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Physiological and Anatomical Basis for Differences in Growth Performance during In Vitro and Ex Vitro Culture of Sea Oat...

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Permanent Link: http://ufdc.ufl.edu/UFE0012804/00001

Material Information

Title: Physiological and Anatomical Basis for Differences in Growth Performance during In Vitro and Ex Vitro Culture of Sea Oats (Uniola paniculata L.) Genotypes
Physical Description: Mixed Material
Copyright Date: 2008

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Holding Location: University of Florida
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Permanent Link: http://ufdc.ufl.edu/UFE0012804/00001

Material Information

Title: Physiological and Anatomical Basis for Differences in Growth Performance during In Vitro and Ex Vitro Culture of Sea Oats (Uniola paniculata L.) Genotypes
Physical Description: Mixed Material
Copyright Date: 2008

Record Information

Source Institution: University of Florida
Holding Location: University of Florida
Rights Management: All rights reserved by the source institution and holding location.
System ID: UFE0012804:00001

Full Text












PHYSIOLOGICAL AND ANATOMICAL BASIS FOR DIFFERENCES IN GROWTH
PERFORMANCE DURING IN VITRO AND EX VITRO CULTURE OF SEA OATS
(Uniolapaniculata L.) GENOTYPES












By

CARMEN VALERO ARACAMA


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


2005

































Copyright 2005

by

Carmen Valero Aracama



































This dissertation is dedicated to my parents, for their unconditional love and
encouragement.















ACKNOWLEDGMENTS

I would like to express my gratitude to my major professors, Dr. Michael E. Kane

and Dr. Sandra B. Wilson, for giving me the necessary guidance, support and

encouragement to reach my goals and for their caring attitude during the course of my

program. They both were great mentors and made genuine efforts towards my

professional success. I would like to thank my committee members, Dr. Joseph C. V. Vu

for all his advice and useful comments during the enzymatic studies, Dr. Charles L. Guy

and Dr. Rebecca L. Darnell for their guidance, support and for serving in my committee.

I would also like to acknowledge the Florida Sea Grant College Program for funding

support to complete this dissertation research. I am very thankful to the Fulbright

Commission for giving me the great opportunity to continue my studies at the University

of Florida. I would also like to thank the College of Agricultural and Life Sciences for

the Alumni Fellowship funding that supported my studies.

My deepest gratitude goes to Nancy L. Philman, for her technical support and help

in the laboratory, but especially for her kindness and friendship. I would also like to

thank Joan Anderson for all her help and kindness doing the enzymatic analyses, Laurie

K. Mecca for her assistance doing the photosynthesis studies in Fort Pierce, Karen Kelly

for her technical support and guidance doing the electron microscopy studies at the

Electron Microscopy Core Laboratory, Dale W. Haskell for his assistance and friendship,

Carolyn Bartuska for her statistical expertise, and John Sawyer, Bart Schutzman and

Richard Phelan for their assistance with computer issues.









All these years would not have been so joyful without the company of all the

students that worked with me in the lab and that have given me support, assistance, help

and friendship: Christine Emshousen, Pete Sleszynski, Gisele Martins, Scott Stewart, Phil

Kauth, Xiu Li Shen, Katia Cerqueira, Ruth Davis, Allison Debatt, Pat Frey, Kathryn

Villazon and Tammy Ju.

I wish to express my gratefulness to all those friends that have been by my side

giving me encouragement through all the years I have been in Gainesville: Hui Cao,

Francis Kok, Gabriela Luciani, Myrian Rybak, Gisele, Jens and Gabriel Schoene, Camila

and Guto Paula, Penny Nguyen, Veronica Emhart, Belkys Bracho, Carola and Vitor Lira,

Kenichi Shibuya, Chiho and Yoshihiro Baba, Mariana Varese, Marisol Davila, Adriana

Castafieda, Juan Pablo Correales, Milena Palenzuela, Mily and Victor Cabrera,

Ramkrishnan, Raquel Rybak and Jawoo and Sunho Koo. I would also like to thank those

who are far away, Olga Tortosa, Chelo Rico, Inma Rico, Conchi Bautista, Elena

Bekyarova, Carmen Agull6, Yumiko Watanabe, Alex Faustino, Silvina Soto, Flavia

Fukushima, Fabiana Imai, Watcharra Chintakovid, Somrak, Robin Durham, Maria

Delgado, Begofia Ricarte, Merry Saez, Francisco Serna, Alejandro and Fumiko Tanaka,

Celine Verissimo, Yulan Xiao, Lok Yee Hin, and Jose Carlos Monzo.

I would like to express my sincere gratitude to my family all around the world, for

their unconditional support and encouragement through the years. Without the life

vision, love and encouragement of my parents I would not have accomplished so many

things in my life. This has helped me make right decisions during my years as a student.

Also I appreciate the support of my brother and sisters and their families, and my

extended family, which are constantly thinking of me and wishing me the best in my









academic as well as personal life. Also, I want to thank my family in law, who has been

very supportive, caring, loving and encouraging since the day we met.

