratitude to Dr. John Zak for his enthusiasm and for
encouraging my love of microbes. I also gratefully thank Dr. Scott Longing for his
advice, and constructive comments. I sincerely thank all three committee members for all
the time and energy they have spent on me throughout the duration of my project. I
gratefully acknowledge Dr. Jason Woodward for his encouragement and
recommendations as well. I also acknowledge Dr. Cynthia McKenney and Mr. Russel
Plowman for their support; I now have a passion for teaching, and a much better
understanding of what it is like to teach college level courses. I want to also thank Mr.
Robby Carlson for his time and technological assistance.
I wish to extend a heartfelt thank you to my lab mates for their time and help. I
extend my gratitude to Niraj Rayamajhi and Bianca Walker for teaching me a multitude
of lab techniques. I want to also thank Jaspreet Kaur and Dr. Eeva Terhonen, specifically
for all there help with data analyses. I thank Pablo Tovar for teaching me how to
troubleshoot, none of my sequences would exist without him. I thank all of them for their
friendship. I gratefully acknowledge the efforts of my professors at Texas Tech
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University, I also thank the Plant and Soil Science Department administrative staff and
chair Dr. Eric Hequet, all your advice and wisdom was well received.
To my parents, Leslie and Kevin Poff, and my siblings Kevin Scott Poff, Crista
Poff and Caroline Poff, thank you for all of your support and love. Last, I want to thank
Daniel Smith for his inspiration, love and encouragement, and for coming with me on this
journey. This would not have been possible without all of these people, I am very lucky
and thankful to have them.
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TABLE OF CONTENTS
ACKNOWLEDGEMENTS .................................................................................... ii
LIST OF TABLES ................................................................................................ vii
LIST OF FIGURES ............................................................................................... ix
I. INTRODUCTION AND BACKGROUND ........................................................ 1
Introduction ....................................................................................................... 1
Orchid classification ..................................................................................... 1
Orchid biology .............................................................................................. 2
Orchid Conservation ..................................................................................... 6
Genus Platanthera ......................................................................................... 8
Platanthera chapmanii .................................................................................. 9
Background ..................................................................................................... 10
In vitro seed germination ...................................................................................... 10
Greenhouse culture of plants ................................................................................ 14
Population augmentation ...................................................................................... 16
Mycorrhizal associations ...................................................................................... 19
Techniques for mycorrhizal identity ..................................................................... 22
Summary of Research Gaps .................................................................................. 24
Objectives of the study.................................................................................... 26
Significance of the study ................................................................................. 26
Literature Cited ............................................................................................... 27
II. COLD-MOIST STRATIFICATION IMPROVES GERMINATION IN A
TEMPERATE TERRESTRIAL NORTH AMERICAN ORCHID ...................... 38
Abstract ........................................................................................................... 38
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Introduction ..................................................................................................... 39
Materials and methods .................................................................................... 43
Seed stratification ....................................................................................... 43
Seed plating and germination assessment .................................................. 44
Data analysis ........................................................................................................ 45
Results ............................................................................................................. 46
Seed germination .................................................................................................. 46
Seedling development............................................................................................ 47
Discussion ....................................................................................................... 48
Literature Cited ..................................................................................................... 52
III. PLATANTHERA CHAPMANII: NUTRIENT SUPPLEMENTATION AND
POPULATION AUGMENTATION .................................................................... 63
Abstract ........................................................................................................... 63
Introduction ..................................................................................................... 64
Materials and Methods .................................................................................... 69
Nutrient supplementation ........................................................................... 69
Population augmentation ............................................................................ 70
Results ............................................................................................................. 72
Nutrient supplementation ...................................................................................... 72
Population augmentation ............................................................................ 73
Discussion ....................................................................................................... 73
Literature Cited ..................................................................................................... 77
IV. DIVERSITY OF MYCORRHIZAE FORMING TULASNELLACEAE IN A
TEMPERATE TERRESTRIAL ORCHID IN EX SITU AND IN SITU
ENVIRONMENTS ............................................................................................... 83
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Abstract ................................................................................................................. 83
Introduction ........................................................................................................... 85
Materials and methods .......................................................................................... 90
Literature cited .................................................................................................... 105
V. CONCLUSIONS ............................................................................................ 127
Literature Cited ............................................................................................. 130
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LIST OF TABLES
2.1
.
Effect of cold-moist stratification (0, 8, or 12 weeks) on seed germination was
experimentally tested in Platanthera chapmanii. An Analysis of Variance
(ANOVA) was conducted (a); Germination of seeds was categorized as
Stage 0 (no further development), Stage 1 (germination; rhizoid development),
or Stage 2 (leaf primordium development). Mean number of seeds that were
plated in an experimental unit, mean number of viable seeds, and mean percent
viability are presented. Mean germination percentages were calculated by
using total number of seeds and number of viable seeds separately. Means
followed by the same letter in each column were statistically similar based on
Fisher’s Least Significant Difference (LSD) test…................................................55
2.2
.
Effect of cold-moist stratification (0, 8, and 12 weeks) on seedling
development after germination and rhizoid development (Stage 1) was
experimentally tested in Platanthera chapmanii. An Analysis of Variance
(ANOVA) was conducted (a); Seed development was categorized as Stage 2
(leaf primordium development) or Stage 3 (root development). Mean number
of seedlings that were categorized as Stage 2 or Stage 3 after a 5 month
exposure to 40-watt florescent bulbs set at a photoperiod of 12 hours.……........57
3.1
.
Effect of nutrient supplementation (0.0x, 0.25x, 0.5x) on Platanthera
chapmanii above ground plant height after 14 weeks of treatment applied
every two weeks. Results of An Analysis of Variance (ANOVA) was
conducted………….......................................................................................……79
4.1 Representative sequence of each of the 18 fungal nrITS-based operational
taxonomic units (OTUs) identified within the roots of Platanthera chapmanii
plants cultured in vitro/greenhouse and those occurring naturally. Each culture
condition was sampled three to four times between 2012 and 2015. The first
letter of an OTU name represents the fungal family to which the OTU
belongs: T, Tulasnellaceae; C, Ceratobasidiaceae. Values in parentheses are
the total number of plants in which a specific OTU was documented. …..…....109
4.2
.
Number of root sections (i.e. sequences) representing each of the 18 fungal
nrITS based operational taxonomic units (OTUs) identified within the roots
of Platanthera chapmanii plants cultured in vitro/greenhouse and those
occurring naturally. Each culture condition was sampled three to four times
between 2012 and 2015. The first letter of an OTU name represents the
fungal family to which the OTU belongs: T, Tulasnellaceae; C,
Ceratobasidiaceae. Values in parentheses are the total number of plants in
which a specific OTU was documented……………………………....………..115
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4.3. Mean pairwise fungal internal transcribed spacer (nrITS) sequence distances
(π ± SE; Nei and Kumar 2000), estimated based on Kimura’s 2-parameter
model, within the fungal family Tulasnellaceae identified in the roots of
Platanthera chapmanii plants that were either cultured in lab / greenhouse
conditions (GF12, GF14, GSu15) or were obtained from a naturally
occurring population (NF12, NF14, NSp15, NSu15).…………………………117
4.4. Mean pairwise distances among fungal nrITS sequences based on Kimura’s
2-parameter model were calculated for fungal communities identified
within the roots of Platanthera chapmanii. Roots of plants raised in vitro
and cultured in greenhouse, and from plants occurring naturally were
sampled. All other mean pairwise distances, except for Platanthera
praeclara (Tovar 2015) and Nervilia nipponica (Nomura et al. 2013) were
calculated by Pandey et al. (2013).…......…………………………………..….121
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LIST OF FIGURES
1.1. A photograph of Mesic to wet pine habitat of Platanthera chapmanii at Watson
Native Plant Preserve, Tyler County, Texas (2014)…… ……….………..……..…36
1.2. A photograph of Platanthera chapmanii during anthesis at Watson Native
Plant Preserve, Tyler County, Texas. Photograph by Jyotsna Sharma (2013)……..36
1.3. A map of the southeast United States showing the geographic range of
Platanthera chapmanii. Areas within Texas, Florida, and Georgia where the
species occurs naturally are shaded in blue blue……………………...……………37
1.4. A photograph of a cross section of Platanthera chapmanii root tissue showing
coils of hyphae, pelotons, within the root cells documented in November 2014.
Scale bar represents 100 µm………………………………………..………………37
2.1. Seed germination and plant development in Platanthera chapmanii
was recorded by using four categories: Stage 0 (no germination), Stage 1
(germination; rhizoid development), Stage 2 (leaf primordium development),
and Stage 3 (root development) ...……………………………. …………..……….58
2.2. Proportion of Stage 2 seedlings of Platanthera chapmanii that reached the
developmental Stage 3 after exposure to light. Duration of exposure to light
(1 to 5 months under 40-watt white florescent bulbs) influenced plant
development to Stage 3. Pre-germination stratification of seeds for 0, 8, or 12
weeks did not influence development from Stage 2 to Stage 3, thus the means
were pooled across the three stratification treatments. Means followed by the
same letter were statistically similar based on Fisher’s Least Significant
Difference (LSD) test…………………….…………..…………………………..…58
3.1 Three photographs of Platanthera chapmanii individuals after planting in 15
cm containers during nutrient supplementation. From left to right; one
individual, one replicate, row of trays…………………………..……….…………80
3.2 A photograph of a typical Platanthera chapmanii individual after being
planted into greenhouse medium….…………..………………………..…………..80
3.3 Two photographs of Platanthera chapmanii individuals. A greenhouse
cultured Platanthera chapmanii individual (a) shown beside a naturally
occurring Platanthera chapmanii individual (b) before planting in native
habitat in southeast Texas during fall 2014……………..……......................……...81
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3.4 A photograph of a fall 2014 Platanthera chapmanii plot with both greenhouse
cultured and native plants relocated in southeast Texas. An arrow is pointing
to one individual………………………..………………...………………………..81
3.5 A photograph of a typical Platanthera chapmanii individual taken directly
out of a culture vessel prior to planting in one of the three locations in
southeast Texas in the spring 2015……...……..……………………...……..……..82
3.6 A photograph of one of the three plots of Platanthera chapmanii individuals
taken directly out of sterile culture and planted in the spring 2015. All
individuals were covered with sphagnum peat moss..……..…………………….....82
4.1 A photograph of root samples collected from in vitro propagated and
greenhouse cultured Platanthera chapmanii individuals before processing
for molecular analysis. ……………………………………………………………119
4.2 A photograph of a cross section of a root of Platanthera chapmanii showing
mycorrhizal hyphal coils, i.e. pelotons, within the cortical cells………...………..120
4.3 Photographic documentation of moniliod cells and fungal hyphae isolated on
potato dextrose agar (PDA). The mycorrhizal fungus was cultured from
roots of Platanthera chapmanii…………………………..……………………….120
chapmanii……………………………………………………………………...145
4.4 Sample-based incidence data, individual-based abundance data and observed
methods were used to construct cumulative, rarefied fungal operational
taxonomic unit (OTU) diversity curves for Platanthera chapmanii
extrapolated to 500 sequences. Operational taxonomic units were built using
122 mycorrhizal fungal sequences and 18 OTUs derived from the roots of
plants that were either cultured in ex situ conditions or were obtained from a
naturally occurring population between 2012 and 2015.…………………....…….121
4.5 A principal component analysis (PCA) scatterplot. Each of the circles
represent one of seven treatments (NF12, NF14, NSp15, NSu15, GF12, GF14,
GSu15) used to obtain mycorrhizal OTUs from the roots of Platanthera
chapmanii plants that were either cultured in lab/greenhouse conditions (GF12,
GF14, GSu15) or were obtained from a naturally occurring population (NF12,
NF14, Nsp15, NSu15) between 2012 and 2015. The PCA shows PC1 and PC2
accounting for 60% of variation in the data. ......……………………..………..….122
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4.6 A maximum likelihood tree of the fungal family Tulasnellaceae built with
operational taxonomic units (OTUs) of nrITS sequences observed in
Platanthera chapmanii roots that were either cultured in lab/greenhouse
conditions (green), obtained from a naturally occurring population (red), or
present in both environments (blue) and orchid mycorrhizal OTUs from
previous publications. The tree was rooted with midpoint method. Bootstrap
values ≤50 were omitted. The tree was built using 1000 bootstrap replicates.
Of the nodes that have two values, the second values are Bayesian probability
values from a Bayesian tree built using 1 million generations…………........…...124
4.7 A maximum likelihood tree of the fungal family Ceratobasidiaceae built with
operational taxonomic units (OTU) clustered using fungal nrITS sequences
observed in Platanthera chapmanii root obtained from a naturally occurring
population (C1) and other orchid mycorrhizal OTUs previously published.
The tree was rooted with a species of Sistotrema. Bootstrap values ≤50 were
omitted. The tree was built using 1000 bootstrap replicates. Of the nodes that
have two values, the second values are Bayesian probability values from a
Bayesian tree built using 1 million generation………………………..…..………126
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CHAPTER I
INTRODUCTION AND BACKGROUND
Introduction
Orchid classification
Orchids belong to the phylum Angiospermae. There are two classes within this
phylum of flowering plants that are important to recognize: 1) monocotyledon (monocot)
and 2) dicotyledon (dicot). Normally, dicot species have a vascular cambium, which is a
tissue system that is responsible for forming woody structures (i.e. bark) on the outside of
a stem and soft tissues on the inside. This allows dicotyledon plants to continue to grow
in diameter. Dicotyledon makes up the larger of the two classes of angiosperms with 267
families that can be divided into 19 suborders (Dressler 1981). Conversely, monocots do
not contain a vascular cambium, meaning their stem diameter growth is usually limited to
one growing season. Because of this, the growth of monocots is somewhat limited. There
is a diversity of forms in which monocots have evolved, many of which are exhibited
within the family Orchidaceae.
Orchidaceae is one of the largest families of flowering plants on earth (Dressler
1981). It has been estimated that there are more than 24,500 species of orchid (Dressler
2005). This number makes up about 10% of all angiosperms (Dressler 2005). Many
orchid species occur in the tropics however species of orchid occur on every continent
including some Antarctic islands (Roberts and Dixon 2003, Clements and Jones 2007),
which gives support to the theory that Orchidaceae originated before the complete
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separation of Pangea approximately 100 mya (Janssen and Bremer 2004, Smith and Read
2008). Because of their wide distribution and habitat range, they are considered by some
to be the most evolved members of the plant kingdom.
The family Orchidaceae is divided into five subfamilies: Apostasiodeae,
Cypripedioideae, Vanillioideae, Orchidioideae, and Epidendrioideae. Epidendrioideae is
considered the largest and most advanced group of orchid species (Singer et al. 2008,
Smith and Read 2008, Royal Botanical Gardens Kew, webpage accessed September
2015). The orchid family may also be divided into epiphytic and terrestrial species;
epiphytic species making up 73% of Orchidaceae and terrestrial making up 27% (Roberts
and Dixon 2008). Unlike epiphytes that anchor themselves on the surfaces of other
plants, terrestrial orchid species establish themselves and grow in the ground, procuring
nutrients from soil (Rassmussen 1995).
Orchid biology
There are some general characteristics that a monocot may exhibit for it to be
considered an orchid. One of the petals of each orchid flower is typically modified to
form a labellum (or 'lip'). Another trait that most orchid flowers exhibit is resupination.
Resupination refers to the rotation of the pedicel when the floral buds are developing,
which leads to the labellum being positioned lower-most when the flower opens (Dressler
1981). Further, almost all orchid species have only one stamen although some species
have two or three. The stamen, instead of being arranged in the middle of the flower
which is common in angiosperms, is arranged on one side of the flower. Secondly, the
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stamen and the pistil are either completely united or at least partially united. This
combination of pistil and stamen is referred to as the column (Roberts and Dixon 2008).
The pollen is usually grouped in two large masses termed pollinia. The rostellum, or a
beaklike modification of the stigma, holds the pollinia and separates the stamen and
gynoecium. This separation minimizes or eliminates self-fertilization. All orchid species
produce microscopic, rudimentary seeds that lack endosperm. Seeds range in size from
200 to1,700 µm, and weigh between 0.3-14 µg. A single capsule sometimes contains
thousands of seeds depending on the orchid species (Smith and Read 2008).
Pollinators of orchid species encompass a variety of Animalia from insects to birds
(Johnson, 1995). This is partially due to the variety of habitats in which orchid
individuals occur combined with the diversity of floral morphologies associated with the
Orchidaceae (Pijl and Dodson 1966). Sexual reproduction is often necessary for the
persistence of orchid species and typically involves an insect vector to carry the pollinia
from one flower to another (Nilsson 1992).
Roots of many orchid taxa share a common characteristic, which is the presence of a
velamen. The velamen is the epidermal layer that is usually spongy and white and may
contain multiple layers of cells (Dressler 1981). The velamen absorbs water, a trait that is
especially favorable for the epiphytic taxa. However, the roots of terrestrial species may
also have a multilayered velamen (Benzing et al. 1982).
It is estimated that 92% of all terrestrial plant species acquire nutrients from some
form of fungal symbiont (Tendersoo et al. 2010). The two general types of mycorrhizae
are endomycorrhizae and ectomycorrhizae. Ectomycorrhizae are characterized by fungal
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growth on the outside of the root cells whereas endomycorrhizae are characterized by
intracellular growth. Ectomycorrhizae are formed commonly by tree species, whereas a
majority of herbaceous species form endomycorrhizae. Approximately 6,000 fungal
species are currently known to form ectomycorrhizae (Barton and Northup 2011).
Ectomycorrhizae do not exhibit intracellular colonization within the roots of the host
plant, but form hyphal networks growing in-between the epidermal and cortical cells.
Additionally, a fungal mantle is formed on the exterior of the root (Smith and Read
2008). Members of this type of mycorrhizae may belong to the fungal phyla
Basidiomycota or Ascomycota (Smith and Read 2008). Arbuscular mycorrhizae (AM),
which are a major group within endomycorrhizae, are the most common type of
mycorrhizae occurring in tracheophytes, pteridophytes, and bryophytes (Smith and Read
2008). Arbuscular mycorrhizae are currently estimated to encompass 120 fungal species
(Barton and Northup 2011). The fungi involved belong to the ancient phylum
Glomeromycota (Smith and Read 2008), which is likely to have originated over 400 mya
(Helgason and Fitter 2005, Schubler et al. 2011).
Arbuscular mycorrhizae are characterized by intercellular growth structures known
as arbuscules and vesicles, and extracellular chlamydospores. Another morphotype of
endomycorrhizae, ericoid mycorrhizae, involve ascomycetes that form hyphal coils in the
root hairs of some Ericales and some bryophytes (Barton and Northup, 2011). Several
mycorrhizal relationships have a mix of traits that are similar to those of
endomycorrhizae and ectomycorrhizae, examples include ectendomycorrhizae, arbutoid
mycorrhizae, and monotropoid mycorrhizae (Smith and Read 2008).
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Orchid mycorrhizae represent yet another morphology of endomycorrhizae. Orchid
mycorrhizae form large hyphal coils (i.e. pelotons) inside the cortical cells of orchid
roots. Often the pelotons occupy a large volume of the cell and are very large when
compared to other types of endomycorrhizal growth (Smith and Read 2008).
At maturity, most of Orchidaceae are at least partially photosynthetic, however,
holomycoheterotrophy is also present in the Orchidaceae. More than 100 orchid species
are achlorophyllous as adults (Dearnaley 2007). The majority of known orchid
mycorrhizae belong to the phylum Basidiomycota. The mycobiont forms pelotons within
the cortical cells of the orchid root. These structures are very large when compared to
other types of endomycorrhizal growth (Smith and Read 2008). In other types of
mycorrhizae, the plant provides carbohydrates for the fungus; however, during the early
stages in the life of an orchid, orchidaceous fungus supplies carbohydrates to the plant
(Smith and Read, 2008). Depending on the way an adult orchid receives carbon, they
may be divided into one of three groups. The three groups include: fully
mycoheterotrophic species, fully autotrophic species and mixotrophic species (Dearnaley
et.al 2012). Of the Basidiomycetes orchid species form symbioses with, most are of the
fungal families Ceratobasidiaceae, Sebacinaceae, and Tulasnellaceae (Otero et al. 2002).
