March 2010 North American Native Orchid Journal

NORTH AMERICAN

NATIVE ORCHID JOURNAL

Volume 16(1) 2010

IN THIS ISSUE:

GROWING CYPRIPEDIUMS IN CONTAINERS

TRANSPLANT METHODS FOR THE ENDANGERED ORCHID

SPIRANTHES PARKSII

PROPAGATION AND CONSERVATION STATUS OF THE NATIVE ORCHIDS OF

THE U.S. ……

and more………….

The North American Native Orchid Journal (ISSN 1084-7332) is a publication

devoted to promoting interest and knowledge of the native orchids of North

America. A limited number of the print version of each issue of the Journal

are available upon request and electronic versions are available to all

interested persons or institutions free of charge. The Journal welcomes

articles of any nature that deal with native or introduced orchids that are

found growing wild in North America, primarily north of Mexico, although

articles of general interest concerning Mexican species will always be

welcome.

NORTH AMERICAN

NATIVE ORCHID JOURNAL

Volume 16 (1) 2010

CONTENTS

NOTES FROM THE EDITOR

1

LOOKING FORWARD

3


Doug Martin, Ph.D.

4

PRELIMINARY RESULTS FOR FIELD ESTABLISHMENT TECHNIQUES OF

CALOPOGON TUBEROSUS

Philip J. Kauth, Michael E. Kane & Timothy R. Johnson

12

A PRACTICAL AND INTEGRATED APPROACH TO NATIVE ORCHID

CONSERVATION AND PROPAGATION AT THE ATLANTA BOTANICAL

GARDEN

Matt Richards & Jenny Cruse Sanders, Ph.D.

25

AN UNDERGRADUATE’S FIRST ADVENTURE INTO FIELD RESEARCH:

AN EPIPHYTIC ORCHID SURVEY IN SOUTHERN FLORIDA

Emily Massey

31


SPIRANTHES PARKSII CORRELL

J. Ryan Hammons, Fred E. Smeins & William E. Rogers

38

PROPAGATION AND CONSERVATION STATUS OF THE NATIVE ORCHIDS OF

THE UNITED STATES (INCLUDING SELECTED POSSESIONS), CANADA,

ST. PIERRE ET MIQUELON, AND GREENLAND

Scott Stewart, Ph.D. & Aaron Hicks

47

SHOULD WE OR SHOULDN'T WE?

ETHICS AND ORCHIDS

The Slow Empiricist

67

RECENT ORCHID LITERATURE OF INTEREST

69

BOOK REVIEWS

72

ASYMBIOTIC TECHNIQUE OF ORCHID SEED GERMINATION

Second revised edition

MICROPROPAGATION OF ORCHIDS,

Volumes 1 & 2 (2nd Edition)

Unless otherwise credited, all graphics were prepared by the authors of the respective papers. The opinions

expressed in the Journal are those of the authors. Scientific articles may be subject to peer review and

popular articles will be examined for both accuracy and scientific content.

Volume 16(1): 1-81 issued January 15, 2010.

Copyright 2010 by the North American Native Orchid Journal

Cover: Calopogon tuberosus var. tuberosus by Stan Folsom

NOTES FROM THE EDITORS

The first issue of the North American Native Orchid Journal for 2010 is

a special issue that is focused on native orchid propagation, cultivation, and

reintroduction. The idea for this special issue was borne from conversations

among Lawrence Zettler, Aaron Hicks, and the Associate Editor about the

need to offer Journal readers an in-depth review of current trends in native

orchid propagation and cultivation work. Recognizing the integration of

orchid reintroduction with propagation and cultivation work, the Associate

Editor has included all three topics in this issue.

This issue presents articles by academic and popular authors about

native orchid propagation, cultivation, and reintroduction. Doug Martin

begins the special issue with detailed advice about cultivating Cypripedium

species in containers, followed by Philip Kauth et al. presenting preliminary

research results from a Calopogon tuberosus reintroduction study. The special

issue continues with Matt Richards and Jenny Cruise Sanders of the Atlanta

Botanical Garden discussing practical and integrated approaches to native

orchid propagation at the Garden. Next, Emily Massey presents her personal

story of orchid field research and reintroduction in southwestern Florida,

followed by Ryan Hammons et al.‖s presentation of translocation work with

the Federally endangered Spiranthes parksii. The issue concludes with a

comprehensive update by Scott Stewart and Aaron Hicks on the propagation

and conservation status of orchid species native to the United States, Canada,

Greenland, Puerto Rico, and selected possessions. This collection of authors

and articles represents a diverse cross section of the cutting edge of orchid

propagation, cultivation, and reintroduction.

The editors wish to thank all the authors and reviewers who made this

special issue possible.

1

The electronic format continues to be well received and we now reach

more than 1800 readers. Back issues from volume 3 (1997) to present are now

available online and you may read the current and back issues at:

http://wiki.terrorchid.org/tow:journals

The current update of the North American Personal Checklist is also

available at that website. The checklist will be updated as needed with new

taxa noted.

Paul Martin Brown, Editor

[email protected]

10896 SW 90th

Terrace, Ocala, FL 34481

36 Avenue F, Acton, Maine 04001 (June- early October)

Scott L. Stewart, Ph.D. Associate Editor

[email protected]

Kankakee Community College

Horticulture & Agriculture Programs

100 College Drive

Kankakee, Illinois 60901

2

http://wiki.terrorchid.org/tow:journals

mailto:[email protected]


LOOKING FORWARD

Future issues scheduled for 2010 of the

North American Native Orchid Journal

will feature such topics as

new taxa in Mesadenus and Corallorhiza,

a very special paper on evolutionary classification

by Richard Bateman from Kew

a new Series

HERE AND THERE

species found in North America and elsewhere

Cypripedium cultivation and hybrids

and much more!

3

Martin: GROWING CYPRIPEDIUMS IN CONTAINERS

4


Doug Martin, Ph.D.

Cypripediums are generally considered hard to grow. However, like all plants, they are

adapted to grow under a particular set of conditions in their natural environment. If the grower can

provide those conditions, the plants will practically grow themselves. Over the years I‖ve developed

methods, most adapted from other growers, which allow me to provide the conditions needed by

many cypripediums. While I still have a lot to learn, particularly about growing seedlings and the

more demanding species, my system seems to work well for the easier growing species and for

hybrids. Not counting seedlings, I now have about two dozen plants of six species and seven hybrids.

I‖ve only lost two mature plants in the last four years, both Cypripedium candidum, one of the more

demanding species. In this article I‖ll describe cypripediums‖ basic cultural requirements and how I

meet them.

GROWING CONDITIONS

While cypripediums can be grown in garden

beds, I prefer to grow mine in containers. This gives

me the ability to experiment with growing conditions.

I can move them to different locations in the yard

with different light levels, and I can easily change the

growing medium.

Containers: For most orchid growers, growing in

containers means using regular flower pots. However,

in the wild cypripedium roots spread out in a circle

from the crown of the plant, as much as two to three

feet in all directions. They also stay shallow, growing

in only the top one to two inches of moist, well

aerated soil (Stoutamire, 1991). To accommodate this

growth habit as much as possible, I use plastic storage

containers that are about 16 in. × 10 in. and 7 in. deep

(41 × 25 × 18 cm), with one blooming-sized plant per

container ( Fig. 1). I‖d like to use larger containers, but

they quickly become too heavy and hard to move.

Fig. 1. One of my standard growing containers for

cypripediums with a single plant of C. parviflorum var.

makasin that I have grown from seed.


5

Although cypripediums can be grown and flowered in regular flower pots, the ones I‖ve seen

don‖t seem to grow as large, or have flowers as large, as plants in the wild or in garden beds. I suspect

that the roots simply don‖t have room to spread. It‖s sort of a cypripedium bonsai. I think that giving

the roots extra room should result in a larger

plant. My largest plant of C. reginae seems to

support this theory, with three inch flowers on

top of two foot tall stalks (Fig. 2).

Cypripediums need consistent moisture at

their roots. To help accommodate this

requirement, my containers mimic the artificial

bogs used to grow other North American native

orchids. I leave the bottoms of the containers

solid and drill holes in the sides about four to five

inches (10 to 16 cm) below the surface of the

medium. The area below the holes acts as a

reservoir and helps to maintain the consistent

moisture level that the roots need. It is not

necessary to grow them this way, but it gives me

a margin of error in watering.

Medium: Because cypripediums need air as well as

consistent moisture at the roots, I use water

retentive growing media that are open, airy, and

free draining. For most cypripediums the

medium should be neutral to slightly acidic. Most

of the mixes I use are 75-80% inert material:

gravel, sand, perlite, Turface, or pumice. The

sand and gravel should be quartz, because too

much limestone in the mix will make it too

alkaline. The remainder of my growing media is organic material. Some cypripedium growers use

peat or chopped tree leaves, but I prefer coir–finely chopped coconut husk fibers. It is moisture

retentive like peat, but isn‖t acidic. I add a handful of ground oyster shells to each container to buffer

the pH of the medium. Two mixes that I‖ve used with good results are:

Medium 1: three parts perlite; one part gravel; one part coir.

Medium 2: four parts perlite; one part coir.

Water: One of the most important considerations when growing cypripediums is that they require

consistent moisture at their roots. The medium should never be allowed to dry out. Cypripediums

are sensitive to water quality, so they should only receive water low in dissolved solids such as rain,

distilled or RO water.

Fig. 2. C. reginae flowering in one of my standard

containers. The flowers are three inches across and the

stems are two feet tall.


6

Fertilizer: Cypripediums are adapted to grow in nutrient-poor soils. They are light feeders and do not

require much fertilizer. In a mostly inorganic medium like I use, they do better with frequent

applications of dilute fertilizer. Any good quality fertilizer will do. I use the Michigan State fertilizer

at about 25 parts per million of nitrogen. I water with it every other week and use it as a foliar spray

on alternate weeks. I fertilize my cypripediums from the time the leaves unfold in the spring until

early September.

Light: Most cypripediums grow in light shade in open forests or under shrubs. They like morning

sun, but must be protected from direct sun during mid-day. I grow mine on the south sides of

deciduous trees where they are in shade after about nine or ten o‖clock in the morning. They get

about 1,800 foot-candles in mid-day. Some like a bit more light, and I move these away from the tree

trunk to where the shade is thinner.

Cypripediums can also be grown indoors under lights. As a light source, I prefer the new T-5

fluorescent bulbs. Regular fluorescent bulbs only produce enough light if they are positioned very

close to the plants. They are only adequate for seedlings and small plants. High pressure sodium and

metal halide lights produce plenty of light but also a lot of heat; the T-5s fall in the middle, with

plenty of light but not too much heat.

Temperature: While some cypripediums are native to northern areas and require cool temperatures

year round, I find that several species and most of the hybrids grow well in my yard in the Kansas

City area. Our summer temperatures are often in the 90°s F (mid 30°s C) and even over 100F (38°

C). In winter, cypripediums require a cold rest period at near freezing temperatures. The Kansas City

area has these conditions from November through March.

I grow all of my seedlings, as well as the cooler growing Asian species, Cypripedium

macranthos, in a basement lightroom where the temperature stays between 72° F (22° C) to 81° F

(27°C). I grow tropical orchids in the lightroom during the winter. When the weather gets warm in

May, the tropicals go outside and the cypripediums go inside. The C. macranthos plants were

decreasing in size every year when I tried to grow them outside with my other cypripediums. Since I

started keeping them in the basement, they are making a comeback.

Humidity: Like all orchids, cypripediums prefer humidity above 50%. However, there does seem to

be some flexibility. My plants often experience humidity levels of about 25% or less outside during

the summer, without any noticeable negative effects.

Potting: Cypripediums should be potted with the roots spread out and the growth buds at or just

above the surface of the medium (Fig. 3). To accomplish this, I fill the container with medium to

about three inches below the top. Then I mound the medium in the center so that it slopes gently

down toward the edges. I place the plant on the mound, spread the roots out evenly and add medium

until the growth buds are just covered and the surface is level throughout. Then I water the plant and

add more medium where it washes down.


7

Pests: Deer and rabbits like to eat

cypripediums, and squirrels like

to dig in the medium. Since we

have a lot of all three, I keep my

cypripediums in cages made

from 1 × 2 in. (2.5 × 5.0 cm)

boards covered with chicken

wire on the sides and top. Four

of the cypripedium containers

fit nicely in a 48 × 18 in. (120 ×

45 cm) cage. Most of my cages

are two feet (60 cm) tall, but the

taller C. reginae need one three

feet (90 cm) tall.

Slugs and snails can be a problem, especially if the container is in contact with the ground

overnight. These pests can devour an entire seedling or eat through the stem of a blooming sized

plant overnight. Because my plants are kept on raised platforms during the growing season, I only

worry about slugs and snails in the spring when I uncover containers that have been buried for the

winter. I treat with slug pellets containing metaldehyde as part of the move to the growing area.

Caterpillars also can reduce a small plant to a leafless stalk seemingly overnight. I check my plants

regularly for signs of caterpillar damage. I treat with a dust designed for caterpillars on roses and

flowering plants.

GROWTH CYCLE

Cypripediums are hardy perennial plants with four distinct seasons in their growth cycle.

They are dormant during the winter and require a cold rest period. In the spring they break

dormancy and grow rapidly, reaching full size and flowering in three to eight weeks, depending on

the species. In the late spring and the summer they grow roots, store food and produce the growth

buds for the following year. In the fall, the stems and leaves wither and die back to the surface of the

medium in preparation for winter dormancy. I treat my plants differently during each of these

seasons and will discuss each in turn.

Winter Care: Cypripediums are dormant in winter so their needs are simple. They need to be kept

cold at temperatures near, but not below, freezing for three to four months. They must not dry out,

and they must be protected from temperature extremes; either very low temperatures that could

freeze the plant solid, or mid-winter warm spells. If the plants warm up, they may break dormancy

and start to grow. The new growths are very tender and will likely be killed when cold temperatures

return. The plants should be prepared for winter after the stems have withered and temperatures

Fig. 3. Growth buds of C. parviflorum var. makasin showing the proper

depth for potting.


8

have gotten cold, but before any hard freezes. Here in Kansas City I prepare my cypripediums for

winter at the end of November.

I prefer to store my plants underground for the winter. I put the containers in a hole deep

enough that the surface of the medium is three

to four inches below grade level, then cover

them with five or six inches of mulch (Fig. 4).

Nature maintains the temperature and moisture

for me. In colder areas, or where the risk of

mid-winter warm spells is greater than here in

the Kansas City area, it may be necessary to dig

deeper and/or use a thicker layer of mulch. It is

important that the hole drains well. I use

inexpensive cypress mulch from the local

hardware store, but some growers use chopped

leaves. I‖ve found that whole leaves pack down

and cause the plants to rot.

Alternatively, cypripediums can be

stored in an unheated garage, basement or

enclosed porch where the temperature stays

near freezing. Mulching the surface of the medium will help maintain the moisture level and protect

the growth buds from light freezes. I‖ve stored other hardy orchids this way and will have to start

doing it with my cypripediums because I‖ve run out of places to dig holes in my yard.

Some growers put their plants, pots and all, in plastic bags and store them in a refrigerator,

not the freezer, through the winter. The medium should be just barely moist. I‖ve discovered the

hard way that if the medium is too wet, the plants will rot.

Spring Care: I move my plants from their winter storage to the growing area when the danger of frost

is low, but when the daytime temperatures are still cool, preferably in the 50°s F (10° to 15° C). The

new shoots are very sensitive to frosts and even a light one will damage the developing leaves and

flower buds. A freeze can kill the entire shoot. However, if the daytime temperatures are too high,

the new shoots expand too rapidly and produce a weak plant.

Based on Kansas City weather, about to first of April I remove the winter mulch, treat for

slugs and snails, and move my cypripediums to their growing area. Once the containers are in their

growing area, the medium warms up and the plants break dormancy. The growth buds formed the

previous year expand and grow to their full size quite quickly. I cover my plants or move them into

the garage if a frost is expected. I also cover them or move them into the garage if strong thunder

storms or tornadoes are forecast. The only other care necessary at this time is to fertilize and to water

if it doesn‖t rain. If the plant is mature, it will bloom now. This all happens in three to eight weeks,

depending on the species. Now is the time when I find out how well I grew my plants last year!

Fig. 4. Cypripedium containers being buried for the winter. I

cover the containers with four to five inches of mulch.


9

Summer Care: Summer is when cypripediums grow into larger and stronger plants, but you won‖t see

the results until the following year. Cypripediums initiate new roots at the base of the new shoot

shortly after flowering. During the summer and into the fall these new roots and the roots from the

previous two years elongate, and the plants store energy in the rhizomes and produce the growth

buds for next year. The entire above-ground portion of the plant for the next growing season -

shoots, leaves and flowers - develops within these growth buds (Fig. 5). The care a cypripedium

receives during summer will determine how strong a plant and how many flowers, if any, it will

produce the next year.

During summer, I continue

watering and fertilizing as in the spring.

The one cultural challenge in the summer is

that cypripediums prefer to have their roots

cool. I mulch the surface of the medium to

protect it and the roots from being heated

by the sun. Of course, this also helps to

keep the medium from drying out. I use

the same mulch used in the winter, and I

am careful not to let the mulch touch the

stem of the plant as this can cause it to rot.

The container can also be set into a

shallow, well draining hole in the ground

to keep the roots cooler. I watch for

caterpillars, slugs and snails, and use an

appropriate treatment immediately if I see

signs of these pests.

Fall Care: In fall, the cypripediums stop

growing and all above-ground growth dies back. I stop fertilizing about the beginning of September. I

keep the medium moist and wait for the plants to go dormant, signaled by the leaves turning brown.

I do any necessary repotting and dividing in late fall before I put the plants in the ground for the

winter. I move my cypripediums to their winter quarters sometime between the first frost and the

first hard freeze.

New Plants: Cypripedium suppliers ship bare-root dormant plants in either the fall or spring. I prefer

to receive them in the fall because spring shipments arrive late in my spring growing season. Most

suppliers are located further north than I am, and by the time they can safely ship plants, the weather

in my area has already warmed up. When I receive plants in the fall, I seal the bare-root plants in

plastic food storage containers with a few drops of water. I then seal the containers in a plastic bag

and put them in the refrigerator until spring. Of course, another option is to pot the plants and put

them with your other cypripediums for the winter.

Fig. 5. C. parviflorum var. makasin growth buds are well

developed by early August. They will continue to grow thicker

over the next one to two months as the roots lengthen and the

plant stores energy in the tubers.


10

RECOMMENDED SPECIES AND HYBRIDS

The genus Cypripedium has 45 species and is quite diverse. Not surprisingly, some are less

demanding in their requirements. Below is a list of some of those species that are considered among

the easiest to grow and that I‖ve had success with. Except as noted, I grow these as described above.

C. parviflorum: The three varieties of the North American yellow lady‖s-slipper are considered to be

among the easiest of the species to grow. The two varieties I have, makasin and pubescens have both

grown well for me.

C. kentuckiense: The largest species in the genus, it grows naturally in sandy stream banks.

Consequently, I use a medium of 50% sand, 30% gravel and 20% peat. However, I would expect it to

grow well in either of my standard mixes.

C. reginae: This species likes a little more light than others, so I grow it where it gets more morning

sun. It grows naturally in more bog-like

conditions than other species. I grow

mine in a mix of three parts Perlite, one

part gravel and two parts ―black peat‖,

which is partially composted peat moss.

C. formosanum: This is often considered

the easiest species to grow. It has rather

long rhizomes and the new growths can

be several inches away from the previous

year‖s, so it requires a large container.

Also, it is prone to breaking dormancy

during a mid-winter thaw, so extra

protection may be needed in winter.

C. macranthos: This species grows well if

protected from temperatures above 80°F

(27°C). I grow these in my basement

lightroom during the hot part of the

summer.

Hybrids: In recent years a number of

cypripedium hybrids have become

available (Fig. 6). Like most orchids,

cypripedium hybrids are easier to grow

than the species and are a good choice for

the beginning grower.

Fig. 6. C. Hilda (C. ×ventricosum × macranthos), one of the

many cypripedium hybrids that are available from commercial growers.