And, most of all, I would like to thank my husband McNair for his love, support,

and understanding through the years. He has always been by my side through this

journey and he is part of this great achievement. Finally, I would like to thank my

unborn baby boy, who has given me all the strength to finish this dissertation.
















TABLE OF CONTENTS



A C K N O W L E D G M E N T S ......... ...................................................................................... iv

LIST OF TABLES .............................................. .. ........ ........... .. x

LIST OF FIGURES ......... ........................................... ............ xi

A B S T R A C T .......................................... ..................................................x v

CHAPTER

1 L ITER A TU R E R E V IE W .............................................................. .. .....................1

Introduction and R ationale ........................................ ........................................ 1
Literature R review .................................................................................. .. ......6
M icropropagation for H habitat R restoration ........................................ .................6
In Vitro Culture Effects in Plant Anatomy................................. ...... ............ ...7
Influence of Exogenous Sugars ..................................... ............... 10
Photoautotrophic Culture.......................................................... ............... 12
Carbon Status during Acclimatization.................... .... ...................... 13
Photosynthetic Rates during Acclimatization ............................................. 14
Cytokinin Carryover Effects on Plantlet Acclimatization............................. 14
C4 Photosynthesis ............................. ........ ............. ..... .... 15
C A M Photosynthesis ........... ......... ...... ......... ........................... ............... 16
R research O objectives .............. .. ...... ...................... .... ..... .... ...... ......... .....17
Comparative Morphology and Anatomy of In Vitro and Ex Vitro Cultured
Sea Oats Genotypes .................. ...... .......... .... .............. 17
Photosynthetic and Carbohydrate Status of Sea Oats Genotypes during In
Vitro and Ex Vitro Culture Conditions........................................ ...............18
Influence of In Vitro Growth Conditions on In Vitro and Ex Vitro
Photosynthetic Rates of Sea Oats Genotypes ...............................................18

2 COMPARATIVE GROWTH, MORPHOLOGY AND ANATOMY OF IN
VITRO AND EX VITRO CULTURED EASY- AND DIFFICULT-TO-
ACCLIMATIZE SEA OATS (Uniolapaniculata L.) GENOTYPES ........................19

In tro d u ctio n .................................................. ................. ................ 19
M materials and M methods ....................................................................... ..................22
C culture C onditions.......... ..... ......................................................... ... .... ....... 22









Description of Treatm ents ........................................................................23
Effect of Stage II duration on in vitro rooting and ex vitro survival of sea
oats genotypes .................................... ......... ...... .......... ..... 23
Comparative Stage II shoot multiplication and growth of sea oats
genotypes .............................. ... .......... ........ ........ ................ 24
Effect of Stage III duration on in vitro rooting, growth and development
and ex vitro survival of sea oats genotypes .........................................24
Comparative anatomy of sea oats genotypes during Stage II, Stage III
and Stage IV culture ........................................ .......................... 25
Statistical A n aly ses.......... ............................................................ .. .... .. .... .. 2 6
R e su lts .................. ................................................ .. .. ..... ........ ............... 2 7
Effect of Stage II Duration on In Vitro Rooting and Ex Vitro Survival of Sea
O ats G enotypes ............................................. .... .. .............. ... 27
In v itro ro o tin g ....................................................................................... 2 7
E x vitro survival .................................................................. ........... .. 27
Comparative Shoot Multiplication and Growth of Sea Oats Genotypes during
Stage II C culture .................. ....... ... .... ................................... ......... 29
Effect of Stage III Duration on In Vitro Rooting and Growth and Ex Vitro
Survival and Acclimatization of Sea Oats Genotypes ...................................35
In vitro rooting and grow th ........................................ ....... ............... 35
Ex vitro survival and acclim atization................................... ... ..................42
Anatomical and Ultrastructural Comparisons ............................................. 42
Optical light microscopy (OLM)......................................................42
Scanning electron microscopy (SEM) ............. ........................................50
Transmission electron microscopy (TEM) .............................................55
D iscu ssion ............... ........... .......................... ............................59
C o n c lu sio n s........................................................................................................... 6 6
Acknowledgements......................... ...............................67

3 PHOTOSYNTHETIC AND CARBOHYDRATE STATUS OF EASY- AND
DIFFICULT-TO-ACCLIMATIZE SEA OATS (Uniolapaniculata L.)
GENOTYPES DURING IN VITRO CULTURE AND EX VITRO
A C C L IM A T IZ A T IO N .................................................................... .....................68

Intro du action ...................................... ................................................ 6 8
M materials and M methods ....................................................................... ..................7 1
C culture C onditions.......... ..... ......................................................... ... .... ... ... 7 1
Photosynthesis Studies ............................................... ............................. 73
Photosynthesis Enzym atic Studies ............................ ................................... 73
Transmission Electron M icrograph Studies ................................ ............... 76
Carbohydrate Studies ................................. .... ...... .... .......... 77
Experimental Designs and Statistical Analyses ..............................................79
R esu lts.................... ...... ... .. ......... ........... ........................ ................. 80
Photosynthetic and Transpiration Status Ex Vitro ...........................................80
Carbohydrate Status in Vitro and Ex Vitro ............................... ............... .80
Chlorophyll and Soluble Protein Contents ............... .................................87
Photosynthetic Enzyme Status in Vitro and Ex Vitro .......................................87


viii









D isc u ssio n ............................................................................................................. 9 2
C o n c lu sio n s........................................................................................................... 9 7
Acknowledge ents .......... .... ........ .. .. ................. ............ 98