Germination of orchid seeds is fairly complex. While the majority of mycorrhizal
relationships between angiosperms and fungi are established after the roots have
developed, orchid seed germination differs in that mycotrophy is often essential for seed
germination and early development (Rasmussen 1995). This is due to the microscopic,
rudimentary embryos in orchid seeds. The undifferentiated embryo contains concentrated
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lipid and protein bodies and very rarely starch grains and glucoprotein bodies
(Rasmussen 1995). These concentrated reserves may be difficult for the embryonic cells
to metabolize, and gluconeogenesis typically occurs only after embryo cells develop a
connection with suitable fungi (Rasmussen 1995, Cribb et al. 2003, Smith and Read
2008). During germination, epidermal cells of a seed lengthen into outgrowths called
rhizoids. Mycorrhizal fungi may enter a seed through these filamentous outgrowths and
form hyphal coils (Rasmussen 1995). The fungi then translocate carbon into the
developing protocorm (i.e. leafless orchid seedling) allowing for differentiation (Smith
and Read 2008). In some cases, even when an orchid seed is in symbiosis with a fungal
species that would normally be compatible, plant development may not be successful
(Dressler 1981). Unlike other angiosperm seedlings, orchid seedlings do not produce a
radicle (i.e. embryonic root). This is because the basal end of the seedling develops
histological features specialized for mycotrophy (Rasmussen 1995).
Orchid Conservation
As stated previously, the Orchidaceae is both a large and widely distributed family.
However, most species of orchid are rare and threatened with extinction (Cribb et al.
2003). All orchid species are protected under the Convention on International Trade of
Endangered Species of Wild Fauna and Flora (CITES).
Globally, habitat loss and degradation is a large threat to Orchidaceae. However,
relying solely on the conservation of rare plant habitat is not feasible to mitigate the loss
of biodiversity the earth is experiencing (Swarts and Dixon 2009). Climate change also
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poses a large threat to many orchid species (Liu et al. 2010). For example, in the Guangxi
Province in China, precipitation is expected to increase and soil moisture levels are
expected to decrease (Liu et al. 2010). This decrease in soil moisture is likely to
negatively affect orchid populations, whereby the species with small populations and
narrow distributions will be most vulnerable to changes (Liu et al. 2010). The study by
Liu et al. 2010 further describes how changes in temperature may drive some orchid
populations to become extinct. The majority of orchid species (72%) in the Yachang
Reserve have populations that occur very close to, or on, mountain tops. If the
temperature in these areas rises, mountain top populations may not survive (Liu et al.
2010).
Orchid wild-collection and illegal trade is another threat that leads to the decline and
extinction of many orchid species (Cribb et al. 2003). Collection is species dependent, but
has led to the decline of many orchid taxa. For example, Cypripedium calceolus was
historically occurring in several countries, but because of wild-collection, it became one
of the rarest plants in the British Isles by the early 2000s (Cribb et al. 2003).
Cruz-Fernandez et al. (2010) suggested that natural ecosystem processes, such as
self-thinning, are intimately related to the persistence of some orchid genera. In this
particular study, they reported absence of a correlation between orchid species richness or
abundance and timber extraction. However, they also reported a positive correlation
between abundance of orchid taxa belonging to the genus Malaxis and abundance of
standing dead trees (Cruz-Fernandez 2010). This provides evidence that some orchid
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species may require natural disturbances such as self-thinning to persist (Coates et al.
2006, Cruz-Fernandez 2010).
Generally, quantifying suitable habitat is an important step for species preservation.
This can be done by developing species-specific modular regression models of areas
using parameters such as, type of vegetation in the area, percent canopy, and soil
moisture and nutrient content. Suitable habitat quantification has been done for some
orchid species, including the federally threatened species Platanthera praeclara and
Isotria medeoloides (Sperduto and Congalton 1996, Wolken et al. 2001). For many
species it is difficult for researchers to assign levels of extinction-threat because sufficient
field data do not exist (Cribb et al. 2003).
Genus Platanthera
A notable genus within the family Orchidaceae, Platanthera, belongs to the
subfamily Orchidoideae, and subtribe Orchidinae (Dressler and Dodson 1960, Efimov
2011). The genus consists of about 200 terrestrial species that are distributed over parts of
North America, Europe, Asia, and North Africa (World Checklist of Selected Plant
Families, Royal Botanical Gardens Kew, webpage accessed October 2015). Most species
in the genus are terrestrial herbs, although a few are humus epiphytes that grow near the
ground (Efimov 2011). Habitats and ecosystems in which Platanthera species occur
include forest, open tundra, and open mesic to wet grasslands. A few species also occur
in the tropical montane rainforests of Borneo, but most inhabit the temperate zone in the
northern hemisphere (Hapeman and Inoue 1997, Efimov 2011). Floral features of this
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genus include wide anthers and can include fringed labella. Flowers may be orange,
yellow, green, white, or purple (Hapeman and Inoue 1997, Efimov 2011).
Platanthera chapmanii
A species within the genus Platanthera, Platanthera chapmanii (Fig. 1) was first
described in 1903 by J.K. Small (Small 1903) and can be distinguished from the similar
looking and closely related Platanthera cristata and Platanthera ciliaris by floral
morphology. The mouth of the nectar spur of P. chapmanii is circular, whereas the
opening of P. cristata and P. ciliaris are more triangular (the nectar spur of P. ciliaris is
also longer). The lobes protruding from the rostellum can be curved in P. chapmanii,
whereas the lobes of P. cristata and P. ciliaris are only slightly curved (Royal Botanical
Gardens Kew, webpage accessed October 2015). These are important distinctions
because P. cristata, P. ciliaris, and P. chapmanii have overlapping geographic
distributions (Liggio and Liggio 1999). Like its close relatives, P. chapmanii flowers in
July to August producing >60 orange flowers on each inflorescence (Fig. 2, Liggio and
Liggio 1999).
Platanthera chapmanii occurs in mesic and wet pine flatwoods, barrens, and
savannas in sandy loam soils in northern Florida, southern Georgia, and southeast Texas
(Fig. 3, Fig. 4). In Georgia the taxon has been reported recently in Camden, Charlton and
Brantley counties although reported population sizes are usually small (i.e. less than ten
individuals) (Richards and Sharma 2014). In Florida, P. chapmanii has been reported
previously in Baker, Clay, Columbia, Duval, Franklin, Jefferson, Liberty, Marion,
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Taylor, Union, and Wakulla counties (Wunderlin and Hansen 2008). Although more
recently it was reported to be restricted to the Apalachicola and Osceola National forests
(Brown 2004). In Texas, the taxon has been historically reported in Tyler, Hardin,
Orange, and Jefferson counties although recent observations have not been made in
Jefferson County (J. Sharma personal observation). One location in Tyler County is host
to the largest known population that hosts up to 260 reproductive individuals (J. Sharma
personal observation). However, the species distribution and habitat range is not
completely known, newly documented populations of the taxon have been recently
reported (Richards and Sharma 2014).
Biology and ecology of P. chapmanii is poorly understood, and there is a lack of
empirically derived protocols for its propagation and reintroduction. A review of
literature is presented below to support the identification of research gaps with respect to
propagation, culture, population augmentation, and the diversity of mycorrhizal fungi
associated with this taxon. Subsequently, the research objectives and hypotheses
associated with each of these subjects are described
Background
In vitro seed germination
Most of what is known about germination and development of orchid seeds has been
developed through in vitro studies (Arditti 1967, Rasmussen and Whigham 1993). Seed
dormancy and other pre-germination requirements are known to occur in seeds of
temperate terrestrial orchid species (Sharma et al. 2003a). Overcoming seed dormancy in
Texas Tech University, Kirsten Poff, August 2016
11
orchid species can, thus, be necessary to obtain germination in vitro (Lauzer et al. 2007).
This is especially true for terrestrial individuals which may have a complex dormancy
pattern (Johansen and Rasmussen 1992, Lauzer et al. 2007). For example, seeds of
Epipactis palustris require a combination of scarification of the testa, an initial incubation
for several weeks at 27°C, followed by cold stratification for 8-12 weeks to initiate
germination. Without these pretreatments germination rate was poor, but once
implemented, germination was increased to 50% (Rasmussen 1992). These complex
patterns of dormancy are likely present in temperate orchid seeds because of the
environmental conditions the seeds are exposed to. The seeds often experience cold and
moist conditions along with some environmental weathering before experiencing a
warming period when winter turns to spring and summer. Accordingly, to germinate
seeds in vitro, these conditions must also be met, although species often vary in their
response to pre-germination treatments.
One method to apply cold-moist stratification to the microscopic orchid seeds is by
first surface sterilizing seeds, placing them in sterile vials containing sterile water and
incubating in the dark at 4-5°C (Zetter et al. 2001, Sharma et al. 2003a). In some cases,
stratification is performed by placing seeds directly onto germination medium then
incubating in the dark at 4-5°C (Richards and Sharma 2014). Period of stratification can
also have an effect on germination success. Species require variable periods of
stratification. This is the case with Platanthera praeclara as well as Platanthera
leucophaea (Zetter et al. 2001, Sharma et al. 2003a). However, not all Platanthera
species require a stratification period to germinate. Platanthera integra, a species native
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12
to the southeastern United States apparently does not have this pre-germination
requirement (Zettler et al. 2000). Although stratification was not tested for P. integra, a
prolonged scarification treatment was recorded to increased germination percentage
(Zettler et al. 2000). Germination without stratification may be possible because of the
ecological requirements of species distributed in climates that do not experience
excessively low minima during the dormant season.
Although germination in the absence of mycorrhizal fungi has not been documented
in nature, it is possible to asymbiotically germinate orchid seeds in vitro by supplying
exogenous sugars and nutrients (Smith 1973, Rasmussen 1995, Lo et al. 2004, Smith and
Read 2008, Godo et al. 2010). Germination of orchid seeds on culture medium containing
salts and sucrose was first recorded in the 1920s (Knudson 1922). By 1967, in vitro
protocols had been established for germinating seeds of some orchid species
asymbiotically in sterile conditions (Arditti 1967). It is now known that orchid species
can vary in their response to the composition of the media used for in vitro germination
requiring empirical testing to identify the most effective germination conditions for each
taxon (Arditti 1967, Olivia and Arditti 1984, Rasmussen and Whigham 1993, Stewart and
Kane 2006). While many orchid species are successfully propagated via this method,
there is still a lack of information concerning germination and development of a
multitude of orchid taxa, especially those that are native to temperate climates (Arditti et
al. 1981, Rasmussen 1995).
Photoperiod is also known to affect orchid seed germination (Stewart and Kane
2006). It is common for seeds to be incubated in the dark during the initial stages of
Texas Tech University, Kirsten Poff, August 2016
13
germination experiments to simulate natural conditions whereby the seeds typically
germinate below the soil surface (Zettler 1994, Sharma et al. 2003a, Stewart and Kane
2006). Occasionally, seeds are exposed to a short photoperiod prior to placing them under
dark conditions. Platanthera integrilabia seeds were reported to germinate best when
exposed to one week of 16 hr photoperiod followed by incubation in the dark
continuously (Zettler 1994). After a leaf primordium begins to develop, protocorms may
be exposed to a light/dark cycle to encourage further development. The terrestrial species
Habenaria macroceratitis produced the highest number of tubers per individual plant
when exposed to a photoperiod of 8 hrs (Stewart and Kane 2006). Similar results were
reported in Calopogon tuberosus var. tuberosus, whereby highest germination was
recorded when seeds were exposed to a photoperiod of 8 hrs (Kauth et al. 2008).
Orchid seeds often exhibit a preference for an optimum germination temperature or
temperature range (Rasmussen et al. 1990). Seeds of Dactylorhiza majalis were reported
to have an optimum germination temperature between 23 and 25°C (Rasmussen et al.
1990). In one case, seeds of Dactylorhiza majalis were more sensitive to temperatures
above their optimum range than below. Dactylorhiza majalis seeds had a higher
germination percentage when exposed to temperatures below 23°C than above 25°C
(Rasmussen et al. 1990).
Platanthera chapmanii has been propagated successfully after a cold-moist
stratification period of about 12 weeks. However, whether a stratification treatment is
necessary or beneficial for inducing germination has not been empirically tested
(Richards and Sharma 2014).
Texas Tech University, Kirsten Poff, August 2016
14
Greenhouse culture of plants
Majority of the plant species cultured in vitro require an acclimatization process to
ensure plant survival ex vitro (Hazarika 2003, Deb and Temjensangba 2006). When
transitioning orchid seedlings from sterile culture conditions into a greenhouse, an
acclimation procedure is often required (McKendrick 2000). It is common for some
plants to perish during the transfer from aseptic in vitro conditions to a greenhouse setting
(Preece and Sutter 1991, Deb and Imchen 2010). This is due to multiple factors.
Humidity is typically lower in the immediate vicinity of individual plants in a greenhouse
while the intensity of light can be higher or lower (Preece and Sutter 1991). There is also
the unavoidable stress of a septic environment (Preece and Sutter 1991). If in vitro
propagated plants are not acclimated appropriately to the greenhouse environment, they
may show signs of stress including wilting, tip necrosis, and death (Preece and Sutter
1991). An orchid plantlet may require several weeks or months to acclimate to
greenhouse conditions (McKendrick 2000, Deb and Temjensangba 2006). Duration of
acclimatization is implicated in the long-term survival of individual plants (Zeng et al.
2012).
The terrestrial orchid Malaxis khasina is reported to be first acclimatized in vitro for
8-10 weeks before transferring to the greenhouse (Deb and Temjensangba 2006). The in
vitro acclimatization may involve a medium transfer from agar-based medium to a mix of
agar and other sterilized media such as coconut husk and forest litter (1:1:1 ratio) (Deb
and Temjensangba 2006). A similar technique involving alternative substrates has been
used for several members of the genera Arachnis and Cleisostoma (Deb and Imchen
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15
2010). After this initial period, plant survival in greenhouse conditions increased (Deb
and Temjensangba 2006, Deb and Imchen 2010).
According to a review by Hazarika (2003), acclimatization of plantlets can be
expensive by constituting up to 60% of production costs, and is time consuming. For
these reasons, sometimes plants are not acclimatized before greenhouse planting.
Additionally, acclimatization may not be equally necessary for every species (M.
Richards pers. comm.).
The medium used for greenhouse culture can be species-specific. Some epiphytic
taxa, such as the medically important Dendrobium tosaense, have been successfully
transferred from sterile agar medium to unsterilized sphagnum moss or tree fern (Lo et al.
2004). Terrestrial species require a weightier medium, although medium texture should
mimic the soil in a taxon's native habitat. Orchid species such as P. chapmanii that occur
in bog habitats with sandy soils typically require a medium that is well drained.
Platanthera chapmanii has been successfully cultured in a medium composed of
builder’s sand, peat moss, milled sphagnum moss, and fine tree fern fiber (Richards and
Sharma, 2014).
Some terrestrial orchid species can be sensitive to conventional fertilizers. Even
when exposed to low concentrations of inorganic (Nitrochalk (England), superphosphate,
magnesium sulphate, potassium sulphate) and organic (hoof and horn, bonemeal and
urea) fertilizers, detrimental effects have been measured (Silvertown et al. 1994). For
example, a significant decrease in flowering was reported in Orchis morio when exposed
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16
to what were considered low concentrations for agriculture (22-88 kg ha-1 N) (Silvertown
et al.1994).
Platanthera chapmanii has been grown in greenhouse conditions (Richards and
Sharma 2014). When in vitro cultured plants were transferred to a greenhouse at the
Atlanta Botanical Garden (ABG), special acclimatization steps or conditions were not
required (M. Richards, per. comm.). While plants of the taxon can be grown in a medium
containing builder’s sand, peat moss, milled sphagnum moss, and fine tree fern fiber
under greenhouse conditions where they produce flowers and capsules subsequently
(Richards and Sharma, 2014), it is not known how sensitive individuals of P. chapmanii
are to fertilization when cultured in a greenhouse. Platanthera chapmanii has been grown
without added nutrients while watered only with dechlorinated water (M. Richards, pers.
comm.). Further, staff at ABG have at times attempted to treat P. chapmanii with a
diluted solution of fertilizer, but whether this nutrient supplementation encouraged or
hindered growth was not documented quantitatively (M. Richards pers. comm.).
Population augmentation
Degradation of rare plant habitat continues largely because of changes in land use
practices and resource utilization (Rochefort 2000). Sphagnum is one example of a
keystone genus in rare plant habitats that is wild-harvested for horticultural peat and fuel
peat (Rochefort 2000). However, conservation of existing rare plant habitat alone is not
sufficient to preserve biodiversity (Rochefort 2000, Cribb et al. 2003). Because such a
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17
disproportional number of orchid species have populations that are in decline, special
emphasis is required for this family of angiosperms (Cibb et al. 2003).
One aspect of restoration ecology, population augmentation, is a method to increase
the size of a population (Batty et al. 2006, Decruse et al. 2013, Richards and Sharma
2014). Orchid plants grown from seeds in vitro and subsequently transplanted in native
habitats is reported as a successful approach for Eulophia cullenii and for Platanthera
chapmanii (Decruse et al. 2013, Richards and Sharma 2014). Another terrestrial species
that was reintroduced successfully in its native habitat is Paphiopedilum wardii (Zeng et
al. 2012). Plants grown from seed were acclimatized and reintroduced into native habitat
in Gaoligong Mountain in Yunnan, new populations in areas where prior documentation
was not recorded were established in Yangchun and Guangzhou in Guangdong (Zeng et
al. 2012). The transplanted plants exhibited survival rates of approximately 50% or
higher and persisted after two years (Zeng et al. 2012). The North American species
Spiranthes brevilabris has had similar success when transplanted into natural habitats
subsequent to in vitro culture (Stewart 2003). Transplant success, however, is not equal
across various taxa. For example, of the 165 Spiranthes brevilabris plants that were
transplanted in the 2003 study, only 17 initiated anthesis after 6 months even though all
165 had survived the first month in the natural habitat (Stewart et al. 2003).
Clonal propagation of members of Orchidaceae may also be carried out for the
purpose of restoration of natural populations (Martin 2003). When propagated clonally
and introduced into the natural habitat, the endangered Ipsea malabarica was
documented with a high survival and flowering rate (Martin 2003). All 50 of the
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18
individuals transplanted survived and initiated anthesis normally (Martin 2003). An
advantage of clonal propagation is that a large number of individuals can be obtained
relatively rapidly that would otherwise take much longer if propagated by using seeds,
however, the genetic diversity is highly compromised in clonal populations (Collins and
Dixon 1992, Martin 2003).
In situ seed sowing, seedling planting, or tuber transplant are additional approaches
to population augmentation (Batty et al. 2006). In one study, 18% of Thelymitra
manginiorum persisted 5 years subsequent to transplanting of seedlings and dormant
tubers (Batty et al. 2006). It is also possible in some cases to sow seeds in situ to facilitate
terrestrial orchid establishment in situ (Huber 2002). This method has been reported to be
successful for Cypripedium kentuckiense (Huber 2002). In some cases, a carrier such as
sugar or cracked corn is mixed with the seeds before sowing in an attempt to recruit new
individuals (Huber 2002).
Field establishment of in vitro raised plants has been reported for P. chapmanii
(Richards and Sharma, 2014). In 2012 and 2013, two-year old plants of P. chapmanii
were transplanted into an existing population. In August 2014, 76% (26 of 34) of the
transplanted P. chapmanii were observed flowering (Richards and Sharma 2014, J.
Sharma pers. comm.). Whether in vitro grown plants can be successfully established
within other natural populations or in potentially suitable habitat currently void of P.
chapmanii plants is not documented. Additionally, whether in vitro raised plants have an
advantage over naturally occurring individuals with respect to transplant success has not
been recorded. If individuals growing naturally are recruited for transplanting and
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19
relocating to another population, they may have an advantage over plants raised in vitro
and then planted into native soil. On the other hand, plants raised in vitro and
acclimatized in a greenhouse may be more robust than naturally occurring individuals
giving the former an advantage.
Mycorrhizal associations
Orchid distribution and abundance patterns depend on a multitude of factors. One
such inclusion could be the distribution and abundance of the suitable mycorrhizal fungal
associates and / or the specificity of the mycorrhizal association (McCormick and
Jacquemyn 2013). Some examples of such correlations include the common species Disa
bracteata and the widespread Pyrorchis nigricans, which have been documented to
associate with diverse and widespread groups of fungi (Bonnardeaux et al. 2007). In the
same study, Bonnardeaux et al. (2007) reported that the most disturbance-tolerant and
rapidly spreading species of orchid were documented as having the broadest fungal webs.
In contrast, some of the more selective and slower-growing orchid species such as
Caladenia falcata and Pterostylis sanguinea form associations with smaller fungal webs
(Bonnardeaux et al. 2007).
Non-photosynthetic orchid taxa (e.g. members of the genus Hexalectris) have been
documented exhibiting high specificity toward their fungal associates (Kennedy et al.