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SOURCES OF CYPRIPEDIUMS

Like their tropical cousins, many cypripedium populations in the wild have been decimated by

over-collection. Please purchase only artificially propagated cypripediums to help reduce the demand

for wild-collected plants. Wild cypripediums have delicate roots that are usually damaged when plants

are collected. Most die within two to three years after being removed from the wild. Cypripediums

require several years to grow to blooming size and reputable growers price them accordingly. Mature

plants sold at bargain prices are almost certainly wild-collected. Many artificially propagated

cypripediums are available from reputable nurseries and commercial orchid growers. Some

companies that I‖ve found to have consistently high quality plants are:

Cyp Haven: http://www.c-we.com/cyp.haven/

Hillside Nursery: http://hillsidenursery.biz/

Itasca Lady Slipper Farm: http://www.ladyslipperfarm.com/

Vermont Lady Slipper Company: http://www.vtladyslipper.com/

Wild Orchid Company: http://www.wildorchidcompany.com/

Spangle Creek Labs: http://www.spanglecreeklabs.com/

SOURCES OF MORE INFORMATION ON GROWING CYPRIPEDIUMS

Most of the suppliers listed above provide cultural information on their websites.

Holger Perner has an excellent chapter on cultivation in The Genus Cypripedium, by Phillip Cribb.

Timber Press, Oregon. 1997.

The cypripedium forum: http://www.cypripedium.de/forum/

Doug Martin, 15523 Johnson Dr., Shawnee, KS 66217 [email protected]

LITERATURE CITED

Stoutamire, W.P. 1991. Central growth cycle of Cypripedium candidum Muhl. root systems in an Ohio Prairie.

Lindleyana 6(4): 235-40.

http://www.c-we.com/cyp.haven/

http://hillsidenursery.biz/

http://www.ladyslipperfarm.com/

http://www.vtladyslipper.com/

http://www.wildorchidcompany.com/

http://www.spanglecreeklabs.com/

http://www.cypripedium.de/forum/


Kauth et al.: PRELIMINARY RESULTS FOR FIELD ESTABLISHMENT TECHNIQUES OF CALOPOGON TUBEROSUS

12

Calopogon tuberosus var. tuberosus

common grass-pink


13

PRELIMINARY RESULTS FOR FIELD ESTABLISHMENT TECHNIQUES

OF CALOPOGON TUBEROSUS

Philip J. Kauth*, Michael E. Kane & Timothy R. Johnson

Abstract

Worldwide habitat loss has led to interest in propagation and reintroduction of orchids. However,

scientific investigation regarding successful field establishment remains poorly understood. Previous research has

indicated that using dormant storage organs and planting seedlings in areas of reduced competition increased

survival of several orchid species. Here we describe methods for establishing Calopogon tuberosus, a North

American terrestrial orchid, on the Florida Panther National Wildlife Refuge. Comparative effects of planting

seedlings and corms on survival and shoot growth were studied. In addition, seedling survival in burned and

unburned plots was studied. While propagule type did not influence survival, date of planting did. A higher

percentage of propagules survived when planted in February 2009 during the early growing season. While more

seedlings were actively growing in the burned plot during April 2009, seedlings in the unburned plot produced

more shoots. The data from this study are being used to develop management plans not only for C. tuberosus, but

also other terrestrial orchids.

Introduction

The worldwide loss of orchid taxa has led to an abundance of research focused on their

conservation, ecology, and reintroduction (Ramsay and Dixon, 2003). Unfortunately, few

reports exist that detail management methods for both orchid populations and their habitat

(Stewart, 2007). Successful establishment of plants into current or former habitats is often the

culmination and goal of orchid conservation research (Batty et al. 2006a). Establishing orchids

in the field is challenging because complex ecological requirements of individual taxa are not

well-understood (Scade et al., 2006).

Successful field establishment of terrestrial orchids has been previously attempted, but

only for a few species (McKendrick, 1995; Ramsay and Stewart, 1998; Stewart et al., 2003;

Batty et al., 2006b; Scade et al., 2006; Yamato and Iwase, 2008), and few studies have

documented field establishment of North American species (Stewart, 2007). Long-term

survival of field-transplanted orchids is often very low, in part because efficient methods for

establishing orchids are lacking (Batty et al., 2006a). The influence of abiotic and biotic factors

on successful field establishment of orchids has not been studied in detail (Scade et al., 2006).

However, field establishment of orchids could be an important tool for both conserving

orchids and furthering our knowledge of orchid ecology (McKendrick, 1995).


14

A major obstacle to field establishment is initial survival of propagules. Only a few

articles highlight techniques for increasing survival of orchid seedlings under in situ conditions

(Batty et al., 2006b; Scade et al., 2006; Smith et al., 2009). Batty et al. (2006b) reported higher

survival of several Australian orchid species when dormant tubers were reintroduced rather

than seedlings. Observations that Thelymitra manginiorum seedlings established more readily

than tubers indicate that field performance of different propagules is species-specific (Smith et

al., 2009). Competition may also be an important factor to consider for successful

establishment (McKendrick, 1995). Dense coverage by native species increased survival of field

transplanted orchid species (McKendrick, 1995; Scade et al., 2006; Yamato and Iwase, 2008),

but areas of greatest vegetation coverage, including weedy species, impeded total survival

(McKendrick, 1995).

Calopogon tuberosus var. tuberosus (Linnaeus) Britton, Sterns & Poggenberg, common

grass-pink, is a corm-forming terrestrial orchid species found throughout eastern North

America from south Florida to Newfoundland, Canada. Calopogon tuberosus is a fairly

common native orchid and is an excellent candidate to examine field establishment methods

because: 1) seeds germinate readily in vitro, 2) seedlings can be produced in several months

(Whitlow, 1996; Kauth et al., 2006; Kauth et al., 2008), and 3) it is a corm-forming species.

Because this species produces corms, the role of propagule type on successful field

establishment can be studied.

The objectives of this study were to: 1) Establish Calopogon tuberosus seedlings at the

FPNWR; 2) Compare survival of seedlings and corms of C. tuberosus; 3) Compare survival

and growth of seedlings in burned and unburned areas; and 4) Recommend management

practices for establishing terrestrial orchids at the FPNWR. The data generated will be used to

recommend management practices for the successful conservation of this species and other

terrestrial orchids worldwide.

Materials and Methods

Field Site

The Florida Panther National Wildlife Refuge (FPNWR) is located in Collier Co.,

Florida and consists of 26,400 acres within the Big Cypress Basin (U.S. Fish and Wildlife

Service 2009). The FPNWR was established in 1989 to protect the Florida panther and the

mosaic of habitats located throughout. As a national wildlife refuge, the FPNWR actively

manages the area with invasive plant removal, prescribed burning, native plant propagation,

and restoration activities.

The FPNWR is divided into 50 fire-management units. Calopogon tuberosus is currently

found in units containing wet prairies (Fig. 1). These grass and sedge dominated communities

at the FPNWR are found between pine flatwoods dominated by Pinus elliotii (Davis, 1943;

Duever et al., 1986). The largest population of C. tuberosus is found in a wet prairie where


15

several hundred plants flower from March through May with peak flowering in mid April.

All field plots were established in this area.

Seed Source and Preparation

Seeds from the FPNWR were collected from mature yellowing capsules in June 2006

and 2007 before capsule dehiscence. Two capsules were collected from at least three plants.

Seed capsules were stored over desiccant in laboratory conditions at 23° C for 2 weeks. Seeds

were subsequently removed from the capsules and placed in glass scintillation vials over silica-

gel desiccant, and stored in darkness at -11° C until use. In all experiments, seeds were surface

sterilized in sterile scintillation vials for 3 min in a solution of 5 mL absolute ethanol, 5 mL

6% NaOCl, and 90 mL sterile dd water. Seeds were rinsed with sterile dd water after surface

sterilization. Solutions were removed with sterile Pasteur pipettes. Seeds were transferred

onto the germination medium with a 10µL sterile inoculating loop.

Fig. 1. Field translocation study at the Florida Panther National Wildlife Refuge. A)

Burned (left) and unburned (right) areas in February 2009. B) Burned (background) and

unburned (foreground) areas. Yellow flags mark one of the transects. C) Transects and

quadrats in the burned area. D) Transect and quadrats in the unburned area in April 2008.

E) Close-up of a quadrat.

B


16

Establishment

Planting occurred in successive years in April 2008 and February 2009. For the 2008

planting, differences in survival of field transplanted seedlings and corms were examined. For

the 2009 planting, the response of planting seedlings in a burned versus unburned area was

studied (Fig. 1A, B, C). For all experiments square quadrats 30 cm × 30 cm were constructed

from PVC piping (1.5 cm diameter). Each quadrat (Fig. 1E) was divided into 16 sections ca.

7.5 cm × 7.5 cm by using 14 gauge coated electrical copper wire. A HOBO H8 Pro weather

station (www.microdaq.com, Ltd., Contoocook, NH) was placed at the site to record daily

temperatures and relative humidity (Fig. 2).

Comparison of propagule type on field survival

Seeds were germinated asymbiotically in vitro beginning March 2007 on P723 medium

supplemented with 1% activated charcoal (PhytoTechnology Laboratories, Shawnee Mission,

KS). 40 mL of medium was dispensed into square 100 × 15 mm Petri plates (Integrid™ Petri

Dish, Becton Dickinson and Company, Franklin Lakes, NJ). Cultures were wrapped in a

single layer of Nescofilm (Karlan Research Products, Santa Rosa, CA, USA) and placed under

Fig. 2. Monthly temperatures recorded at the field transplant site in the Florida

Panther National Wildlife Refuge from April 2008-March 2009. Average

temperatures represent the mean daily high or low over the entire month. Data

were collected with a HOBO H8 Pro series weather station.


17

a 12 h photoperiod at 25° C. After 8 weeks culture (May 2007), seedlings were transferred to

PhytoTech Culture Boxes (PhytoTechnology Laboratories, Shawnee Mission, KS) containing

100 mL P723 medium. After an additional 30 weeks culture, corms were chilled at 10°C in

darkness from October 2007 to January 2008. This was accomplished by removing the shoots

and roots from the seedlings, and transferring corms to fresh P723 medium in PhytoTech

Culture Boxes. After the chilling period, corms were again transferred to fresh P723 medium

in PhytoTech Culture Boxes for an additional 12 weeks under a 12 h photoperiod. Seedlings

were subsequently moved to greenhouse conditions April 2008. Seedlings were planted in 9-

cell pack trays (Model #IKN0809, Hummert International, Earth City, MO) containing

Fafard 2 soilless potting mix (Conrad Fafard, Inc., Agawam, MA). Seedlings were covered

with clear vinyl humidity domes to prevent desiccation, and placed under 50% shade cloth

and a natural photoperiod. Average light levels were 300 µmol m-2 s-1 measured at 12 noon,

and average temperatures ranged from 21.6 ± 2° C to 29.3 ± 3° C. After one week humidity

domes were removed and seedlings were watered as needed.

Three 10 m transects (Fig. 1D) were establish April 23, 2008. Each transect contained

four quadrats 2.5 m apart. A randomized block design was used to plant propagules. Corms

and seedlings were assigned randomly to a quadrat and quadrat section. Sixteen propagules

were used in each quadrat (8 seedlings and 8 corms per quadrat). A total of 192 propagules

were planted. Propagules were irrigated with distilled water upon initial planting. Data were

collected on 20 May 2008, 9 July 2008, 27 February 2009, and 23 April 2009.

Seedling survival in a burned and unburned field plot

Seeds were germinated in vitro starting January 2008 on BM-1 Terrestrial Orchid

Medium (PhytoTechnology Laboratories, Shawnee Mission, KS) supplemented with 1%

activated charcoal. 40 mL of medium was dispensed into square 100 × 15 mm Petri plates.

Cultures were placed under a 12 h photoperiod at 25°C. After 8 weeks culture (March 2008),

seedlings were transferred to PhytoTech Culture Boxes containing 100 mL BM-1 medium.

After an additional 30 weeks culture, corms were transferred to new PhytoTech Culture

Boxes containing 100 mL BM-1 medium and chilled at 10°C from October 2008 to December

2008. Seedlings were moved to greenhouse conditions December 2008 until ready for field

establishment February 2009. Greenhouse transfer procedures were similar to those

previously described. Average light levels were 253 µmol m-2 s-1 measured at 12 noon, and

average temperatures ranged from 20.8 ± 2.3°C to 28.8 ± 2.8°C.

In January 2009, the wet prairie was burned except the area where Calopogon tuberosus

field plots were previously established in 2008. This presented a unique opportunity to

compare the effects of planting seedlings in the burned and unburned areas. Two 10 m

transects were established in both the burned area and unburned area. Three quadrats were

allocated to each transect. Sixteen seedlings were planted in each quadrat for a total of 48

seedlings per transect and 192 seedlings for the experiment.


18

Data Collection and Statistical Analysis

For both experiments, survival of all seedlings was recorded. Two different categories

were classified in determining propagule survival. Percentage of actively growing green shoots

was recorded. Percentage of emergent shoots was recorded when shoots were present, but not

necessarily actively growing (i.e. shoots were yellow and brown due to senescence). Seedling

leaf measurements were recorded before the February 2009 experiment, and again in April

2009. Shoot emergence data were analyzed using proc glimmix, logistic regression, and least-

square means in SAS v9.1.

Results

Comparison of Propagule Type on Field Survival

Propagule type (F = 0.50, p = 0.48) did not influence survival, but date (time of data

recorded) was significant (F = 20.4, p < 0.0001). At the initial data collection in May 2008, a

higher proportion of seedlings (43.8%) had actively growing shoots compared to corms

(32.3%) (Fig. 3). After 1 month of field establishment, less than 50% of all propagules had

actively growing shoots regardless of treatment. In July 2008, natural leaf senescence had

occurred so that no shoots were actively growing. A higher proportion of emergent shoots

were observed on seedlings (22.9%) compared to corms (12.5%).

Fig. 3. Survival of Calopogon tuberosus propagules at the Florida Panther National Wildlife

Refuge. Histobars represent the mean response of three separate transects each with four

quadrats containing 16 propagules. A total of 96 propagules were planted per treatment for a

total of 192 propagules.


19

Fig. 4. Survival of Calopogon tuberosus seedlings in a burned and unburned plot at the

Florida Panther National Wildlife Refuge. A) Percentage of plants with actively growing

shoots marked by the presence of a growing green shoot. B) Percentage of plants with either

have actively growing shoots or previously emerged shoots that senesced. Histobars

represent the mean of two transects with three quadrats containing 16 seedlings. Ninety-six

seedlings were planted in each treatment for a total of 192 total seedlings.


20

Data collected during February 2009 occurred during the early growing season in

south Florida. The number of green shoots was higher on corms (12.5%) compared to

seedlings (10.4%), but this difference was not significant (Fig. 3). In April 2009, no significant

difference was observed between the survival of corms (6.25%) and seedlings (8.33%), and the

presence of shoots further declined. At this time, one seedling in the early flowering stage

established from a corm propagule was observed No shoots were observed in June 2009.

Percent of total survivorship of all combined propagules were as follows: 38.0% (May 2008),

18.9% (July 2008), 11.4% (February 2009), and 7.3% (April 2009).

Seedling Survival in a Burned and Unburned Field Plot

Burning significantly influenced percent of emerged shoots (F = 48.7, p < 0.0001),

while the unburned plot influenced the percentage of actively growing shoots (F = 4.32, p =

0.04). Two months after field establishment, the number of actively growing shoots declined

in both plots. Three percent and 11% of actively growing shoots were observed in the burned

and unburned areas, respectively (Fig. 4B). However, senesced shoots were visible on seedlings

in the burned plot, but none in the unburned plot (Fig. 4A). Total survivorship was 7.3%

when combining all data. Of the actively growing shoots, shoot lengths were recorded in

April 2009 (Table 1). Shoot lengths on all seedlings with actively growing shoots in the

burned plot increased, while three of the eleven recorded leaf measurements in the unburned

plot decreased (Table 1).

Treatment Transect Quadrat # Seedling # Height

(Feb 2009)

Height

(April 2009)

Unburned 1 1 14 85 28

1 2 3 25 66

1 3 3 100 75

1 3 6 60 90

1 3 13 90 108

2 1 14 70 92

2 2 4 62 220

2 2 9 76 35

2 2 10 63 72

2 3 5 85 125

2 3 12 26 102

Burned 1 1 13 52 91

1 2 14 10 165

2 1 3 95 140

Table 1. Shoot lengths recorded for actively growing Calopogon tuberosus seedlings in February and April 2009.

All measurements are in mm. Seedlings were measured in February under greenhouse conditions prior to

transplant, and the April data collection was on seedlings after field transplant on the Florida Panther National

Wildlife Refuge.


21

Discussion

This is the first study of the field establishment of Calopogon tuberosus, and one of the

only scientifically documented orchid field establishment studies in North America (Stewart

et al., 2003; Zettler et al., 2007). However, conclusive results were not obtained due to the

short-term nature of the study, and more results are likely after several years of monitoring.

Absence of an actively growing shoot did not indicate propagule death since corms may have

been present beneath the soil surface. Their presence beneath the soil was not confirmed in

order to minimize soil disturbance. In addition, shoots on field-transplanted seedlings may

have senesced naturally because senescence naturally occurs in late May through early June.

Field establishment of orchids may depend on propagule type such as seedlings or

storage organs. Dormant storage organs, depending on species, were found to successfully

survive initial field establishment compared to seedlings (Debeljak et al. 2002; Batty et al.

2006b). Dormant storage organs may be able to survive drought conditions better than

seedlings (Batty et al. 2006b); however, results are species specific. Caladenia arenicola and

Diuris magnifica established more readily in the field when dormant tubers were planted

rather than seedlings, but Thelymitra manginiorum established more readily from seedlings

(Batty et al., 2006a). However, no C. arenicola propagules and only 10% of D. magnifica tubers

survived into the third growing season. 70% and 35% of T. manginiorum seedlings and tubers,

respectively, survived into the third growing season (Batty et al., 2006a). Likewise, Smith et al.

(2009) found that 2-3 year old plants (35%) established more readily in the field compared to

tubers (11%) after 4 years.

In this study, no statistical differences in shoot emergence were observed between

corms (6.25%) and seedlings (8.33%) after the first year. The low rates of shoot emergence

may have been caused by propagule death or dormancy of corms. Terrestrial orchids can

remain dormant for several years (Kery and Gregg 2004) thus long-term monitoring is

necessary to observe propagule survival. Throughout 2008-2009, south Florida experienced

drought conditions that may have contributed to propagule death. The juvenile state of the

propagules (1 year old) may have resulted in poor field establishment as well. Tuber size

influenced the survival of several Australian orchids with larger tubers increasing survival

compared to smaller tubers (Batty et al., 2006a; Smith et al., 2009). Likewise, larger Calopogon

tuberosus corms or more mature plants may have increased survival in the present study due to

greater storage reserves that tubers can utilize to sustain drought conditions and initiate

growth (Batty et al., 2006a).

The influence of competition, shading, and weed coverage influences the establishment

of orchids in the field (McKendrick, 1995; Scade et al., 2006). This is the first report

comparing field establishment of an orchid in a burned and unburned are in North America.

The effects of establishing orchids in burned plots have apparently not been studied, but

smoke was shown to be effective at promoting germination of several Australian plant species


22

(Flematti et al., 2004). Although this study investigated seed germination, the study could

explain a higher percentage of actively growing shoots in the burned plot. The effects of

smoke on emergence of Calopogon tuberosus shoots may warrant investigation.

The influence of competition on plant establishment has also been examined. In the

present study, more actively growing shoots of Calopogon tuberosus were observed on

seedlings in the unburned area during April 2009. Shoots on the seedlings in the burned area

were brown and senesced with the exception of three plants. The surrounding native grasses

in the unburned area likely shaded the seedlings providing increased survivorship and soil

moisture. Seedlings in the unburned area did not receive any level of shading and likely caused

seedling desiccation. Shading led to increased survival of several other terrestrial orchids

(McKendrick, 1995, 1996; Scade et al., 2006; Yamato and Iwase, 2008), but areas of dense shade

and competition can lead decreased seedling survivorship (McKendrick, 1995; Yamato and

Iwase, 2008). Fire is a necessary natural disturbance in many ecosystems (Duncan et al., 2008)

including wet-prairies in south Florida. Competition with weeds and invasive species during

field establishment often reduces the successful field establishment of seedlings (Moyes et al.

2005). Native perennials were established readily in a burned grassland and dolomite glade

areas. In addition, reduced weedy species, and prevented forest succession (Moyes et al., 2005;

Duncan et al., 2008).