4 INFLUENCE OF IN VITRO GROWTH CONDITIONS ON IN VITRO AND EX
VITRO PHOTOSYNTHETIC RATES OF EASY- AND DIFFICULT-TO-
ACCLIMATIZE SEA OATS (Uniolapaniculata L.) GENOTYPES......................100

Introduction .................................................................................................. 100
M materials and M methods ........................................... ....................................... 102
In V itro Culture Conditions .................. ......... ......... ................. .... ........... 102
In Vitro Growth and Net Photosynthetic Rates............................................. 104
Ex vitro Greenhouse Conditions, Growth, Survival, Photosynthetic Rates and
Transpiration Rates .................... ........ ............ ...... .............. .. 105
Experimental Design and Statistical Analysis.......................................106
R esu lts ................. .................... ...................................................... 10 8
In V itro Survival and G row th....................................... ......................... 108
In V itro N et Photosynthetic R ate ..................... ................... ............... .... 108
Ex Vitro Survival, Transpiration, Photosynthesis and Growth .......................111
D discussion ............................. .. ........................... .............. ............... 116
In Vitro Survival, Growth and Photosynthetic Rates .............. ..................116
Effect of In Vitro CO2 Enrichment........... ................... ................... .............. 117
Ex Vitro Acclimatization................ ......... ................... 118
C o n c lu sio n s......................................................................................................... 12 1
A know ledgem ents ........... .... ........... .. .............. ................. .. 122

5 CONCLUSIONS ......................................... ....................123

APPENDIX

EFFECT OF META-TOPOLIN DURING MULTIPLICATION, ROOTING AND
ACCLIMATIZATION OF EASY- AND DIFFICULT-TO-ACCLIMATIZE SEA
OATS (Uniolapaniculata L.) GENOTYPES ........................................ ............... 128

In tro d u ctio n .................................................. ............................... 12 8
M materials and M methods ........................................... ....................................... 129
R e su lts .............................. ..................................................... 1 3 1
In Vitro Shoot Multiplication ................. ............. ...................131
Carry Over Effect on Vitro Rooting..... ................ ..... ............131
Carry Over Effects on Ex Vitro Acclimatization ...........................................132

LIST OF REFERENCES ......... ...................................... ...... .... ............... 140

B IO G R A PH ICA L SK ETCH ......... ................. ...................................... .....................157















LIST OF TABLES


Tablege

2-1 Comparative shoot number, leaf number and leaf length of EK 11-1 and EK 16-3
sea oats genotypes after 4, 8 and 12 weeks Stage II culture. ................................33

2-2 Comparative shoot number, leaf number and leaf length of EK 11-1 and EK 16-3
sea oats genotypes after 3, 6 and 9 weeks Stage III culture...................................38

2-3 Comparative root number and root length ofEK 11-1 and EK 16-3 sea oats
genotypes after 3, 6 and 9 weeks Stage III culture............... ....... ............... 39

4-1 Environmental conditions during Stage III rooting for EK 11-1 and EK 16-3 sea
oats genotypes. ......................................................................107

4-2 In vitro shoot and root dry weights of EK 11-1 and EK 16-3 sea oats genotypes. In
vitro culture conditions were: PA (photoautotrophic), PM (modified
photomixotrophic) PME (modified photomixotrophic with CO2 enrichment), and
control (conventional photom ixotrophic) ........................................... ..................110

4-3 Net photosynthetic rate per dry weight (Pnw) during in vitro rooting ofEK 11-1 and
EK 16-3 sea oats genotypes cultured under PA (photoautotrophic), PME (modified
photomixotrophic with CO2 enrichment), PM (modified photomixotrophic with
ambient CO2) and control (conventional photomixotrophic) conditions.............112

4-4 Ex vitro shoot and root dry weights and longest leaf length of EK 11-1 and EK 16-
3 sea oats genotypes. In vitro culture conditions included: PA (photoautotrophic),
PM (modified photomixotrophic) PME (modified photomixotrophic with CO2
enrichment), and control (conventional photomixotrophic)................................115















LIST OF FIGURES
Figure page

2-1 Comparative morphological differences in shoot and root development of EK 11-1
and EK 16-3 sea oats genotypes after 4, 8 and 12 weeks Stage II culture followed
by 6 w eeks Stage III culture. ........................................ ......................................28

2-2 Comparative survival percentage after 4 weeks under Stage IV culture of Stage II
unrooted, and Stage III rooted sea oats microcuttings after 4, 8 or 12 weeks Stage II
cu ltu re ......... ...... ............ ...................................... ........................... 30

2-3 Comparative morphological differences in shoot multiplication of EK 11-1 and EK
16-3 sea oats genotypes after 4, 8 and 12 weeks Stage II culture..........................31