2011). However, there is growing documentation of photosynthetic species of orchid also
exhibiting high specificity. The genus Cypripedium is one such example; species within
this genus have been reported to associate largely with members of a narrow clade of
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20
Tulasnellaceae (Shefferson et al. 2005). Because of the specificity of these relationships,
the geographical range of species within the genus Cypripedium could be restricted by
the availability of the mycobionts (Shefferson et al. 2005). There is not always a
connection between narrow distribution of an orchid species and high mycorrhizal
specificity. The endemic North American species Piperia yadonii has been recorded as
associating with a diversity of fungi from three fungal families: Ceratobasidiaceae,
Tulasnellaceae, and Sebacinaceae (Pandey et al. 2013) despite being restricted to a single
County in California, USA.
The diversity of mycorrhizal fungi in and around the rhizosphere may affect the
fungal species that colonize orchid roots. However, in some cases there are many root-
exclusive species. Investigation of potential mycorrhizal fungi of the terrestrial orchid
Neottia ovata showed 68 total species of mycorrhizal fungi, 21 of which were exclusively
present in the roots of the orchid (Jacquemyn et al. 2015).
Although a majority of the known orchid mycorrhizae belong to the phylum
Basidiomycota, some are Ascomycetes (Dearnaley 2007). Because mycorrhizae are
indispensable in orchid development, their documentation and conservation is a part of
orchid species conservation. The loss of suitable fungi could lead to the further decline of
orchid populations (Sharma el al. 2003b).
There is documentation of mycobionts associating with species in the genus
Platanthera. It is common to find Tulasnellaceae and Ceratobasidiaceae in the roots of
Platanthera species including Platanthera praeclara and Platanthera leucophaea
(Currah et al. 1990, Zettler and Hofer 1998, Sharma et al. 2003b). Platanthera
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21
leucophaea has been recorded associating largely with Ceratobasidiaceae in nine
populations in Michigan and Illinois (Zettler and Piskin 2011). Across the nine
populations, 75 plants were sampled. Eighty-eight percent of the fungal isolates were
grown from these 75 plants were observed as Ceratobasidiaceae. These strains of
Ceratobasidiaceae were recovered from various stages of growth including protocorms,
seedlings and mature plants of P. leucophaea (Zettler and Piskin 2011). Although results
from this study suggested that Ceratobasidiaceae is a common associate to P. leucophea,
abundance and distribution of specific strains within Ceratobasidiaceae were not recorded
(Zettler and Piskin 2011). A recent range-wide study of P. praeclara across its
geographic range, spanning from Monitoba (Canada) to Missouri, showed
Ceratobasidiaceae being dominant in most populations (Tovar 2015). Species of
Platanthera are not strictly documented as being dominated by Ceratobasidiaceae.
Protocorms of Platanthera holochila, a Hawaiian endemic species, were documented to
associate with strains of Tulasnellaceae (Zettler et al. 2011). Platanthera chapmanii is
known to associate with Tulasnellaceae (Richards and Sharma 2014). However, the
results from Richards and Sharma (2014) are preliminary and additional fungal data are
necessary. It is not known how specific P. chapmanii mycorrhizal associations are
compared to other terrestrial orchid taxa.
Temporal variation of mycorrhizae is not well documented for terrestrial orchid
species. Throughout a growing season, mycorrhizal associations within a population may
vary (Ercole et al. 2014). The meadow orchid Anacamptis morio has been recorded with
mycorrhizal differences over several seasons; Tulasnella being more common in the
Texas Tech University, Kirsten Poff, August 2016
22
autumn and winter and in the summer Ceratobasidium was more common (Ercole et al.
2014). Platanthera praeclara was tested in a similar way, however the results of the 2015
study were only partially conclusive (Tovar 2015). Tovar (2015) observed an overall
change in mycorrhizal community from one year to the next however whether these
changes were significant were not recorded. In addition, whether terrestrial orchid species
maintain mycorrhizal specificity when raised in vitro and cultured in a greenhouse (with
or without nutrient supplementation) is not fully documented (Richards and Sharma
2014). In addition to revealing whether or not an orchid species is highly specific to its
mycorrhizal associations, this could be important for species grown and cultured in vitro
for restoration and re-introduction into native habitats.
Techniques for mycorrhizal identity
Much information on orchid mycorrhizae has been generated from in vitro isolation
of fungi (Zettler et al. 2001, Sharma et al. 2003b, Zhu et al. 2008). This technique is
performed by plating sections of surface sterilized roots or individual pelotons on an
agar-based medium. With this technique, fungal isolates may be identified or used for
symbiotic germination. However, it is often difficult to accurately identify the fungal
isolates because contaminants and endophytes may be isolated and mistaken for
mycorrhizae (Taylor and McCormick 2007, Zhu et al. 2008, Dearnaley et al. 2012). In
addition, fungi from inactive pelotons may be excluded when using this method, or one
peloton may contain several different fungal taxa making it difficult to isolate a single
fungus (Zhu et al. 2008). Although there are multiple protocols that can be followed to
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23
reduce these issues, molecular techniques can be more reliable than culture-based fungal
identification methods (Zhu et al. 2008).
It is common to use DNA-based barcoding to identify fungi (Schoch et al. 2012).
The nuclear ribosomal internal transcribed spacer (ITS) region is a universal DNA
barcode marker for fungi and is useful for identifying mycorrhizae (Schoch et al. 2012).
Orchid-fungus specific primers have been designed for the ITS region of the fungal
nuclear DNA, including primers that are specific for Tulasnellaceae, Thelephoraceae and
generalized basidiomycete fungi (Taylor and McCormick 2008). Before 2008, it was
difficult to amplify Tulasnellaceae using standard primers because the family exhibits
accelerated evolution of the nuclear ribosomal operon (Taylor and McCormick 2008).
The pair ITS4-1 and ITS1-OF were designed to help amplify basidiomycete fungal DNA.
If the target DNA is Tulasnellaceae, the primer ITS4-Tul can be paired with ITS1 or ITS5
to amplify this notoriously difficult to amplify group of fungi (Taylor and McCormick
2008). The primer pairs have been designed for Sanger sequencing and have proven
useful in identifying orchid mycorrhizal species (Nontachaiyapoom et al. 2010,
Tendersoo et al. 2010, Roche et al. 2010, Bailarote et al. 2012, Jacquemyn et al. 2012,
Pandey et al. 2013). In addition, Sanger sequencing may offer higher resolution than 454-
pyrosequencing but this is not always the case (Tedersoo et al. 2010). In Sanger
sequencing, longer sequences can be achieved; next generation sequencing often
sequences ITS-2 region resulting in shorter sequences (around 300 bp) then Sanger
sequencing. In addition, because of the amount of data provided by next generation
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24
sequencing, it may be difficult to distinguish between endophytes and peloton-forming
mycorrhizal fungi (Tovar 2015).
Summary of Research Gaps
Minimal data are available on the biology and ecology of Platanthera chapmanii
(Small 1903, Liggio and Liggio 1999, Brown 2004, Richards and Sharma 2014). Some of
the understudied areas include dormancy breaking techniques, seed viability, and
germination rate. Overall, it appears that response of seeds to stratification is variable
across species, and generalizations may not be possible (Zettler et al. 2000, Zettler et al.
2001, Sharma et al. 2003a, Lauzer et al. 2007). This may be especially true because of the
local climatic adaptations of temperate species. Although asymbiotic germination is
documented for the species, empirical data for P. chapmanii seed viability and optimal
cold stratification period or its necessity is not known (Richards and Sharma 2014).
Greenhouse culture of P. chapmanii has been successful without an acclimatization
period (M. Richards pers. comm.). However, nutrient supplementation has not been
tested. Terrestrial orchid taxa can be sensitive to commercial fertilizers, even in diluted
amounts but this is not recorded for many species (Silvertown et al. 1994).
Augmenting the existing populations or establishing new populations of terrestrial
Orchidaceae can be a successful method of conservation as it can directly increase
numbers of individuals in a population (Cribb et al. 2003, Batty et al. 2006, Decruse et al.
2013, Richards and Sharma 2014). Augmentation of a natural population in Texas
[Watson Native Plant Preserve (WNPP)] was conducted by using individuals grown in
Texas Tech University, Kirsten Poff, August 2016
25
vitro and cultured in a greenhouse (Richards and Sharma 2014). An unanswered question
though is whether this method is repeatable at additional sites. In addition, it is not known
if the plants need to be cultured in a greenhouse prior to outplanting or if they can be
established in the field directly after sterile culture.
Peloton-forming fungi from the family Tulasnellaceae have been identified from the
roots of P. chapmanii occurring naturally in the population at WNPP (Richards and
Sharma 2014). Whether the species is specific in its association with this single fungal
family is not known because of the limited, preliminary data (Richards and Sharma
2014).
Soil microbial communities could have an effect on associations of P. chapmanii.
Roots of greenhouse acclimatized individuals of the taxon could have different peloton-
forming species associating with them then naturally occurring individuals. Greenhouse
medium is different from native soil, plants growing in a greenhouse may be associating
with smaller fungal webs than those at WNPP.
Population augmentation has been proven somewhat successful with P. chapmanii
(Richards and Sharma 2014). In addition, knowledge concerning P. chapmanii
development and mycorrhizal associations will be helpful in the conservation of the
species. Three specific research questions concerning the biology of P. chapmanii were
developed.
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26
Objectives of the study
1) Evaluate the effect of 8 and 12 week cold-moist stratification pre-germination
treatments on seed germination and plant development in Platanthera chapmanii.
2) Evaluate the effect of supplemental nutrients on the plant height of in vitro raised
Platanthera chapmanii plants and compare above-ground emergence of in vitro /
greenhouse cultured plants of Platanthera chapmanii with naturally-occurring,
relocated Platanthera chapmanii plants after transplanting within naturally
occurring populations.
3) Document the diversity of mycorrhizae forming fungi of Platanthera chapmanii
in response to time and growing environment.
Significance of the study
This study will allow an evaluation of how effective the pre-germination treatment of
cold-moist stratification is on P. chapmanii seeds. It will also estimate the effectiveness
of nutrient supplementation and lend knowledge to how easily natural populations of P.
chapmanii can be augmented using lab raised individuals. The importance of mycorrhizal
associations in terrestrial orchids is well documented. Data from this study will lend
identity to some associations of P. chapmanii. All of the information produced through
this study will allow for the better conservation of the species P. chapmanii.
Texas Tech University, Kirsten Poff, August 2016
27
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Figure 1.1. A photograph of Mesic to wet pine habitat of Platanthera chapmanii at
Watson Native Plant Preserve, Tyler County, Texas (2014).
Figure 1.2. A photograph of Platanthera chapmanii during anthesis at Watson Native
Plant Preserve, Tyler County, Texas. Photograph by Jyotsna Sharma (2013).
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Figure 1.3. A map of the southeast United States showing the geographic range of
Platanthera chapmanii. Areas within Texas, Florida, and Georgia where the species
occurs naturally are shaded in blue.
Figure 1.4. A photograph of a cross section of Platanthera chapmanii root tissue
showing coils of hyphae, pelotons, within the root cells documented in November 2014.
100 µm.
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CHAPTER II
COLD-MOIST STRATIFICATION IMPROVES GERMINATION IN A
TEMPERATE TERRESTRIAL NORTH AMERICAN ORCHID
Abstract
Seed dormancy is a common evolutionary adaptation in temperate plant taxa.
Dormancy mechanisms can prevent seeds from germinating at inopportune times, such as
a cold period. The influence of pre-germination stratification treatments on in vitro seed
germination and seedling development in Platanthera chapmanii, a rare temperate
terrestrial orchid native to the southeastern United States is reported. Seeds were
subjected to either 0, 8, or 12 weeks of cold-moist stratification at 5°C. Mean seed
viability was 89%. Nine months after plating, seeds exposed to 8 and 12 weeks of
stratification resulted in higher germination (Stage 1; 32% and 35%, respectively) in
comparison to 25% germination in non-stratified seeds. Once a protocorm developed a
leaf primordium (i.e., reached Stage 2), development to Stage 3 (root development) was
independent of the pre-germination treatments. Exposure to artificial lights for 3, 4, and 5
months resulted in 32%, 44%, and 63% of the Stage 2 seedlings developing into Stage 3
photosynthetic root-bearing seedlings. The results indicate that in vitro seed germination
in this temperate terrestrial orchid can be improved by using cold-stratification. Further,
leaf- and root-bearing seedlings can be obtained through the methods reported herein.
Key words: Asymbiotic germination, cold-moist stratification, seed dormancy, sterile
culture, plant conservation, Orchidaceae.
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Introduction
Seed dormancy is a consequence of evolutionary adaptation and an important
survival mechanism in many plant species (McMahon et al. 2011). Dormancy prevents
seed germination until adverse conditions abate because seedlings that develop in
unfavorable environmental conditions may perish without reproducing. Types of
dormancies exhibited across the plant kingdom vary by species and are a result of the
specific adaptations of a species to its local climatic conditions. Typically, temperate
species tend to manifest seed dormancy during cold periods. Within the temperate
biomes, seeds of species native to colder latitudes may be adapted to longer dormancy
periods compared to those that occur in the warmer temperate regions. Further, the
mechanism of dormancy may be physical or physiological, and sometimes a complex
combination of dormancies may occur in seeds in response to evolutionary pressures
(Baskin and Baskin 1998). For example, physical dormancy is a type of exogenous
dormancy that requires scarification for water to pass through the impermeable layers of
a seed coat (Baskin and Baskin 1998). Seeds with other dormancies, such as
physiological or chemical dormancy, may be permeable to water but require a metabolic
change to occur before germination (Baskin and Baskin 1998). Understanding the
dormancy mechanisms in seeds has implications for plant reproductive ecology, biology,
and propagation.
The microscopic dust-like seeds in the family Orchidaceae, the largest
angiosperm family on earth with an estimated 25,000 to 30,000 species distributed across
the planet, represent a variety of complex seed dormancy mechanisms (Johansen and
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Rasmussen 1992, Rasmussen 1995, Lauzer et al. 2007). When used for in vitro
propagation, orchid seeds can require pre-germination treatments to overcome physical
and / or physiological dormancy (Johansen and Rasmussen 1992, Rasmussen 1995,
Zettler et al. 2001, Sharma et al. 2003, Lauzer et al. 2007). For example, seeds of
Platanthera praeclara Sheviak and M.L. Bowles, which is native to the midwestern U.S.,
germinated in vitro only after they were exposed to 4 or 6 month cold-moist stratification
periods (Sharma et al. 2003). Seeds of Platanthera leucophaea (Nutt.) Lindl., a sister
species, have also been documented to require at least 2 months of cold stratification to
germinate (Stoutamire 1996). Another study on in vitro seed germination of P.
leucophaea documented that non-stratified seeds showed ≤ 5% germination, whereas 8
week and 16 week stratification increased germination to <20% and >30%, respectively
(Bowles et al. 2002). Conversely, seeds of Platanthera integra (Nutt.) A. Gray, a species
native to the southeastern United States responded to scarification with an ethanol: 5.25%
sodium hypochlorite (NaOCl) (Clorox): deionized water (1:1:1,v:v:v) solution.
Germination percentage and protocorm development increased to 14.5% and 27.2% in
seeds scarified for 1 or 2 hours respectively, in comparison to 1.6% and 6% with shorter
(1 min or 30 min, respectively) scarification treatments (Zettler et al. 2000). Given that P.
integra is native to acidic bog habitats in warmer temperate zones, its seeds may have
developed a physical dormancy mechanism instead.
Considering that the family Orchidaceae exemplifies evolutionary advances in
angiosperms and that a majority of orchid taxa are rare, and routinely require specialized
in vitro propagation techniques (Dressler 1981), knowledge of species-specific
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propagation protocols is necessary to produce propagules for both research and
conservation. However, empirically developed propagation protocols exist for relatively
few species. This lack of knowledge is especially evident in the temperate terrestrial
orchid taxa native to North America, perhaps because of a perceived lack of their
commercial value. At the same time, conservation threats (i.e. changes in land use) to rare
plants are increasing globally (Swarts and Dixon 2009). In fact, little is known about the
biology and ecology of most orchid species. Platanthera chapmanii (Small) Luer, a rare
species native to the southeastern United States, faces similar knowledge gaps. This
severely understudied taxon has a geographic range limited to northern Florida, southeast
Georgia, and southeast Texas (Poole et al. 2007). Because of conversion from native
longleaf pine (Pinus palustris Mill.) to industrial pine forest and urban development, the
quality of habitat in which P. chapmanii naturally occurs continues to decline (Gilliam
and Platt 2006). Populations of P. chapmanii are often small with ≤10 individuals, and
the only large population with ≥100 flowering individuals occurs in southeast Texas
(Richards and Sharma 2014). Anthesis time for P. chapmanii is between late July and
early August when individual plants produce single inflorescences with ≥60 orange
flowers (Liggio and Liggio 1999). The taxon is assumed to be an obligate outcrossing or
facultative outcrossing species, as is the case with most species of Platanthera (Argue
2012). According to a multi-species study conducted on the coastal plain of Florida and
Alabama, P. chapmanii has been documented to be pollinated by several species of long-
tongued butterflies including Phoebis sennae (Linnaeus), Papilio troilus (Linnaeus),
Papilio palamedes (Drury), and Papilio marcellus (Cramer) (Argue 2012). In southeast
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Texas, Papilio palamedes has been documented carrying P. chapmanii pollinia (J.
Sharma, pers. obs.). If pollination and fertilization is successful, each flower can
potentially produce a capsule containing thousands of dust-like seeds. Capsule
dehiscence typically occurs in October, and subsequently seeds are presumed to
experience cold and moist conditions during the winter before environmental conditions
change to facilitate germination and seedling development. Information on reproductive
biology and natural recruitment is not available for this taxon. However, a preliminary
study reported in vitro propagation, culture, and outplanting of laboratory raised plants
into the wild for population augmentation (Richards and Sharma 2014). Experimental
data on germination and development however do not exist.
The objective of this study was to quantify the influence of cold stratification on
in vitro seed germination and plant development in a North American terrestrial
temperate orchid, P. chapmanii. Considering that the seeds of P. chapmanii are exposed
to average minima as low as -9°C at 32°N and -7°C at 29°N (USDA 2016), it was
hypothesized that non-stratified seeds of P. chapmanii will exhibit a lower germination
percentage in comparison to cold stratified seeds. Further, it was expected that seeds
stratified for 8 weeks at 5°C will yield similar germination and plant development as
those stratified for 12 weeks at 5°C; this expectation was based on the relatively short
(~10-40 days below 0°C) cold period the species experiences across its natural
distribution (NOAA 2016).
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Materials and methods
Seed stratification
Seeds were collected from multiple capsules in October 2014 from haphazardly
selected individuals of P. chapmanii at a population in southeast Texas. A maximum of
one seed capsule was collected from each selected plant. Capsules were placed on a filter
paper at room temperature at approximately 40% RH to allow them to desiccate further
and dehisce. Seeds were then collected and placed in a 1.5ml glass vial. The vial
containing the seeds was stored over silica gel desiccant at -20°C until further use.
Seeds were prepared for the 8 and 12 week cold-moist stratification treatments by
first surface sterilizing them with a 0.6% NaOCl solution for 3 min. Seeds were then
rinsed in sterile ultrapure water and approximately equal portions were placed in each of
two 2ml safe-lock microcentrifuge tubes (Eppendorf, Hamburg, Germany) containing
approximately 1-1.5ml sterile ultrapure water. The vials were inverted several times,
wrapped in aluminum foil, and stored at 5°C for their respective stratification periods.
The timing of initiating the stratification treatments was staggered (8 and 12) to allow for
the seeds in all three treatments to be plated at one time.
Once the stratification treatments had been applied, the seeds from both 8 and 12
week treatments were again surface sterilized by submerging in a 0.6% NaOCl solution
for an additional 6 min prior to plating on sterile nutrient medium. At this time, the seeds
in the 0 week cold-moist stratification treatment were surface sterilized by submerging in
a 0.6% solution of NaOCl for 12 min. The difference in total NaOCl exposure time from
9 min (8 and 12 week cold-moist stratification) to 12 min (0 week cold-moist
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44
stratification) prior to plating of seeds was to ensure that the embryos softened by cold-
moist stratification were not damaged during the second surface sterilization procedure.
Seed plating and germination assessment
After the seeds were subjected to their respective pre-germination treatments, they
were plated onto sterile P723 medium (Phytotechnology Laboratories, Overland Park,
Kansas) contained in sterile single-use Stericon-4 237 ml polystyrene containers
(Phytotechnology Laboratories, Overland Park, Kansas) in February 2015.