While the results of this study are preliminary due to the short-term monitoring of the

plots, the techniques employed can be applied to other orchid species worldwide. More

definitive results may be observed after another growing season when seedlings in the burned

area may re-emerge. Due to the drought conditions the past 2 years in south Florida,

additional irrigation may have improved propagule survival. In addition, using symbiotically

grown seedlings or inoculating soil with mycorrhizal fungi may have improved seedling

survival as well (Batty et al., 2006a; Scade et al., 2006; Smith et al., 2009).

Conclusions

Based on the success of research with other terrestrial orchids, the following should

also be considered to successfully establish Calopogon tuberosus seedlings in the field: 1) Using

more mature seedlings to younger seedlings may provide sufficient carbohydrate reserves to

survive initial planting. 2) When planting dormant corms, larger corms should be planted. 3)

Propagules should be planted at the beginning of the growing season. 4) Due to frequent

drought conditions, the effects of supplemental irrigation could be studied. 5) Plots should be

monitored for several years to observe successful field establishment.

Philip J. Kauth*, Michael E. Kane, Timothy R. Johnson

Plant Restoration, Conservation, and Propagation Biotechnology Program, Environmental Horticulture

Department, University of Florida , PO Box 110675, Gainesville, FL 32611, USA.

*Corresponding author: email [email protected]



23

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Batty, A.L., M.C. Brundrett, K.W. Dixon, and K. Sivasithamparam. 2006a. New methods to improve symbiotic

propagation of temperate terrestrial orchid seedlings from axenic culture to soil. Australian Journal of

Botany 54: 367-74.

Batty, A.L., M.C. Brundrett, K.W. Dixon, and K. Sivasithamparam. 2006b. In situ seed germination and

propagation of terrestrial orchid seedlings for establishment at field sites. Australian Journal of Botany 54:

375-81.

Davis, J.H. 1943. The Natural Features of Southern Florida. The Florida Geological Survey, Tallahassee, FL.311.

Debeljak, N., M. Regvar, K.W. Dixon, and K. Sivasithamparam. 2002. Induction of tuberisation in vitro with

jasmonic acid and sucrose in an Australian terrestrial orchid, Pterostylis sanguinea. Plant Growth

Regulation 36: 253-60.

Duever, M.J., J.E. Carlson, J.F. Meeder, L.C. Duever, L.H. Gunderson, L.A. Riopelle, T.R. Alexander, R.L.

Myers, and D.P. Spangler. 1986. The Big Cypress National Preserve. National Audubon Society, New

York.

Duncan, R.S., C.B. Anderson, H.N. Sellers, and E.E. Robbins. 2008. The effect of fire reintroduction on endemic

and rare plants of a southeastern glade ecosystem. Restoration Ecology 16: 39-49.

Flematti, G.R., E.L. Ghisalberti, K.W. Dixon, and R.D. Trengove. 2004. A compound from smoke that

promotes seed germination. Science 305: 977

Kauth, P. J., W.A. Vendrame, and M.E. Kane. 2006. In vitro seed culture and seedlings development of

Calopogon tuberosus. Plant Cell, Tissue and Organ Culture 85: 91-102.

Kauth, P. J., M.E. Kane, W.A. Vendrame, and C. Reinhardt-Adams. 2008. Asymbiotic germination response to

photoperiod and nutritional media in six populations of Calopogon tuberosus var. tuberosus

(Orchidaceae): evidence for ecotypic differentiation. Annals of Botany 102: 783-93.

Kery, M., and K. Gregg. 2004. Demographic analysis of dormancy and survival in the terrestrial orchid

Cypripedium reginae. Journal of Ecology 92: 686-95.

McKendrick, S.L. 1995. The effects of herbivory and vegetation on laboratory-raised Dactylorhiza praetermissa

(Orchidaceae) planted into grassland in Southern England. Biological Conservation 73: 215-20.

McKendrick, S.L. 1996. The effects of shade on seedlings of Orchis morio and Dactylorhiza fuchsii in chalk and

clay soil. New Phytologist 134: 343-52.

Moyes, A.B., M.S. Witter, and J.A. Gamon. 2005. Restoration of native perennials in a California annual

grassland after prescribed spring burning and solarization. Restoration Ecology 13: 659-66.

Ramsay, M.M. and J. Stewart. 1998. Re-establishment of the lady's slipper orchid (Cypripedium calceolus L.) in

Britain. Botanical Journal of the Linnaean Society 126: 173-81.

Ramsay, M.M. and K.W. Dixon. 2003. Propagation science, recovery, and translocation or terrestrial orchids. In

K. W. Dixon, S. P. Kell, R. L. Barrett, and P. J. Cribb [eds.], Orchid Conservation. Natural History

Publications (Borneo), Kota Kinabalu, Sabah, Malaysia.

Scade, A., M.C. Brundrett, A.L. Batty, K.W. Dixon, and K. Sivasithamparam. 2006. Survival of transplanted

terrestrial orchid seedlings in urban bushland habitats with high or low weed cover. Australian Journal

of Botany 54: 383-89.

Smith, Z. F., E.A. James, M.J. McDonnell, and C.B. McLean. 2009. Planting conditions improve translocation

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Botany 57: 200-209.

Stewart, S.L. 2007. Integrated conservation of Florida Orchidaceae in the genera Habenaria and Spiranthes: model

orchid conservation systems for the Americas. PhD Dissertation, University of Florida, Gainesville, FL,

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Stewart, S.L., L.W. Zettler, J. Minso, and P.M. Brown. 2003. Symbiotic germination and reintroduction of

Spiranthes brevilabris Lindley, an endangered orchid native to Florida. Selbyana 24: 64-70.

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24

Whitlow, C.E. 1996. Mass production of Calopogon tuberosus, pp.5-10. In C. Allen [ed.], North American Native

Terrestrial Orchids: propagation and production. North American Native Terrestrial Orchid Conference,

Germantown, Maryland.

Yamato, M. and K. Iwase. 2008. Introduction of asymbiotically propagated seedlings of Cephalanthera falcata

(Orchidaceae) into natural habitat and investigation of colonized mycorrhizal fungi. Ecological Research

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Richards & Sanders: A PRACTICAL AND INTEGRATED APPROACH TO NATIVE ORCHID CONSERVATION AND

PROPAGATION AT THE ATLANTA BOTANICAL GARDEN

25

A PRACTICAL AND INTEGRATED APPROACH TO NATIVE ORCHID

CONSERVATION AND PROPAGATION AT THE

ATLANTA BOTANICAL GARDEN

Matt Richards & Jenny Cruse Sanders Ph.D.

In recent years interest has grown in the propagation of rare orchids native to North

America. For the Atlanta Botanical Garden it stems originally from the need to reproduce

valuable collections held in public trust for display and education interests in the Fuqua

Orchid Center (Fig. 1). Now the focus has grown to include work on propagating natives

specifically for conservation purposes. These effective methods of propagation and production

have not always been a result of scientific research backed by statistical analysis, but still have

contributed greatly to our ongoing commitment to sustain and improve in situ orchid

populations in the Southeastern United States (Fig. 2). As the program continues to grow, the

Garden maintains its project driven approach to assist in the protection of habitat as well as to

produce plants suitable for conservation work while developing a balance between scientific

research, practical propagation and ensuing horticultural practices.

For decades, the Garden‖s conservation program has been forged by following through

project driven goals and objectives. Work has been focused on forming relationships between

private-landowners, federal, state, and local agencies. Needs are defined to outline the

roadmap towards reaching each particular goal. The Georgia Plant Conservation Alliance was

formed in 2005 and is inclusive of many levels of contributors in the native plant conservation

community. As a founding member, the Garden has united with other botanical gardens,

institutions, universities, individuals, governmental and non-governmental agencies to provide

a service to the broad scheme of plant conservation (Fig. 3). Through this alliance, many of

the collaborators have been able to facilitate rare plant conservation without the obstacles

presented by having a formal organization. All parties involved with each specific project have

identified their particular talents, resources, and capabilities they can contribute to the overall

project. These services may include diplomatic relations, propagation and production, on the

ground labor, or motivating an army of dedicated volunteers to see the project through. The

Atlanta Botanical Garden has found its niche in rare plant propagation and horticultural

excellence (Fig. 4). All participants contribute in some way or another, and all of the work is



26

done in close collaboration in large part due to the coordination efforts carried out by the

GPCA Conservation Coordinator, Jennifer Ceska at the State Botanical Garden of Georgia.

Orchids in Georgia continue to require field surveys to assess the current status of state

element occurrence records. We also attempt to identify suitable habitat and additional

populations of native species. The Georgia Orchid Initiative through the Atlanta Botanical

Garden ultimately aims to resurvey all orchid species known to occur in state and catalog

localities with GPS and GIS technologies. Through tissue culture, field surveys, horticultural

experiments, and ex situ propagation, the orchid research team is taking an integrated and

collaborative approach to better understand the geography, natural history and reproductive

biology of native orchids. This initiative hopes to provide an increased understanding of the

biology and geographic distribution of native terrestrial orchids in Georgia (Fig. 5).

The Garden often works with the GADNR (Georgia Department of Natural

Resources) to identify the needs for orchid surveys and species recovery in the state. We have

decided it would make most sense to concentrate our efforts using the state ranks, surveying

the element occurrence records first of S1, S2 and S3 species, and then move through the list

to S5 respectively. In addition, peripheral habitat is identified using satellite imagery, soil

maps, GIS technologies, and ground reports to better assess the status of the species. Once

populations have been located, we begin work on assessing the habitat threats, and potential

management strategies (if any) (Fig. 6). For some of these orchid populations there is a need to

safeguard populations through propagation and ex situ collections, especially those included in

the S1/S2 G1/G2 rarity status. This propagated material is grown in an organized and indexed

manner so that it may be used responsibly to augment populations or for future

reintroduction into conservation lands (if deemed necessary and appropriate). One example

would be that of Platanthera chapmanii (Chapman’s fringed orchid) (Fig. 7). Known

historically from parts of south Georgia, the species had not been documented in the state for

nearly a century. In September of 2009 a GPCA member, Dr. Richard Carter of Valdosta

State University confirmed extant populations of the species he had seen earlier in 2006. Soon

after the discovery was documented, seed was collected from two populations for in vitro

propagation. At the time of this writing, stage-3 germination had been achieved (Fig. 8). With

some good horticulture and a bit of luck, the future of this population of rare orchids is better

protected. The need to propagate the species was prioritized by assessment of habitat. The

habitat was surveyed by members of the GPCA and considered to be in extreme danger of

alteration, subject to road improvements, herbicide applications, and timber activities.

Measures will soon be taken to develop further conservation strategies.

Additionally, Platanthera integrilabia (monkey-face orchid) occurs in a handful of

counties in the State of Georgia. We have worked together with the GADNR to survey

extant populations, additional habitat on conservation lands, and any new 'undiscovered



27

populations' of the species. Since 2007, surveys have been active for the species. Meanwhile,

propagation has been ongoing, safeguarding populations on private land in ex situ collections

produced from seed with the hope of augmenting these populations one day (Fig. 9).

Although one of the rarest orchids in North America, it is oddly very easily propagated from

seed. The seed is sterilized using 90 ml RO water, 10 ml Clorox, and .01 ml Tween 20 shaking

vigorously for 10 minutes. The seed is poured into a sterile filter and rinsed several times using

sterile RO water. The seed is allowed to air dry within a laminar flow hood, and then

sprinkled onto sterile media. This species is easily germinated on ½-strength P668 from

PhytoTech Labs. After approximately 5 months, the seedlings are transferred to ¾-strength

P668. Although it will germinate and grow on various other media, the cost and simplicity of

this procedure effectively produces reproductive plant material in a short amount of time (Fig.

10). The process is typically adapted to the natural bio-rhythm of the species. The seed is

harvested and sown at the time it would naturally fall to the ground. It is placed in dark

cabinets after sowing, and then into cold storage during the winter months. During this time,

the seedlings continue to develop. The ensuing spring allows the seedlings to emerge into

photosynthetic growth under artificial lighting while still in sterile culture. Soon after the

emergence of the first shoot, the plants are transferred onto fresh media and allowed more

space to grow. When the plants are large enough to be transferred into pots and soilless media

in our greenhouses, they are typically overwintered in vitro and potted up immediately

following 120 days at ca. 38 F. By following the natural biorhythm of the plant, it becomes

easier to transition them between growing stages and eventually back into their native habitat.

This method would typically allow for outplanting at the time the species goes dormant.

The propagation of our native orchids has been well documented in the past and it is

common to find available scientific and popular articles regarding propagation of many of our

natives. There are also many books that have been published on the subject of orchid

propagation. The Garden approaches each species independently, researches what has been

done by others in the past with the species and then develops a plan to tackle the first obstacle

of germination. We sow our orchids using various methods, some are green capsule, some are

dry seed, some are sterilized in different fashions, and many different media are used. Since

2002, records have been kept defining the work done with each orchid species. Although

much of this remains unpublished, the database is a critical tool used daily in the Tissue

Culture Lab. We can refer back many years to varying treatments and procedures used at

propagating hundreds of species. With this information available, our staff can better develop

an experiment that could eventually lead to successful propagation. Generally speaking, we

sow most seed on several different media, and develop a plan to simulate natural bio-rhythm

of the plant to what extent is possible. Once germination has been achieved, we will select

several replate media for trial. Once the best media for replating is determined through

observation and analysis of growth, the plants are transferred to quickly produce healthy

plants suitable for transfer to ex vitro culture. Again, each species is treated independently and



28

our talented and experienced staff develops the appropriate soil amendments to achieve

desired growth and continues to refine the protocol for each species during greenhouse

culture.

Orchids are one of the most charismatic groups of plants, and yet few people are aware

that so many species of orchids are native to Georgia. At the Atlanta Botanical Garden we

reach a large audience with approximately 350,000 visitors per year, and we can inform this

audience about native orchid species (Fig. 11). In 2007 we educated more than 14,500 students

through educational programs (tours, outreach, afterschool, camps) developed for

kindergarten through eighth grade, and 4971 adults through tours, lectures, training programs

and family programming. Information learned through the Georgia Orchid Initiative will be

made available to teachers as a resource for teaching about native plant species. Display

plantings and interpretive signage will also be developed in the conservation garden, the

children‖s garden, and the garden discovery carts will inform visitors about our research on

native orchids in Georgia and North America (Fig. 12).

Matt Richards & Jenny Cruse Sanders Ph.D., Atlanta Botanical Garden, 1345 Piedmont Ave NE,

Atlanta, Georgia 30309 [email protected]; [email protected]

www.atlantabotanicalgarden.org

www.atlantabotanicalgarden.org/site/conservation/native_plants

www.uga.edu/gpca/

The Atlanta Botanical Garden (Garden) has served the Southeastern region as both a horticultural resource and a

place of enjoyment since 1976. The Garden has two facilities under its stewardship – 30 acres in the heart of

Midtown Atlanta and 185 acres in Gainesville, Georgia (Smithgall Woodland Garden). Ranked as one of the top

ten botanical gardens in the United States, the Garden develops and maintains plant collections for display,

education, research, conservation, and enjoyment. The Garden offers stunning garden displays and exceptional

education programs for people of all ages. Many of its collections of rare and endangered plant species cannot be

seen anywhere else in the world, and its conservation work, both nationally and internationally, is critical to

preserving our natural heritage.

The Fuqua Orchid Center opened to the public in 2002 providing an exciting opportunity to further develop

and display its already distinguished orchid collection. The display glasshouses maximize and augment the

existing tropical lowland orchid collections and provide specialized facilities for new collections of orchids that

grow at high elevations. Back-up greenhouse facilities for orchid care and a Tissue Culture Lab for plant

propagation are also included in this center as are greenhouse facilities to propagate, and safeguard rare indexed

plant populations of the southeastern United States for conservation purposes. Outdoors, and adjacent to the

Fuqua Orchid Center is the Conservation Garden that highlights native bog habitats of the southeastern United

States including coastal plain, cataract, and mountain bogs. For more information on programs, hours of

operation, events, and classes, please visit www.atlantabotanicalgarden.org.



http://www.atlantabotanicalgarden.org/

http://www.atlantabotanicalgarden.org/site/conservation/native_plants

http://www.uga.edu/gpca/

http://www.atlantabotanicalgarden.org/



29

Fig. 1 (left) Atlanta Botanical Garden Conservation Garden D. Lentz

Fig. 2 (right) Cypripedium kentuckiense (ivory-lipped lady‖s-slipper) out-planting S. Larson

Fig. 3 (left) GPCA members at 2009 fall meeting Okefenokee Swamp. J. Ceska

Fig. 4 (right) ABG Conservation Greenhouse facility. M. Richards

Fig. 5 (left) New

population of

Listera smallii

(Small‖s twayblade)

discovered in 2008.

B. Wilson

Fig. 6 (right) Survey

of Platanthera spp.

emerging after late

winter burn in a coastal plain bog. L. Kruse



30

Fig. 7 (left)

Platanthera

chapmanii

R. Carter.

Fig. 8 (right)

Stage 3

germination of

Platanthera

chapmanii

M. Richards

Fig. 9 (left) Platanthera

integrilabia growing in vitro

R. Gagliardo

Fig. 10 (right) Platanthera

integrilabia produced from

seed flowering in the

conservation greenhouse.

M. Wenzel

Fig. 11 (left)

Visitors reading an

interpretive display of

Epipactis gigantea (stream

orchid), a Flagship Species

for the North American

Region Orchid Specialist

Group. M. Richards

Fig. 12 (right) Hybrid

swarm of Platanthera

grown from seed at ABG

on display in the Fuqua

Orchid Center. M. Richards

Massey: AN UNDERGRADUATE‖S FIRST ADVENTURE INTO FIELD RESEARCH:


31

AN UNDERGRADUATE’S FIRST ADVENTURE INTO FIELD

RESEARCH:


Emily Massey

My interest in orchids began when I was an undergraduate student at Illinois College

(IC), a small liberal arts institution located in Jacksonville, Illinois. There, I worked with Dr.

Lawrence Zettler in the Orchid Recovery Program. This program focuses on the propagation,

study, and reintroduction of threatened and endangered orchid species. While at IC, I

participated in a number of studies with several different orchids. These projects included two

studies involving symbiotic seed germination. In the first study, we examined crossing effects

on seed viability, germination, and protocorm growth in Platanthera leucophaea (Nuttall)

Lindley, the eastern prairie fringe orchid. Seed germination, propagation, and reintroduction

of Epidendrum nocturnum Jacquin, the night-fragrant epidendrum was examined in our

second study (Massey et al., 2007). I also participated in a study to asymbiotically propagate

several epiphytic south Florida orchids such as E. amphistomum A. Richard, the dingy-

flowered star orchid; E. rigidum Jacquin, the rigid epidendrum; Polystachya concreta (Jaquin)

Garay & Sweet, the yellow helmet orchid; Prosthechea

cochleata (Linnaeus) W.E. Higgins var. triandra (Ames), the

Florida clamshell orchid; and Vanilla phaeantha

Reichenbach f., the oblong-leaved vanilla orchid.

All of these studies were conducted in the

laboratory, except for the reintroduction of Epidendrum

nocturnum, which is an endangered Florida epiphyte with

night fragrant flowers that are believed to be pollinated by a

species of hawkmoth. Although I liked lab work, it was this

study that introduced me to field research and it was one of

the best experiences I had while working with Dr. Zettler.

The project took place in the fall of 2005 at the Florida

Panther National Wildlife Refuge (FPNWR). The FPNWR

is located 20 miles east of Naples in Collier County,

Florida and was established as a safe haven for the

diminishing Florida panther population and other

threatened and endangered animal and plant species.

Fig. 1 Image of me and the two

other students (William Kutosky,

and Kris McDonald) reintroducing

E. nocturnum at the Florida Panther

National Wildlife Refuge in 2005

S.L. Stewart



32

For about a week, two of my lab mates and I visited the refuge to reintroduce

Epidendrum nocturnum seedlings propagated in our lab. At the time of the reintroduction, the

cool, murky water at most sites was waist and chest deep for my 5‖2‖‖ stature (Fig. 1). You

definitely had to be careful where you were walking, or I should say feel around where you

were stepping, because you sure could not see through the water beneath your feet. This was

due to the presence of tannins that darken the water into a coffee-like brew. This was a very

intimidating project and I was a little apprehensive at first. We were venturing out into the

wilderness, with the possibility of running into an alligator or worse, and we were on foot.