2-4 Comparative Stage II shoot dry weights of EK 11-1 and EK 16-3 sea oats
genotypes after 4, 8 and 12 weeks culture. ................................... ............... 32

2-5 Comparative relative leaf length frequency per plantlet of sea oats genotypes after
week 4, week 8, and week 12 Stage II culture. .............................. ......... ...... .34

2-6 Comparative shoot dry weights, and root dry weights of sea oats genotypes after
w eeks 3, 6 and 9 Stage III culture. ........................................ ....................... 36

2-7 Comparative morphological differences in rooting and shoot multiplication of EK
11-1 and EK 16-3 sea oats genotypes after 8 weeks Stage II followed by 3, 6 and 9
w weeks Stage III culture ............................. ...... ........................... .... ............37

2-8 Comparative relative leaf length frequency per plantlet of sea oats genotypes after
week 3, week 6, and week 9 Stage III culture .......................................................40

2-9 Comparative relative root length frequency per plantlet of sea oats genotypes after
week 3, week 6, and week 9 Stage III culture .......................................................41

2-10 Comparative ex vitro shoot dry weights, and root dry weights of sea oats genotypes
after weeks 3, 6 and 9 Stage III culture followed by 4 weeks Stage IV culture.. ....43

2-11 Comparative morphological differences in shoot multiplication of EK 11-1 and EK
16-3 sea oats genotypes previously cultured for 8 weeks in Stage II and 6 weeks in
Stage III, at weeks 0, 3, and 6 Stage IV culture. ............. ........................ .......... 44











2-12 Comparative histological leaf sections of EK 11-1, and EK 16-3 sea oats genotypes
in greenhouse-produced leaves. ........................................ ......................... 47

2-13 Comparative histological leaf sections of EK 11-1 and EK 16-3 genotypes at week
4, week 8 and week 12 Stage II culture......................................... ............... 48

2-14 Comparative histological leaf sections of EK 11-1 and EK 16-3 genotypes at week
3, week 6 and week 9 Stage III culture ........................................ ............... 49

2-15 Comparative SEM of adaxial epidermis of EK 11-1, and EK 16-3 genotypes in
greenhouse-produced leaves.......................................................... ............... 51

2-16 Comparative SEM of stomata on adaxial epidermis of EK 11-1, EK 16-3 genotypes
in greenhouse-produced leaves. ........................................ ......................... 52

2-17 Comparative SEM of stomata on adaxial epidermis of EK 11-1, and EK 16-3
genotypes at week 4, week 8 and week 12.................................... .................53

2-18 Comparative SEM of stomata on adaxial epidermis of EK 11-1, and EK 16-3
genotypes at week 3, week 6, and week 9 Stage III culture..............................54

2-19 Comparative TEM of chloroplasts of EK 11-1, and EK 16-3 genotypes after Stage
IV acclimatization in the greenhouse. Mesophyll and bundle sheath cell
chloroplasts are show n. .......................... ...................... ............ .... ...... ...... 56

2-20 Comparative TEM of chloroplasts of EK 11-1 and EK 16-3 genotypes at week 4,
week 8, and week 12 Stage II culture. Mesophyll and bundle sheath cell
chloroplasts are show n. ........................... ................. ...... ...... ...... 57

2-21 Comparative TEM of chloroplasts of EK 11-1 and EK 16-3 genotypes at week 3,
week 6, and week 9 Stage II culture. Mesophyll and bundle sheath cell chloroplasts
are sh o w n .......................................................................... 5 8

3-1 Effect of in vitro culture conditions on ex vitro survival ex vitro transpiration rate
per leaf area, and ex vitro net photosynthetic rate per leaf area (Pni) of EK 11-1 and
EK 16-3 genotypes during Stage IV culture. ................................... ..................... 81

3-2 Comparative shoot starch content and root starch content of EK 11-1 and EK 16-3
genotypes during in vitro Stage III and after microcuttings were rooted for 6 weeks
Stage III and transferred to Stage IV ...................................... ....... ............... 83









3-3 Comparative TEM of chloroplasts of EK 11-1, and EK 16-3 genotypes after 3
weeks, 6 weeks and 9 weeks Stage III culture conditions in bundle sheath cells....84

3-4 Comparative shoot sucrose content, shoot hexose content, and shoot total soluble
sugar content of EK 11-1 and EK 16-3 genotypes during in vitro Stage III and after
microcuttings were rooted for 6 weeks Stage III and transferred to Stage IV.........85

3-5 Comparative root sucrose content, root hexose content, and root total soluble sugar
content ofEK 11-1 and EK 16-3 sea oats genotypes during in vitro Stage III and
after microcuttings were rooted for 6 weeks Stage III and transferred to Stage IV.86

3-6 Comparative Achlorophyll content per shoot g fresh weight, and TSP content per
shoot g fresh weight ofEK 11-1 and EK 16-3 sea oats genotypes during in vitro
Stage III and after microcuttings were rooted for 6 weeks Stage III and transferred
to Stage IV ..................... ... ......... ..... ....... ..... .. ............... ..........89