Approximately 80 ml of medium was used per vessel and between 100 and 500 seeds
were spread onto each vessel. Each of the three cold-moist stratification treatments was
replicated 30 times with an experimental unit defined as one culture vessel. After the
seeds were plated, a dissecting microscope was used to count and record the total number
of seeds within each of the 90 vessels. At the same time, a count of viable seeds was
performed. Within the 30 containers representing the 0 week stratification treatment, each
seed was observed for presence of a healthy embryo (defined as a clear, hyaline, rounded
embryo) to be categorized as a viable seed (Figure 2.1). However, water imbibition
instead was used as a measure of viability for seeds in the remaining 60 plates which
contained seeds that were cold-moist stratified for 8 or 12 weeks. A swollen embryo
(indicating imbibition) was counted as a viable embryo (Figure 2.2).
A visual assessment of germination and development was performed by inspecting
all seeds in each experimental unit every 30 days. Germinating seeds and developing
seedlings were categorized in one of the three categories, i.e., Stage 1, 2, or 3 (Figure
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2.4). Stage 1 was defined as germination and the presence of one or more rhizoids; Stage
2 as the presence of a leaf primordium on the developing protocorm; and Stage 3 as the
presence of at least one root.
Once a seed reached Stage 2, it was transferred to new containers with fresh P723
medium. Sterile Magenta GA-7 vessels (Sigma-Aldrich, St. Louis, Missouri), with
approximately 50 ml of P723 medium were used for the transfer. The newly transferred
protocorms were then placed on a culture rack and exposed to 40-watt white florescent
light bulbs set at a photoperiod of 12 hours. The developing seedlings were subsequently
examined every 30 days for further development (Figure 2.5. Nine months after the seed
plating, young plants were individually examined for root development. After this, they
were placed in autoclaved PTcon 947 ml culture vessels (Phytotechnology laboratories,
Overland Park, Kansas) with approximately 150-200 ml of autoclaved P723 medium for
further development. The experimental data collection for this study was considered
complete at this time.
Data analysis
A one-way Analysis of Variance (ANOVA) was performed with Stage 0, Stage 1,
Stage 2, and Stage 3 germination as the dependent variables and stratification treatment
as the independent variable. The means were separated using Fisher’s Least Significant
Difference (LSD) test. To test whether each treatment received a similar number of seeds,
ANOVA was performed using the total number of seeds plated in each experimental unit.
Texas Tech University, Kirsten Poff, August 2016
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A similar procedure was also used to test the differences in viability among the plated
seeds.
Further, a two-way ANOVA was performed with stratification period and
duration of exposure to light as the two independent variables and Stage 3 proportions as
the dependent variable. Means were separated by using Fisher’s LSD test.
All statistical analyses were performed using RStudio 0.99.842 (RStudio Team
2015) using the agricolae package with α = 0.05.
Results
Seed germination
Among the approximate 22,348 individual seeds used across the three
stratification treatments, mean viability was 89% (Table 2.1). Results from an ANOVA
and Fisher’s LSD test showed that P. chapmanii seeds exposed to the 0 week
stratification treatment had lower Stage 1 germination percentage than the 8 and 12 week
cold-moist stratification treatments. When the total number of seeds sown was used as the
denominator for calculating percent of seeds that reached Stage 1 germination, the 0
week stratification treatment had lowest germination (mean = 22.8%; p = 0.00), whereas
the means were statistically similar for the 8 and 12 week treatments (28.7% and 30.7%,
respectively) (Table 2.1). Similar results were observed when the number of viable seeds
was used as the denominator to calculate germination percentages. In this case, mean
germination in the 0 week cold-moist stratification treatment was 25.4% (p = 0.00),
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whereas 32.4% and 35.1% germination among the 8 and 12 week treatments, respectively
was observed. Finally, the highest percentage of un-germinated seeds was observed in the
0 week stratification treatment (77.3% of all plated seeds and 74.6% of viable seeds)
while both 8 and 12 week stratification treatments had lower percentages of un-
germinated seeds (Table 2.1).
Seedling development
When data for seedling development from Stage 2 to Stage 3 were analyzed using
a two-way ANOVA including stratification and duration of exposure to light, there were
no interactive effects or effect of stratification treatments on the means. However, an
influence of duration of exposure to light on seedling development was observed.
Even though not significantly different (p = 0.27), the absolute means of Stage 3
seedlings ranged from 16.4% (12 week cold-moist stratification) to 22.1% (0 week cold-
moist stratification across the three stratification treatments) (Table 2.2).
In response to exposure to light, seedlings that reached Stage 2 required at least 1
month of exposure to artificial lights to develop roots (i.e., to reach Stage 3; Figure 2.6,
Figure 2.7). As the duration of exposure to lights increased from 1 to 5 months, the mean
percentage of Stage 3 seedlings increased significantly (Figure 2.6, Figure 2.7). While the
means for 1 and 2 month exposure were similar, 32%, 44%, and 63% of Stage 2
seedlings reached Stage 3 after 3, 4, and 5 months, respectively (Figure 2.6, Figure 2.7).
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Discussion
As in many plant species, seed dormancy in temperate terrestrial orchids is
common (Johansen and Rasmussen 1992, Rasmussen 1995, Lauzer et al. 2007). The
duration and type of seed dormancy, however, is often species- and climate- dependent.
Temperate terrestrial species sometimes require a cold-moist stratification period of at
least 8 weeks to initiate germination (Rasmussen 1992, Sharma et al. 2003). Although
Richards and Sharma (2014) reported propagation of P. chapmanii from seed after ≥12
weeks of exposure to cold-moist stratification conditions, germination percentages were
not quantified in their study; the resulting plants however, were reported to survive for >3
years. Further, the authors documented reproduction and survival of the artificially
propagated plants both in cultivation in a greenhouse environment and in the native
habitat of the species (Richards and Sharma 2014). In the present study, it is reported that
cold-moist stratification improves asymbiotic in vitro germination among P. chapmanii
seeds when compared to non-stratified seeds. An increase of approximately 10% was
observed when seeds were stratified for either 8 or 12 weeks; however, differential
influence of stratification on plant development past Stage 1 (germination and rhizoid
development) was not observed. While lower than the mean germination obtained after
treating seeds with stratification, up to 25.4% germination in non-stratified seeds was
observed. Similarly, Zettler et al. (2000) reported a low (15%) germination in non-
stratified P. integra seeds collected in North Carolina; however, seeds in their study were
not plated on nutrient-rich asymbiotic medium. Cold-moist stratification treatments were
not included in their study, hence it is not known whether P. integra, which is native to
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similar habitats and climate as P. chapmanii, has similar pre-germination stratification
requirements. Another congeneric species native to southeastern U.S., Platanthera
clavellata (Michx.) Luer, however, yielded much higher (47%) germination in non-
stratified seeds collected from Tennessee and South Carolina (Zettler and Hofer 1998).
Altogether, these two species have an overlapping distribution with P. chapmanii in
southeast Texas and northern Florida, although, P. clavellata extends also to northern
latitudes in Quebec and Ontario (USDA 2016). The germination percentage for non-
stratified seeds reported in P. clavellata is higher than those observed both in P.
chapmanii and P. integra. These data confirm that results from any individual species
should not be broadly applied to even the congeners from similar habitats, and that
species-specific studies are necessary to understand the nuances within each taxon. It is
also clear that additional and alternative pre-germination treatments should be examined
for P. chapmanii such as scarification or different light / dark periods. At the same time,
it is possible that cold stratification may improve the germination among P. integra and
P. clavellata seeds. On the other hand, testing the efficacy of symbiotic fungi in
improving germination in P. chapmanii could also be considered.
The similarity between the germination percentages obtained from 8 and 12 week
stratification treatments in the study suggests that stratification periods longer than 8
weeks may not be necessary to improve germination. In fact, it is possible that a
stratification period between 0 and 8 weeks could optimize germination in P. chapmanii.
In southeast Texas, where the seeds for this experiment were collected, the average
minima for the coldest month (January) ranged from -2°C to 4°C between 2012 and
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2016, whereas the average maxima ranged from 15°C to 21°C between 2012 and 2016
(NOAA 2016). Considering this, continuous cold-moist stratification at 5°C for 8 weeks
might have been excessive. Whether a shorter stratification duration, or other pre-
germination treatment combinations, would increase germination beyond 35.1%
(maximum observed in this study) remains to be empirically tested, however.
Additionally, because germination rates can vary among disjunct populations of the same
species, germination studies with seeds from additional populations of P. chapmanii
could help to further clarify differences in germination in relation to provenance.
Although the species has a wide range from east to west (81° W to 94° W), P. chapmanii
populations are disjunct, small and occur north to south within a relatively narrow
latitudinal zone between 29° N and 32° N (NOAA 2016). Seeds used in the current study
were collected from a single population, though it is the largest documented population of
the species and potentially the most genetically diverse. However, even this relatively
large population may contain reduced genetic variation considering the long inter-
population distances. In some plants, germination percentages correlate positively with
genetic diversity and population size as in the perennial prairie species Silene regia Sims
(Menges 1991). Similarly, the North American species Ipomopsis aggregata (Pursh) V.E.
Grant exhibited reduced germination in seeds from populations with ≤100 individuals
than seeds collected from larger populations (Heschel and Paige 1995). Conversely, a
study on the perennial rock plant Draba aizoides Pall. Ex M. Bieb showed that
populations with lower genetic variation exhibited high germination rates when compared
to populations with higher genetic variation (Vogler and Reisch 2013). Combined with
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51
population genetic diversity analyses, range-wide germination studies should be pursued
to elucidate provenance differences and to assist with conservation of P. chapmanii.
The effect of climate change is reported to be more severe on rare plants with
fragmented populations. According to a review by Walther et al. (2002), the vegetative
growth and flowering in multiple plant species in Germany is occurring progressively
earlier in the year since the 1960s. The capacity of orchid seeds from temperate regions to
germinate in the absence of stratification, as documented in this and other studies, could
be an increasingly useful evolutionary adaptation as the climate changes (Canadell and
Noble 2001). This strategy could allow natural recruitment and time for adaptation under
milder climatic conditions.
Stratification treatments also may influence growth and development beyond
germination in temperate orchid taxa. For example, seeds of P. praeclara stratified for 6
months and cultured symbiotically developed roots after 60 days of culture. In
comparison, the 4 month stratification period did not yield root-bearing seedlings
(Sharma et al. 2003). Considering that plants developed to Stage 3 (root-bearing,
photosynthetic seedlings) in the study consistently across the three stratification
treatments, it is evident that plant development up to 9 months beyond germination
(Stage 1) is independent of the pre-germination stratification period in P. chapmanii.
While additional reproductive biology and recruitment studies must be conducted for P.
chapmanii, the results provide an effective and efficient protocol for generating plants of
the species for experimental and conservation applications.
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Argue, C.L. 2012. The pollination biology of North American orchids. Volume 1:
Springer publishing, New York, New York.
Baskin, C.C., and J.M. Baskin. 1998. Seeds: ecology, biogeography, and evolution of
dormancy and germination. Academic Press, New York, New York.
Bowles, M.L., Jacobs, K.A., Zettler, L.W., and T.W. Delaney. 2002. Crossing effects on
seed viability and experimental germination of the federal threatened Platanthera
leucophaea. Rhodora 104:14-30.
Canadell, J., and I. Noble. 2001. Challenges of a changing earth. Trends in Ecology and
Evolution 16: 664–666.
Dressler, R.L. 1981. The orchids: natural history and classification. Harvard University
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Gilliam, F.S., and W.J. Platt. 2006. Conservation and restoration of the Pinus palustris
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Heschel, M.S. and K.N. Paige. 1995. Inbreeding depression, environmental stress, and
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Johansen, B., and H. Rasmussen. 1992. Ex situ conservations of orchids. Opera Botanica
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Lauzer, D., Renaut, S., St-Arnaud, M., and D. Barabe. 2007. In vitro asymbiotic
germination, protocorm development and plantlet acclimation of Aplectrum
hyemale (Muhl Ex Willd.) Torr.(Orchidaceae). The Journal of the Torrey
Botanical Society 134: 344-348.
Ligio, J., and A.O. Liggio. 1999. Wild orchids of Texas. The University of Texas Press,
Austin, Texas.
McMahon, M.J., Kofranek, A.M., and V.E. Rubatzky. 2011. Plant science. Prentice Hall,
Upper Saddle River, New Jersey.
Menges, E.S. 1991. Seed germination percentage increases with population size in a
fragmented prairie species. Conservation Biology 5:158-164.
NOAA, NCEI. 2016. Monthly Summaries Map (https://gis.ncdc.noaa.gov/maps, 29
March 2016). NCEI GIS Agile Team, Asheville, NS 28801-5001 USA.
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Poole, J. M., Carr, W. R., Price, D. M., and J.R. Singhurst. 2007. Rare plants of Texas.
Texas A&M University Press, College Station, Texas.
Rasmussen, H.N. 1992. Seed dormancy pattern in Epipactis palustris (Orchidaceae):
requirements for germination and establishment of mycorrhiza. Physiologia
Plantarum, 86(1):161-167.
Rasmussen, H.N. 1995. Terrestrial orchids: from seed to mycotrophic plant. Cambridge
University Press, Cambridge, England.
Richards, M. and J., Sharma. 2014. Review of Conservation Efforts for Platanthera
chapmanii in Texas and Georgia. The Native Orchid Conference Journal 11:1-11.
RStudio Team. 2015. RStudio: Integrated development for R. RStudio, Inc., Boston, MA
URL http://www.rstudio.com.
Sharma, J., Zettler, L.W., Van Sambeek, J.W., Ellersieck, M.R., and C.J. Starbuck. 2003.
Symbiotic seed germination and mycorrhizae of federally threatened Platanthera
praeclara (Orchidaceae). American Midland Naturalist 149:104-120.
Small, J. K. 1903. Flora of the Southeastern United States. Small J.K., New York, New
York.
Stoutamire, W.P. 1996. Seeds and seedlings of Platanthera leucophaea (Orchidaceae). p.
55-61. In: C. Allen (ed.). Proceedings of the North American Native Terrestrial
Orchid-Propagation and Production Conference. National Arboretum,
Washington, D.C.
Swarts, N.D., and K.W. Dixon. 2009. Terrestrial orchid conservation in the age of
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USDA, NRCS. 2016. The PLANTS Database (http://plants.usda.gov, 29 March 2016).
National Plant Data Team, Greensboro, NC 27401-4901 USA.
Vogler, F. and C. Reisch. 2013. Vital survivors: low genetic variation but high
germination in glacial relict populations of the typical rock plant Draba
aizoides. Biodiversity and Conservation 22:1301-1316.
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J.M., Hoegh-Guldberg, O. and F. Bairlein. 2002. Ecological responses to recent
climate change. Nature 416:389-395.
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Zettler, L.W. and C.J. Hofer. 1998. Propagation of the little club-spur orchid (Platanthera
clavellata) by symbiotic seed germination and its ecological
implications. Environmental and Experimental Botany 39:189-195.
Zettler L.W., Stewart S. L., Bowles M. L., Jacobs K. A. 2001. Cold assisted symbiotic
germination of the federally threatened orchid, Platanthera leucophaea (Nuttall)
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Zettler, L.W., Sunley, J.A., and T.W. Delaney. 2000. Symbiotic seed germination of an
orchid in decline (Platanthera integra) from the Green Swamp, North Carolina.
Castanea 65:207-212
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Table 2.1. Effect of cold-moist stratification (0, 8, or 12 weeks) on seed germination was
experimentally tested in Platanthera chapmanii. An Analysis of Variance (ANOVA) was
conducted (a); Germination of seeds was categorized as Stage 0 (no further
development), Stage 1 (germination; rhizoid development), or Stage 2 (leaf primordium
development). Mean number of seeds that were plated in an experimental unit, mean
number of viable seeds, and mean percent viability are presented. Mean germination
percentages were calculated by using total number of seeds and number of viable seeds
separately. Means followed by the same letter in each column were statistically similar
based on Fisher’s Least Significant Difference (LSD) test.
2.1a.
Sum of Squaresy Mean Square f value p valuex
zstrat_stage 0t
0.48 0.01 15.36 0.00
strat_stage 0v 0.59 0.01 17.82 0.00
strat_stage 1t 0.45 0.01 13.71 0.00
strat_stage 1v 0.56 0.01 16.21 0.00
strat_stage 2t 0.12 0.00 12.20 0.00
strat_stage 2v 0.15 0.00 11.46 0.00
Total seeds 839462 11499 0.91 0.34
Viable seeds 658411 9019 0.95 0.33
Viability 0.14 0.00 0.86 0.36
z’t’ = proportions calculated by using total number of seeds that were plated; ‘v’ =
proportions calculated by using only the number of viable seeds yFactor level df = 1; residual df = 73 xα = 0.05
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2.1b.
z A single vessel containing multiple orchid seeds served as an experimental unit y’t’ = percentages calculated by using total number of seeds that were plated; ‘v’ = percentages calculated by using only the number of
viable seeds x Proportions were converted to percentages for presentation in the tables
w α = 0.0
Stratification
(# of weeks)
nz total seeds
(#)
viable seeds
(#)
viability
(%)
Stage 0t y
(%)
Stage 0v
(%)
Stage 1t
(%)
Stage 1v
(%)
Stage 2t
(%)
Stage 2v
(%)
0 26 282 251 89.5w ax 77.3 b 74.6 b 22.8 b 25.4 b 11.6 b 13.0 b
8 26 361 321 89.2 a 71.3 a 67.7 a 28.7 a 32.4 a 14.1 a 15.7 a
12 23 223 198 88.3 a 68.4 a 64.0 a 30.7 a 35.1 a 15.5 a 17.4 a
p-value 0.34 0.33 0.36 0.00 0.00 0.00 0.00 0.00 0.00
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Table 2.2. Effect of cold-moist stratification (0, 8, and 12 weeks) on seedling
development after germination and rhizoid development (Stage 1) was experimentally
tested in Platanthera chapmanii. An Analysis of Variance (ANOVA) was conducted (a);
Seed development was categorized as Stage 2 (leaf primordium development) or Stage 3
(root development). Mean number of seedlings that were categorized as Stage 2 or Stage
3 after a 5 month exposure to 40-watt florescent bulbs set at a photoperiod of 12 hours.
2.2a
yFactor level df = 1; residual df = 125 xα = 0.05
2.2b
z A single vessel containing multiple orchid seeds served as an experimental unit yα = 0.05
df Sum of
Squaresy
Mean
Square
f value p valuex
zstrat_stage3 125 8.29 0.07 1.24 0.27
Stratification
(# of weeks)
nz Stage 2
(#)
Stage 3
(%)
0 45 21 22.1 ay
8 46 34 16.5 a
12 36 28 16.5 a
p-value 0.27
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Figure 2.1. Two photographs of Platanthera chapmanii seeds prior to cold-moist
stratification at lower and higher magnification.
Figure 2.2. Two photographs of Platanthera chapmanii seeds after cold-moist
stratification showing imbibition at lower and higher magnification.
1 mm
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Figure 2.4. Seed germination and plant development in Platanthera chapmanii was recorded by using four categories: Stage 0 (no
germination), Stage 1 (germination; rhizoid development), Stage 2 (leaf primordium development), and Stage 3 (root development).
1 mm 2 mm 1 mm 1 cm
Stage 3 Stage 0 Stage 1 Stage 2
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Figure 2.5. A photograph of Platanthera chapmanii seedlings at Stage 2 (leaf primordia
development) after being exposed to light for approximately 3 weeks.
1 cm
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Figure 2.6. Proportion of Stage 2 seedlings of Platanthera chapmanii that reached the
developmental Stage 3 after exposure to light. Duration of exposure to light (1 to 5
months under 40-watt white florescent bulbs) influenced plant development to Stage 3.
Pre-germination stratification of seeds for 0, 8, or 12 weeks did not influence
development from Stage 2 to Stage 3, thus the means were pooled across the three
stratification treatments. Means followed by the same letter were statistically similar
based on Fisher’s Least Significant Difference (LSD) test.
0 a0.02 a
0.32 b
0.44 c
0.63 d
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
1 2 3 4 5
Pro
port
ion o
f S
eedlin
gs
Time under lights (Months)
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Figure 2.7. Proportion of Platanthera chapmanii Stage 2 seedlings that reached plant
development Stage 3 when subjected to cold-moist stratification for 0, 8, or 12 weeks.
Duration of exposure to light (1 to 5 months under 40-watt white florescent bulbs)
influenced plant development to Stage 3. Means followed by the same letter in each
column were statistically similar based on Fisher’s Least Significant Difference (LSD)
test.