However, the longer we worked, the less this seemed to matter. I may have had to wring out

my clothes every night of the trip, but this field research experience was one of the most

memorable moments of my life. I had many other new experiences as well. I got to take my

first spin in a swamp buggy, which is basically a very large, open-air vehicle resembling a

monster truck without a top (Fig. 2).

Another new experience and probably one of the more amusing moments of this

excursion took place when a local news reporter and cameraman came out to the refuge to

capture our efforts. The cameraman must have known what he was getting into because he

showed up wearing boots and worn clothing. Perhaps the reporter should have consulted

with him before he dressed that morning because he wore shiny dress shoes, khaki pants, a

very neat button down shirt, and a tie. Needless to say the reporter was a bit out of his

element, but he was a good sport about it. With a smile, he waded out into the swampy water

after a few minor wardrobe adjustments (i.e., rolled up his pant legs a good three or four

inches and donned a pair of borrowed boots) to film a portion of the piece.

Fig. 2. One of the many swamp buggies in the fleet.



33

We eventually reintroduced 43 Epidendrum nocturnum seedlings back into the wild.

Unfortunately, this part of Florida had just been damaged by Hurricane Wilma. This stripped

many of the trees‖ upper canopy, exposing the seedlings to higher light levels and fewer than

10% of our reintroduced seedlings remained one year later. Another issue was that we had

little idea of what trees to affix these seedlings and what microhabitat conditions they needed

(i.e., epiphytic assoc-

iates, location on the

tree, and the light levels

required). The locations

we selected for these

plants were based on

observations made by

the staff at the refuge

and students performing

research on site. The

FPNWR is home to

about 27 orchid species

in 17 genera with many

of these species being

threatened or endanger-

ed. It is possible that the

survival of many of

these species hinges on

the habitat (i.e., tree

species in the area), the microhabitat (i.e., substrate of establishment and epiphytic associates),

and other factors (i.e., light level to which they are exposed). However, little to no data has

been collected on the orchid microhabitats of these species at the refuge.

This leads us to the study at hand. My project surveyed an area of the FPNWR for

epiphytic orchids and the mircohabitats associated with them. The site was classified as a

slough transitioning to a floodplain swamp and was believed to consist mainly of pop ash

(Fraxinus caroliniana), pond apple (Annona glabra), and baldcypress (Taxodium distichum) for

epiphytic orchids and catalogued the microhabitats associated with them. Some of the species

I surveyed were orchids that I had worked with in the Orchid Recovery Program back in

Illinois. Again, I was working in some of the same sites I had visited two years ago, but the

terrain was slightly different. For one, the atmosphere was very different. The cooler fall

weather had been replaced by the hot and very humid summer months. The site was no

longer flooded and I could see where I was stepping most of the time. Despite this, I still

encountered some obstacles. About once a week, I experienced tiny paper cuts on my exposed

arms and legs, cuts that were the direct result of the very tall and sharp saw-grass (Cladium

jamaicense), which in some spots was taller than me. The saw-grass was also an area of concern

because alligators often find this habitat to be conducive for nest building. I encountered

3

Fig. 3. Image of one transect at the study site.



34

many more mosquitoes than I had in the fall and began each day by spraying myself with bug

spray. Fortunately the only animals I came across were deer and a couple of harmless snakes.

The data were collected in June and July of 2007 by another student, Cabrina

Hamilton, and me. Data were collected for this project along 30 transects, 140 m long and 10

m apart for a total area sampled of 42,000 m2

(Fig. 3). An orchid was counted in the survey if

it was within 1 meter from the ground

(Fig. 4). A midday light measurement was also

collected for each plant using a Sper Scientific, Broad Range LUX/FC meter (840022) and

recorded in Lux. Some other data collected consisted of the phorophyte (i.e., a plant on which

epiphytes grow) for each orchid, the substrate on which the orchid was established (i.e., moss,

bark of host tree, lichens, or a combination of any two), the diameter of the part of the tree

closest to the orchid was measured in centimeters, and the orientation of the orchid in regards

to substrate tilt (i.e., located on the trunk, an angled or a horizontal limb, or on a fallen tree)

along with the directionality of the orchid (i.e., facing N, E, W, S, NE, NW, SE, and SW).

The orchid‖s epiphytic associates were measured (i.e., vascular plants like bromeliads and ferns

and non-vascular organisms like lichens and mosses). We also subjectively determined the

percentage of the area in the microhabitat they comprised and estimated the number of

species present.

Figs. 4-7. Mature orchids sampled at the survey site. Campylocentrum pachyrrhizum is an example of a

leafless orchid [5], Prosthechea cochleata var. triandra an orchid with leaves and visible pseudobulbs [6],

and Epidendrum amphistomum an orchid with leaves and no visible pseudobulbs [7].

4

5 6 7



35

The orchids themselves were divided into three categories based on their

morphological differences: leafless (Fig. 5)

(i.e., Campylocentrum pachyrrhizum (Reichenbach

f.) Rolfe, crooked-spur orchid; ribbon orchid and Harrisella porrecta (Reichenbach f.)

Fawcett & Rendle, the leafless harrisella), orchids with leaves and visible pseudobulbs (Fig. 6)

(i.e., Encyclia tampensis (Lindley) Small, the Florida butterfly orchid; P. concreta; and P.

cochleata var. triandra), and orchids with leaves and no visible pseudobulbs (Fig. 7) (i.e.,

Epidendrum amphistomum, E. nocturnum, and E. rigidum). They were further subdivided into

their stages of development. The plants without leaves were separated by the number of green

roots: seedlings (>3 green roots), juveniles (3-5 green roots), and mature plants (with

flowering or fruiting bodies or >5 green roots). The plants with leaves were separated into

seedling (plant ≤0.5 cm), juvenile (plant ≥0.5 cm and ≤10 cm), and mature (flowering or

fruiting bodies or plant ≥10 cm) plants. The number of green roots (leafless orchids) and the

number of green leaves were counted (orchids with leaves). If a plant was in flower or fruiting,

then we also counted the number of flowers and capsules. During the study the only orchids

in flower were E. amphistomum (Fig. 8)

and P. concreta (Fig. 9)

and the only orchid seen in

fruit was E. amphistomum.

We sampled 419 orchids in total with a majority of the orchids surveyed being

juveniles with fewer mature plants and seedlings. Of the mature plants, E. amphistomum were

fruiting (2) and flowering (7). Polystachya concreta was also in flower (1). Most of the orchids

surveyed were found on pop ash (Fraxinus caroliniana) (371) with 100%, 89%, and 88% of

them being leafless, leaves with pseudobulbs, and leaves without pseudobulbs respectively.

Leafless orchids were observed on trunks or branches <51 cm in diameter whereas orchids

with leaves and visible pseudobulbs, as well as orchids with leaves and no visible pseudobulbs

were noted on trunks and branches between 11-110 cm. All of the seedlings sampled occurred

on moss, and it appeared that the juvenile and mature plants were either on moss or a

combination of moss and bark. A majority of the epiphytic orchids without leaves were

found on horizontal substrates, whereas orchids with leaves were affixed to branches/trunks

at a 45 degree angle or a vertical position. Moreover, the majority of the orchids were oriented

on substrates that received little direct sunlight (N, NE position). Many of the epiphytic

5 7

8 9

Figs. 8, 9. Mature orchids in flower during the study: Epidendrum amphistomum [8] and Polystachya concreta [9].

4

8

5 6 7 8 9

9



36

associates consisted of mosses and ferns (e.g., resurrection fern, Pleopeltis spp.), as well as

bromeliads, vines and occasionally lichens (Massey et. al., 2008).

Taken together, it appears the orchids at this site are established on moss or a

combination of moss and bark of the phorophyte, which is largely pop ash (F. caroliniana)

with all of the seedlings being established on moss. These orchids were often facing in a

northerly or northeasterly direction and either on branches or trunks tilted at an angle or

vertical for the orchids with leaves and horizontal for the leafless orchids.

Currently, the Orchid Recovery Program at Illinois College is propagating many of

the orchid species surveyed in this study with the hopes of reintroducing them. Despite the

thoroughness of this study, more research is needed before we can give orchids reintroduced

in this area a fighting chance. For instance, the data in this study seem to indicate that the

epiphytic orchids grow prominently on pop ash (F. caroliniana). We are unsure if this result is

due to there being more trees of this species in the area or if there is a physical (i.e., bark

texture, moisture capabilities, or the arrangement of the canopy cover) or chemical association

between the orchids surveyed and this species of tree. A future study could survey the trees in

this area or analyze the bark of all of the species of trees indicated in this study to better

understand this relationship. Also it seemed that despite the orchids growing in close

proximity to lichens few of them were established on or very close to lichens. Perhaps there is

a reason for this dissociation. I suggest further studying of the orientation and tilt for these

and other orchid species and more data collection of the phorophytes and branch diameter,

especially for the orchids with leaves. In addition to these factors, studies regarding the stage

of the orchid best suited for reintroduction should be assessed. Our data indicate that juveniles

(i.e., plant ≥0.5 cm and ≤10 cm) were highly abundant at the site so perhaps plants should

only be reintroduced if they are at a juvenile or mature plant stage.

Further study of these threatened and endangered orchids is needed. Many of these

species are in danger of being poached, having their habitats destroyed by humans and

hurricanes, and having their territory encroached upon by exotic species. Hopefully, this

study will promote future research aimed at improving the survival of both Florida orchids

and other threatened and endangered species through reintroduction or better protection and

management of their habitats.

Acknowledgements

Many people were influential from the conception of this study to completion of this manuscript.

Cabrina Hamilton for her aid in data collection, Dr. Lawrence Zettler for asking me to join his lab and fostering

my love of research, Larry Richardson and the U.S. Fish and Wildlife Service for allowing me to come and work

on the Florida Panther Refuge, Dr. Scott Stewart for helping with the formation of the study, Illinois College

and the Charles and Dorothy Frank Scholarship for funding my study, Dr. Elizabeth Rellinger for her all of her

patience in helping me with statistical analysis, the refuge staff for their assistance during the survey, and for

everyone who spent their precious time reviewing this article. I kindly thank all of them for their support.



37

Emily Massey, Department of Environmental Horticulture, University of Florida, PO Box 110675,

Gainesville, FL, 32611. [email protected]

As for my future plans, this study was a great experience and while I still enjoy laboratory work, it really

sparked my interest in field research too. Currently, I am a graduate student enrolled at the University of Florida

earning my Master‖s degree in Environmental Horticulture. My proposed project examines water relationships,

specifically the affects of stress, and growth in two tree species. Although this is not based in ecological

restoration, this study will provide a good basis for future research. Eventually I plan on returning to more

ecologically based projects and securing a research position.

Literature Cited:

Massey, E.E., K. Hamilton, S.L. Stewart, L.W. Richardson, and L.W. Zettler. 2008. Substrate preferences of

epiphytic orchids (seedlings, juveniles, mature plants) within the Florida Panther National Wildlife

Refuge. Illinois State Academy of Science 101: 62-63.

Massey, E.E. and L.W. Zettler. 2007. An expanded role for in vitro symbiotic seed germination as a conservation

tool: Two case studies in North America (Platanthera leucophaea and Epidendrum nocturnum).

Lankesteriana 7(1-2): 303-08.


Hammons, Smeins & Rogers: TRANSPLANT METHODS FOR SPIRANTHES PARKSII

38

Spiranthes parksii Navasota Ladies’-tresses

Grimes County, Texas


39


SPIRANTHES PARKSII CORRELL

J Ryan Hammons, Fred E. Smeins & William E. Rogers

ABSTRACT

Spiranthes parksii (Navasota ladies‖ tresses) is an endangered terrestrial orchid endemic to the Post Oak

Savanna ecosystem in central-east Texas. Methods of whole plant transplantation are needed to conserve

individuals that will be destroyed by development activities. A soil-intact and a bare-root method were evaluated.

Spiranthes parksii and its congener, S. cernua can be distinguished when in flower, but are indistinguishable from

one another based on morphology of their leaf rosettes. Unknown leaf rosettes of S. parksii or S. cernua were

transplanted into areas where S. parksii and S. cernua were known to co-occur. Compared to percent production

of leaf rosette and flower production of undisturbed individuals on-site, transplanted individuals by both

methods have been successful.

INTRODUCTION

Spiranthes parksii, Navasota ladies’-tresses, is an endangered orchid endemic to central-

east Texas within the Post Oak Savanna Ecoregion where it co-occurs with its congener S.

cernua which has a broad distribution across eastern North America (Pelchat, 2005; Brown,

2008). Spiranthes parksii has also been found further east in the Pineywoods Ecoregion,

however, vegetation documented at these occurrences was similar to the Post Oak Savanna,

and not typical of the Pineywoods (Bridges & Orzell, 1989). The Post Oak Savanna Ecoregion

is dominated by native bunchgrasses and forbs with scattered clumps of trees and shrubs,

primarily post oak (Quercus stellata) (TPWD, 2009). Other common woody species are

blackjack oak (Quercus marilandica), black hickory (Carya texana), American beautyberry

(Callicarpa americana), yaupon (Ilex vomitoria), farkleberry (Vaccinium arboreum), winged

elm (Ulmus alata), eastern redcedar (Juniperus virginiana), and water oak (Quercus nigra)

(Brezanson, 2009). Common grass species are little bluestem (Schizachyrium scoparium), other

bluestems (Andropogon spp.), Indiangrass (Sorghastrum nutans), purpletop (Tridens flavus),

curly threeawn (Aristida desmantha), and longleaf spikegrass (Chasmanthium sessilifloraum).

This system was originally maintained as a savanna by frequent fires and grazing by bison,

and with their absence, tree/shrub species increase and grasses/forbs decrease (TPWD, 2009).

Within the Post Oak Savanna, Spiranthes parksii typically occurs on sparsely vegetated areas

along the upper reaches of ephemeral and intermittent drainages. Individuals are also found

away from drainages along game/livestock trails and/or in small herbaceous openings at a

tree/shrub dripline where a herbaceous patch meets a tree/shrub community (Hammons,

2008; USFWS, 2009). Spiranthes cernua, nodding ladies’-tresses, also occurs in these habitats.


40

A solid waste landfill is needed for Bryan/College Station, Texas and surrounding

areas. During construction, an estimated 379 Spiranthes parksii plants will be destroyed. In

order to meet mitigation requirements, the United States Fish and Wildlife Service (USFWS)

Biological Opinion required 57 hectares of deed restricted areas be purchased around the

landfill footprint to protect and conserve S. parksii plants that occurred in those areas and to

serve as recipient sites for transplanted individuals. As well, the Biological Opinion permits

research to develop procedures for successful transplantation of at-risk plants to protected

areas. It is our goal to explore soil-intact and bare-root methods of transplantation.

METHODS

Both Spiranthes parksii and S. cernua are perennial and produce a leafless inflorescence

during mid-fall (Oct.-Nov.). A basal rosette of leaves is produced between November and

April, which is followed by a dormant underground stage until the next flowering season.

Identification of the two species is apparent during flowering; however, they cannot be

differentiated during the leaf rosette stage of growth.

All transplantations occurred at the end of leaf rosette growth to minimize disturbance

during the growing period. Additionally, transplantation occurred when soil moisture was at

field capacity. All were placed in deed restricted areas where other Spiranthes parksii/S. cernua

flowering individuals were previously documented. Plant locations were marked in the field

with survey flags and GPS positions so they could be re-visited to monitor survival.

Additionally, several hundred undisturbed S. parksii/S. cernua leaf rosettes were marked in the

same area to monitor survival compared with transplants. All transplanted individuals and

between 22 and 540 undisturbed leaf rosettes were monitored for flowering and leaf rosette

production each year after transplantation.

Root Tuber Distribution and Bare-Root Transplantation

Based on size and length of rosette leaves, six small and four large individuals were

excavated in spring 2007. Length of each leaf and root tuber was measured, and each were

summed to give total leaf length and total root tuber length to 1) determine if leaf size and

root tuber size are correlated, and 2) determine the size and extent of root tubers so that root

tubers would not be damaged during transplantation. For this study, a root tuber constitutes

any underground structure growing from the bud zone. Soil was removed from the root

tubers, individuals were wrapped in a wet paper towel and transplanted to deed restricted

areas within two hours following excavation (Fig. 1).

In spring 2008, an additional cohort of 57 Spiranthes cernua/S. parksii leaf rosettes and

two known S. parksii were re-located and transplanted. In spring 2009, 14 known S. parksii

individuals were transplanted. Of these, six had <5 cm of one root tuber taken for

examination of mycorrhizal fungi infection and isolation in the laboratory.


41

Soil-intact Transplantation

In spring 2007, a 20 cm diameter PVC pipe was used to excavate individuals while

keeping the soil intact around root tubers. The PVC pipe was cut into 15 cm lengths and

beveled at the bottom so it could be hammered into the soil around a leaf rosette. A shovel

was then placed underneath the PVC pipe so that soil within the PVC pipe could be

excavated. After excavation, plants were transplanted to deed restricted areas within

approximately two hours. A hole was carefully dug in the deed restricted areas to fit the

diameter and depth of the transplant inside the PVC pipe. After placing the transplant and

PVC pipe in the pre dug hole, the PVC pipe was removed and soil was fed into the cracks

around the transplant to fill any large air spaces (Fig. 2).

Fig. 1. Methodology for bare-root transplantation. a) shovel buried deep beneath plant and soil slightly raised, b)

individual carefully taken out of soil with most soil removed so measurements could be taken, c) root tubers

wrapped in a wet paper towel, and d) stored for transport to deed restricted areas.

RESULTS

Root Tuber Demographics and Bare-Root Transplantation

For the 10 bare-root transplants in spring 2007, total leaf length for the small

individuals ranged from 5 to 11 cm, while total leaf length for the large individuals ranged

from 22 to 32 cm. The number of root tubers per individual ranged from 2 to 8. Total leaf

length and total root tuber length were positively correlated (R2

= 0.84; p= .000). The

maximum depth of a root tuber from the base of the stem was 9 cm, while the maximum

lateral distance was 8 cm. Root tubers were found to be both exhausted and not exhausted in

S. parksii/S. cernua individuals, as noted by Wells et al. (1991; Fig. 3).


42

Fig. 2. Methodology for soil-intact transplantation. a) PVC section centered around plant and hammered into

ground, b) shovel slid underneath PVC section to be lifted out, c) transplants placed for transportation, and d)

hole dug to fit PVC, transplant placed in pre-dug hole, PVC removed, and soil fed into cracks where PVC was to

rid of any air spaces.

Fig. 3. Spiranthes rosette individual that does not have remnants of an exhausted root tuber (left) and one with

two exhausted root tubers (right).

With the exception of leaf rosette production in 2008, subsequent production of the 10

bare-root transplants have had a higher percent production than undisturbed Spiranthes

cernua/S. parksii individuals also originally found in spring 2007 (Fig 4). Individual plants


43

show no consistent pattern. One individual remained dormant for 2 flowering and 2 leaf

rosette stages, but emerged as a leaf rosette in fall 2009. Another has formed a flowering stalk

and leaf rosette for all stages of growth monitored thus far. However, none flowered as S.

parksii.

Soil-Intact Transplants – Spring 2007

Flower and leaf rosette production of soil-intact transplants has been similar to

undisturbed Spiranthes parksii/S. cernua leaf rosettes on site, and has surpassed percent

production of undisturbed individuals (Fig. 4). These plants have also exhibited considerable

variability. Some have remained dormant for as many as four growing seasons before

emerging as a flowering stalk or leaf rosette. Five flowered as S. parksii, of which two have

flowered all three consecutive years. Other individuals have flowered as S. cernua or remain

unknown as to the species due to herbivory before identification could be confirmed.

Fig. 4. Percent production of S. parksii/S. cernua transplanted and undisturbed leaf rosettes (spring 2007) each

growing season post-transplantation. Numbers in bars represent the number of individuals observed each

growing season.

Bare-Root Transplants – Spring 2008

Percent leaf rosette and flower production of these transplants have been consistently lower

than undisturbed leaf rosettes on site (Fig. 5). However, one individual flowered as Spiranthes

parksii in 2008, and other individuals are still producing vegetatively including one of the

known S. parksii. Sixteen appeared to be destroyed by feral hogs during winter of 2008 after

transplantation. Despite this disturbance, three individuals transplanted the area emerged as

leaf rosettes in 2009.


44

Fig. 5. Percent production of S. parksii/S. cernua transplants and undisturbed leaf rosettes each growing season

post-transplantation. Numbers in or above bars represent the number of individuals observed each growing

season.