3-7 Comparative rubisco activity per shoot g fresh weight, PEPC activity per shoot g
fresh weight, and PEPC/Rubisco ratio ofEK 11-1 and EK 16-3 sea oats genotypes
during in vitro Stage III and after microcuttings were rooted for 6 weeks Stage III
and transferred to Stage IV ........... .................. .......... ............... ............... 90

3-8 Comparative PEPC activity per shoot mg total soluble protein (TSP), rubisco
activity per shoot mg TSP ofEK 11-1 and EK 16-3 sea oats genotypes during in
vitro Stage III and after microcuttings were rooted for 6 weeks Stage III and
transferred to Stage IV ......................... .... .............. ...................... .....91

4-1 Six-week old in vitro cultures of EK 16-3 and EK 11-1 sea oats genotypes under
photoautotrophic [PA], modified photomixotrophic enriched [PME], modified
photomixotrophic [PM] and conventional photomixotrophic [control] culture
con edition s. ....................................................................... 10 9

4-2 Effect of in vitro culture conditions on net photosynthetic rate per plant (Pnp) of EK
11-1 and EK 16-3 genotypes during Stage III culture. ........... ...............113

4-3 Effect of in vitro culture conditions on ex vitro survival, ex vitro transpiration rate
per leaf area, and ex vitro net photosynthetic rate per leaf area (Pni) of EK 11-1 and
EK 16-3 genotypes during Stage IV culture. ........................... ...............114

A-i Effect of cytokinin type and concentration after 4 weeks Stage II culture on shoot
cluster dry weight of EK 11-1 and EK 16-3 sea oats genotypes............................ 133

A-2 Effect of cytokinin type and concentration after 4 weeks Stage II culture on leaf
number ofEK 11-1 and EK 16-3 sea oats genotypes ........................................ 134

A-3 Effect of cytokinin type and concentration after 4 weeks Stage II culture on number
of harvestable shoots (> 30 mm) of EK 11-1 and EK 16-3 sea oats genotypes.....135









A-4 Effect of cytokinin type and concentration after 4 weeks Stage II followed by 6
weeks Stage III culture on rooting ofEK 11-1 and EK 16-3 sea oats genotypes. .136

A-5 Effect of cytokinin type and concentration supplemented in Stage II (4 weeks)
followed by 6 weeks Stage III on ex vitro survival ofEK 11-1, and EK 16-3 sea
oats genotypes. ......................................................................137

A-6 Effect of cytokinin type and concentration after 4 weeks Stage II followed 6 weeks
Stage III and 6 weeks Stage IV on ex vitro leaf length of EK 11-1 and EK 16-3 sea
oats genotypes. ......................................................................138

A-7 Effect of cytokinin type and concentration after 4 weeks Stage II followed 6 weeks
Stage III and 6 weeks Stage IV on ex vitro shoot number of EK 11-1 and EK 16-3
sea oats genotypes ........................................................................... .... 139














Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy

PHYSIOLOGICAL AND ANATOMICAL BASIS FOR DIFFERENCES IN GROWTH
PERFORMANCE DURING IN VITRO AND EX VITRO CULTURE OF SEA OATS
(Uniolapaniculata L.) GENOTYPES

By

Carmen Valero Aracama

December 2005

Chair: Michael E. Kane
Cochair: Sandra B. Wilson
Major Department: Environmental Horticulture

Sea oats (Uniolapaniculata L.), a dune species native to the southeastern U.S., is

commonly used for dune stabilization and restoration in Florida. A micropropagation

protocol was developed for sea oats commercial production. However, significant

variability in shoot production, rooting and ex vitro survival among sea oats genotypes

was observed. Understanding the morphological, anatomical, and physiological basis for

differences among genotypes would allow development of efficient micropropagation

protocols to produce diverse sea oats genotypes for dune stabilization. Growth and

development of two sea oats genotypes with differing acclimatization capacities were

compared at morphological and anatomical levels as a function of the duration of

multiplication and rooting stages using light and electron microscopy. During in vitro

and ex vitro stages, changes in photosynthetic rates were monitored, and carbohydrate

levels and photosynthetic enzymes activities were measured. Additionally, sea oats









photosynthetic capacity in vitro and ex vitro and during acclimatization was evaluated

using in vitro photoautotrophic and photomixotrophic culture conditions. During the

rooting stage, the easy-to-acclimatize genotype (EK 16-3) developed short but numerous

roots and "grass-like" leaves with fully expanded blades. Conversely, the difficult-to-

acclimatize genotype (EK 11-1) developed few long roots and short and thick "lance-

like" leaves without expanded blades. During in vitro development of "grass-like"

leaves, EK 16-3 plantlets exhibited increases in activities of the two photosynthetic

enzymes phosphoenolpyruvate carboxylase (PEPC) and ribulose 1,5-bisphosphate

carboxylase/oxygenase (rubisco), and chlorophyll and total soluble protein content.