0 a 0 a 0 a0.02 a 0.02 a 0.03 a
0.46 b
0.27 b
0.21 b
0.53 c
0.35 c
0.46 c
0.64 d 0.63 d 0.62 d
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0 8 12
Pro
port
ion o
f seedlin
gs
Stratification treatment (# of weeks)
Month 1 Month 2 Month 3 Month 4 Month 5
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CHAPTER III
PLATANTHERA CHAPMANII: NUTRIENT SUPPLEMENTATION AND
POPULATION AUGMENTATION
Abstract
There is lack of protocols describing greenhouse culture of temperate terrestrial
orchid species along with the protocols of their field establishment. Platanthera
chapmanii is a rare species of temperate terrestrial orchid native to the southeastern
United States. Its geographic range is restricted to fragmented populations in Georgia,
Florida and southeast Texas. The current preliminary study attempts to measure the
effects of supplemental nutrients on plant height in P. chapmanii individuals being
cultured in a greenhouse. In addition, it compares above-ground emergence during
flowering season of in vitro / greenhouse cultured plants to naturally occurring
individuals after being transplanted into native habitat in the fall and spring seasons.
Plants were first propagated in vitro and then planted in containers in a greenhouse
setting. The plants were exposed to 0.00x, 0.25x, and 0.50x concentration of commercial
solution of supplemental nutrients once every other week. After a period of 14 weeks,
there was no difference between the three treatments in terms of plant height (p = 0.14).
When the preliminary data were collected in August 2015 for all the plants in the
transplanting experiment, none of the plants, in any treatment were above ground. These
data are a good basis for future studies on the culture and outplanting of temperate
terrestrial orchid species. As anthropogenic changes in land-use continue, restoration
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ecology is becoming an increasing important means for temperate terrestrial orchid
conservation.
Introduction
With evidence of a serious extinction crisis mounting, the need to conserve
biodiversity is growing (Canadell and Noble 2001, IUCN 2009). Up to 70% of plant
species assessed by the International Union for the Conservation of Nature are threatened
with extinction (IUCN 2015). Destruction and degradation of natural systems are largely
responsible for this. According to Brooks et al. (2002) about 50% of the world's vascular
flora is restricted to 25 regional hotspots. Of these hotspots, at least two-thirds have
experienced anthropogenic changes in land-use causing the degradation and destruction
of rare plant habitat (Brooks et al. 2002). With this in mind, it is unlikely that
conservation of all plant species can be accomplished by reservation and preservation
alone (Swarts and Dixon 2009).
Orchidaceae is the largest family of flowering plants with species estimates
between 25,000-35,000 (Dressler 1981). Not only is it the largest, but also one of the
most diverse and widespread (Dressler 1981, Swarts and Dixon 2009). Although the
majority of orchid species are considered rare, the family is severely understudied
(Dressler 1981). Terrestrial species encompass one-third of orchid species and up to half
of the total extinct species in the family (Swarts and Dixon 2009, IUCN 1999).
Temperate terrestrial orchids of North America are under continuing threat because of
changes in land-use and conservation efforts that involve restoration of the North
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American terrestrial orchid habitats are few. One aspect of restoration ecology,
population augmentation, is a method that can be utilized to increase the size of rare plant
populations directly (Batty et al. 2006, Decruse et al. 2013, Richards and Sharma 2014).
Propagation and culture protocol knowledge is necessary to produce propagules for
restoration purposes.
Most of what is documented on orchid propagation and culture is developed for
horticulturally or medically important genera (Griesbach 2002, Park et al. 2002, Lo et al.
2004). In such protocols, an acclimatization period is often used for orchid species
propagated in vitro to ensure survival in the field (Hazarika 2003, Deb and Temjensangba
2006). This usually involves slowly transitioning orchid protocorms or seedlings from
sterile culture conditions into a greenhouse, where the plants are cultured for some time
before they become robust enough to be planted ex vitro (McKendrick 2000). It is
common for some plants to die during the transition between sterile conditions and
greenhouse setting because of a sharp change in abiotic factors (e.g. humidity and
temperature) (Preece and Sutter 1991, Deb and Imchen 2010). Once the plant is in the
greenhouse, depending on the species, an orchid seedling may require several weeks or
months to acclimate to the new environmental conditions (McKendrick 2000, Deb and
Temjensangba 2006). Duration of acclimatization is implicated in the long-term survival
of individual plants (Zeng et al. 2012).
Nutrient supplementation during greenhouse culture is documented for some
horticultural epiphytic species but very rarely for terrestrial species (Wang and Gregg
1994, Wang 1996). The threatened terrestrial orchid Bletia purpurea, after being
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66
transferred from aseptic conditions to a greenhouse setting, was successfully treated with
150 ppm N-P-K balanced liquid fertilizer (Peter’s 20-20-20, The Scott’s Company,
Marysville, OH). Whether supplementing with nutrients during greenhouse culture aided
the growth of the individuals remains unstudied (Dutra et al. 2008). For some terrestrial
species, conventional fertilizers may hinder growth, even when exposed to low
concentrations. For example, significant decreases in flowering in populations of Orchis
morio was measured after the naturally occurring plants were exposed to what were
considered low concentrations of organic and inorganic fertilizers (22-88 kg ha-1 N)
(Silvertown et al. 1994).
Growing and culturing orchid species in vitro and subsequently transplanting
them into native habitat has been reported as a successful approach for some temperate
terrestrial species (Stewart et al. 2003, Zeng et al. 2012, Decruse et al. 2013, Richards
and Sharma 2014). The species Paphiopedilum wardii was established successfully into
native habitat after being cultured in vitro (Zeng et al. 2012). In the study by Zeng et al.,
plants grown from seed were first acclimatized, then used to augment a population in
Gaoligong Mountain in Yunnan. New populations of P. wardii were established in areas
where no prior documentation was recorded in Yangchun and Guangdong (Zeng et al.
2012). Not only did the field established plants exhibit survival percentages ≥50, the
populations persisted after two years (Zeng et al. 2012). The temperate terrestrial North
American species Spiranthes brevilabris has had similar success when transplanted into
its native habitat after in vitro culture (Stewart et al. 2003). Of the 165 Spiranthes
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brevilabris individuals transplanted, survival was 100% after a month in the field,
although only 17 initiated anthesis after 6 months (Stewart et al. 2003).
Clonal propagation of certain members of Orchidaceae for the purpose of native
population restoration is also possible (Martin 2003). The endangered Ipsea malabarica
was documented with a high survival and flowering rate after being introduced into
native habitat. In the 2003 study, all 50 individuals transplanted survived and initiated
anthesis normally (Martin 2003). An advantage of clonal propagation is that a large
number of individuals can be obtained relatively rapidly, this would otherwise take much
longer if propagated by using seeds. However, the genetic diversity is highly
compromised in clonal populations and is therefore often not used for ecological
purposes (Collins and Dixon 1992, Martin 2003).
Platanthera chapmanii (Small 1903), a temperate terrestrial species that is
relatively quick growing has had some success with field establishment (Richards and
Sharma 2014). The taxon is native to the southeastern United States, it’s geographic
range is limited to highly fragmented populations in southern Georgia, northern Florida
and southeastern Texas (Liggio and Liggio 1999). In 2012 and 2013, acclimatized two-
year old plants of P. chapmanii were transplanted into an existing population. In August
2014, 76% (26 of 34) of the transplanted P. chapmanii were observed flowering
(Richards and Sharma 2014, J. Sharma pers. comm.). In vitro asymbiotic propagation and
greenhouse culture may be an efficient way to augment native populations, and establish
new populations of certain temperate terrestrial orchid species. Before field establishment
can be attempted, it is thought that individuals should undergo greenhouse
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acclimatization and culture so that they are robust enough to survive ex vitro. Whether or
not supplementing terrestrial orchids with nutrients during greenhouse culture would be
beneficial is not well reported.
The objective of this study was to quantify the effect of nutrient supplementation
on P. chapmanii above ground plant height when cultured in a greenhouse setting. Also,
to quantify the effect of greenhouse acclimatization, plant source, and planting date on
plant emergence after outplanting P. chapmanii into native habitat. Commercial fertilizer
may be detrimental to some species of terrestrial orchids. Platanthera chapmanii has
been cultured in a greenhouse setting with the assistance of fertilizer and without (M.
Richards, pers. comm.). Effects of this nutrient supplementation have not been quantified.
Because of this knowledge, it was expected that plants treated with a very low
concentration of fertilizer to benefit from the added nutrients and plants treated with
higher concentrations to have detrimental effects. Considering the preliminary study by
Richards and Sharma (2014), fairly good survival percentage for both fall and spring
plantings were expected. Because of what is known about acclimatization, the more
robust greenhouse acclimatized individuals should have higher survival percentage than
the individuals planted directly from aseptic culture. Further, naturally occurring P.
chapmanii individuals that are relocated will most likely have a lower rate of plant
emergence than those individuals that were raised and cultured in vitro because of how
robust the greenhouse cultured plants are.
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Materials and Methods
Nutrient supplementation
To test the effect of nutrient supplementation on growth of two-year old P.
chapmanii plants, 64 in vitro grown individuals were used in this experiment.
Experimental treatments included: 1) biweekly application of 0.5x dilution of the
recommended concentration of Scotts Miracle-Gro 24-8-16 (Scotts Company LLC.,
Marysville, Ohio), 2) biweekly application of 0.25x dilution of the same fertilizer as
above, and 3) no supplemental nutrient solution. Twenty-one or 22 individuals were
haphazardly assigned to each of the three experimental treatments.
Plants cultured under sterile conditions were used in this experiment. The vessels
containing the plants were stored at 5°C from November 2014 to April 2015. At the time
of transferring into greenhouse conditions, plants were removed from culture vessels and
rinsed with reverse osmosis water to remove sterile culture medium. Plants were then
planted directly into 15 centimeter wide plastic greenhouse containers filled with a
soilless medium composed of 44% sphagnum peat moss (Premier Tech Horticulture Ltd,
Quebec, Canada), 26% milled sphagnum moss (Mosser Lee, Millston, Wisconsin), 20%
all-purpose coarse grain builders sand (Quikrete, Atlanta, Georgia), and 10% small tree
fern fiber (repotme, Georgetown, Delaware) v:v:v:v. The Containers were distributed
across 12 trays that were 28 cm long, 56 cm wide and 6.3 cm deep with an approximately
2.5 cm deep layer of reverse osmosis water. Nutrient treatments described above were
commenced after giving the plants about 3-4 weeks to acclimate to greenhouse
conditions. Height of each plant was measured every 14th day for 14 weeks for the
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duration of the experiment. Because the plants were placed in trays in groups of 5 or 6,
each tray served as an experimental unit.
Population augmentation
Platanthera chapmanii seeds collected in 2009 from a large population in southeast
Texas were germinated and propagated in sterile conditions by Atlanta Botanical Garden.
After sterile germination and propagation, the plants were cultured in a greenhouse until
they were excavated and shipped to Texas Tech University. The plants were stored at 5°C
from November 2014 until their planting. Platanthera chapmanii were stored with
sphagnum moss and kept moist with deionized water to prevent drying until they were
used for experimental purposes. In November 2014 the fall transplanting experiment was
initiated. Twelve of the in vitro propagated plants and twelve naturally occurring
individuals of P. chapmanii from a population in southeast Texas were used for
transplanting into an experimental location in Big Thicket National Preserve (BTNP).
Approximately 8 kg of soil were collected from the immediate vicinity of the plants
collected to serve as inoculum at the recipient site at BTNP. It was ensured that soil was
excavated without disturbing or destroying any non-target plants of P. chapmanii. All
experimental activities were conducted under valid permits obtained from the state of
Texas.
At BTNP, two experimental plots were established. The locations of these plots were
determined after examining the area for suitable habitat. Six from each source (in vitro
raised or those excavated from the naturally occurring population) were planted into each
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of the two experimental plots. To plant an individual, a mixture of sphagnum peat moss
(Scott’s Miracle-Gro company, Marysville, OH) and natural soil was used in about a 1:1
(v:v). The sphagnum moss that the lab-raised plants were shipped from ABG in, and the
8 kg of soil from the native population was also distributed across the 24 plants. In vitro
raised plants were labeled with a single metal tag engraved G1 through G12, while the
native plants were labeled N1 through N12.
The experimental plots for spring 2015 were established in mid-March. These
experimental plots were located in close proximity to the fall 2014 plots. The same
protocol as above was followed for the two spring 2015 plots. The metal label codes were
modified to GS1 through GS12, and NS1 through NS12 to denote greenhouse spring and
native spring respectively.
Three more plots were made in the spring of 2015 for one hundred P. chapmanii individuals
that were planted directly from sterile culture into native soil. The location of each plot was
chosen by looking for areas with little shade and soil that was saturated with water. The
plots were constructed by digging holes approximately 15 cm apart. Individuals were
planted by mixing sphagnum peat moss and natural soil in each hole in a ratio of about
1:1, v:v. The soil was packed lightly around each seedling and a thin layer of sphagnum
was placed on top of each seedling. Each plot was watered thoroughly. Planting for all
three plots was performed in March 2015.
The first plot included 34 seedlings. This plot was located in southeast Texas close to
Big Thicket National Preserve. This specific area was chosen because it is known to be
able to support P. chapmanii. It is the same area in which the native adult individuals of
P. chapmanii were relocated from.
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The second plot included 35 plants and was planted in close proximity to the fall and
spring greenhouse acclimated/native relocated plots. This area was chosen because it is
known to support P. chapmanii, had relatively little shade and the soil was water-
saturated.
The third plot included 31 seedlings and was planted inside BTNP at an area along
the Pitcher Plant Bog Trail. The area was chosen in a similar fashion as the previous plant
plots, but there are no P. chapmanii individuals that are known to occur at this specific
location.
In early August 2015 presence or absence and flowering or vegetative data were
collected from all 7 plots for analysis. All metal labels were located and the emergence or
non-emergence of each plant was recorded.
Results
Nutrient supplementation
When change in P. chapmanii above ground plant height was evaluated,
according to an analysis of variance (ANOVA) there was no significant difference
between the three treatments after a duration of 14 weeks (p = 0.14, Table 3.1). The
changes in plant height averages ranged from 1.7 cm in the zero nutrient treatment to 4.8
cm in the 0.5x nutrient treatment but variation in the data was too high for the differences
to be significant.
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Population augmentation
When above ground emergence was compared across the treatments during data
collection in August, zero plants were above ground. When the native and lab raised
plants were compared over the two planting dates using a Fisher’s Exact Test, there was
no significant differences in above ground emergence for the groups (p = 1).
Discussion
Rare plant habitat is being degraded globally (Brooks et al. 2002). With the
disappearance of habitat restoration ecology is becoming increasingly more important for
species conservation (Swarts and Dixon 2009). Developing protocols for the propagation
and culture of plants in vitro is critical to produce propagules for restoration measures.
The ability for researchers to propagate and culture rare plants in vitro for field
establishment is a proven way to directly increase the population numbers of some rare
species (Batty et al. 2006, Decruse et al. 2013, Richards and Sharma 2014). Because
protocols exist for a relatively small number of rare plants, and an even smaller number
of temperate terrestrial orchid species, the current study is relevant for restoration
ecologists and conservation enthusiasts.
The asymbiotic germination and propagation of P. chapmanii has been previously
documented (Richards and Sharma 2014). The current study is helpful for elaborating on
greenhouse culture conditions favored by the study species. Commercial fertilizer used in
this experiment was diluted to 0.5x and 0.25x concentrations. Because it is not
documented whether fertilizer has a positive effect on P. chapmanii growth, extracaution
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was taken with protocol development. The rare plant is known to grow without nutrient
supplementation. A diluted solution of fertilizer was used for the treatments since the
study was preliminary. Because of a lack of documentation on greenhouse culture of
temperate terrestrial orchids native to North America, acclimatization procedures were
not available to us. Individuals were planted directly from aseptic culture vessels into
orchid bog mix. By the end of the experiment, many of the plants, regardless of
treatment, had tip necrosis. Perhaps the decrease in humidity and increase in temperature
from the culture vessels to the green house had an adverse effect. In addition, the trays
used during the experiment were not optimal. The 56 by 28 by 6.3 cm trays used to
increase humidity and soil water saturation made the experiment more self-sufficient.
Because the trays were fairly large they only needed to be refilled with water every other
to every two days depending on weather. However, because more plants were placed in
each of these large trays, this decreased the amount of true replicates in the experiment
drastically from 64 to 12. The current preliminary study was more focused on the
successful culture and growth of P. chapmanii plants in a greenhouse setting then forcing
a treatment effect on the rare species. For future studies, it is recommended that plants be
acclimatized more slowly into greenhouse conditions then placed in individual trays. This
may be helpful if environmental conditions (e.g. temperature, light intensity and
humidity) are especially different than in vitro germination and propagation conditions.
The greenhouse culture of these rare plants is critical so that by the time they are
transplanted to the field, they are robust enough to survive natural conditions.
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Platanthera chapmanii individuals used in these experiments all originated from
seeds collected at the largest known population of the species in southeast Texas. Being
the largest population, it is most likely the most genetically diverse. In addition, there
may be an overabundance of necessary mycorrhizal fungi at this location. Although,
these are speculations as no soil microbial or genetic analyses have been performed. In
the study by Richards and Sharma (2014) P. chapmanii individuals germinated and
propagated asymbiotically and cultured in a greenhouse setting were successfully
introduced into the largest population in southeast Texas. During the current study, an
attempt to augment another population was made, as well as augmenting this large
population. The population that was attempted to augment with robust, greenhouse
acclimatized plants had only one naturally occurring individual. Although most
documented populations of P. chapmanii across its range have ≤10 individuals, only one
documented individual would be considered a small population. In the current study, the
majority of adult plants transplanted in the fall of 2014 had emerged in March 2015,
including the naturally occurring individual. In August there were no above ground
growth from any of the plots. The naturally occurring individual was also not above
ground. Data collected in August 2016 may be helpful in determining whether or not the
plants in the experiment survived. Although, some species of terrestrial orchids have been
recorded as remaining dormant for multiple years (Rasmussen 1995).
Of the individuals that were planted directly into native soil from in vitro
conditions, none were above ground at any of the three plot areas. Even the plot that was
meant to augment the largest population of individuals did not have any above ground
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growth during the August data collection. These plants may be in a dormant state, or the
direct transition from sterile conditions to native soil may have been too big of a shock.
Additionally, the soil may have been lacking compatible mycorrhizae which the
mixotrophic species may require to become established (Rasmussen 1995). Platanthera
chapmanii is a rare species whose habitat is in decline (Gilliam and Platt 2006). Results
from these preliminary studies should be used to help develop protocols for the
propagation of P. chapmanii and similar species of temperate terrestrial orchid, for
conservation efforts.
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Literature Cited
Brooks, T.M., Mittermeier, R.A., Mittermeier, C.G., Gustavo, A.B.D.F., Rylands, A.B.,
Konstant, W.R., Flick, P., Pilgrim, J., Oldfield, S., Magin G., and C. Hilton-
Taylor. 2002. Habitat loss and extinction in the hotspots of biodiversity.
Conservation Biology, 16: 909–923.
Canadell, J, and I. Noble. 2001. Challenges of a changing earth. Trends in Ecology and
Evolution, 16: 664–666.
Dressler, R.L. 1981. The orchids: natural history and classification. Harvard University
Press, Cambridge, Massachusetts and London, England.
Dutra, D., Johnson, T.R., Kauth, P.J., Stewart, S.L., Kane, M.E. and L. Richardson. 2008.
Asymbiotic seed germination, in vitro seedling development, and greenhouse
acclimatization of the threatened terrestrial orchid Bletia purpurea. Plant Cell,
Tissue and Organ Culture, 94(1):11-21.
Gilliam, F.S., Platt, W.J. 2006. Conservation and restoration of the Pinus palustris
ecosystem. Applied Vegetation Science, 9:7-10.
Griesbach, R.J. 2002. Development of Phalaenopsis Orchids for the Mass-Market. p.
458–465. In: J. Janick and A. Whipkey (eds.), Trends in new crops and new uses.
ASHS Press, Alexandria, VA.
IUCN. 1999. IUCN guidelines for the prevention of biodiversity loss due to biological
invasion. Species, 31–32: 28–42.
IUCN. 2009. Extinction crisis continues apace. International News Release.
http://www.iucn.org/?4143/Extinction-crisis-continues-apace
Liggio, J., and A.O. Liggio. 1999. Wild orchids of Texas. The University of Texas Press,
Austin, Texas.