Spiranthes parksii Bare-Root Transplants – Spring 2009

Four of the 14 (28%) flowered and four (28%) produced a rosette of leaves in fall 2009

following transplantation. One of the six which had <5 cm of a root tuber taken flowered,

while three produced a rosette of leaves.

DISCUSSION

Transplanted individuals for both methods appeared in subsequent years. Bare-root

transplants from spring 2007 have had a higher percentage of post-production than those in

spring 2008 and soil-intact in spring 2007. However, this could be due to a low sample size.

While percentages are consistently lower for bare-root transplants in spring 2008, one of 59

has flowered as Spiranthes parksii and some are still persisting vegetatively. Soil-intact

transplants have produced the best results for S. parksii since five individuals have flowered as

S. parksii, of which two have flowered all three flowering seasons monitored. Additionally,

percent production of these has been higher than undisturbed plants in the last three growing

seasons. Bare-root transplantation of 14 known S. parksii from spring 2009 which flowered at

least once in the previous two years have produced inflorescences and leaf rosettes after

transplantation, including those that had <5 cm of a root tuber removed.


45

Pileri (1998) noted that after excavating five Spiranthes cernua plants to analyze the

root tubers for mycorrhizal infection, all but one plant that was destroyed by a small mammal

survived transplantation by reappearing the next year. She also noted that they were better

able to survive when transplanted during the vegetative or early reproductive phases.

However, others believe, or have found, that bare-root transplanting of terrestrial orchids is

unsuccessful (Ferry, 2008; Steinauer, 2008). In this study, S. cernua (spring 2007) and S. parksii

(spring 2008 and spring 2009) responded positively to bare-root transplantation. The three S.

cernua that flowered after bare-root transplantation in spring 2007 were of the larger leaf

rosettes. The success of these could be due to large underground root tubers which could be

used to offset the effects of disturbance caused by transplantation. As well, mature S. parksii

transplanted in spring 2009 could also be using underground reserves to offset the effects of

transplantation.

Previous efforts of soil-intact transplantation of Spiranthes parksii using 15 cm diameter

irrigation pipe at the TMPA Gibbons Creek Lignite Mine conservation areas yielded positive

results (Parker 2006). However, quantitative data and long-term observations were not made.

Efforts of soil-intact transplanting in other terrestrial orchids have been unsuccessful, as with

Isotria medeoloides (Brumbeck, 1996). In this study, both Spiranthes parksii and S. cernua

responded well to this method. In fact, compared to undisturbed leaf rosettes at the study site,

percent production of soil-intact transplants have been greater in the last three growing

seasons. This might be due to placement of transplants since they were placed in areas of ideal

habitat of S. parksii/S. cernua. Undisturbed leaf rosettes may be persisting in areas which have

become unfavorable for flowering due to woody encroachment. However, quantitative data

would need to be collected to verify this.

While all transplants were placed in areas where Spiranthes parksii/S. cernua occurred,

placement could possibly be influencing post-production since microhabitats vary greatly

within a savanna patchwork. Additionally, initial size of leaf rosettes prior to transplantation

could affect post-production. However, detailed analysis of microhabitats and plant sizes

would need to be conducted to pursue these hypotheses.

CONCLUSIONS

While both methods of transplantation have yielded positive post-production in

individuals, if given the time and labor, the soil-intact method would be preferred. Not only

has this method yielded higher survival, but the intact soil may contain tubers of plants other

than the target individual. Upon digging up one Spiranthes parksii for bare-root transplanting

in spring 2009, another individual was found dormant as a root tuber. This was also seen

when taking soil samples around individual plants. Upon returning to the laboratory to sieve

soil samples, a Spiranthes spp. root tuber was found.

Comparison of transplanted individuals to undisturbed plants of the same species is

critical in giving accurate results of success or failure. If given only the results of transplanted

individuals in this study, one might conclude individuals are dying due to transplantation.


46

However, transplanted and undisturbed individuals have both declined and/or fluctuated in

subsequent production after transplanting was initiated. Long-term monitoring of these

individuals is crucial to clarify life history characteristics and environmental variables that

influence the persistence of undisturbed and transplanted individuals.

ACKNOWLEDGEMENTS

We would like to thank the Brazos Valley Solid Waste Management Agency (BVSWMA) for funding and HDR,

Inc. for assistance with this research. Individuals to thank are Linda Langlitz, Josh Grace, Martha Ariza, and

Trey Witcher for their assistance in transplanting and monitoring of individual plants.

J Ryan Hammons, Fred E. Smeins & William E. Rogers

Department of Ecosystem Science and Management, Texas A&M University, College Station, Tex.

[email protected]

LITERATURE CITED

Brown, P.M. & S.N. Folsom. 2008. Field Guide to the Wild Orchids of Texas. Gainesville: University Press of

Florida.

Bezanson, D. 2000. Natural Vegetation Types of Texas and Their Representation in Conservation Areas. The

University of Texas at Austin. http://www.abisw.org/bezanson/

Bridges, E.L. and S.L. Orzell. 1989. Additions of Noteworthy Vascular Plant Collections from Texas and

Louisiana, with Historical, Ecological and Geographical Notes. Phytologia 66: 12-69.

Brumback, W.E. and C.W. Fyler. 1996. Small Whorled Pogonia (Isotria medeoloides) Transplant Project. In Falk,

D.A., C.I. Millar, and M. Olwell. 1996. Restoring Diversity: Strategies for Reintroduction of Endangered

Plants. Washington, D.C.: Island Press,

Ferry, R.J. 2008. Relocating Terrestrial Orchid Plants. North American Native Orchid Journal 14: 179-82.

Hammons, J.R. 2008. Demographic, Life Cycle, Habitat Characterization and Transplant Methods for the

endangered orchid, Spiranthes parksii Correll. M.S. Thesis, Department of Rangeland Ecology and

Management, Texas A&M University, College Station, Texas.

Parker, K.M. 2006. Personal communication. Texas Ecological Services, College Station, Texas.

Pelchat, C. 2005. Spiranthes parksii Correll – Navasota Ladies‖ Tresses. McAllen International Orchid Society

Journal 6: 9-15.

Pileri, V.S. 1998. Root morphology, distribution of mycorrhizae, and nutrient status of the terrestrial orchid

Spiranthes cernua. M.S. Thesis, Department of Biology, University of Nebraska at Omaha, Omaha,

Nebraska.

Steinauer, G. 2008. Transplanting a Rare Orchid. Nebraska Game and Parks Commission Annual Report of the

Wildlife Conservation Fund.

Texas Parks and Wildlife Department. Accessed 2009. Post Oak Savanna and Blackland Prairie Wildlife

Management. http://www.tpwd.state.tx.us/landwater/land/habitats/post_oak/

United States Fish and Wildlife Service. 2009. Navasota Ladies‖-Tresses (Spiranthes parksii) 5-Year Review:

Summary and Evaluation. Austin Ecological Services Field Office, Austin, Tex.


http://www.abisw.org/bezanson/

http://www.tpwd.state.tx.us/landwater/land/habitats/post_oak/

Stewart & Hicks: PROPAGATION AND CONSERVATION STATUS OF NATIVE ORCHIDS

47

PROPAGATION AND CONSERVATION STATUS OF THE NATIVE

ORCHIDS OF THE UNITED STATES (INCLUDING SELECTED

POSSESIONS), CANADA, ST. PIERRE ET MIQUELON,

AND GREENLAND

Scott Stewart, Ph.D. & Aaron Hicks

INTRODUCTION

Conservation of biodiversity has become a primary biological, economic, and

humanistic concern as the global community faces the sixth great extinction event in the

Earth's history (Canadell and Noble, 2001). The implementation of conservation efforts must

begin with careful planning, otherwise we risk the Johnny Appleseed effect of biodiversity

conservation—we scatter our efforts into the wind and whichever efforts result in fruitful

conservation we consider successful and all others we consider unproductive. This naive

approach to the conservation of biodiversity runs the risk of missing important biotic

components of global biodiversity.

This paper is an attempt to gather verifiable propagation, cultivation, and conservation

status information on the native orchids of the United States, Canada, and associated foreign

lands in one document that may be used to guide future orchid conservation efforts in these

regions. We have chosen to include all orchid species and varieties considered native to the

United States, Canada, Puerto Rico, the U.S. Virgin Islands, Guam, Saint Pierre et Miquelon

islands, and Greenland. Species and varieties considered as introduced, exotic, escaped from

cultivation, and waifs have been excluded. Color and growth forms have also been excluded as

their taxonomic status and genetic stability are often controversial. All propagation and

conservation data has been verified through scientific publications and personal

communications with experts in orchid propagation, cultivation, and conservation. In general,

the most recent taxonomic checklist proposed by Brown (2009) has been used throughout.

The current work represents the first effort to gather such a volume of specific information

for such a large number of species and widespread geographic area, and should be considered a

working draft. The authors invite comments and additional verifiable data from readers.


48

GEOGRAPHIC ANALYSIS

Although national boundaries are political rather than biological, a brief analysis may

be enlightening with respect to the conservation of orchid species. From a financial

standpoint, the geographic areas reported here encompass a significant proportion of global

productivity (Table 1).

Table 1—Gross domestic products of major regions in current work. Data summarized from World Bank,

World Development Indicators, and CIA World Factbook.

Country Gross Domestic Product (US $ billions)

United States 14200 (2008)

Canada 1400 (2008)

Puerto Rico 67.9 (2001)

US Virgin Islands 2 (1993)

Guam 2.5 (2005 est.)

Total 15672

With approximately 410 taxa, this comes out to about $38.3 billion in average

domestic productivity per species in the geographic regions surveyed. A conservative figure

for the number of species globally is approximately 24,000 taxa, with the regions surveyed

here making up only 1.7% of the total. With global productivity calculated at $60.6 trillion in

2008 (World Bank), a figure of $2.52 billion in productivity per species is reached, a

substantially smaller figure. To paraphrase the International Union for Conservation of

Nature (IUCN), if orchid growers cannot pull a plant back from the brink of extinction, what

hope is there for other plant families? To extend this statement, if economic powerhouses

with all their resources

cannot preserve their

own species—which are

relatively few in number

when compared to the

global diversity—what

hope is there for other

orchids?

Hawaii

Despite the abun-

ance of hybrids and

introduced species, the

true native orchid taxa of

Hawaii are limited to

three species, one of

which (Platanthera holo-

chila, puahala-a-kane;

Fig. 1) is listed as Fig. 1. Platanthera holochila (puahala-a-kane) in natural habitat in Hawaii.

L. Zettler


49

threatened under the U.S. Endangered Species Act. There has been some propagation work

with this species—symbiotic germination efforts using mycobionts isolated from local

Hawaiian populations were unsuccessful while symbiotic efforts using non-Hawaiian

mycobionts have been successful, and there was no desire to introduce non-native mycobionts

from outside the islands. There has been reasonable success in propagating the species using

asymbiotic methods (McDonald et al., 2006; L. Zettler, personal communication).

Guam

Of the species native to Guam, only a small number have been successfully propagated

and brought into horticultural cultivation. Little is known of the plants of the island,

although the genera represented should be considered generally straightforward in asymbiotic

culture systems. It seems likely they would present few difficulties in terms of artificial

propagation. Nothing is known of the natural mycobionts or symbiotic culture requirements

of the orchids of Guam.

Western United States and Canada

This region presents myriad natural biomes: ranging from desert, to Pacific rain forest,

to true Arctic environments in the northernmost portions of Alaska. Several species native to

this region are showy (Cypripedium californicum, California lady's-slipper; Fig. 2) and

warrant additional propagation effort. Even in the arid desert states orchids are surprisingly

well-represented, found in all but three and ten counties of Arizona and New Mexico

respectively (Coleman, 2002).

Fig. 2. Cypripedium californicum (California lady‖s-slipper) photographed in southern Oregon, U.S.A.

S.L. Stewart


50

Federally protected species of interest in this region include Piperia yadonii (Yadon's

piperia), which has eluded attempts at cultivation, although there has been some recent

interest in the ecology and propagation of the species (George et al., 2009; Sharma et al., 2007;

R. Buck, personal communication). A locally endemic species, Spiranthes delitescens (Canelo

Hills ladies'-tresses), has proven to be remarkably easy to grow in culture (Hicks, 2007),

producing large numbers of plants from friable callus when stressed in sterile tissue culture.

Seedlings have flowered in cultivation at University of Arizona. A total of four populations

are known, although a fifth has been reported (M. Falk, personal communication).

Another species of interest in the region is Spiranthes infernalis (Ash Meadows ladies'-

tresses), known from a cluster of populations over an area of approximately 28 acres in Nye

County, Nevada. Estimates as to its total population numbers vary, but the species‖ global

population is estimated in the low one thousands, possibly as low as 1107 (Morefield, 2001).

There has reportedly been an effort to propagate the species at Royal Botanic Gardens at Kew.

Central United States and Canada

With the change of the American prairie, Platanthera leucophaea (eastern prairie

fringed orchid; Fig. 3) has dwindled in numbers and is currently listed as Federally

threatened. Asymbiotic efforts to propagate the species have met with limited success

(Stoutamire, 1996); however, symbiotic

propagation efforts have been highly successful

(Stewart, 2000; Zettler, 1999; Zettler et al., 2005;

Zettler et al., 2001). Also afforded Federally

threatened status, P. praeclara has been the subject

of extensive asymbiotic and symbiotic

propagation efforts. Asymbiotic efforts with P.

praeclara have been met with reasonable success

after lengthy culture periods and multiple cold

treatments of in vitro seed (From and Read,

1998). Sharma et al. (2003) reported the successful

symbiotic propagation of the species.

Also native to this region is the Federally

threatened Texas endemic Spiranthes parksii

(Navasota ladies’-tresses). There has been some

success in propagating this species from seed by

researchers at the Atlanta Botanical Garden.

Wilson (web page) reports the species has proven

to be remarkably easy to cultivate using

asymbiotic techniques. Additional asymbiotic and

symbiotic propagation efforts are currently

underway (R. Hammons, personal com-

munication).

Fig. 3. Platanthera leucophaea (eastern prairie fringed

orchid) photographed in southern Wisconsin, U.S.A.

P. M. Brown.


51

Eastern United States and Canada, excluding Florida

The eastern United States are home to a number of Platanthera species that are state-

listed as endangered, in many cases. Many of these species have proven to be resistant to

traditional propagation techniques, and success has been sporadic to nonexistent. Similar to P.

leucophaea, many species have been severely impacted by a variety of anthropogenic factors

such as urbanization, agriculture, and fire suppression.

In addition to the many Platanthera species of this region, Isotria medeoloides (small

whorled pogonia), known from the eastern United States, is listed as Federally threatened,

and had been the focus of many ecological and horticultural studies. The species has resisted

numerous attempts at artificial propagation; however, new efforts by the Smithsonian

Environmental Research Center and the U.S. Park Service are planned (J. O'Neill, personal

communication).

Florida

With more than a hundred species

known from Florida alone, it is practical to

treat the state as a separate entity for

conservation purposes. In addition to the

intrinsic diversity present in Florida is the

urban development the state has undergone,

in conjunction with the influx of invasive

species that further threatens the state's

native species. Perhaps of greatest interest is

the leafless Dendrophylax lindenii (ghost

orchid; Fig. 4), which is an endangered

species under state law. However, the plant

has proven to be remarkably easy to grow in

vitro, provided seeds can be reliably

produced. Unfortunately, mortality is high

amongst deflasked plantlets and growth is

slow; it seems likely there is some aspect of

the species' cultivation that remains cryptic

such that it may eventually be brought into

cultivation—although this is a theme that has

been repeated for many decades without

realization (Correll, 1978).

Afforded similar state-level protection

is Cyrtopodium punctatum (cigar orchid),

which forms large (to 1.5 meter), robust

plants whose populations have been

decimated by poaching and habitat

Fig. 4. Dendrophylax lindenii (ghost orchid)

photographed in southwestern Florida, U.S.A.

S. L. Stewart


52

alteration. However, the species forms large capsules producing similarly large quantities of

seeds that germinate and grow on a variety of media with no special requirements in

asymbiotic culture. From a mechanical standpoint, the roots are not like those of most

orchids, forming tangled mats in vitro, resulting in damage when deflasked. From this, it may

be best to plant seedlings in individual tubes. Considerable research effort has been focused on

the asymbiotic propagation, reproductive biology, and potential for reintroduction of the

cigar orchid (Dutra, 2008; Dutra et al., 2009a,b). Preliminary symbiotic germination and

reintroduction of the species has been successful (Stewart and Richardson, 2008; S. Stewart,

unpublished data).

Another state-listed endangered species is Basiphyllaea corallicola (Carter's orchid),

which is easy to propagate asymbiotically, forming new pseudobulbs with fresh shoots every

few months. Similarly protected, both Epidendrum nocturnum (night-fragrant epidendrum)

and Macradenia lutescens (Trinidad macradenia) offer no difficulties in vitro. A symbiotic

propagation protocol has even been developed for E. nocturnum (Zettler et al., 2007) and

preliminary reintroductions have taken place (Stewart, 2008). Another species whose numbers

have declined to the point where it has been listed as endangered by the state is Tolumnia

bahamensis (Florida's dancing lady); germination is straightforward, while subsequent culture

is made difficult by the usual cultural quirks within the Oncidiinae in that differentiation is

slow, resulting in large clusters of protocorms without roots. These problems usually resolve

with subsequent replating, resulting in large numbers of plants that eventually form stout

seedlings with good roots. Mortality is high when deflasked. Bletia purpurea (pine-pink) is

another Floridian species whose populations face decline as urbanization in southern Florida

increases. The germination, in vitro culture, and transfer to the greenhouse of the species is

quite easy (Dutra et al., 2008); however, the species is rarely seen in the commercial

marketplace.

Although not afforded protection, Eulophia alta (wild coco) is a strong grower in

sterile flask, although others have noted that it germinates and grows quite readily from seed

sown directly in rich earth. Johnson et al. (2007) presented a side-by-side comparison of

asymbiotic versus symbiotic germination this species, demonstrating preliminary evidence of

an advantage during symbiotic germination and subsequent in vitro seedling development.

Several other Florida native species are considered endangered, and lack active

conservation efforts, such as Beloglottis costaricensis (Costa Rican ladies'-tresses),

Bulbophyllum pachyrachis (rat-tail orchid), Cyclopogon cranichoides (speckled ladies'-tresses),

Epidendrum amphistomum (dingy-flowered star orchid), Ionopsis utricularioides (delicate

ionopsis), Liparis elata (tall twayblade), Mesadenus lucayanus (copper ladies'-tresses),

Polystachya concreta (pale-flowered polystachya), Vanilla barbellata (worm-vine), and Vanilla

mexicana (thin-leaved vanilla), among others. The State of Florida affords some 54 taxa

endangered status, 16 more are protected as threatened, and two more—Encyclia tampensis

(Florida butterfly orchid) and Epidendrum magnoliae var. magnoliae (green-fly orchid; syn.

Epidendrum conopseum)—are protected as commercially exploited.


53

A number of Florida native orchids are also found throughout the Greater and Lesser

Antilles—including Cuba, Puerto Rico, and the U.S. Virgin Islands. Cuba is not treated in this

paper. While the degree of this wider geographic distribution can be seen in comparing the

native orchid flora of Florida to that of Puerto Rico, other islands in the Antilles may also

possess some species known from Florida. For example, Dendrophylax lindenii, Cyrtopodium

punctatum, Eulophia alta, and Bletia purpurea are all known from Cuba and Florida (Llamacho

and Larramendi, 2005).

Puerto Rico

Much of the primary forest of Puerto Rico was removed early in the 20th century and

it is difficult to state with certainty that any species of orchid were lost. However, two

endemic species—Lepanthes caritensis (Carite babyroot orchid) and the Federally endangered

Lepanthes eltoroensis (Luquillo Mountain babyroot orchid)—are known only from very

small populations (Tremblay et al., 1998) and a third Puerto Rican orchid, Cranichis ricartii

(Puerto Rico helmet orchid), is Federally endangered and may be extirpated. Relatively little

is known about the propagation of most orchids found in Puerto Rico.