These increases were correlated with higher net photosynthetic rates in EK 16-3 than EK

11-1 plantlets after ex vitro transfer. Both genotypes accumulated carbohydrate and

starch reserves during in vitro rooting, which were depleted during the transition from

photomixotrophic to photoautotrophic mode of nutrition. However, due to the lack of

production of photosynthetically competent leaves on EK 11-1 plants coupled with rapid

exhaustion of carbohydrate reserves, acclimatization and survival ex vitro for this

genotype were low. Conversely, rapid production of photosynthetically competent leaves

resulted in 100% survival ex vitro in EK 16-3 plantlets. In conclusion, the major cause

for low acclimatization was the in vitro formation of leaves with abnormal anatomy

correlated with limited photosynthetic capacity.














CHAPTER 1
LITERATURE REVIEW

Introduction and Rationale

Florida's beach and dune systems have significant economic and environmental

value. Seventy-five percent of Florida's population resides in the coastal counties where

beach related tourism alone contributes more than $19 billion to the state's economy

annually. Besides providing a unique wildlife habitat, Florida's coastal sand dunes serve

as a natural defense to mitigate the destructive action associated with tropical storms,

hurricanes and human activity (Crewz, 1987; Dahl and Woodard, 1977; Woodhouse,

1982). During the 2004 hurricane season, nearly all of the state's sandy beach shoreline

was adversely affected. According to the Florida Department of Environmental

Protection, it was estimated in 2005 that about 365 of Florida's 825 miles of sandy

beaches were in a critical state of erosion. Destabilization and erosion of coastal beach

and dune vegetation systems by natural forces or man-made activities have increased the

risk of catastrophic ecological as well as economic damage following storm events.

Continued protection and restoration of dune areas and high-energy beaches are

necessary to prevent additional losses.

Control of coastal erosion is accomplished in Florida by installing mechanical

structures such as fences, or by dredging sand from the offshore zone for beach re-

nourishment. However, dune stabilization by planting bare areas with native or introduced

dune species is the most cost-effective accepted practice to control erosion. The root and

rhizome system of those species holds particles of sand in place, while the above ground









vegetation retards wind and water driven erosion and promotes sand accretion

(Woodhouse, 1982).

The most effective dune species used for beach and dune stabilization are perennial

grasses, including American beachgrass (Ammophila breviligulata Fern.) and sea oats

(Uniolapaniculata L.). In the southeastern United States, sea oats is the primary native

dune grass used for beach and dune stabilization and restoration because it dominates the

foredune zone in the dune-strand ecosystems of the South and Atlantic and Gulf coasts

(Wagner, 1964; Brown and Smith, 1974). Additionally, sea oats has the ability to rapidly

colonize and establish, and exhibits high tolerance to heat, drought, salt spray and

occasional inundation by salt water (Wagner, 1964; Woodhouse, 1982). Growth of sea

oats is stimulated by sand burial, which occurs frequently on the dune. The linear grass

blades of sea oats help trap sand on the dune by raising the laminar boundary layer of the

wind velocity profile, causing sand deposition. Regrowth occurs even after rapid

deposition of sand up to 1 m thick. Plant reproduction occurs by seed and by rhizome

extension, which allows rapid plant distribution to help stabilize the surface of the dune.

Due to repeated coastal erosion, demand for sea oats planting materials has

significantly increased. Currently, the planting of a one-mile-long twenty-foot-wide

length of eroded beach in Florida requires 22,000 sea oats plants at a total cost of $40,000

(0. Bundy, pers. comm.). An estimated 24-36 million sea oats would be required to

revegetate the entire beach coastline of Florida (Sylvia et al., 2003). Sea oats is

propagated in commercial nurseries from field-collected seeds. However, sea oats is not

regarded as a prolific seed producer (Hester and Mendelssohn, 1987; Bachman and

Whitwell, 1995; Burgess et al., 2002; Burgess et al., 2005). Of the six to eight fertile









florets in each spikelet, most have embryos that have aborted and only a few florets

actually set seeds (Wagner, 1964; Burgess et al., 2002). Consequently, dwindling natural

stands have made it necessary for the State of Florida to impose restrictions on field

harvesting of sea oats plants and seeds. These restrictions significantly limit the natural

sources of sea oats and the ability of Florida's native plant nurseries to meet the demand

for plant materials. Concerns regarding the potential introduction of un-adapted ecotypes

at revegetation sites have also limited the use of seed material obtained from distant

geographic sources. In Florida, this second issue has been addressed by restricting the

collection of plant source materials to within populations local to the targeted planting

site. Therefore, alternative propagation methods are needed for mass production of

diverse sea oats genotypes.

Vegetatively, sea oats is a rhizomatous, herbaceous perennial. Buds at the base of

the vegetative shoot can become tillers (offshoots) or rhizomes (underground stems) that

eventually will develop aboveground shoots (Wagner, 1964). Miller et al. (2003)

developed a system to induce tiller formation from rhizome fragments collected from the

beach after a hurricane takes place. For this system, rhizome fragments needed to be

collected immediately after the storm event, and replanted within 7 days to obtain 45%

transplant success after rhizome reburial. Difficult access to the damaged areas and

short-term viability of transplants were limiting factors to the use of this revegetation

system.