Lo, S.F., Nalawade, S.M., Kuo, C.L., Chen, C.L. and H.S. Tsay. 2004. Asymbiotic
germination of immature seeds, plantlet development and ex vitro establishment
of plants of Dendrobium tosaense makino—A medicinally important orchid. In
Vitro Cellular & Developmental Biology-Plant, 40(5):528-535.
Park, S.Y., Murthy, H.N. and K.Y. Paek. 2002. Rapid propagation of Phalaenopsis from
floral stalk-derived leaves. In Vitro Cellular & Developmental Biology-
Plant, 38(2):168-172.
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Small, J. K. 1903. Flora of the Southeastern United States. Small J.K., New York, New
York.
Swarts, N.D., and K.W. Dixon. 2009. Terrestrial orchid conservation in the age of
extinction. Annals of Botany, 104(3):543-556.
Wang, Y.T., and L.L. Gregg. 1994. Medium and fertilizer affect the performance of
Phalaenopsis orchids during two flowering cycles. HortScience, 29(4):269-271.
Wang, Y.T. 1996. Effects of six fertilizers on vegetative growth and flowering of
Phalaenopsis orchids. Scientia Horticulturae, 65(2):191-197.
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Table 3.1. Effect of nutrient supplementation (0.0x, 0.25x, 0.5x) on Platanthera
chapmanii above ground plant height after 14 weeks of treatment applied every two
weeks. Results of an Analysis of Variance (ANOVA) are presented.
ANOVA df Sum of Squaresy Mean Square f value p valuex
NutriSup_PlantHeight 9 39.57 10.78 2.46 0.14
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Figure 3.1. Three photographs of Platanthera chapmanii individuals after planting in 15
cm containers during nutrient supplementation. From left to right; one individual, one
replicate, row of trays.
Figure 3.2. A photograph of a typical Platanthera chapmanii individual after being
planted into greenhouse medium.
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Figure 3.3. Two photographs of Platanthera chapmanii individuals. A greenhouse
cultured Platanthera chapmanii individual (a) shown beside a naturally occurring
Platanthera chapmanii individual (b) before planting in native habitat in southeast Texas
during fall 2014.
Figure 3.4. A photograph of a fall 2014 Platanthera chapmanii plot with both
greenhouse cultured and native plants relocatedin southeast Texas. An arrow is pointing
to one individual.
a b
1 cm
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Figure 3.5. A photograph of a typical Platanthera chapmanii individual taken directly
out of a culture vessel prior to planting in one of the three locations in southeast Texas in
the spring 2015.
Figure 3.6. A photograph of one of the three plots of Platanthera chapmanii individuals
taken directly out of sterile culture and planted in the spring 2015. All individuals were
covered with sphagnum peat moss.
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CHAPTER IV
DIVERSITY OF MYCORRHIZAE FORMING TULASNELLACEAE IN A
TEMPERATE TERRESTRIAL ORCHID IN EX SITU AND IN SITU
ENVIRONMENTS
Abstract
Variation in mycorrhizal fungal diversity and specificity within the Orchidaceae is
of special interest considering the intimate involvement of orchid associated fungi in
germination and development of individuals. Generally, only a few plants from a few
populations are sampled once to document orchid mycorrhizal fungi without explaining
temporal and spatial variation in the associations. Further, use of mycorrhizal fungi is
recommended for propagating orchid plants for conservation activities, however the
availability and diversity of orchid mycorrhizal fungi in ex situ growing environments is
not documented. The first comparison of mycorrhizal associations of plants of a
temperate terrestrial orchid from in situ and ex situ environments is reported. The fungal
nuclear ribosomal internal transcribed spacer (nrITS) region was amplified and
sequenced from roots collected between 2012 and 2015 at multiple phenological stages
from plants cultured ex situ in laboratory and greenhouse and from the natural habitat.
Across seven sampling events, 122 sequences and 18 operational taxonomic units
(OTUs) were identified, 17 of which represented the fungal family Tulasnellaceae and
one belonged to the Ceratobasidiaceae. Two of the 18 OTUs were shared by individuals
from both growing environments. Of the OTUs originating from the ex situ environment,
eight were exclusive and were genetically closely related. Plants growing in situ also
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hosted eight exclusive OTUs, seven of which belonged to the Tulasnellaceae. Temporal
variation in abundance-based diversity was supported by a principal component analysis
(PCA) but was not by Fisher’s exact test or Kruskal-Wallis. Although the mycorrhizal
fungi from in situ and ex situ conditions segregated in different OTUs within the
Tulasnellaceae, 13 of the 17 OTUs clustered within a single clade in the phylogram. Two
of the 18 OTUs consisted of individual sequences originating from both sources. Overall,
little variation among the mycorrhizal fungi associated with juveniles, plants in anthesis,
and plants entering dormancy was detected. The data suggest that P. chapmanii prefers
OTUs within a few narrow clades of Tulasnellaceae regardless of its phenological stage
or growing environment
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Introduction
Symbioses involve the coexistence of two dissimilar species. Microbial symbioses
are common and have at times led to the development of new cellular structures and
physiological processes in both symbionts (Barton and Northrup 2011). The theory of
endosymbiosis is a good example of this (Rost et al. 2006). In biology, symbioses
describe interactions ranging from parasite-host interactions to interactions in which both
organisms benefit mutually. Symbiotic interactions have been reported across the six
taxonomic kingdoms and include a large diversity of organisms (Ricklefs 2010).
Mycorrhizal associations describe the symbiotic relationship between the roots of
a plant and fungi. It is estimated that 90% of terrestrial plant species form mycorrhizal
associations of some type (Smith and Read 2008). Mycorrhizal relationships can be
mutualistic as the case in vesicular-arbuscular mycorrhizae, the most common
mycorrhizal association. In a mutualistic mycorrhizal relationship, fungi receive
carbohydrates from the phytobiont and in return provide nutrients, especially
phosphorous (Smith and Read 2008, Barton and Northrup 2011). Orchid mycorrhizae,
however, are considered less mutualistic and more parasitic (Taylor et al. 2002). The
family Orchidaceae is estimated to contain approximately 35,000 species (Dressler 1981),
of which two-thirds are epiphytic or lithophytic while the remaining third are terrestrial.
For an orchid seed to germinate and develop in nature, it must first be colonized by a
compatible fungal symbiont (Rasmussen 1995). Orchid plants receive both nutrients (e.g.
phosphorous) and carbohydrates from their fungal partners (Smith and Read 2008). The
intracellular hyphal coils, i.e. pelotons, occur in the cortical cells of roots and are
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characteristic of orchid mycorrhizae. Pelotons are known to be digested and utilized by
the orchid (Senthikumar and Krishnamurthy 1998). Orchid mycorrhizae are unusual in
that mycorrhizal symbiosis is considered necessary for germination and early
development in a majority of orchid taxa, whereas most other types of mycorrhizae form
after early plant development (Rasmussen 1995, Smith and Read 2008). Orchids produce
microscopic seeds with undifferentiated embryos and a small amount of concentrated
nutrient reserve. A germinating orchid seed is often considered fully mycotrophic until it
acquires photosynthetic capability; adults are typically mixotrophic utilizing both
photosynthesis and mycotrophy simultaneously to acquire carbon. Conversely, non-
photosynthetic orchid taxa remain mycotrophic throughout their life and are thus
categorized as holomycoheterotrophic.
A majority of fungi that form orchid mycorrhizae belong to the phylum
Basidiomycota with few belonging to Ascomycota (Dearnaley 2007). Globally, a large
majority of the reported orchid mycorrhizae belong to the basidiomycete families
Tulasnellaceae, Sebacinaceae, and Ceratobasidiaceae (Shefferson et al. 2005, Dearnaley
2007). It is thought that orchid-fungal associations may generally be broad in
photosynthetic orchid species, especially when they are compared to
holomycoheterotrophic genera such as Hexalectris (McCormick et al. 2004, Taylor and
Bruns 2002, Pandey et al. 2013). However, exceptions include several species in the
photosynthetic genus Cypripedium, which exhibit high specificity by associating only
with narrow clades within the Tulasnellaceae (Shefferson et al. 2005; Shefferson et al.
2007).
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In addition to carbon acquisition strategies, geographic range, and plant species
abundance may also play a role in symbiont specificity and diversity (McCormick et al.
2004). Despite their wide geographic ranges, several orchid taxa are reported to associate
with relatively narrow clades of fungi. For example, three fairly widespread
photosynthetic genera native to North America, Liparis lilifolia, Goodyera pubescens,
and Tipularia discolor were documented associating with a low diversity of fungi
(McCormick et al. 2004). In this study, 53 fungal isolates from root fragments and
protocorms collected between 1997 and 2001 yielded lower or equal mycorrhizal fungal
diversity in comparison to the nonphotosynthetic orchid Cephalanthera austinae which is
known to exhibit high specificity (McCormick et al. 2004). The use of widely distributed
fungi by an orchid might enable wide plant distributions regardless of mycorrhizal
specificity. When 13 plants of Pheladenia deformis were sampled from 9 locations across
≥2000 km in Western Australia and Victoria, all except one of the 26 fungal isolates
grouped in a single operational taxonomic unit (OTU) within the genus Sebacina (Davis
et al. 2015). However, conclusions about specificity of orchid mycorrhizal partners may
be inaccurate when only cultured fungi are included in the analyses. And while it is
difficult to closely compare studies that utilize different techniques, broad comparisons
among studies may be made based on the genetic similarity of OTUs.
The narrowly distributed and endemic orchid Piperia yadonii which was shown to
be generally non-specific in its mycorrhizal associations, although it exhibits variation in
associations because of habitat (Pandey et al. 2013). Along with geographic variation,
evidence for seasonal variation in mycorrhizal associations exists for some
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photosynthetic terrestrial species although this subject largely remains understudied
(Ercole et al. 2014). Six different individuals were sampled at five sampling events
within a single population of Anacamptis morio. The species was recorded associating
with the genus Tulasnella in the autumn and winter whereas Ceratobasidium was much
more common in the summer (Ercole et al. 2014).
Considering that a majority of orchid taxa are rare, conservation activities,
including propagation and reintroduction of in vitro propagated plants, are utilized to
improve the conservation status of a species. Because mycorrhizal fungi are inseparable
from orchid biology, it is necessary to consider the mycorrhizal specificity and the ability
of propagated plants in forming partnerships with suitable fungi. Some options to
maximize fungal compatibility in the propagated plants include: (1) introducing
asymbiotically propagated seedlings directly into their natural habitats; (2) first
corroborating and isolating fungi that the orchid taxon prefers in its natural habitat and
then utilizing the isolates in symbiotic propagation; (3) generating asymbiotically
propagated seedlings that are further raised in greenhouse conditions and then
introducing them in the wild habitat. There is some concern with the third option that the
orchid may form mycorrhizal associations with fungal strains that are not available in the
natural habitat and that ‘foreign’ strains could be introduced into natural habitats as a
result. While the orchid plants may succeed after introduction, the mycorrhizal fungal
strains might disturb the microbial community dynamics in the soil otherwise. Because
the transplant success of asymbiotic seedlings is low, and symbiotically propagated plants
are also exposed to greenhouse environments and thus likely become mycorrhizal with
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additional fungi even if sterile medium is used initially, the third option appears equally
viable. However, the most important step that researchers can include is to confirm the
similarity or differences in the fungi that form mycorrhizae with orchids in cultivation
and in nature to inform the re-introduction decisions. Such comparisons, however, are
very limited so far. One study reported that orchid plants, including P. chapmanii,
propagated in sterile culture and subsequently grown in greenhouse environments can
become mycorrhizal with orchid mycorrhizal fungi (Richards and Sharma 2014). The
organic components (peat, milled sphagnum, and fine fern fiber) of the substrates used
for cultivating Platanthera species native to bog habitats likely contain saprophytic fungi
including those that can form orchid mycorrhizae. Whether the orchid plants utilize the
same or similar mycorrhizal symbionts in an ex situ environment that they associate with
in their native habitats, however, has not been studied empirically. In a preliminary study,
Richards and Sharma (2014) reported peloton forming fungi belonging to the
Tulasnellaceae within the roots of asymbiotically cultured plants of P. chapmanii that
were cultivated in a peat-based substrate for >1 year. This bog-orchid substrate was
composed of 44% peat moss, 26% milled sphagnum, 20% sand, and 10% fine tree fern
fiber (v:v:v:v). The same study by Sharma and Richards (2014) also reported
Tulasnellaceae in the roots of naturally occurring plants. However, the phylogenetic
relationship between peloton forming fungi from plants cultivated in the greenhouse
directly after sterile in vitro culture and those that occur naturally in their wild habitats
were not reported. The temperate terrestrial orchid taxon, P. chapmanii, was used to test
the hypothesis that plants obtained via asymbiotic germination methods and subsequent
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greenhouse culture will have a different suite of mycorrhizal fungi when compared to
plants of the same species occurring in their natural habitat. Platanthera chapmanii is a
suitable model for testing the study questions because it responds relatively quickly to
both in vitro and greenhouse culture conditions. To test the hypothesis, influence
of growing environment (i.e., greenhouse cultivated plants that were never exposed to the
natural habitat and those occurring in their natural habitat) on mycorrhizal fungal
community composition was quantified. Further, whether the mycorrhizal communities
within the roots of P. chapmanii exhibit phenological variation within each of the two
growing environments (i.e., ex situ or native habitat) was tested.
Materials and methods
Study species
Platanthera chapmanii is a temperate terrestrial orchid native to North America.
The rare perennial occurs in mesic and wet pine flatwoods, barrens, and savannas in
sandy loam soils. Its disjunct populations occur in southern Georgia, northern Florida and
southeastern Texas within the United States (Liggio and Liggio 1999; Poole et al. 2007;
Richards and Sharma 2014). Populations are often small with ≤10 individuals, and so far
only one population is known to host ≥100 plants (Richards and Sharma 2014).
Individuals of P. chapmanii typically emerge above-ground between late February and
early March, and flower from late July to early August when they produce a single
raceme with ≥60 orange flowers. Seeds dehisce from mid to late October when above-
ground organs begin to senesce.
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Root collection and surface sterilization
Roots were collected between 2012 and 2014 from ex situ cultured plants and
naturally occurring plants (Figure 4.1, Richards and Sharma 2014). Each sampling event
was considered an experimental treatment.
In a preliminary study in November 2012, roots representing greenhouse culture
conditions (GF12) and natural habitat conditions (NF12) were sampled to develop
species-specific methods for molecular identification of orchid mycorrhizal fungi. In
November 2014, up to 30 cm root tissue from each of five ex situ cultured plants (GF14)
and each of six individuals from a natural population in southeastern Texas (NF14) was
collected. To compare the fungal species composition in fall (November) and spring, up
to 12 cm root tissue was collected from each of the four sampled individuals from the
same native population in March 2015 (NSp15). Greenhouse cultured plants were not
sampled in March 2015 because the group of plants that were sampled in November 2014
were being vernalized at 4°C in a refrigerator at this time and would not have had the
opportunity to associate with new fungi between November 2014 and March 2015 while
being stored at 4°C.
Subsequently, in August 2015, 15 greenhouse cultured plants were sampled (GSu15)
by collecting up to 10 cm of tissue from each individual. At this time, between 12 and 18
cm of root tissues from eight plants at the same native population (NSu15) that was
previously sampled in November 2012, November 2014, and March 2015 was collected.
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At each sampling event, roots were stored on ice immediately after collection and
transported to the laboratory. Within 48 hours of collection, roots were washed free of
soil and other debris and examined for the presence of pelotons (Figure 4.2). Peloton
containing roots were surface sterilized by exposing the tissues to: 1) a 30s rinse in 70%
ethanol, 2) a 30s rinse in 0.6% sodium hypochlorite, and 3) a 30s rinse in 70% ethanol.
The root pieces were then washed with sterile ultrapure water until they were free of the
residues of ethanol and sodium hypochlorite. After surface sterilization, the epidermis
was removed before the roots were divided into smaller (~3 cm) pieces for further
processing. Finally, individual root segments were finely minced and stored at -80oC until
DNA was extracted. Additionally, culture of peloton-forming fungi from roots collected
from the native habitat in August 2015 was attempted. The root fragments that were used
to culture fungi on nutrient medium were surface-sterilized and processed similarly
except instead of ultralow freezing, the finely macerated tissue was suspended in molten
potato dextrose agar (PDA) contained within a 14 cm petri dish. The plates were
examined every 1 to 3 days for growth of fungi with characteristics of orchid mycorrhizal
fungi. Hyphae from actively growing fungi were collected and cultured in individual
plates to obtain pure cultures.
DNA extraction, PCR amplification and PCR product cleaning
Deoxyribose nucleic acid (DNA) was extracted from each sample by using the
DNeasy Plant Mini Kit (Qiagen, Germany) and protocol with a few modifications. Prior
to lysing the tissues, samples were dehydrated with liquid nitrogen, and immediately
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lysed in a tissuelyser for 3 min at 30 disruptions / sec. After this initial lysis, 400 µl of
3.3% solution of polyvinylpyrrolidone (PVP) in AP1 lysis buffer was added to each
sample. Further, incubation time during the cell disruption step was increased to 2 hours
at 65°C during which the samples were shaken and vortexed every 30 minutes.
Deoxyribonucleic acid was not extracted from the cultured fungi; instead, fungal
mycelium from pure cultures was examined and presence of moniliod cells was
confirmed prior to subjecting the fungus to direct Polymerase Chain Reaction (PCR)
(Figure 4.3).
Polymerase chain reaction of the nuclear ribosomal internal transcribed spacer region
(nrITS) region was accomplished using the primer pairs ITS1-OF / ITS4-OF or ITS1 /
ITS4-Tul (Taylor and McCormick 2008, Sigma-Aldrich, Missouri, USA). Each 25 µl
reaction was prepared using Promega GoTaq Flexi DNA polymerase reagent kit using 4
µl of DNA (Promega, Wisconsin, USA). The thermocycling profile included: initial hold
for 2 mins at 95°C followed by 35 cycles of: denaturation of 45s at 94°C, annealing at
either 52°C or 58°C for 45 seconds, and extension at 72°C for 2 minutes. After the 35
cycles, the reactions underwent a final extension at 72°C for 5 minutes (Pandey et al.
2013). A positive and a negative control for each set of PCR reactions that were prepared
and run together was used.
A 2% agarose gel electrophoresis was carried out to verify amplification by using 4
ul of PCR product. Ethidium bromide was used as the florescent dye, and a 1 kb ladder
was used to estimate fragment size. Samples that showed a clear, single band between
600-800 bp were cleaned using DNA Clean and Concentrator 5 kit (Zymo Research,
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Irvine, USA). The concentration (ng/µl) and quality of the cleaned product was measured
in NanoDrop 2000c (Thermo Scientific, USA). Samples that showed multiple bands or
bands that were wide and / or unclear underwent a gel extraction protocol after
performing electrophoresis with 21 to 25 µl PCR product. The desired bands (~600-800
bp) were then extracted from the gel and cleaned using the Genelute gel extraction kit
(Sigma-Aldrich, St. Louis, Missouri). Sequencing reactions were prepared and sent for
sequencing at the DNA Analysis Facility on Science Hill at Yale University (New Haven,
CT).
Data analyses
Editing and assembly of the raw sequences was performed with CodonCode
Aligner version 6.0.2 (CodonCode Corporation, Centerville, Massachusetts). Sequences
were trimmed at both ends by removing 25 bp sections that had ≥3 bases with phred
scores below 20. If the resulting trimmed sequences were shorter than 400 bp the entire
sequence was excluded from the analyses. Subsequently, the following procedures were
performed on the 122 sequences that passed quality control filters.
The taxonomic identity of each sequence was determined at family level by using
BLAST (NCBI Genbank, http://www.ncbi.nlm.nih.gov/genbank/). The FASTX-Toolkit
version 2.0 was used to trim sequence ends so that the final length of all sequences was
455 bp (Gordon and Hannon 2010). SWARM version 2.5 was used to build OTUs (d =
12) without data dereplication (Dales 2000). The sequence that was most abundant in an
OTU was used as the representative sequence for that OTU.
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To estimate sequence distances within the Tulasnellaceae, sequences were first
aligned using MUSCLE in MEGA version 6 (Tamura et al. 2013). Mean pairwise
distances among all sequences within the family were calculated using Kimura’s 2-
parameter model and a gamma distribution. Further, pairwise distances were calculated
separately for sequences originating from plants from the greenhouse environment, the
natural environment, and each sampling event within the two categories.
Cumulative, rarefied OTU diversity curves were constructed by including all 18
OTUs in EstimateS version 9.1.0 with sample-based incidence, individual-based
abundance and observed methods (Colwell 2006). Shannon and Simpson diversity
indices were calculated for each treatment with RStudio version 8 using the Phyloseq
package (RStudio Team).
