Platanthera chapmanii: culture, population augmentation ...

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

Texas Tech University, Kirsten Poff, August 2016

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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


3

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).


5

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


6

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


8

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


9

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,


10

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


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


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


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).


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


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


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


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


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


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


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


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


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


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


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


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.


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.


27

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36

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).


37

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.


38

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.


39

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


40

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


41

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


42

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).


43

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


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


45

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.


46

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),


47

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).


48

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


49

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


50

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


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.


52

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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

ecosystem. Applied Vegetation Science 9:7-10.

Heschel, M.S. and K.N. Paige. 1995. Inbreeding depression, environmental stress, and

population size variation in scarlet gilia (Ipomopsis aggregata). Conservation

Biology 9:126-133.

Johansen, B., and H. Rasmussen. 1992. Ex situ conservations of orchids. Opera Botanica

113:43–48.

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.

https://gis.ncdc.noaa.gov/maps/ncei/summaries/monthly


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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.



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.


extinction. Annals of botany 104:543-556.

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.

Walther, G.R., Post, E., Convey, P., Menzel, A., Parmesan, C., Beebee, T.J., Fromentin,

J.M., Hoegh-Guldberg, O. and F. Bairlein. 2002. Ecological responses to recent

climate change. Nature 416:389-395.

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54

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)

Lindley. American Midland Naturalist 145:168-17

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


55

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


56

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


57

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


58

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


59

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


60

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


61

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)


62

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


63

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


64

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


65

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


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


67

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


68

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.


69

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


70

duration of the experiment. Because the plants were placed in trays in groups of 5 or 6,

each tray served as an experimental unit.


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


71

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.


72

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.


73


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


74

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.


75

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


76

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.


77

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


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


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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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.


79

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.


81

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


82

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.


83

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


84

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


85

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


86

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).


87

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


88

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


89

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


90

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.


91

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.


92

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


93

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,


94

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.

http://www.ncbi.nlm.nih.gov/genbank/


95

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


96

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,


97

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).


98

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


99

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


100

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


101

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


102

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


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.


104

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.


105

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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.


110

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


tccgcgttgtgagtctaacaccagttgtaacacttttacaaccggtagcgctggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaag

tccaccacttatacgtgaatcatcgagttgttgaacgcactgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtatcccttcgggagtcttttcgttaagaccc

gagttcggagtcctcggtcttcggatcgtgttctcttagatgcgtcgcgccgatcgcctgatgggtcactctaatgcctgagcgtggagtccctcggagctgagaggcgcttgacc

gagtgttgagctcgcgtcgccaagtccgcacgtcttggacgtcggtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaa


ttttacaaccggtagcgatggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagttgttga

acgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtattccttcgggagtcctttttccaaaggaccggagttgggattcttggttctttggatcgtgttct

cttagatgtgtcacaccgatcgcctgatgggtcctctaatgcctaagcgtggagttcctgaaagtctgagacgtgcttgaccgggtcttgagctcgcgtcaccaagtctgcctaaca

agcagtactacaacacatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccaggatccctcag


111

Fungal OTU

Trimmed Sequence


ttttacaaccggtagcgatggtcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagttgttgaa

cgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtaatccttcgggagtccttttaactaaggacccgagttcggagtcctcggtcctctggatcgtgtt

ctcttagatgcgtcgcaccgatcgcctgatgggtcctctaatgcctaagcgtggagttccttcagagtccgaaacgtgcttgaccgggtgttgagctcgcgtcaccaagtctgccta

accagcagtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaactaggatccctca


aaactttttacaaccggtagcgatggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtcccccacttatacgtgaatcatcgagt

tgttgaacgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtaatccttcgggagtccttttaactaaggacccgagttcggagtcctcggtcctctgg

atcgtgtttttttagatgcgtcgccccgatcgcctgatgggtcctctaatgcctaagcgtggagttccttcagagtccgaaacgtgcttgaccgggtgttgagctcgcgtcaccaagt

ctgcctaaccagcagtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaactaggat

T8 >PchN1-1-ITS4-Tul

aaactttttacaaccggtagcgatggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatcatcgagt

tgttgaacgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtattccttcgggagtccttttccaaggacccgagttcggagtcctcggtcccacctgg

atcgtgttctctcagatgcgtcgcaccgatcgcctgatgggtcctctaatgcctaagcgtggagttccttcagagtcccacacgtgcttgaccgagtcttgagctcgcgtcaccaagt

ccgccttaaccagcggtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaactaaccagga

T9 >PchN3-1-ITS4-Tul

agttgtataaactttttacaaccggtagcgatggatcccttggcacgtcattcgatgaagaccgttgcaaattgcgataaagtgatgtgatgcgcaagtccaccacttatacgtgaatc

atcgagttgttgaacgcattgcaccgcgccctaatccggctgcggtatgcccctttgagcgtcattgtattccttcgggagtccttttccaaggacccgagttcggagtcctcggtcc

cacctggatcgtgttctctcagatgcgtcgcaccgatcgcctgatgggtcctctaatgcctaagcgtggagttccttcagagtcccacacgtgcttgaccgagtcttgagctcgcgtc

accaagtccgccttaaccagcggtactacaacgcatgacctcattggggtaggacaacccgctagacttaagcatattaatcagcggaggaaaagaaact


112

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


113

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


114

Fungal OTU Trimmed Sequence

C1 >PCHSN3-1-ITS4-OF

tgtgagacggattgaccctctcgggggacggtccgtctattatacataaactccaataattaaatttgaatgtaatttgatgtaacgcatctttgaactaagtttcaacaacggatctctt

ggctctcgcatcgatgaagaacgcagcgaaatgcgataagtaatgtgaattgcagaattcagtgaatcatcgaatctttgaacgcaccttgcgctccttggtattcctcggagcatg

cctgtttgagtatcatgaaattctcaaaataaatcttttgttaactcgattgattttattttggacttggaggtctgcagattcacgtctgctcctcttaaatttattagctggatctctgtgacat

cggttccactcggcgtgataagtatcactcgctgaggacactgtaaaaggtggccggagttactgaagaaccgcttctaatagtccattg


115

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.


116

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


117

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


118

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


119

Figure 4.1. A photograph of root samples collected from in vitro propagated and

greenhouse cultured Platanthera chapmanii individuals before processing for molecular

analysis.


120

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


121

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


122

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.


123

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


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.


125

JQ972064_Uncultured_Ceratobasidiaceae_Piperia_yadonii

C1


AY634119_Uncultured_Ceratobasidiaceae_Epipactis_gigantea

HM141020_Uncultured_Ceratobasidiaceae_Goodyera_pendula


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


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.


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.


128

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


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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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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,

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