Micropropagation, the rapid in vitro production of plants on a defined sterile

medium in culture vessels, has many advantages over conventional plant propagation

methods in horticulture, agronomy and forestry fields (Debergh and Zimmerman, 1991;









Jeong et al., 1995; Hartmann et al., 2002). The use of this technology for multiple

applications is currently expanding worldwide (Conner and Thomas, 1981).

Micropropagation has also been used for commercial production of plants used for

habitat restoration (Kane et al., 1993; Kane and Philman, 1997; Seliskar and Gallagher,

2000), and therefore, it could be utilized for sea oats propagation. This technology also

provides the opportunity to select and rapidly produce diverse ecotypes with ecologically

valuable characteristics, particularly from localized sites.

In 1997, a project funded by the U.S. Department of Commerce to Dr. Michael

Kane, Environmental Horticulture Department/University of Florida, was initiated to 1)

examine genetic diversity and population structure in four Florida sea oats populations

using genetic markers; 2) use micropropagation technology to clonally multiply

representative genotypes from populations at these four sites; and 3) conduct reciprocal

transplant studies at the sites using micropropagated genotypes to determine effects of

genotype and geographical source on survival and growth following outplanting. As a

consequence of that project, more than 28 sea oats genotypes in culture had been

established (M.E. Kane, unpublished data). These genotypes were specifically collected

from four sites in Florida: two Atlantic (Anastasia State Recreation Area [AN] and

Sebastian Inlet State Recreation Area [SI]), and two Gulf coast populations (Egmont Key

National Wildlife Refuge [EK] and St. George Island State Park [SG]).

The micropropagation shoot culture protocol was initially developed using a single

sea oats genotype (Philman and Kane, 1994). However, significant problems were

encountered when attempting to acclimatize Stage II unrooted microcuttings or Stage III

rooted microcuttings to ex vitro conditions (Stage IV). Leaves produced during Stage II









were not expanded but more cylindrical in cross section. Survival of microcuttings under

ex vitro conditions was very low. Microcuttings that rooted for 28 days in vitro (Stage

III) exhibited only slightly higher survival ex vitro. When transferred to ex vitro

conditions, these rooted plantlets began to exhibit new root and leaf development, but

quickly stopped growing and died.

In subsequent studies, the severity of this problem varied when applied to different

genotypes. In most genotypes, survival of rooted microcuttings ex vitro was significantly

increased when microcuttings were rooted on Stage III medium for at least 6 to 8 weeks.

During this extended culture period, morphological and anatomical changes of sea oats

shoots and roots occurred during in vitro growth and development, and a greater root

mass and more "grass-like" leaves with expanded blades typically developed. Although

the physiological basis for increased survival is unknown, it is conceivable that these

"grass-like" leaves were more photosynthetically competent than the "lance-like" leaves

present after only 3 weeks in Stage III culture. Furthermore, carbohydrate reserves in sea

oats possibly became depleted during Stage IV acclimatization.

For the purpose of maintaining genetic diversity, it is critical to be able to

micropropagate a wide range of sea oats genotypes. However, commercial utilization of

sea oats micropropagation could be limited by the poor survival rates of certain

genotypes during acclimatization. Despite the extraordinary potential of

micropropagation and all of its advantages, many problems still exist (Desjardins et al.,

1995). The stimulation of vegetative growth in vitro, which is desirable in terms of

production efficiency, induces several anatomical, morphological and physiological

changes that seriously affect the performance of plantlets after ex vitro transfer.









Understanding the causes of aberrant morphology and physiology in vitro is necessary to

increase growth and survival of plantlets during ex vitro acclimatization.

Literature Review

Micropropagation for Habitat Restoration

Multiple studies have been published on the feasibility of applying

micropropagation techniques to commercial production of aquatic, dune, and marsh

species for habitat restoration (Straub et al., 1988; Cook et al., 1989; Li and Gallagher,

1996; Kane and Philman, 1997; Rogers et al., 1998; Kane et al., 1999; Seliskar and

Gallagher, 2000). Yet, few studies have been focused on applying micropropagation

techniques for sea oats commercial production, especially production of diverse

genotypes. Hovanesian and Torres (1986) reported indirect shoot regeneration from

callus derived from in vitro germinated sea oats seeds. However, micropropagation

systems based upon adventitious shoot production from callus are inherently more

unreliable, both in terms of genetic stability and shoot regeneration rate, than shoot

multiplication from axillary meristems (Pierik, 1987). Philman and Kane (1994)

developed a micropropagation protocol for sea oats using tillers explants that induced

axillary shoot regeneration for in vitro multiplication. However, significant variability in

growth and development (especially during acclimatization) was observed when applied

to 28 different sea oats genotypes collected from four populations sampled in Florida's

Gulf and Atlantic coasts.

Random amplified polymorphic DNA (RAPD) genetic analyses of adult sea oats

plants and seedlings from those four populations indicated significant genetic variations

between the two coastal communities, with the most variation occurring within the Gulf

coast genotypes (Ranamukhaarachchi et al., 1999). These genetic differences were









attributed to the increased environmental stresses of the Gulf coast. Florida's Gulf coast

experiences a greater frequency of storms and hurricanes than does its Atlantic coast.