U.S. Virgin Islands

Twenty-six orchid species are known from the U.S. Virgin Islands, all of which are

also found in Puerto Rico (Ackerman, 1995). St. Thomas is the most species-rich with 23

species, followed by St. John (13 species), and St. Croix (9 species). Taken as a whole, 29

species are known from both the U.S. and British Virgin Islands. Only Eulophia alta and

Erythrodes hirtella (false helmet orchid) are known from the British Virgin Islands and not

from the U.S. Virgin Islands. Ackerman (1995) commented that additional species may have

been present in both the U.S. And British Virgin Islands at one time, but rapid urbanization

on the populated islands may have caused some species to become extirpated from the islands.

As in Puerto Rico and Florida, the U.S. Virgin Islands also have a number of exotic orchids

that have escaped cultivation and become established on various islands, including

Spathoglottis plicata (Philippine ground orchid; St. Thomas), Vanilla mexicana (Mexican

vanilla; St. Croix), and V. planifolia (commercial vanilla; St. Croix, St. John, and St.

Thomas).

HORTICULTURAL NOTES

Several of the genera of interest here remain recalcitrant to existing propagation

techniques. Genera such as Corallorhiza remain cryptic in their in vitro cultural needs,

although Jay O'Neill (personal communication) notes that sporadic germination on

commercial media has occurred, while semi-wild field plantings have been met with good

success, including one recruit growing to flower. The reliance of mycotropic interactions for

the seed germination and plant development of saprophytic terrestrial genera (i.e.,

Corallorhiza, Hexalectris) may be the root cause for their difficulty in in vitro propagation.

The genus Cypripedium is by far the most successful in terms of propagation and

commercialization, with a number of species being made available for sale on a regular basis.

In fact, the majority of the North American species are available through a number of


54

reputable companies as mature plants, and it is expected that this market has considerable

potential for gardeners that would like to engage in the horticultural cultivation of these

species. Several species have been presented as candidates for commercial propagation; prices

and availability have improved

over the past several years. The

genus is desirable and

conspicuous, making it an ideal

subject for commercialization in

order to potentially reduce the

collection of plants from the

wild.

Cyrtopodium punctatum is

perhaps one of the most

underrepresented species with

respect to propagation. It is so

fecund and such a strong grower

that it would seem to be an ideal

candidate for commercial

exploitation in its native state of

Florida. The decimation of wild

populations of the species

implies its desirability could

prove useful to nurseries that are

willing to produce specimens for

local growers. Similarly,

Eulophia alta, Epidendrum

nocturnum, and Encyclia tamp-

ensis are readily propagated;

Eulophia alta could probably be

introduced into popular culture

much in the same way as the

other two have. Occasionally

available from commercial

vendors, Epipactis gigantea

(stream orchid) is an ideal

candidate for western gardens.

Both Calypso bulbosa var. americana (eastern fairy-slipper) and C. bulbosa var.

occidentalis (western fairy-slipper; Fig. 5) are diminutive, but showy, native orchids with

tremendous horticultural potential. The species has frustrated many individuals attempting in

vitro propagation with sporadic germination and limited subsequent growth. Ashmore (1999)

has reported the successful in vitro asymbiotic propagation and ex vitro cultivation of both

Calypso varieties, and numerous man-made Calypso hybrids. The scaling-up of such work

Fig. 5. Calypso bulbosa var. occidentalis (western fairy-slipper)

photographed in southern Oregon, U.S.A.

S.L. Stewart


55

would be required before Calypso would be regularly available to gardeners; however, such

propagation, hybridization, and horticultural selection work are important first-steps toward

commercialization of these showy native orchids.

The genus Goodyera has been popular for many years as a plant for culture in terraria;

other allied members of the genus have been available as jewel orchids in the trade for years,

with few—if any—from artificial propagation. Propagation of jewel orchids for the purposes

of horticulture is most commonly by division. It seems likely most, if not all, of the native

Goodyera species in the horticultural trade have been wild-collected. It would be desirable to

provide for this genus in artificial propagation as well.

The genus Habenaria is almost unknown with respect to commercial asymbiotic seed

propagation, as with members of the genus Listera, Malaxis, Hexalectris, Piperia, Platanthera,

and Spiranthes. There is little to say about this otherwise unrelated group other than to say

that new techniques, media, and mycobionts will have to be developed to master this group of

plants. The exception would be the genus Spiranthes, which has proven to be prolific in sterile

culture for the most part. The propagation of Habenaria species has received considerable

attention in recent years, particularly as a part of habitat or integrated conservation efforts in

Florida (Stewart, 2007; Stewart and Kane, 2006a, b; Stewart and Zettler, 2002).

Tropical epiphytic genera, including Bulbophyllum, Dendrobium, Encyclia,

Epidendrum, Lepanthes, Maxillaria, Oncidium, Pleurothallis, and so forth, have a better

outlook than terrestrial species, if for no better reason conventional commercial orchid

propagation laboratories are better suited to the propagation of these plants, many of which

can be produced in large numbers if the economic incentives are present, and fresh, clean

material can be produced.

CONCLUDING REMARKS

In total, there are six species afforded Federal protection as endangered species, the

highest level of protection: Cranichis ricartii, Lepanthes eltoroensis, Piperia yadonii, Platanthera

holochila, Spiranthes delitescens, and Spiranthes parksii. Three species—Isotria medeoloides,

Platanthera praeclara, and P. leucophaea—are considered Federally threatened. Of these, only

four—Spiranthes delitescens, S. parksii, Platanthera praeclara, and P. leucophaea—have been

artificially produced in substantial quantities. Several other species are limited to small

colonies in single geographic areas (Platanthera pallida, Spiranthes infernalis), exist as

exceedingly small populations (Lepanthes caritensis), have cryptic germination requirements

(Calypso bulbosa, Isotria medeoloides, Platanthera spp.), are achlorophyllous (Corallorhiza spp.),

or survive transfer from sterile tissue culture rarely or not at all (Dendrophylax lindenii, some

Cypripedium spp.). Indeed, listing or upgrading some species from threatened to endangered

status is probably merited, although from a conservation perspective this makes attempts to

propagate these species markedly more difficult.

While most species that have been targeted in propagation efforts have been brought

into cultivation, a few remain refractory to existing techniques—or the material provided for


56

propagation has been so scant that insufficient experiments could be run. A few, such as

Platanthera species, have sporadic success that is reminiscent of that with Paphiopedilum

species; if this is the case, perhaps we will see enhanced germination with subsequent

generations, with each passage through sterile flask (or symbiotic culture) selecting those seeds

that are likely to germinate under such conditions. It is our hope that, given the resources of

the geographic region discussed here, all species could eventually be propagated and made

available for culture in public and private gardens, or reintroduction if the need arises.

Asymbiotic and symbiotic propagation status and current conservation status for

species found within the geographic region of the United States, Canada, Puerto Rico, the

U.S. Virgin Islands, Guam, Greenland, and Saint Pierre et Miquelon is found in Table 2.

ACKNOWLEDGEMENTS

The authors would like to thank the numerous individuals who contributed personal

communications and information for this paper. Paul Martin Brown was instrumental during

editing and revision of this work.

Scott Stewart, Ph.D., Director, Horticulture & Agriculture Programs, Kankakee Community College,

Kankakee, IL 60901 [email protected]

Aaron Hicks, The Orchid Seedbank Project, P.O. Box 7042, Chandler, AZ 85246, [email protected]

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Dutra, D., T.R. Johnson, P.J. Kauth, S.L. Stewart, M.E. Kane, and L. Richardson. 2008. Asymbiotic seed

germination, in vitro development, and greenhouse acclimatization of the threatened terrestrial orchid

Bletia purpurea. Plant Cell, Tissue and Organ Culture 94:11-21.

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Lindley. American Midland Naturalist 145: 168-75.


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Table 2—Propagation and conservation statuses for orchids species native to survey regions. Geographic range

key: US=United States (including Alaska), HI=Hawaii, CAN=Canada, SPM=St. Pierre et Miquelon,

G=Greenland, PR=Puerto Rico, VI=U.S. Virgin Islands, and GU=Guam. Propagation status key: 1=available

commercially on a regular basis, 2=has been available commercially in recent past, 3=is routinely maintained in

laboratory culture, 4=has been produced experimentally in laboratory culture, 5=experimental culture has had

limited success, and 6=experimental culture has been unsuccessful. Conservation status key: F=Federally

protected as endangered, f=Federally protected as threatened, S=state protected as endangered, s=state protected

as threatened, o=state protected by other designation. Propagation data from botanic gardens have been capped

at ―3‖ as their role generally does not involve the production of plants for commercial ventures; nd = no data.

Taxa

Geographic Range In vitro

Propagation -

Asymbiotic

In vitro

Propagation -

Symbiotic

Conservation

Status Reference(s)

Amerorchis rotundifolia US, CAN,G nd nd S, s 11

Anoectochilus sandvicensis HI nd nd nd 22

Aplectrum hyemale US, CAN 2,5 4 S,s,o 3,11,14,23,24

Arethusa bulbosa US, CAN, SPM nd nd S, s, o 11

Basiphyllaea corallicola US, PR 3, 4 nd S 2, 11, 23

Beloglottis costaricensis US nd nd S 11

Bletia patula US, PR 4, 5 nd nd 11, 14, 23

Bletia purpurea US 2, 4 6 S 2, 6, 11, 24

Brachionidium ciliolatum PR nd nd nd 11

Brachionidium sherringii PR nd nd nd 11

Brassavola cucullata PR, VI 2, 3 nd nd 2, 8, 11, 24

Brassavola nodosa PR 1, 2, 3 nd nd 1, 2, 11, 23, 24

Brassia caudata US 3 nd S 11, 24

Broughtonia domingensis PR nd nd nd 11

Bulbophyllum guamense GU 2 nd nd 9, 20, 23

Bulbophyllum longiflorum GU 3 nd nd 9, 20

Bulbophyllum pachyrhachis US nd nd nd 18

Bulbophyllum profusum GU 6 nd nd 9, 20, 23

Calanthe triplicata GU 3 nd nd 9, 20, 24

Calopogon barbatus US 3 nd nd 11, 24

Calopogon multiflorus US 3, 6 nd S 11, 14

Calopogon oklahomensis US 3 nd nd 11, 24

Calopogon pallidus US 3 nd nd 11, 24

Calopogon tuberosus var. simpsonii US 3, 4 5 S, o 6, 11

Calopogon tuberosus var. tuberosus US, CAN, SPM 1, 3, 5 5 S, o 1, 2, 6, 11, 13, 14, 24

Calopogon × fowleri US nd nd nd 19

Calopogon × goethensis US nd nd nd 19

Calopogon × obscurus US nd nd nd 19

Calopogon × simulans US nd nd nd 19

Calopogon × vulgaris US nd nd nd 19

Calypso bulbosa var. americana US, CAN 4 nd S, s, o 11, 23

Calypso bulbosa var. occidentalis US, CAN 4 nd S, s 11, 14

Campylocentrum micranthum PR 6 nd nd 2, 11

Campylocentrum pachyrrhizum US, PR 6 nd S 11, 23

Campylocentrum pygmaeum PR nd nd nd 11

Cephalanthera austiniae US, CAN nd nd nd 11, 18

Cleistesiopsis bifaria US 5, 6 nd nd 11, 14, 19, 24

Cleistesiopsis divaricata US 3 nd nd 11, 19, 24

Cleistesiopsis oricamporum US nd nd nd 19

Cleistesiopsis × ochlockoneensis US nd nd nd 19


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Taxa Range Asymbiotic Symbiotic Status Reference(s)

Cochleanthes flabelliformis PR 4, 6 nd nd 11, 23, 24

Coeloglossum viride var. virescens US, CAN nd nd nd 18

Coeloglossum viride var. viride US, CAN nd nd nd 18, 19

Coelogyne guamensis GU nd nd nd 9, 20

Comparettia falcata PR 3, 5 nd nd 2, 11, 23

Corallorhiza bentleyi US nd nd nd 11

Corallorhiza maculata var. maculata US, CAN, SPM nd 6 o 11, 22

Corallorhiza maculata var. mexicana US nd nd nd 18, 19

Corallorhiza maculata var. occidentalis US, CAN nd nd nd 11

Corallorhiza maculata var. ozettensis US nd nd nd 11

Corallorhiza mertensiana US, CAN nd nd nd 11

Corallorhiza odontorhiza var. odontorhiza US, CAN 5 4 S, s, o 3, 11

Corallorhiza odontorhiza var. pringlei US, CAN nd nd nd 11

Corallorhiza striata var. striata US, CAN nd nd S, o 11

Corallorhiza striata var. vreelandii US, CAN nd nd nd 11

Corallorhiza trifida US, CAN, SPM, G nd nd S, s, o 11

Corallorhiza wisteriana US nd nd S, o 11

Corymborkis forcipigera PR nd nd nd 11

Corymborkis veratrifolia GU nd nd nd 9

Cranichis muscosa US, PR 3 nd S 11, 24

Cranichis ricartii PR nd nd F, S 11

Cranichis tenuis PR nd nd nd 8

Cyclopogon cranichoides US, PR 6 nd nd 11, 23

Cyclopogon elatus US, PR, VI 6 nd S 8, 11, 23

Cyclopogon miradorense PR nd nd nd 11

Cypripedium acaule US, CAN, SPM 1, 2, 3 4 S, o 1, 5, 11, 16, 24

Cypripedium arietinum US, CAN 1, 2 nd S, s, o 5, 11, 16

Cypripedium californicum US 1, 5 nd nd 1, 5, 11, 14

Cypripedium candidum US, CAN 1, 3, 5 nd S, s, o 1, 5, 11, 14

Cypripedium fasciculatum US 2, 6 nd o 5, 11, 14

Cypripedium guttatum US, CAN 1 nd nd 5, 11, 14

Cypripedium kentuckiense US 1, 3, 5 nd S, o 5, 11, 13, 14, 15,16, 17, 24

Cypripedium montanum US, CAN 1, 5 nd nd 11, 14, 16

Cypripedium parviflorum var. makasin US, CAN 1, 4 nd S, o 1, 5, 11

Cypripedium parviflorum var. parviflorum US 1, 3, 6 nd S, s, o 11, 14, 15, 17, 24

Cypripedium parviflorum var. pubescens US, CAN, SPM 1, 3, 6 nd S, s, o 1, 5, 11, 14, 15, 16, 17, 24

Cypripedium passerinum US, CAN 1 nd nd 11, 16

Cypripedium reginae US, CAN 1, 4, 5 nd S, s, o 1, 5, 11, 14, 15, 16, 17

Cypripedium yatebeanum US 1 nd nd 5, 11

Cypripedium × alaskanum US 1 nd nd 11, 16

Cypripedium × andrewsii nm. andrewsii US, CAN 1, 3 nd nd 1, 11, 15, 16

Cypripedium × andrewsii nm. landonii US, CAN nd nd nd 11

Cypripedium × columbianum US, CAN nd nd nd 11

Cypripedium × herae US, CAN nd nd nd 19

Cyrtopodium macrobulbon US 6 nd nd 2, 11, 18, 19

Cyrtopodium punctatum US, PR 1, 3, 4 4 S 2, 6, 11, 14, 24

Dactylorhiza aristata var. aristata US 5 nd nd 11, 14

Dactylorhiza aristata var. kodiakensis US nd nd nd 11

Dactylorhiza praetermissa var. praetermissa CAN 1 nd nd 16, 18, 19

Deiregyne confusa US nd nd nd 11

Dendrobium guamense GU nd nd nd 9, 20

Dendrobium phillippense GU nd nd nd 9, 20


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Dendrobium scopa GU nd nd nd 9, 20

Dendrophylax lindenii US 1, 3 5 S 2, 6, 11, 12, 24

Dendrophylax porrectus US, PR nd nd nd 18, 19

Dichaea hystricina PR nd nd nd 11

Dichaea latifolia PR nd nd nd 11

Dichaea pendulata PR 3 nd nd 1, 8

Dichromanthus cinnabarinus US nd nd nd 11

Dichromanthus michuacanus US nd nd nd 19

Didymoplexis fimbriata GU 6 nd nd 9, 20, 23

Dilomilis montana PR nd nd nd 11

Domingoa haematochila PR nd nd nd 11

Elleanthus cordidactylus PR nd nd nd 11

Eltroplectris calcarata US, PR 2, 6 nd S 2, 11, 23

Encyclia gravida PR nd nd nd 11

Encyclia isochila PR nd nd nd 11

Encyclia pygmaea US, PR nd nd S 11

Encyclia rufa US 3, 4 nd nd 2, 11, 23

Encyclia tampensis US 1, 3 4 o 2, 11, 22, 24

Epidendrum acunae US nd nd S 11

Epidendrum anceps PR, VI 2, 3 nd nd 8, 11, 23, 24

Epidendrum amphistomum US, PR, VI 3 nd S 11, 18, 22, 24

Epidendrum antillanum PR nd nd nd 11

Epidendrum boricuarum PR nd nd S 11

Epidendrum ciliare PR, VI 2, 3 nd nd 8, 11, 23, 24

Epidendrum floridense US 3, 6 nd nd 11, 23, 24

Epidendrum jamaicense PR 6 nd nd 11, 23

Epidendrum magnoliae var. magnoliae US 3 4 o 11, 22, 24

Epidendrum magnoliae var. mexicanum US 3 nd nd 11, 24

Epidendrum miserrimum PR nd nd nd 11

Epidendrum mutelianum PR nd nd nd 11

Epidendrum nocturnum US, PR 1, 3, 4 3, 4 S 2, 6, 11, 22, 24

Epidendrum ramosum PR nd nd nd 11

Epidendrum rigidum US, PR 3, 4 nd S 11, 22, 24

Epidendrum secundum PR 1 nd nd 11, 23

Epidendrum strobiliferum US nd nd S 11

Epidendrum tridens PR nd nd nd 11

Epidendrum vincentinum PR nd nd nd 8

Epipactis gigantea US, CAN 1, 4 nd o 2, 11, 16, 24

Eria rostriflora GU nd nd nd 9, 20

Erythrodes hirtella PR nd nd nd 11

Erythrodes plantaginea PR nd nd nd 11

Eulophia alta US, PR 2, 4, 5 4 nd 1, 2, 6, 11, 14, 24

Eulophia macgregorii GU nd nd nd 9, 20

Eulophia pulchra GU nd nd nd 9, 20

Eurystyles ananassocomos PR nd nd nd 11

Galeandra bicarinata US nd nd S 11

Galearis spectabilis US, CAN 5, 6 6 S, s, o 1, 3, 11, 22, 24

Geodorum densiflorum GU nd nd nd 9, 20

Goodyera oblongifolia US, CAN 1 nd S, o 11, 16

Goodyera pubescens US, CAN 4, 5, 6 4 S, o 2, 3, 11, 14, 22, 24

Goodyera repens US, CAN, SPM 6 nd S, o 11, 14

Goodyera tesselata US, CAN 6 nd S, s, o 11, 23


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Govenia floridana US 6 6 S 6, 11, 24

Govenia utriculata PR nd nd S 11

Gymnadeniopsis clavellata var. clavellata US 3 4, 5 nd 18, 19, 22, 24

G. clavellata var. ophioglossoides US nd nd nd 18, 19

Gymnadeniopsis integra US 5 5 S, s 11, 22, 24

Gymnadeniopsis nivea US 5 nd S, s 11, 24

Habenaria alata PR, VI nd nd nd 8, 11

Habenaria amalfitana PR nd nd nd 11

Habenaria distans US, PR 5 5 S 6, 11

Habenaria eustachya PR nd nd nd 11

Habenaria macroceratitis US 2, 4 4 nd 6, 11, 23

Habenaria monorrhiza PR, VI 6 nd nd 8, 11, 23

Habenaria odontopetala PR 2, 3 3 nd 6, 11, 23

Habenaria quinqueseta US 3 3 nd 6, 11, 24 ×

Habenaria repens US, PR 2, 3 4 nd 6, 11, 14, 24

Hapalorchis lineatus PR nd nd nd 11

Helleriella punctulata PR nd nd nd 11

Heterotaxis sessilis US nd nd nd 19

Hexalectris spicata var. arizonica US nd nd nd 18, 19

Hexalectris spicata var. spicata US 3 nd S, s, o 11, 24

Hexalectris grandiflora US nd nd nd 11

Hexalectris nitida US nd nd S 11

Hexalectris revoluta var. colemanii US nd nd nd 19

Hexalectris revoluta var. revoluta US nd nd nd 19

Hexalectris warnockii US nd nd o 11

Ionopsis satyrioides PR nd nd nd 11

Ionopsis utricularioides US, PR, VI 4 nd S 8, 11, 23, 24

Isochilus linearis PR 2, 6 nd nd 2, 11, 23

Isotria medeoloides US, CAN 6 6 f, S, s, o 3, 11, 22

Isotria verticillata US, CAN 6 nd S, s, o 11, 14, 24

Jacquiniella globosa PR nd nd nd 11

Jacquiniella teretifolia PR nd nd nd 11

Koellensteinia graminea PR 6 nd nd 11, 23

Leochilus puertoricensis PR nd nd nd 11

Lepanthes caritensis PR 6 nd nd 2, 11

Lepanthes dodiana PR nd nd nd 11

Lepanthes eltoroensis PR nd nd F, S 11

Lepanthes rubipetala PR nd nd nd 8

Lepanthes rupestris PR nd nd nd 11

Lepanthes sanguinea PR nd nd nd 11

Lepanthes selenitepala PR nd nd nd 11

Lepanthes veleziana var. retusicolumna PR nd nd nd 11

Lepanthes veleziana var. veleziana PR nd nd nd 11

Lepanthes woodburyana PR nd nd nd 11

Lepanthopsis melanantha US, PR 5, 6 nd S 11, 23, 24

Liparis elata US, PR, VI 5 nd S 8, 11, 24

Liparis guamensis GU nd nd nd 9, 20

Liparis hawaiensis HI nd 6 nd 22

Liparis liliifolia US, CAN 3 4 S, s 3, 11, 24

Liparis loeselii US, CAN 4 nd S, s, o 11, 23

Liparis saundersiana PR nd nd nd 11

Liparis vexillifera PR nd nd nd 11


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Liparis × jonesii US nd nd nd 11