A principal component analysis (PCA) was performed based on the abundances
of OTUs in each of the seven treatments using a correlation cross-products matrix in SAS
version 9.4 (SAS Institute Inc. 2013). A Fisher’s exact test and Kruskal-Wallis tests were
performed to evaluate whether the abundances of the OTUs were significantly different
among the treatments. Phylograms were generated separately for Tulasnellaceae and
Ceratobasidiaceae by using the self-generated OTU data and previously published
sequences of orchid mycorrhizal fungi from NCBI GenBank
(http://www.ncbi.nlm.nih.gov/genbank/) to place fungal OTUs from P. chapmanii in the
context of known orchid mycorrhizal fungi. Reference sequences and OTU representative
sequences were aligned by using MUSCLE in MEGA version 6. Maximum-likelihood
trees were then constructed in MEGA using Kimura’s 2-parameter model, as it was
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estimated to be the best fit for the data. Support values were estimated via 1000 bootstrap
replications. Bayesian trees were also constructed for both families using MrBayes
version 3.2.6 with 1 million generations until the standard deviation between two runs
was <0.01. Trees from the initial 250,000 generations were discarded (Huelsenbeck and
Ronquist. 2001). The Ceratobasidiaceae maximum likelihood and Bayesian trees were
rooted with Sistotrema sp. Because of the accelerated diversification within the
Tulasnellaceae nuclear ribosomal region, the conserved 5.8s region was extracted from
the full ITS sequences by using ITSx software version 1.0.11 (Bengtsson-Palme et al.
2013) before phylograms were constructed. Similar tree building protocol was followed
as described above except the 5.8s Tulasnellaceae tree was midpoint rooted due to a lack
of a defensible related outgroup. Using FigTree version 1.4.2, nodes were arranged in an
increasing order and topology was transformed to a cladogram (Rambaut 2007). Because
the topologies were similar for both types of trees for each fungal family, Bayesian
probability values along with bootstrap values ≤50 were included in the maximum-
likelihood trees.
Results
Eighteen fungal OTUs representing two fungal families, Tulasnellaceae and
Ceratobasidiaceae, were identified from the roots of P. chapmanii across all sampling
events. Of the 18 OTUs, 17 belonged to the Tulasnellaceae and one to the
Ceratobasidiaceae (Table 4.1; Table 4.2). Tulasnellaceae was represented by 121 (99.2%)
of the 122 sequences. Among the 18 OTUs, 5 were singletons. Across all treatments,
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44.4% of the OTUs were observed in multiple plants (Table 4.2). None of the sampled
plants hosted fungi belonging to multiple families (Table 4.2). Ceratobasidiaceae was
observed only in one individual in the NSp15 treatment, while the NSu15 treatment had
the highest fungal richness with eight OTUs, three of which were exclusive to that
treatment. Treatments GF12 and NF12 were represented exclusively by one OTU each.
The most frequently encountered OTU (T8) was observed in 11 naturally occurring
individuals representing three sampling events (NF14, NSp15, and NSu15).
The mean pairwise sequence distance among P. chapmanii mycorrhizal fungal
sequences in this study was 0.094 ± 0.031. The mean pairwise sequence distance among
the ex situ cultured plants (π = 0.064 ± 0.013) was lower than the mean distance observed
among the Tulasnellaceae obtained from plants growing in their native habitat (π = 0.116
± 0.036) (Table 4.3). Of the greenhouse treatments, GF14 had the largest mean pairwise
distance (π = 0.047 ± 0.009), whereas among the sampling events from naturally
occurring populations, the largest mean pairwise distances were within NSp15 (0.228 ±
0.065) and NSu15 (0.209 ± 0.035). These two sampling events also represented the
highest mean pairwise distances across all treatments (Table 4.3). When compared to
other orchid species, the mean pairwise sequence distance among mycorrhizal fungal
sequences of P. chapmanii from plants growing in their natural habitat (π = 0.116 ±
0.036) was larger than some widely distributed temperate terrestrial orchids (e.g.
Cypripedium japonicum and Cypripedium candidum) and smaller than others (e.g.
Piperia yadonii and Ophrys fuciflora) (Table 4.4).
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The diversity curves estimated a higher OTU count at a larger sample size of up
to 500 sequences. At the extrapolated value of 500 sequences, the slopes of both the
sample-based and individual-based OTU diversity curves started to level but the slope did
not reach zero (Figure 4.4). Both Shannon and Simpson indices showed that NSp15 and
NSu15 hosted the highest fungal diversity with Shannon index values of 1.30 and 1.55,
respectively, and Simpson index values of 0.72 and 0.71, respectively. Of the ex situ
treatments, GSu15 was the most diverse (Shannon’s index of 1.14 and Simpson’s index
of 0.63).
The PCA showed that 60% of the variation among treatments was explained by
PC1 and PC2 (Figure 4.5). Principal component one was negatively correlated with T3,
T4, T5, T6, and T7 (all eigenvector values = -0.33) and most positively correlated with
T13, T14, and T15 (all eigenvector values = 0.26). Principal component three accounted
for 20.5% of the variation in the data and was most negatively correlated with T1 (all
eigenvector values = -0.22) and most positively correlated with T16 (all eigenvector
values = 0.41). Four of the seven treatments clustered together in the center of the
scatterplot (GF12, NSp15, NF12, and NF14) while the remaining three (GF14, NSu15,
and GSu15) separated from the cluster as individual points (Figure 4.5). Results from the
Fisher’s exact test and Kruskal-Wallis analysis of variance did not support differences in
OTU abundances among the seven treatments. The Fisher’s exact test yielded a P-value
of 1, and the P-value from the Kruskal-Wallis test was 0.423.
Maximum likelihood and Bayesian trees showed similar relationships among the
fungal OTUs. The bootstrap values that exhibited weak branch support values (≤50) were
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not considered. With this criterion, thirteen of the 17 OTUs representing both sources of
plants (GF12, GF14, GSu15, NF14, NSp15, and NSu15) grouped together in the same
clade. These OTUs grouped with Tulasnella sp. from the European orchid A. morio and
Tulasnellaceae from the North American species Goodyera pubescens, Cymbidium
faberi, Platanthera Praeclara, and Tipularia discolor among others (Figure 4.6). Apart
from the main clade that included 13 OTUs, two OTUs representing mycorrhizal fungi
from the greenhouse environment grouped together in a separate monophyletic clade next
to uncultured Tulasnellaceae from Epidendrum firmum. The remaining two OTUs
representing fungi from naturally occurring plants separated in the farthest clade closer to
uncultured Tulasnellaceae from Paphiopedilum dianthum and E. firmum (Figure 4.6).
The single Ceratobasidiaceae OTU segregated in a node next to a species of uncultured
Ceratobasidiaceae from the endemic North American orchid Piperia yadonii (Figure 4.7).
Discussion
Mycorrhizal specificity among the Orchidaceae is variable across the family
depending on the life history or distribution of a taxon (Arditti et al. 1990; Masuhara et
al. 1993; McKendrick et al. 2002; Selosse et al. 2002; Otero et al. 2004; Pandey et al.
2013; Bonnardeaux et al. 2014; Ercole et al. 2014; Jacquemyn et al. 2015). Considering
that there are approximately 35,000 taxa distributed across the planet, broad
generalizations are difficult to make and are likely inappropriate until a majority of the
taxa are studied extensively. Majority of the orchid species are rare in their occurrence
and thus are the target of conservation efforts including augmentation and restoration of
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natural populations. Because orchid mycorrhizal fungi are necessary for germination and
development of plants, consideration of fungal specificity and maintaining the integrity of
local genotypes becomes significant. While orchid mycorrhizal fungi are free-living
saprophytic fungi with cosmopolitan distributions, orchid-fungus partnerships can be
highly specific (Selosse et al. 2002; McCormick et al. 2004; Shefferson et al. 2005;
Shefferson et al. 2007; Nomura et al. 2013) or general (Pandey et al. 2013; Bonnerdeaux
et al. 2014) depending on the life history of an orchid taxon and its adaptive traits.
It is reported that a rare terrestrial temperate orchid with small, disjunct
populations distributed across a wide geographic area in the southeastern United States is
specific in its mycorrhizal relationships with fungi from a few clades within the
Tulasnellaceae (Tables 1, 2, 3, Figure 4.6) regardless of the growing substrate and
environment. Only a single OTU belonging to the Ceratobasidiaceae was recovered
across all sample sources and events (Table 4.2). Further, the orchid mycorrhizal
relationship remained specific over time through phenological stages and years in both
the naturally occurring plants and in plants that acquired mycorrhizal fungi during
greenhouse culture after sterile in vitro germination and development (Table 2, Figure
4.6). Because seeds from the same natural population that was studied for variation in
natural fungal diversity were used, the possibility that the genotype or ecotype of the
orchid was a factor in dictating the mycorrhizal diversity associated with P. chapmanii in
two disparate growing environments can be largely ruled out. It is possible though that if
other natural populations of P. chapmanii are studied similarly, regional variation in
mycorrhizal diversity and specificity patterns might emerge. The rarefaction curves
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generated with data originating from the experiment predicted that more OTUs would be
discovered by increasing the sample size, which could increase the possibility of
recovering additional OTUs from the Tulasnellaceae or fungi from other lineages (Figure
4.4). Considering that the fungal OTUs associated with P. chapmanii roots in all seven
treatments, including the greenhouse environment approximately 1160 km from the
natural population in Texas, showed phylogenetic similarity, it is unlikely that additional
sampling of the same population will increase mycorrhizal diversity beyond the
Tulasnellaceae. It is, however, possible that the particular plant genotype (or ecotype)
used to test the hypothesis is more specific toward the Tulasnellaceae from the clades
identified, and that other natural populations (in GA, for example) and their progeny form
mycorrhizae with fungi from additional lineages. Spatial variation in mycorrhizal
diversity has been previously documented in other temperate terrestrial orchids
(Masuhara and Katsuya 1994; Taylor and Bruns 1999; Shefferson et al. 2005; Pandey et
al. 2013). Platanthera chapmanii was observed with small mycorrhizal diversity
differences and high specificity regardless of growing environment.
When mycorrhizal fungal sequences obtained from the naturally occurring P.
chapmanii were compared with laboratory and greenhouse cultured, higher diversity was
observed in naturally occurring fungi (Table 4.3). Although differences in pairwise
distances exist among the seven treatments, the differences are slight and among small
numbers (Table 4.3). When compared to other temperate terrestrial orchids with a wide
geographic range including Platanthera praeclara, Anacampis laxiflora, Ophrys
fuciflora, the mean pairwise sequence distance within the Tulasnellaceae sequences from
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the naturally occurring P. chapmanii was average to low (Tables 3, 4). Similarly, a
congeneric species, Platanthera praeclara, which has a wide geographic range in North
America associated with fungal species of Tulasnellaceae. Mean pairwise distance (0.135
± 0.006) among P. praeclara-associated Tulasnellaceae was higher than in P. chapmanii
(0.116 ± 0.036; Table 4.4). A narrowly endemic relative, Piperia yadonii, also exhibited
higher mean pairwise distance within the family Tulasnellaceae (0.231 ± 0.026) than
fungi from either P. chapmanii or P. praeclara while it also associated with fungi from
Ceratobasidiaceae and Sebacinaceae (Pandey et al. 2013). The majority of the species
mentioned above are considered as having high specificity towards their mycorrhizal
associations. Since fungi from P. chapmanii follow the same pattern, there is evidence for
high mycorrhizal specificity in the taxon.
Although a narrow phylogenetic breath of fungi was observed among all the
sampling treatments, temporal variation in OTU richness and evenness among sampling
events was observed in this study (Table 1, Figure 4.4). The abundance-based PCA also
suggests temporal variation in the fall and summer sampling events from the ex situ
environment (GF14 and GSu15), whereby these two treatments segregated independently
from the cluster of all treatments except NSu15 (Figure 4.5). Five exclusive OTUs were
observed (T3, T4, T5, T6, T7) in GF14, whereas GSu15 had two (T16, T17) of the four
OTUs exclusive to the sampling event while the remaining two were shared with NSu15.
Similarly, NSu15 segregated from all other samples originating from the natural
population (NF12, NF14, and NSp15) and was the most OTU-rich (8 OTUs) treatment
hosting three exclusive OTUs (T12, T13, T14). The clustering observed in the
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103
abundance-based PCA in addition to the variation among the treatments observed in the
mean pairwise distances indicate slight temporal variation within each environment
(Table 1, Figure 4.5). This is also supported by both Shannon and Simpson indices; the
fall treatments hosting lower diversity (GF12, NF12, GF14, and NF15) clustered together
while the spring and summer samples hosting higher diversity (GSu15, NSp15 and
NSu15) separated. However, because 13 of the 17 Tulasnellaceae OTUs clustered
together in the same clade of the maximum likelihood tree, this variation appears to be
among closely related species of fungi (Figure 4.6). Additionally, the abundance of the 18
OTUs was different among the seven sampling events according to the Fisher’s exact test
(p = 1.00) and the Kruskal-Wallis (p = 0.42). Altogether, strong evidence for sampling-
event based difference was not detected. This was not the case with Tovar 2015 or Ercole
et al. 2014. Ercole et al. (2014) observed large seasonal differences of fungi associating
with A. morio. Tovar (2015) observed some spatial and temporal differences in fungi
associating with P. praeclara. Both studies showed much larger temporal variation than
was observed in P. chapmanii. Based on the data, P. chapmanii exhibits lower temporal
and spatial variation of mycorrhizal fungal diversity than either A. morio or P. praeclara.
Because of a lack of documentation, further temporal comparisons to P. chapmanii
cannot be made at this time.
The current paradigm suggests that high mycorrhizal specificity is not limited to
nonphotosynthetic orchids but is also present in many photosynthetic terrestrial orchids
(Selosse et al. 2002; McCormick et al. 2004; Shefferson et al. 2005; Shefferson et al.
2007; Nomura et al. 2013). With some exceptions (Pandey et al. 2013; Bonnerdeaux et al.
Texas Tech University, Kirsten Poff, August 2016
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2014) temperate terrestrial orchids have been documented associating with fungi that
exhibit narrow phylogenetic breadth (Selosse et al. 2002; McCormick et al. 2004;
Shefferson et al. 2005; Shefferson et al. 2007; Nomura et al. 2013). In the current study, a
high abundance of mycorrhizal fungi from the same or closely related clades within the
Tulasnellaceae was observed regardless of whether the species was cultured ex situ or
sampled from its natural habitat. With some evidence for temporal and growing
environment variation, the observed orchid mycorrhizal diversity within the roots of P.
chapmanii generally remains low across phenological stage and year of sampling. The
data also clearly suggest that that the mycorrhizal fungi that P. chapmanii prefers in its
natural habitat are widely distributed and available in peat-based greenhouse substrate.
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Table 4.1. Representative sequence of each of the 18 fungal nrITS-based operational taxonomic units (OTUs) identified within the
roots of Platanthera chapmanii plants cultured in vitro/greenhouse and those occurring naturally. Each culture condition was sampled
three to four times between 2012 and 2015. The first letter of an OTU name represents the fungal family to which the OTU belongs:
T, Tulasnellaceae; C, Ceratobasidiaceae. Values in parentheses are the total number of plants in which a specific OTU was
documented.
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Fungal OTU
Trimmed sequence
T1 >PCG1_ITS4_TUL
tacaaccggtagcgctggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagttgttgaa
cgcattgcaccgcgccctaaaccggctgcggtatgcccctttgagcgtcattacatccttcgggagtctccttttctggagacccgagttcggagtcctcggtcccttgggatcgtgt
tctctcagatgcatcgcgccgatcgctttgatgggtactctaatgcctgagcgtggagtccctctgaagtcgagacgcgtttgaccgggtggtgagcccgtgtcggcaagtccacg
tccgctgcgacgtcggtactacaaccacatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccaggatc
T2 >PCN1_ITS4_TUL
tacgtgtcttgtagactctgatgataagaaatacaaccagtagcactggatccctcggcatgccattcgatgaagaccgtagcgagttgcgataagcgatgtgatatgcgagtccca
aactgatacgtgaaccatcggatcgtcgaacgcactgcaccgaggattgcccatccacggtataccacattgagtgtcattattcgttcgtctctgacgagttcggggtccacggcc
ttgccgcgttccctcagattgaagtctgtggcgtcaacctgaccttgctagtgtctgtcgagccccctttgacgagttcactgggtacgctacgtccgcaccacaggtcggtctggc
cgggacgcttgcgtccaaccgttctctaatgatgacctcacggtggtaagattacccgctaaacttaagcatattaatcagcggaggaaaagaaactaacaagg
T3 >PchG1-1-ITS4-Tul
taacacttttacaaccggtagcgctggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgag
ttgttgaacgcactgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtatcccttcgggagtcttttcgttaagacccgagttcggagtcctcggtcttcggatc
gtgttctcttagatgcgtcgcgccgatcgcctgatgggtcactctaatgcctgagcgtggagtccctcggagctgagaggcgcttgaccgagtgttgagctcgcgtcgccaagtcc
gcacgtcttggacgtcggtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccagg
T4 >PchG3-5-ITS4-Tul
tccgcgttgtgagtctaacaccagttgtaacacttttacaaccggtagcgctggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaag
tccaccacttatacgtgaatcatcgagttgttgaacgcactgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtatcccttcgggagtcttttcgttaagaccc
gagttcggagtcctcggtcttcggatcgtgttctcttagatgcgtcgcgccgatcgcctgatgggtcactctaatgcctgagcgtggagtccctcggagctgagaggcgcttgacc
gagtgttgagctcgcgtcgccaagtccgcacgtcttggacgtcggtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaa
T5 >PchG3-6-ITS4-Tul
ttttacaaccggtagcgatggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagttgttga
acgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtattccttcgggagtcctttttccaaaggaccggagttgggattcttggttctttggatcgtgttct
cttagatgtgtcacaccgatcgcctgatgggtcctctaatgcctaagcgtggagttcctgaaagtctgagacgtgcttgaccgggtcttgagctcgcgtcaccaagtctgcctaaca
agcagtactacaacacatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccaggatccctcag
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Fungal OTU
Trimmed Sequence
T6 >PchG4-1-ITS4-Tul
ttttacaaccggtagcgatggtcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagttgttgaa
cgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtaatccttcgggagtccttttaactaaggacccgagttcggagtcctcggtcctctggatcgtgtt
ctcttagatgcgtcgcaccgatcgcctgatgggtcctctaatgcctaagcgtggagttccttcagagtccgaaacgtgcttgaccgggtgttgagctcgcgtcaccaagtctgccta
accagcagtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaactaggatccctca
T7 >PchG4-5-ITS4-Tul
aaactttttacaaccggtagcgatggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtcccccacttatacgtgaatcatcgagt
tgttgaacgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtaatccttcgggagtccttttaactaaggacccgagttcggagtcctcggtcctctgg
atcgtgtttttttagatgcgtcgccccgatcgcctgatgggtcctctaatgcctaagcgtggagttccttcagagtccgaaacgtgcttgaccgggtgttgagctcgcgtcaccaagt
ctgcctaaccagcagtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaactaggat
T8 >PchN1-1-ITS4-Tul
aaactttttacaaccggtagcgatggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagt
tgttgaacgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtattccttcgggagtccttttccaaggacccgagttcggagtcctcggtcccacctgg
atcgtgttctctcagatgcgtcgcaccgatcgcctgatgggtcctctaatgcctaagcgtggagttccttcagagtcccacacgtgcttgaccgagtcttgagctcgcgtcaccaagt
ccgccttaaccagcggtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccagga
T9 >PchN3-1-ITS4-Tul
agttgtataaactttttacaaccggtagcgatggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatc
atcgagttgttgaacgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtattccttcgggagtccttttccaaggacccgagttcggagtcctcggtcc
cacctggatcgtgttctctcagatgcgtcgcaccgatcgcctgatgggtcctctaatgcctaagcgtggagttccttcagagtcccacacgtgcttgaccgagtcttgagctcgcgtc
accaagtccgccttaaccagcggtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaact
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Fungal OTU Trimmed Sequence
T10 >PCHSN1-1-ITS4-OF
ccgtgttactcctcgtgagcacacgctaaagagcgatatacgtgtcttgtagactctgatgataagaaatacaaccagtagcactggatccctcggcatgccattcgatgaagaccg
tagcgagttgcgataagcgatgtgatatgcgagtcccaaactgatacgtgaaccatcggatcgtcgaacgcactgcaccgaggattgcccatccacggtataccacattgagtgt
cattattcgttcgtctctgacgagttcggggtccacggccttgccgcgttccctcagattgaagtctgtggcgtcaacctgaccttgctagtgtctgtcgagccccctttgactgagttc
actgggtacgctacgtccgcaccacaggtcggtctggccgggacgcttgcgtccaaccgttctctaatgatgacctcacggtggtaagattacccgctaaact
T11 >PchNs3-3-ITS4-TUl
tacaaccggtagcgatggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagttgttgaa
cgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtattccttcgggagtccttttacaaggacccgagttcggagtcctcggtcctcttctggatcgtgt
tctcttagatgcgtcgcaccgatcgcctgatgggtcctctaatgcctaagcgtggagttccttcagagtccgagacgtgcttgaccgggtcttgagctcgcgtcaccaagtctgccg
taacaagcagtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccaggatccctca
T12 >PchNsu2-3-ITS4-TUl
actttttacaaccggtagcgctggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtaaatcatcgagttgt
taaacgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtatcccttcgggagtcctttacaaggacccgagttcggagtcctcggtccgacctggatc
gtgttctcttagatgcgtcgcaccgatcgcctgatgggtcctctaatgcctaagcgtggagttccttcagggtccgacacgtgcttgaccgggtcttgagcctgtgtcaccaagtctg
cccaacaagcagtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccaggatccc
T13 >PchNsu8-1-ITS4-TUL
taaagaacgttccgcattgtgagtctaacaccagttgtaaacttttacaaccggcagcgctggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtg
atgcgcaagtccaccacttatacgtgaatcatcgagttgttgaacgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtattccttcgggagtctttcctt
gcgaaagacccgagttcggagtcctcggtcttttggatcgtgttctcttagatgcgtcgcgccgatcgcctgatgggtactctaatgcctgagcgtggagtccctctgggtttgagac
gcgcttgaccggccattgggctcgcgtcaccaagtccacatcctttgggatgctggtactacaacgcatgacctcatcggggtaggacaacccgctaga
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Fungal OTU Trimmed Sequences
T14 >PchNsu8-4-ITS4-TUL
atggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagttgttgaacgcactgcaccgcg
ccctaaaccggctgcggtatgcccctttgagcgtcattgcattccttcgggagtctcctttcgagagacccgagttcggagtcctcggtcctctggggaccgtgttctctcagatgcg
tcgcgccgatcgcccgatgggtcgctctcatgcctgagcgtagagtccctctggagtcgagacgcgctggaccgggcgtttgggcccgccgtcgccaagtccgacccgatgct
tttctgcatgggtttcggtactacaaccacatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccaggatcc
T15 >PchNSu8-9-ITS4-TUL
tttacaaccggtagcgatggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagttgttga
acgcattgcaccgcgccctaaaccggctgcggtatgcccctttgagcgtcattgtattccttcgggagtcctttttacaaaggacccgagttcggagtcctcggtcctctggatcgtg
ttctcttagatgcgtcgcaccgatcgcctgatgggtctctaatgcctaagcgtggagttccttcagagtccgagacgtgcttgaccgggtgttgagctcgcgtcaccaagtctgcctc
acaagcagtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccaggatccctcag
T16 >Pch.253-1-ITS4-TUL
caaccggcagcgctggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagttgttgaacg
cattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtattccttcgggagtctttccttgcgaaagacccgagttcggagtcctcggtcttttggatcgtgttct
cttagatgcgtcgcgccgatcgcctgatgggtactctaatgcctgagcgtggagtccctctgggtttgagacgcgcttgaccggccattgggctcgcgtcaccaagtccacatcct
ttgggatgctggtactacaacgcatgacctcatcggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccaggatcccc
T17 >Pch.255-1-ITS4-TUL
caaccggcagcgctggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagttgttgaacg
cattgcaccgcgccctaaaccggctgcggtatgcccctttgagcgtcattgtattccttcgggagtctttccttgctgaaagacccgagcttggagtcctcggtcctttggatcgtgtt
ctctcagatgcgtcgcgccgatcgcctgatgggtactctaatgcctgagcgtggagtcccttcgagtttgagacgcgcttgaccggccgttgggctcgcgtcaccaagtccgcgt
cctcctggacgtcggtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccaggatccc
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Fungal OTU Trimmed Sequence
C1 >PCHSN3-1-ITS4-OF
tgtgagacggattgaccctctcgggggacggtccgtctattatacataaactccaataattaaatttgaatgtaatttgatgtaacgcatctttgaactaagtttcaacaacggatctctt
ggctctcgcatcgatgaagaacgcagcgaaatgcgataagtaatgtgaattgcagaattcagtgaatcatcgaatctttgaacgcaccttgcgctccttggtattcctcggagcatg
cctgtttgagtatcatgaaattctcaaaataaatcttttgttaactcgattgattttattttggacttggaggtctgcagattcacgtctgctcctcttaaatttattagctggatctctgtgacat
cggttccactcggcgtgataagtatcactcgctgaggacactgtaaaaggtggccggagttactgaagaaccgcttctaatagtccattg
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Table 4.2. Number of root sections (i.e. sequences) representing each of the 18 fungal nrITS based operational taxonomic units
(OTUs) identified within the roots of Platanthera chapmanii plants cultured in vitro/greenhouse and those occurring naturally. Each
culture condition was sampled three to four times between 2012 and 2015. The first letter of an OTU name represents the fungal
family to which the OTU belongs: T, Tulasnellaceae; C, Ceratobasidiaceae. Values in parentheses are the total number of plants in
which a specific OTU was documented.