Listera auriculata US, CAN nd nd S, s 11

Listera australis US, CAN nd nd S, s, o 11

Listera banksiana US, CAN nd nd nd 11

Listera borealis US, CAN 6 nd o 11, 23

Listera convallarioides US, CAN, SPM nd nd S, s, o 11

Listera cordata var. cordata US, CAN, SPM nd nd S, s, o 11

Listera cordata var. nephrophylla US, CAN nd nd nd 11

Listera smallii US 6 nd S, s, o 11, 24

Listera ×veltmanii US, CAN nd nd nd 11

Luisia teretifolia GU 6 nd nd 9, 20, 23

Lycaste barringtoniae PR nd nd nd 11

Macradenia lutescens US 2 nd S 2, 11

Malaxis abieticola US nd nd nd 21

Malaxis bayardii US, CAN nd nd S, o 11

Malaxis brachypoda US, CAN nd nd S, s, o 11

Malaxis corymbosa US nd nd o 11

Malaxis diphyllos US, CAN nd nd nd 11

Malaxis major PR nd nd nd 11

Malaxis massonii PR nd nd nd 11

Malaxis paludosa US, CAN nd nd S 11

Malaxis porphyrea US nd nd nd 11

Malaxis soulei US nd nd o 11

Malaxis spicata PR, US nd nd nd 11

Malaxis unifolia US, CAN, SPM 5 nd S, s, o 11, 24

Malaxis wendtii US nd nd nd 11

Maxillaria acutifolia PR nd nd nd 11

Maxillaria coccinea PR 3 nd nd 2, 11

Maxillaria parviflora US, PR nd nd S 11

Mesadenus lucayanus US, PR, VI nd nd S 11

Microthelys rubrocallosa US nd nd nd 19

NervillIa platychila GU nd nd nd 9, 20

Nervillia aragoana GU nd nd nd 9, 20

Nervillia jacksoniae GU nd nd nd 9, 20

Nidema ottonis PR nd nd nd 11

Oncidium altissimum PR, VI nd nd nd 8, 11

Oncidium floridanum US 5 5 S 6, 11, 24

Oncidium meirax PR nd nd nd 11

Pelexia adnata US, PR 5 nd S 11, 14

Phreatia minutiflora GU nd nd nd 9

Phreatia thompsonii GU nd nd nd 9, 20

Piperia candida US, CAN nd 6 nd 6, 11

Piperia colemanii US nd nd nd 11

Piperia cooperi US nd nd nd 11

Piperia elegans subsp. decurtata US nd nd nd 11

Piperia elegans subsp. elegans US, CAN nd 5 nd 6, 11

Piperia elongata US, CAN nd 5 nd 6, 11

Piperia leptopetala US nd nd nd 11

Piperia michaelii US nd nd nd 11

Piperia transversa US, CAN nd 6 nd 6, 11

Piperia unalascensis US, CAN nd 5 nd 6, 11

Piperia yadonii US nd nd F 11


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Platanthera aquilonis US, CAN nd nd nd

Platanthera blephariglottis US, CAN, SPM 3, 5 nd S, s, o 11, 14, 18, 24

Platanthera brevifolia US nd nd nd 11

Platanthera chapmanii US 2, 5 nd nd 11, 23, 24

Platanthera chorisiana US, CAN nd nd s 11

Platanthera ciliaris US, CAN 2, 3, 5 5 S, s 1, 11, 14, 22, 23, 24

Platanthera conspicua US 3 nd nd 18, 24

Platanthera convallariifolia US nd nd nd 21

Platanthera cristata US 5 4, 5 S, s, o 11, 22, 24

Platanthera dilatata var. albiflora US, CAN nd nd nd 18

Platanthera dilatata var. dilatata US, CAN, SPM 5, 6 nd S, s, o 1, 11, 14, 24

Platanthera dilatata var. leucostachys US, CAN nd nd nd 11

Platanthera flava var. flava US, CAN 5 nd S, s, o 11, 24

Platanthera flava var. herbiola US, CAN nd nd S, s, o 11

Platanthera grandiflora US, CAN, SPM 6 nd S, s, o 11, 24

Platanthera holochila HI 4, 5 6 F 22

Platanthera hookeri US, CAN 5 nd S, s, o 11, 24

Platanthera huronensis US, CAN nd nd o 11

Platanthera hyperborea G nd nd nd 25

Platanthera integrilabia US 3, 5 4 S, s 11, 14, 22, 24

Platanthera lacera US, CAN, SPM 3, 6 3 o 11, 24

Platanthera leucophaea US, CAN 3, 5 3 f, S, s, o 6, 11, 14, 22, 24

Platanthera limosa US nd nd o 11

Platanthera macrophylla US, CAN nd nd s 11

Platanthera obtusata subsp. obtusata US, CAN, SPM nd nd o 11

Platanthera obtusata subsp. oligantha US nd nd nd 11

Platanthera orbiculata CAN, SPM, US nd nd S, s, o 11

Platanthera pallida US nd nd nd 11

Platanthera peramoena US 4 nd S, s 11, 23, 24

Platanthera praeclara US, CAN 3, 6 3 f 1, 11, 14

Platanthera psycodes US, CAN, SPM 4, 6 nd S, s, o 1, 11, 14, 24

Platanthera purpurascens US nd nd nd 11

Platanthera shriveri US nd nd nd 19

Platanthera sparsiflora US nd nd o 11, 18

Platanthera stricta US, CAN nd nd o 11

Platanthera tescamnis US nd nd nd 19

Platanthera tipuloides var. behringiana US nd nd nd 11

Platanthera yosemitensis US nd nd nd 19

Platanthera zothecina US nd nd nd 11

Platanthera × andrewsii US, CAN nd nd nd 11

Platanthera × apalachicola US 5 6 nd 6

Platanthera × beckneri US nd nd nd 19

Platanthera × bicolor US 3 nd nd 19, 24

Platanthera × canbyi US nd nd nd 19

Platanthera × channellii US nd nd nd 19

Platanthera × correllii US nd nd nd 19

Platanthera × enigma US, CAN nd nd nd 19

Platanthera × estesii US nd nd nd 11

Platanthera × evansiana US nd nd nd 19

Platanthera × folsomii US nd nd nd 19

Platanthera × hollandiae CAN nd nd nd 19

Platanthera × keenanii US nd nd nd 19


64


Platanthera × kelleyi US nd nd nd 19

Platanthera × lassenii US nd nd nd 11

Platanthera × lueri US 3 nd nd 19, 24

Platanthera × osceola US 5 6 nd 19, 24

Platanthera × reznicekii CAN nd nd nd 19

Platanthera × smithii US nd nd nd 19

×Platanthopsis vossii US nd nd nd 19

Platythelys sagreana US nd nd nd 18

Platythelys querceticola US nd nd nd 11

Pleurothallis appendiculata PR nd nd nd 11

Pleurothallis aristata PR nd nd nd 11

Pleurothallis domingensis PR nd nd nd 11

Pleurothallis gelida US, PR 4, 6 nd S 11, 23, 24

Pleurothallis obovata PR nd nd nd 11

Pleurothallis pruinosa PR nd nd nd 11

Pleurothallis pubescens PR nd nd nd 11

Pleurothallis racemiflora PR 5 nd nd 2, 11

Pleurothallis ruscifolia PR nd nd nd 11

Pleurothallis wilsonii PR nd nd nd 11

Pogonia ophioglossoides US, CAN, SPM 2, 3 nd S, s, o 11, 23, 24

Polystachya concreta US, PR, VI 2, 3 nd S 2, 8, 11, 23

Polystachya foliosa PR, VI 2, 3 nd nd 2, 8, 11, 23

Ponthieva brittoniae US nd nd S 11

Ponthieva racemosa US, PR, VI 5, 6 nd S 6, 8, 11, 24

Ponthieva ventricosa PR nd nd nd 11

Prescottia oligantha US, PR, VI 5 nd nd 8, 14

Prescottia stachyodes PR, VI nd nd nd 8

Prescottia pellucid PR nd nd nd 8

Prosthechea boothiana var. erythronioides US 3 nd nd 19, 24

Prosthechea cochleata var. triandra US, PR, VI 1 4 nd 6, 8, 19

Prosthechea pygmaea US, PR nd nd S 11

Pseudorchis straminea CAN nd nd nd 11

Psilochilus macrophyllus PR nd nd nd 11

Psychilis kraenzlinii PR 4 nd nd 11, 23

Psychilis krugii PR nd nd nd 11

Psychilis macconnelliae PR, VI nd nd nd 8

Psychilis monensis PR 6 nd nd 8, 23

Pteroglossaspis ecristata US 3 nd nd 18, 24

Pteroglossaspis pottsii US 5 nd nd 19, 24

Rhynchophreatia micrantha GU nd nd nd 9

Sacoila lanceolata US, PR, VI 2, 3, 4, 5 6 nd 1, 8, 24

Sacoila paludicola US nd nd nd 18, 19

Sacoila squamulosa US nd nd nd 18, 19

Scaphyglottis modesta PR nd nd nd 11

Schiedella arizonica US nd nd nd 18, 19

Spiranthes amesiana US nd nd nd 11

Spiranthes brevilabris US 4 3, 4 S 6, 11

Spiranthes casei var. novaescotiae CAN nd nd nd 18

Spiranthes casei var. casei US, CAN nd nd nd 18

Spiranthes cernua US, CAN 2, 3, 5 4 nd 1, 2, 6, 14, 19, 24

Spiranthes delitescens US 3 4 F 2

Spiranthes diluvialis US, CAN 6 nd nd 18, 23


65


Spiranthes eatonii US nd nd nd 11

Spiranthes floridana US 4 6 nd 6, 11

Spiranthes infernalis US nd nd nd 11

Spiranthes lacera var. gracilis US, CAN 4, 6 nd nd 14, 24

Spiranthes lacera var. lacera US, CAN nd nd nd 18

Spiranthes longilabris US nd 3, 4 s 11, 22

Spiranthes lucida US, CAN nd nd S, s, o 11

Spiranthes magnicamporum US, CAN 2, 3, 5 4 S, s, o 1, 11, 14, 24

Spiranthes ochroleuca US, CAN nd nd S, s, o 11

Spiranthes odorata US 1, 3, 4 4 S, o 1, 6, 11, 24

Spiranthes ovalis var. erostellata US, CAN 3 4 nd 18, 22, 24

Spiranthes ovalis var. ovalis US nd nd nd 18

Spiranthes parksii US 4, 5 nd F, S 11, 24

Spiranthes praecox US 5 nd nd 11

Spiranthes romanzoffiana US, CAN, SPM 5 nd S, s, o 2, 11, 14

Spiranthes stellata US nd nd nd 19

Spiranthes sylvatica US 5 nd nd 18

Spiranthes torta US, PR, VI nd nd nd 8, 18

Spiranthes tuberosa US 6 5 S, s, o 6, 11, 23

Spiranthes vernalis US 2, 3 4 S, s, o 1, 11, 24

Spiranthes × borealis US, CAN nd nd nd 11

Spiranthes × eamesii US nd nd nd 19

Spiranthes × folsomii US nd nd nd 19

Spiranthes × intermedia US nd nd nd 11

Spiranthes × itchetuckneensis US nd nd nd 11

Spiranthes × meridionalis US nd nd nd 11

Spiranthes × simpsonii US, CAN nd nd nd 11

Stelis perpusilliflora PR nd nd nd 11

Stelis pygmaea PR nd nd nd 11

Stenorrhynchos speciosum PR 4 nd nd 11, 24

Taeniphyllum marianense GU nd nd nd 9, 20

Tetramicra canaliculata PR, VI 2, 3 nd nd 2, 8, 11, 23

Tipularia discolor US 3, 5, 6 nd S, s, o 3, 11, 14, 24

Tolumnia bahamensis US 1, 3 4 S 2, 11, 24

Tolumnia prionochila PR, VI 2 nd nd 8, 11, 23

Tolumnia variegata PR, VI 1, 3 4 nd 8, 11, 24

Trichocentrum carthagenense US nd nd nd 18

Trichocentrum maculatum US nd nd S 11

Trichosalpinx dura PR nd nd nd 11

Triphora amazonica PR nd nd S 11

Triphora craigheadii US nd nd S 11

Triphora gentianoides US 4 nd nd 11, 24

Triphora hassleriana PR nd nd nd 11

Triphora latifolia US nd nd S 11

Triphora rickettii US 6 nd nd 11, 23

Triphora surinamensis PR nd nd nd 11

Triphora trianthophoros var. texensis US nd nd nd 18

Triphora trianthophoros var. trianthophoros US 3 nd nd 18, 24

Tropidia polystachya US, PR 6 nd S 11, 24

Vanilla barbellata US, PR, VI nd nd S 8, 11

Vanilla claviculata PR nd nd nd 11

Vanilla dilloniana PR, US nd nd S 11


66


Vanilla mexicana US, PR, VI nd nd S 8, 11

Vanilla phaeantha US nd nd S 11

Vanilla poitaei PR nd nd nd 11

Wullschlaegelia aphylla PR Nd nd nd 11

Zeuxine fritzii GU Nd nd nd 11, 20

References for Table 2:

1. From, M. 2009. Personal communication with A. Hicks.

2. Hicks, A. 2009. Personal communication.

3. O'Neil, J. 2009. Personal communication with A. Hicks.

4. Sheviak, C. 2009. Personal communication with A. Hicks.

5. Steele, W. 2009. Personal communication with A. Hicks.

6. Stewart, S. 2009. Personal communication.

7. Whitten, M. 2009. Personal communication with A. Hicks.

8. Ackerman, J.D. 1995. An orchid flora of Puerto Rico and the Virgin Islands. New York Botanical Garden Press.

9. Raulerson, L. 2006. Checklist of plants of the Mariana Islands. University of Guam Herbarium Contribution 37: 1-69.

10. Stone, B.C. 1970. Flora of Guam. University of Guam Press.

11. USDA PLANTS Database: http://plants.usda.gov/.

12. Hermann, P. 2009. Personal communication with A. Hicks.

13. Whitlow, C. 2009. Personal communication with A. Hicks.

14. Stoutamire, W. 2009. Personal communication with A. Hicks.

15. Vermont Ladyslipper web page: http://www.vtladyslipper.com/.

16. Fraser's Thimble Farms web page: http://www.thimblefarms.com/.

17. Zielinski, R. 2009. Personal communication with A. Hicks.

18. Brown, P.M. & S.N. Folsom. 2003. The Wild Orchids of North America, north of Mexico. University Press of Florida.

19. Brown, P.M. 2009. Personal checklist of the wild orchids of North America, north of Mexico.

20. Raulerson, L. and A.F. Rinehardt. 1992. Ferns and Orchids of the Mariana Islands. American Printing Corporation.

21. Flora of North America web page: http://www.fna.org/.

22. Zettler, L. 2009. Personal communication with S. Stewart.

23. Meyers, T. 2009. Personal communication with A. Hicks.

24. Richards, M. 2009. Personal communication with A. Hicks.

25. Brown, P.M. 2009. Personal communication with S. Stewart.

http://plants.usda.gov/

http://www.vtladyslipper.com/

http://www.thimblefarms.com/

http://www.fna.org/

Empiricist: SHOULD WE OR SHOULDN'T WE?

67

SHOULD WE OR SHOULDN'T WE? ETHICS AND ORCHIDS

The Slow Empiricist

I've written about how the orchid enthusiast likes to play with creating a new and

better specimen that meets some criteria of esthetics that the orchid world deems desirable. I

have opted for leaving the natives as they are and enjoying them for their unique and

irreplaceable beauty.

Does that stance mean that I am opposed to the idea of reintroducing a native orchid

to a habitat that once contained these plants? Not necessarily. If the plants were lost to the

area because of man's interference such as clearing the land and building on the site, as Disney

World in Florida did, when it obliterated thousands of yellow fringeless orchids to put in a

parking lot, then I feel that it is perfectly acceptable to reintroduce the plants to the area if

possible.

The 'if possible' raises interesting philosophical and ethical questions. How ethical is it

to play with these orchids and take the chance that you can introduce them to a spot if you

fail? I remember a few years ago an attempt was made to plant seedlings of an orchid along a

likely spot in Goethe State Forest in Dunnellon, Florida. Many of the plantlets did not

survive. The plants that were reintroduced in areas where there were other plants did well and

are still surviving after seven years or so. The ones that failed were put in likely habitat but

something was missing and they declined and disappeared. More work was needed to make

such an attempt possible. Like all experiments, how ethical is it to let living things, even

plants, open to the possibility of failure and their ultimate demise?

Then there is the possibility of the reintroduced plants taking over the area. If the

yellow fringeless orchids mentioned above were put back in an area adjacent to the Disney lot

that had other native species in it and the reintroduced plants really took and flourished like

they had before the bulldozers obliterated them would they annihilate the other natives in the

area? Would this be acceptable if the annihilated species weren't orchids?

Although it might seem farfetched, suppose the reintroduced orchids could hybridize

with other native orchids growing nearby and produce stunning new crosses or even a new

species. It would be exciting and probably very valuable in the long run and I'm not sure of

the consequences such an occurrence would make.

Empiricist: SHOULD WE OR SHOULDN'T WE?

68

I have never been unhappy about introduced species such as the Epipactis helleborine,

the broad leaved helleborine, coming into the United States and colonizing areas because it

doesn't drown out the competition. Or at least so far it hasn't. Another orchid from overseas

is Zeuxine, the lawn orchid, that has populated Florida from north to south and isn't invasive.

It isn't like the Brazilian pepper that has overgrown south Florida pushing out desirable

species in its relentless search for new places to grow. Does this mean we could take a native

of say Western Asia and introduce it successfully here in the U.S. and feel justified if it takes

off like the Brazilian pepper?

Being able to introduce native species into the landscape does raise some concerns for

fragile habitats if it becomes common practice for orchids to be propagated by anyone,

especially if they proliferate wherever they are introduced. Are amateur gardeners well

enough schooled to be able to go to a local nursery and buy orchids that have been proven to

survive in an area and put them willy-nilly wherever they want?

I am especially wary of those individuals who profess to want to improve a species

and start that process of refining the native orchid to produce a 'superior' plant. But, of

course, I don't care for all the hybridizing that goes on amongst the orchid hobbyists. I know

this is highly unlikely in my time but who really knows

what man's meddling might produce?

So should we or shouldn't we? I say let's proceed

with caution and keep in mind all the possibilities of

our actions. We could be creating a monster that will

comeback to haunt us or we could be creating a blessing

in repopulating an area with species that should never

have been obliterated in the first place.

Your Slow Empiricist

Spiranthes brevilabris, short-lipped ladies‖-tresses

Goethe State Forest, Levy Co., Florida

P.M. Brown


69


From: 2009. American Journal of Botany 96: 2022-30.