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Plant source Lab / Greenhouse Natural Population
Fall 2012 Fall 2014 Summer 2015 Fall 2012 Fall 2014 Spring 2015 Summer 2015
OTU GF12 GF14 GSu15 NF12 NF14 NSp15 NSu15
Tulasnellaceae
T1 8 (1) 0 0 0 0 0 0
T2 0 0 0 6 (1) 0 0 9 (2)
T3 0 20 (4) 0 0 0 0 0
T4 0 1 (1) 0 0 0 0 0
T5 0 6 (2) 0 0 0 0 0
T6 0 3 (1) 0 0 0 0 0
T7 0 1 (1) 0 0 0 0 0
T8 0 0 0 0 13 (5) 2 (1) 18 (5)
T9 0 0 0 0 2 (2) 0 1 (1)
T10 0 0 0 0 0 2 (1) 0
T11 0 0 9 (6) 0 0 1 (1) 5 (2)
T12 0 0 0 0 0 0 1 (1)
T13 0 0 0 0 0 0 1 (1)
T14 0 0 0 0 0 0 2 (1)
T15 0 0 4 (3) 0 0 0 2 (1)
T16 0 0 3 (2) 0 0 0 0
T17 0 0 1 (1) 0 0 0 0
Ceratobasidiaceae
C1 0 0 0 0 0 1 (1) 0
Total sequences 8 31 17 6 15 6 39
Total OTUs 1 5 4 1 2 3 8
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Table 4.3. Mean pairwise fungal internal transcribed spacer (nrITS) sequence distances
(π ± SE; Nei and Kumar 2000), estimated based on Kimura’s 2-parameter model, within
the fungal family Tulasnellaceae identified in the roots of Platanthera chapmanii plants
that were either cultured in lab / greenhouse conditions (GF12, GF14, GSu15) or were
obtained from a naturally occurring population (NF12, NF14, NSp15, NSu15).
Platanthera chapmanii mycorrhizal community Tulasnellaceae
n π-distance ± SE
Sequences from all sources
121
0.094 ± 0.031
Sequences from greenhouse cultured plants 56 0.064 ± 0.013
GF12 8 0.000 ± 0.000
GF14 31 0.047 ± 0.009
GSu15 17 0.037 ± 0.008
Sequences from naturally occurring plants 65 0.116 ± 0.036
NF12 6 0.000 ± 0.000
NF14 15 0.001 ± 0.001
NSp15 5 0.228 ± 0.065
NSu15 39 0.209 ± 0.035
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Table 4.4. Mean pairwise distances among fungal nrITS sequences based on Kimura’s 2-
parameter model were calculated for fungal communities identified within the roots of
Platanthera chapmanii. Roots of plants raised in vitro and cultured in greenhouse, and
from plants occurring naturally were sampled. All other mean pairwise distances, except
for Platanthera praeclara (Tovar 2015) and Nervilia nipponica (Nomura et al. 2013)
were calculated by Pandey et al. (2013).
Taxon
Tulasnellaceae
Ceratobasidiaceae
Geographic
range
n π-distance n π-distance
Platanthera chapmanii 65 0.116 ± 0.036 Wide
Platanthera praeclara 69 0.135 ± 0.006 238 0.039 ± 0.003 Wide
Anacamptis 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
Nervilia nipponica 9 0.157 ± 0.007 Wide
Chiloglottis trapeziformis 12 0.006 ± 0.002 Wide
Goodyera foliosa 7 0.062 ± 0.009 Wide
Goodyera tesselata 5 0.044 ± 0.006 Wide
Cypripedium japonicum 18 0.033 ± 0.006 Wide
Cypripedium candidum 7 0.003 ± 0.002 Wide
Goodyera hachijoensis 5 0.103 ± 0.009 Moderate
Cypripedium calceolus 12 0.005 ± 0.002 Moderate
Hexalectris grandiflora 6 0.023 ± 0.003 Moderate
Piperia yadonii 58 0.231 ± 0.026 71 0.077 ± 0.006 Restricted
Chiloglottis aff. jeanesii
14
0.014 ± 0.003
Restricted
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Figure 4.1. A photograph of root samples collected from in vitro propagated and
greenhouse cultured Platanthera chapmanii individuals before processing for molecular
analysis.
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Figure 4.2. A photograph of a cross section of a root of Platanthera chapmanii showing
mycorrhizal hyphal coils, i.e. pelotons, within the cortical cells.
Figure 4.3. Photographic documentation of moniliod cells and fungal hyphae isolated on
potato dextrose agar (PDA). The mycorrhizal fungus was cultured from roots of
Platanthera chapmanii.
100 µm
100 µm
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Figure 4.4. Sample-based incidence data, individual-based abundance data and observed
methods were used to construct cumulative, rarefied fungal operational taxonomic unit
(OTU) diversity curves for Platanthera chapmanii extrapolated to 500 sequences.
Operational taxonomic units were built using 122 mycorrhizal fungal sequences and 18
OTUs derived from the roots of plants that were either cultured in ex situ conditions or
were obtained from a naturally occurring population between 2012 and 2015.
0
5
10
15
20
25
30
35
40
0 100 200 300 400 500
# o
f O
TU
s
# of Sequences
Observed cumulative OTU diversity
Series2
Individual-based rarefied cumulativeOTU diversity
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Figure 4.5. A principal component analysis (PCA) scatterplot. Each of the circles
represent one of seven treatments (NF12, NF14, NSp15, NSu15, GF12, GF14, GSu15)
used to obtain mycorrhizal OTUs from the roots of Platanthera chapmanii plants that
were either cultured in lab/greenhouse conditions (GF12, GF14, GSu15) or were obtained
from a naturally occurring population (NF12, NF14, Nsp15, NSu15) between 2012 and
2015. The PCA shows PC1 and PC2 accounting for 60% of variation in the data.
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T9 T11 T8 T7 T6
T15
FJ613162_Tulasnella_calospora_Cymbidium_faberi KU664578_Tulasnella_sp._Goodyera_pubescens
AY373290_Tulasnella_bifrons_Tipularia_discolor T14 DQ388045_Tulasnella_calospora_Pleurothallis_lilijae JX649080_Uncultured_Tulasnellaceae_Dactylorhiza_fuchsii
T13 T17 DQ068773_Epulohiza_sp._Platanthera_praeclara T16 KJ495969_Tulasnella_sp._Anoectochilus_formosanus T1 T12 T4 JX998854_Uncultured_Tulasnellaceae_Epidendrum_firmum T3 AJ549127_Tulasnellaceae_Orchis_morio AY634122_Uncultured_Tulasnellaceae_Epipactis_gigantea AY373299_Tulasnella_sp._Tipularia_discolor JF691517_Uncultured_Tulasnellaceae_Agraecum_ramosum EU218891_Epulorhiza_anaticula_Corallarhiza_sp. AY373297_Tulasnella_danica_Goodyera_pubescens
AY373302_Tulasnella_sp._Tipularia_discolor AY373310_Tulasnella_sp._Tipularia_discolor
JQ994433_Uncultured_Tulasnellaceae_Piperia_yadonii AY373304_Tulasnella_sp._Tipularia_discolor JQ994441_Uncultured_Tulasnellaceae_Piperia_yadonii DQ178073_Uncultured_Tulasnella_Stelis_hallii JQ994406_Uncultured_Tulasnellaceae_Piperia_yadonii HM802322_Uncultured_Tulasnella_Singularybas_oblongus DQ388046_Tulasnella_asymmetrica_Thelymitra_luteocilium GQ241845_Uncultured_Tulasnellaceae_Paphiopedilum_dianthum T2 T10
AY373291_Tulasnella_deliquescens_Goodyera_pubescens
AY373276_Tulasnella_sp._Goodyera_pubescens
JX998909_Uncultured_Tulasnellaceae_Epidendrum_firmum AY373296_Tulasnella_tomaculum_Goodyera_pubescens JN655638_Tulasnella_sp._Pseudorchis_albida JF926488_Uncultured_Tulasnella_Orchis_purpurea AB369929_Epulorhiza_sp._Cypripedium_macranthos JF926484_Uncultured_Tulasnella_Orchis_purpurea GU066935_Uncultured_Tulasnellaceae_Orchis_mascula GU066934_Uncultured_Tulasnellaceae_Orchis_mascula EF433953_Uncultured_Tulasnellaceae_Cypripedium_guttatum DQ925640_Unculured_Tulasnellaceae_Cypripedium_reginae
T5
JX649085_Uncultured_Tulasnellaceae_Anacamptis_morio
EU583714_Uncultured_Tulasnellaceae_Orchis_simia GQ907249_Uncultured_Tulasnellaceae_Orchis_anthropophora GQ907273_Uncultured_Tulasnellaceae_Orchis_anthropophora
2.0
62/94
71/95
89/90
69
85 99/92
65
77 66
63
100/100
1/1
98/62 68/1
77/1 95/1
99/94
Texas Tech University, Kirsten Poff, August 2016
124
Figure 4.6. A maximum likelihood tree of the fungal family Tulasnellaceae built with
operational taxonomic units (OTUs) of nrITS sequences observed in Platanthera
chapmanii roots that were either cultured in lab/greenhouse conditions (green), obtained
from a naturally occurring population (red), or present in both environments (blue) and
orchid mycorrhizal OTUs from previous publications. The tree was rooted with midpoint
method. Bootstrap values ≤50 were omitted. The tree was built using 1000 bootstrap
replicates. Of the nodes that have two values, the second values are Bayesian probability
values from a Bayesian tree built using 1 million generations.
Texas Tech University, Kirsten Poff, August 2016
125
JQ972064_Uncultured_Ceratobasidiaceae_Piperia_yadonii
C1
JQ972091_Uncultured_Ceratobasidiaceae_Piperia_yadonii
AY634119_Uncultured_Ceratobasidiaceae_Epipactis_gigantea
HM141020_Uncultured_Ceratobasidiaceae_Goodyera_pendula
JQ972104_Uncultured_Ceratobasidiaceae_Piperia_yadonii
FJ788724_Uncultured_Ceratobasidiaceae_Pterygodium_catholicum
EU218895_Ceratohiza_sp._Goodyera_repentis
DQ068771_Ceratobasidiaceae_Platanthera_praeclara
GQ405535_Ceratobasidium_sp._Pterostylis_sp.
JF273479_Ceratobasidiaceae_Erycina_pusilla
AF503970_Ceratobasidiaceae_Tolumnia_variegata
HQ914117_Ceratobasidium_sp._Plectorrhiza_tridentata
DQ834419_Thanatephorus_sp._Vanilla_planifolia
EU218892_Thanatephorus_ochraceus_Corallorhiza_sp.
KF151202_Rhizoctonia_sp._Aa_achalensis
KF151201_Rhizoctonia_sp._Aa_achalensis
AJ549180_Ceratobasidiaceae_Orchis_laxiflora
AJ549123_Ceratobasidiaceae_Dactylorhiza_incarnata
GU066936_Uncultured_Ceratobasidiaceae_Orchis_morio
AF345558_Sistotrema_Dactylorhiza_majalis
2.0
54/97
89/97
100/99
98/98
77/93
64/83
99/97
70/97
88/100
Texas Tech University, Kirsten Poff, August 2016
126
Figure 4.7. A maximum likelihood tree of the fungal family Ceratobasidiaceae built with
operational taxonomic units (OTU) clustered using fungal nrITS sequences observed in
Platanthera chapmanii root obtained from a naturally occurring population (C1) and
other orchid mycorrhizal OTUs previously published. The tree was rooted with a species
of Sistotrema. Bootstrap values ≤50 were omitted. The tree was built using 1000
bootstrap replicates. Of the nodes that have two values, the second values are Bayesian
probability values from a Bayesian tree built using 1 million generation.
Texas Tech University, Kirsten Poff, August 2016
127
CHAPTER V
CONCLUSIONS
Much is unknown concerning the molecular and reproductive ecology of
temperate terrestrial orchid species. The rare species Platanthera chapmanii was used in
this study because of its wide geographic range, relatively quick growth and
development, and its known ability to be asymbiotically propagated. Seed dormancy is a
survival mechanism that can be evolutionarily advantageous for many temperate orchid
species. Many temperate terrestrial species are known to require pre-germination
treatments (e.g. cold-moist stratification) to induce germination. Based on an Analysis of
Variance and Fisher’s Least Significant Difference test, P. chapmanii seeds that were
treated with cold-moist stratification conditions at 5°C for 8 or 12 weeks had a higher rate
of germination than seeds that are not cold-moist stratified. After germination, however,
development up to nine months was independent of the pre-germination treatment. This
is most likely because of dormancy mechanisms the species has adapted to prevent
germination at inopportune times. Many species of temperate terrestrial orchids exhibit
extremely low germination rates (≤5%) when they do not undergo cold-moist
stratification (Bowles et al. 2002). The southern distribution of P. chapmanii may be the
reason for a relatively high germination rate without the cold-moist stratification
treatment (25%). This capacity to germinate in the absence of stratification could be a
useful adaptation as climate continues to change.
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Rare terrestrial orchids are thought to be sensitive to commercial fertilizers, and
the nutrient supplementation study described above was designed with this in mind
(Silvertown et al. 1994). There were no significant differences in plant height between
the three nutrient treatments (0.00x, 0.25x, and 0.50x). It is possible that significant
differences would exist if other response variables including chlorosis and tip necrosis
were measured, or if the concentration of fertilizer was increased. Differences in shoot
height was highly variable among treatments, and hence an increase in replications could
also add more confidence in the data. From the population augmentation studies, strong
conclusions cannot yet be drawn.
In this research, Platanthera chapmanii was documented to form mycorrhizal
pelotons with fungi from the families Tulasnellaceae and Ceratobasidiaceae. Of the 122
high quality fungal nuclear ribosomal internal transcribed spacer (nrITS) sequences, 121
were of Tulasnellaceae. The only Ceratobasidiaceae sequence was from a naturally
occurring P. chapmanii individual that was sampled in spring 2015. Mean pairwise
distance of the sequences from greenhouse sources were smaller than the mean pairwise
distance of sequences from naturally occurring plants, but OTU richness was the same. In
addition, a majority of the OTUs (13) clustered together on the same clade of the
maximum likelihood tree independent of treatment. Because the majority of sequences
were obtained with Tulasnellaceae specific primers, results may have been biased
towards Tulasnellaceae. However, high quality sequences could not be generated using
primers that were designed for other families within Basidiomycota including
Sebacinaceae and Ceratobasidiaceae indicating that the results represent the mycorrhizal
Texas Tech University, Kirsten Poff, August 2016
129
preference of the P. chapmanii reliably. Since the pairwise distances between sequences
were fairly small, and the OTUs clustered closely on the maximum likelihood tree, it is
suggested that the taxon is specific towards associations with narrow clades of the
Tulasnellaceae.
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130
Literature Cited
Bowles, M.L., Jacobs, K.A., Zettler, L.W., and T.W. Delaney. 2002. Crossing effects on
seed viability and experimental germination of the federal threatened Platanthera
leucophaea. Rhodora 104:14-30.
Silvertown, J., Wells, D.A., Gillman, M., Dodd, M.E., Robertson, H., and K.H. Lakhani.
1994. Short term and long term effects of fertilizer application on the flowering
population of the green-winged orchid Orchis morio. Biological Conservation,
69:191-197.
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