Genetic diversity of the endangered and narrow endemic Piperia yadonii

(Orchidaceae) assessed with ISSR polymorphisms

Sheeja George, Jyotsna Sharma, and Vern L. Yadon

Abstract: Highly endangered plants that are also narrow endemics are generally found

to be genetically depauperate and thus are exceedingly

susceptible to ecological and

anthropological threats that can

lead to their extinction. Piperia yadonii is restricted to

a

single California county within a biodiversity hotspot. We used

nine primers to

generate intersimple sequence repeat (ISSR)

data to assess its genetic diversity and

structure. Within each

population, 99% of the loci were polymorphic, expected

heterozygosity

was low, and a majority of the loci were shared with few other

populations. Forty percent of the total variation could be attributed

to population

differentiation while the rest (60%) resides within

populations, and the genetic

distances between populations were

independent of the corresponding geographical

distances. High

divergence among populations is likely due to fragmentation

and

limited gene flow. Each population contains several private

loci, and ideally, each

should be protected to preserve the

overall diversity of the species. Because P. yadonii

currently

retains a modest amount of genetic variation among individuals

within

populations, preserving and expanding the habitat at

each site to allow natural

expansion of populations would be

additional strategies for its conservation before

populations

become too small to persist naturally.

From: 2009. Annals of Botany 104(3): 543-56

Terrestrial orchid conservation in the age of extinction

Nigel D. Swarts and Kingsley W. Dixon

Background: Conservation through reserves alone is now considered unlikely to

achieve protection of plant species necessary to mitigate direct losses of habitat and the

pervasive impact of global climate change. Assisted translocation/migration represents

new challenges in the face of climate change; species, particularly orchids, will need

artificial assistance to migrate from hostile environments, across ecological barriers

(alienated lands such as farmlands and built infrastructure) to new climatically buffered

sites. The technology and science to underpin assisted migration concepts are in their

infancy for plants in general, and orchids, with their high degree of rarity, represent a

particularly challenging group for which these principles need to be developed. It is

likely that orchids, more than any other plant family, will be in the front-line of

species to suffer large-scale extinction events as a result of climate change.


70

Scope: The South West Australian Floristic Region (SWAFR) is the only global

biodiversity hotspot in Australia and represents an ideal test-bed for development of

orchid conservation principles. Orchids comprise 6 % of all threatened vascular plants

in the SWAFR, with 76 out of the 407 species known for the region having a high

level of conservation risk. The situation in the SWAFR is a portent of the global crisis

in terrestrial orchid conservation, and it is a region where innovative conservation

solutions will be required if the impending wave of extinction is to be averted. Major

threatening processes are varied, and include land clearance, salinity, burning, weed

encroachment, disease and pests. This is compounded by highly specialized pollinators

(locally endemic native invertebrates) and, in the most threatened groups such as

hammer orchids (Drakaea) and spider orchids (Caladenia), high levels of mycorrhizal

specialization. Management and development of effective conservation strategies for

SWAFR orchids require a wide range of integrated scientific approaches to mitigate

impacts that directly influence ecological traits critical for survival.

Conclusions: In response to threats to orchid species, integrated conservation

approaches have been adopted (including ex situ and translocation principles) in the

SWAFR with the result that a significant, multidisciplinary approach is under

development to facilitate conservation of some of the most threatened taxa and build

expertise to carry out assisted migration to new sites. Here the past two decades of

orchid conservation research in the SWAFR and the role of research-based approaches

for managing effective orchid conservation in a global biodiversity hotspot are

reviewed.

From: 2009. Botanica Helvetica 119: 69-76.

Mathematical inference of the underground clonal growth of Epipactis helleborine

(L.) Crantz (Orchidaceae, Neottieae)

Anna Jakubska-Busse, Malgorzata Dudkiewicz, Pawel Jankowsi, and Radoslaw Sikora

Abstract: Epipactis helleborine (L.) Crantz (Orchidaceae, Neottieae) can spread by

sexual or vegetative propagation. The choice of strategy likely depends on the

environmental conditions. The rhizome is the organ of vegetative reproduction; hence,

it is crucial to understand its development. Unfortunately, it is hardly possible to

investigate rhizome morphology directly, since E. helleborine is a protected species in

most European countries. The goal of our investigation was to infer the growth

patterns of underground parts of an orchid population from long-term annual

observations of its aboveground shoots. We implemented the Minimum Spanning Tree

method to determine a likely set of underground connections between shoots and to

simulate the annual growth of new rhizomes. Furthermore, we modeled the spatial

distribution of shoots with a density kernel estimator to compare the density gradients

with the direction of growth of the rhizomes. Observed shoot numbers fluctuated

between 72 and 183 from year to year. Our results suggest that (1) vegetative

reproduction prevails in the studied population, (2) the population consists of about a

dozen clones with a diameter of up to 6 m, (3) rhizomes produce up to five new shoots

at one branch end per year, (4) rhizomes develop in the direction of decreasing


71

population density, and (5) nodes of rhizomes may produce new offshoots after up to

7 years of dormancy.

From: 2008. Floriculture, Ornamental and Plant Biotechnology 5: 375-91.

Techniques and applications of in vitro orchid seed germination

Philip J. Kauth, Daniela Dutra, Timothy R. Johnson, Scott L. Stewart, Michael E.

Kane, and Wagner Vendrame

Abstract: In nature orchid seeds germinate only following infection by mycorrhizal

fungi that provide the developing embryo with water, carbohydrates, minerals, and

vitamins. Orchid seeds were first germinated at the base of wild-collected potted

orchids, but germination was unreliable and seedling mortality rates were high. In

vitro germination techniques, which were developed in the early 1900s, have resulted

in more reliable germination and propagation of many orchid taxa. The earliest in

vitro orchid seed germination techniques utilized mycorrhizal fungi found in nature to

simulate germination and seedling development. In 1922 Lewis Knudson germinated

orchid seeds in vitro by sowing seeds on sterile nutrient medium amended with

sucrose. This technique is known as asymbiotic seed germination since no fungal

mycobiont is used to promote germination. For both symbiotic and asymbiotic orchid

seed germination, many conditions must be address such as photoperiod, temperature,

and mineral nutrition. In the case of symbiotic germination, another important factor

is fungal compatibility. In recent years, the limitations that seed dormancy poses to the

germination of orchid seed have also been examined. In this chapter techniques and

applications of asymbiotic and symbiotic seed germination will be discussed in relation

to photoperiod, temperature, nutrition, seed dormancy, and fungal mycobionts.

72

BOOK REVIEWS

ASYMBIOTIC TECHNIQUE OF ORCHID SEED GERMINATION

SECOND REVISED EDITION

Aaron J. Hicks, Chapter 7 by Scott Stewart, PhD.

The Orchid Seedbank Project.

2009. Raven Roost Books. 185 pp.

ISBN 0-9673049-3-8 $54.00

http://www.ravenroostbooks.com/?page=shop/flypage&

product_id=1464&keyword=Hicks&searchby=author&o

ffset=0&fs=1&CLSN_513=1259942277513a4de039774047

3a50e1

Literature on orchid seed germination is abundant

but scattered throughout scientific publications not readily

available to the general public. Several books have been

published that describe basic instructions for orchid seed

sowing including Orchids from Seed by P.A. Thompson, Home Orchid Growing by R.T.

Northern, and Growing Orchids from Seed by P. Seaton and M. Ramsay. In writing Asymbiotic

Technique of Orchid Seed Germination, Mr. Hicks hoped to provide a manual that will “clarify

the entire flasking process for all readers.” There is no doubt that Mr. Hicks wanted to create

a comprehensive guide involving as much information on orchid seed germination as possible.

In doing so, I believe the book actually provided too much unnecessary information that

could potentially leave some confused and overwhelmed.

Upon receiving the book, I was impressed with its quality. The black and white and

glossy cover caught my eye immediately, and the scanning electron microscopy image of the

orchid seeds was an excellent choice. The paper used in the book‖s production was good

quality. I was also impressed with the figures and diagrams throughout the book. I was

especially pleased that many figures were in color, which enhanced the overall quality of the

book. Mr. Hicks also did a good job at providing a list of companies that sell tissue culture

equipment and supplies.

I was pleased with many aspects of the book. The section in Chapter 3 on pollination

was well written and useful, and the figures were an excellent addition. Likewise the sections

on harvesting and storing orchid seed were welcomed because many hobbyists have questions

concerning these topics. Mr. Hicks also does a thorough job describing aseptic sowing areas,

and includes information for constructing a glove box. In Chapter 4 the techniques for sowing

orchid seed were described quite thoroughly. I was pleased that Mr. Hicks included

descriptions of using both loose, mature seed and green capsules. Mr. Hicks gives the readers

several options to surface sterilize seed, and provides two excellent figures that helped to

clarify the text. Appendix I, which was the most useful section of the book, outlined the

actual flasking technique that was easy-to-follow.

http://www.ravenroostbooks.com/?page=shop/flypage&product_id=1464&keyword=Hicks&searchby=author&offset=0&fs=1&CLSN_513=1259942277513a4de0397740473a50e1




73

Dr. Stewart‖s contribution on symbiotic germination was a welcomed edition since

this technique is popular for conservation purposes. In addition, no other book to my

knowledge provides steps outlining symbiotic germination. Dr. Stewart provided thorough

background and history on the orchid seed/fungal relationship. The glossary at the end of the

chapter is a necessary addition for those not familiar with the scientific terms. Although the

highly scientific writing may be daunting for the hobbyist, this chapter would be a necessary

edition to any orchid seed scientist‖s library.

I found that much of the information presented was unnecessary. Related information,

such as that regarding the green capsule technique, was scattered throughout the book making

it difficult to read at times. Chapter 6 on advanced techniques could be absorbed into other

chapters. Mr. Hicks devotes several pages in Chapter 6 to media modifications, which I

believe could be included in the media section in Chapter 3. Having an entire chapter solely

devoted to media would have been a welcomed revision. Also, the section in Chapter 6 on

osmotic strength and phenolics could have been edited out. I also did not find the section in

Chapter 6 on new directions in seed disinfection useful since Mr. Hicks already described

several techniques in Chapter 4. That being said, I also felt that there were too many

techniques discussed for disinfecting seeds, many of which would not be used. I was very

puzzled why Mr. Hicks even mentioned using chlorine gas as a sterilization technique

considering this is incredibly dangerous. Presenting two or three useful techniques would

make the text less confusing. I also would have recommended combining Chapters 1 and 2

and increasing the information on the unique biology and anatomy of orchid seeds.

Overall I thought the book was a good attempt at providing a comprehensive guide to

orchid seed germination. However, the book could have been simpler with less information.

Mr. Hicks even states in Appendix I that much of the information is of little or no practical

value for the beginner. I agree with his statement. I would recommend the book to hobbyists

and growers who want a book with enormous amounts of information on the subject.

Philip Kauth, Ph.D.

Plant Restoration, Conservation, and Propagation Biotechnology Program, Environmental Horticulture

Department, University of Florida , PO Box 110675, Gainesville, FL 32611, USA. [email protected]


74

MICROPROPAGATION OF ORCHIDS, VOLUMES 1 & 2 (2ND

EDITION)

Joseph Arditti.

2008. Wiley-Blackwell Publishing, Hoboken. Hardcover.

Vol. I (pages 1–756), Vol. II (pages 757–1523). $475

http://www.wiley.com/WileyCDA/WileyTitle/productCd-

1405160888.html

More than 30 years ago, the first volume of Joseph

Arditti, PhD‖s, Orchid Biology, Reviews and Perspectives series

was published. In the initial volume, an appendix was added

that served as a manual to growing orchids via clonal

propagation and tissue culture — a relatively new and

exciting branch of orchid biology at that time. Largely

because of the appendix, demand for the first volume of Orchid Biology continued, even after

going out of print in 1990. Consequently, Arditti and fellow colleague Robert Ernst, PhD,

embarked on a mission to expand the appendix and publish it as a separate book,

Micropropagation of Orchids. Published in 1993, the initial volume of Micropropagation of

Orchids combined the Orchid Biology appendix with more recent procedures and methods

leading up to 1990. What resulted was a massive (682 page), informative book that also

remained in demand after its printing. Although Arditti officially retired in 2001, he was

compelled to write a second edition of Micropropagation of Orchids, largely because of a wealth

of new publications on the subject that surfaced between 1990 and 2000. Hence, the 2nd

edition of Micropropagation of Orchids was subsequently born.

As predicted, the 2nd edition is even more massive than the first, encompassing two

volumes. Both are housed by a handsome, durable sleeve that measures 10½ inches (26 cm) in

height, 3½ inches (9 cm) in width and 8 inches (20 cm) in depth.

Volume I encompasses the set‖s first three chapters. In typical fashion, Chapter 1 is a

historical account of orchid micropropagation, complete with informative text, black-and-

white portraits of noteworthy specialists and other select images. The chapter concludes with

some predictions and potential breakthroughs that lie ahead. Those who have read the first

Micropropagation of Orchids will find that Chapters 2 and 4 have been revised and rewritten.

In Chapter 2 (General Outline of Techniques and Procedures), new information was added

that should be useful to many readers, ranging from the educated beginner to the seasoned

specialist. Many of the chapter‖s 74 pages contain various tables and figures that nicely

supplement the text. Among the topics addressed include media components, plant growth

regulators, pH, stock solutions, media preparation, sterile technique and culture conditions

(e.g., illumination), among others. Given that the author was unaware of published

information on the techniques aimed at how to deal with/handle internal culture

contamination, the second edition of Micropropagation of Orchids, like the first, does not

address the issue. However, Chapter 2 does provide a section on anticontaminants

(antibiotics) and their use. The remaining 600-plus pages of Volume I are dedicated to Chapter

http://www.wiley.com/WileyCDA/WileyTitle/productCd-1405160888.html

http://www.wiley.com/WileyCDA/WileyTitle/productCd-1405160888.html

75

3, Methods for Specific Genera. In this chapter, 59 genera of terrestrial and epiphytic orchids

alike are addressed, beginning with Acampe and ending with Lycaste. Well-known genera such

as Cattleya, Dendrobium and Encyclia are discussed along with the mixed company of lesser-

known examples, as well as hybrid-derived genera. For each, tables outlining various media

are provided. Unfortunately but understandably, the font size of each table is extremely small

(about like reading the label of a pinned insect specimen in an entomology collection). Those

who have bifocals will be well-served.

In Volume II, Chapter 3 continues to march through an additional 52 genera,

beginning with Malaxis and ending with Zygopetalum. Taken together, a total of 111 genera

are outlined by this huge chapter spanning both volumes. In Chapter 4, the author provides a

brief summary of some broad generalizations about orchid tissue culture that many readers

should appreciate, given the often unpredictable nature of the practice, and the suffocating

wealth of new information. Volume II nears completion with a succulent references section

that consumes 76 pages. A few of the newer references are accompanied by links to the

Internet.

The book concludes with a series of eight useful appendices: 1) General Information

on Supplies, Equipment, Terms, and Reagents; 2) Sources of Supplies and Equipment; 3) Sites

of Interest on the World Wide Web; 4) Light; 5) Formulary; 6) various units and other values

used in micropropagation; 7) Additional Information; and 8) Plant Preservative Mixture. A

glossary and index aptly follow.

Aside from being an excellent resource like its predecessor, I especially liked the

various historical insets that were strategically dispersed throughout Chapter 3 (e.g., Origin of

the term protocorm, page 254; use of vanilla by Aztec Emperor Montezuma, page 1291). This

approach was both informative and captivating, and served as a reminder of why we go to

such extremes in our quest to propagate these plants.

In closing, Micropropagation of Orchids, 2nd edition, does not cater to those interested

in related fields of study (e.g., bioengineering, cytogenetics, molecular biology, seed

germination), as the author had intended. Instead, the book lives up to its billing as a useful

resource strictly dedicated to micropropagation. As Arditti states in the book‖s preface, new

methods will undoubtedly surface in the coming years, but this edition of Micropropagation of

Orchids will be his last treatment. Secure a copy for your personal library before it is out of

print once and for all.

Lawrence W. Zettler, Ph.D.

Illinois College, Jacksonville, Illinois. [email protected]

This review previously appeared in ORCHIDS magazine from the American Orchid Society and is reprinted by

permission.


76

77

FROM THE SWAMPS OF SOUTH FLORIDA TO THE WILDS OF

NORTHERN ALASKA….

TO WINDSWEPT NEWFOUNDLAND AND THE BIG BEND OF

WEST TEXAS

WILD ORCHIDS…. from the University Press of Florida

by Paul Martin Brown & Stan Folsom

Ordering information from University Press of Florida www.upf.com or 1-800-226-3822 or for signed and inscribed copies from the authors at [email protected]

http://www.upf.com/


78

Native orchids are increasingly threatened by pressure from population growth and development but,

nonetheless, still present a welcome surprise to observant hikers in every state and province. Compiled and illustrated by long-time orchid specialist Paul Martin Brown, this pocket guide to the woodland and bog rein orchids forms part of a series that will cover all the wild orchids of the continental United States and Canada. Brown provides a description, general distributional information, time of flowering, and habitat requirements for each species as well as a complete list of hybrids and the many different growth

and color forms that can make identifying orchids so challenging.

Lady's-slippers in Your Pocket A Guide to the Native Lady's-slipper Orchids, Cypripedium, of the United States and Canada 34 color photos, 2008 $9.95, 1-58729-655-1, 978-1-58729-655-0

Ladies'-tresses in Your Pocket A Guide to the Native Ladies'-tresses Orchids, Spiranthes, of the United States and Canada 30 color photos, 2008 $9.95, 1-58729-656-X, 978-1-58729-656-7

Grass-pinks and Companion Orchids in Your Pocket A Guide to the Native Calopogon, Bletia, Arethusa, Pogonia, Cleistes, Eulophia, Pteroglossaspis, and Gymnadeniopsis

Species of the Continental United States and Canada 43 color photos, 3 drawings, 2008 $9.95, 1-58729-700-0, 978-1-58729-700-7

Twayblades and Adder’s-mouth Orchids in Your Pocket A Guide to the Native Liparis, Listera, and Malaxis Species of the Continental United States and Canada 49 color photos, 3 drawings, 2008 $9.95, 1-58729-702-7, 978-1-58729-702-1

Fringed Orchids in Your Pocket A Guide to Native Platanthera Species of the Continental United States and Canada 47 color photos, 3 drawings, 2009 $9.95, 1-58729-812-0, 978-1-58729-812-7

Woodland and Bog Rein Orchids in Your Pocket A Guide to Native Platanthera Species of the Continental United States and Canada 50 color photos, 3 drawings, spring 2010 $9.95, 1-58729-8627, 978-1-58729-862-2

Each laminated guide is 16 3/4 x 16 7/8 inches and folds to 4 1/8 x 9 inches.

OR FROM THE AUTHORS AT [email protected]


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GGeenneerraa OOrrcchhiiddaacceeaarruumm

The six volumes of Genera Orchidacearum will provide a complete, robust classification of the orchids,

descriptions of individual species, and cultivational information. The

series, superbly illustrated with color photographs and line drawings

of all the genera, will be an indispensable reference tool for scientists

and for orchid breeders, collectors and enthusiasts. Order from

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Epidendroideae (Part 2)

Alec M. Pridgeon, Phillip Cribb, Mark W. Chase, Finn N.

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The first reference work of the world's orchid genera that reelects

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Alec M. Pridgeon, Phillip Cribb, Mark W. Chase, Finn N. Rasmussen

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This fourth volume in the series treats the first 210 genera of the largest

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Volume 3:

Orchidoideae (Part 2), Vanilloideae

Alec M. Pridgeon, Phillip J. Cribb, Mark W. Chase, Finn N.

Rasmussen

$195.00 Hardback, Jul 2003

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This third volume describes the remaining 105 genera of subfamily

Orchidoideae and all 15 genera of subfamily Vanilloideae.


Volume 2

Orchidoideae (Part 1)

Alec M. Pridgeon, Phillip J. Cribb, Mark W. Chase, Finn N. Rasmussen

$300.00 Hardback, Mar 2001

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Volume 1:

General Introduction, Apostasioideae, Cypripedioideae

Alec M. Pridgeon, Phillip J. Cribb, Mark W. Chase, Finn Rasmussen

$250.00 Hardback, Nov 1999

ISBN13: 9780198505136 ISBN10: 0198505132

Genera Orchidacearum, Volume 6

in preparation - due 2010




http://www.nhbs.com/genera_orchidacearum_volume_6_tefno_161410.html

http://www.nhbs.com/info/book-status-explanations.html

Documents

March 2010 North American Native Orchid Journal