Dune overwash and waves have created low-relief foredunes at St. George (SG) and a

severely eroded foredune at Egmont Key (EK). Variations in seed weight, temperature

requirements for seed production and germination, and seedling growth between the Gulf

of Mexico and Atlantic populations further support genetic as well as phenotypic

divergences (Seneca, 1972; Colosi, 1979; Hester and Mendelssohn, 1987).

Preliminary in vitro observations using sea oats genotypes collected from the Gulf

coast indicate that genotypes collected from areas closer to the shoreline exhibit lower

survival rates during ex vitro acclimatization than those collected further away from the

shoreline. Ranamukhaarachchi et al. (1999) were reluctant to extrapolate ecotypic

differences from the genetic analysis due to an absence of reciprocal transplant studies.

Other possible contributions to low survival rates of sea oats plantlets during ex vitro

acclimatization include morphological, anatomical, biochemical and physiological

changes due to the tissue culture environment.

In Vitro Culture Effects in Plant Anatomy

Plants that develop in vitro acclimate to the environment in which they grow.

Starting from the aerial environment of the vessel headspace, this system is characterized

by a saturating atmosphere with water vapor, with very low vapor pressure deficit

(Brainerd and Fuchigami, 1981; Fujiwara and Kozai, 1995), relatively low light

intensities (photosynthetic photon flux, PPF; 12-70 [tmol m-2 s-1), relatively high and

constant temperature (25 + 3 C) (Fujiwara and Kozai, 1995), low CO2 concentration

during photoperiod (90 [tmol mol1) (PospiSilova et al., 1988), high CO2 concentration









during dark period (3000-9000 [tmol mol1) (De Proft et al., 1985), and high C2H4

concentration (1.2 [tmol mol1) (Kozai and Kubota, 2005). These characteristics, except

for the light intensity, are largely due to the low number of air exchanges of the vessels

and the relatively small air volume of the vessel headspace.

High relative humidity and low CO2 concentration in the vessel limit plant gas

exchange (Pospisilova et al., 1992). Leaves produced in vitro have a thin cuticle layer

and abnormal stomatal function stomataa remain open even when subjected to sudden

environmental changes; Sutter et al., 1992), which may exacerbate water stress during

acclimatization to ex vitro conditions, and structural changes that may affect CO2 fixation

mechanisms and light harvesting apparatus (alterations in the chloroplast ultrastructure).

Despite the self-evident importance of the gaseous environment for tissue cultures, it is

an often neglected component during the design of a micropropagation system. This

shortcoming can have unacceptable consequences for culture performance because of the

strong physiological impact of the gases involved, notably 02, CO2 and C2H4, on plant

growth and development (Jackson et al., 1994).

Significant changes in leaf morphology and anatomy, especially in epidermal

characteristics caused by in vitro culture conditions, have been reported (Pospisilova et

al., 1999). Sweetgum (Liquidambar styraciflua L.) plantlets cultured in vitro had a less

developed cuticle, as compared to the well developed cuticle in leaves of transplanted and

field grown plants (Wetzstein and Sommer, 1982). Gilly et al. (1997), in studying the

cuticle formation of ivy (Hedera helix L.) plants in vitro and after transferring to ex vitro

conditions observed a progressive activation of cuticle biosynthesis as the plants adapted

to the ex vitro environmental conditions.









Cuticle and epicuticular wax biosynthesis plays an important role in preventing

water loss when plants are transferred to ex vitro conditions (Sutter and Langhans, 1982).

In vitro light levels and relative humidity appear to be critical during the development of

epicuticular waxes, stomata and epidermal cells (Capellades et al., 1990), although Gilly

et al. (1997) also observed that cuticle formation is not exclusively dependent on stress

conditions, but is also genetically programmed.

As a result of the tissue culture environment, in-vitro produced shoots and plantlets

are also reduced in size compared to greenhouse-produced plants (Donnelly and Vidaver,

1984a). The relatively high concentration of cytokinins during the multiplication stage

tends to inhibit apical dominance (Murashige, 1974). Additionally, in some species,

vascular connections are reduced, thin, and poorly structured (Leshem, 1983).

Oftentimes, the low light environment in vitro produces leaves that resemble shade leaves

or hydrophytic leaves ex vitro (Lee et al., 1988). Also, leaves produced in vitro usually

have low chlorophyll content (Grout and Aston, 1977), low percent dry matter or

hyperhydrated shoots (Ziv, 1990), restricted leaf area expansion (Kozai et al., 1992), and

poorly structured spongy and palisade tissues (Donnelly et al., 1985). These

morphological and anatomical characteristics result in low photosynthetic ability

associated with low activities of the photosynthetic enzyme rubisco (ribulose-1,5-

bisphosphate carboxylase/oxygenase) and increased activity of PEPC

(phosphoenolpyruvate carboxylase) (Hdider, 1994; Hdider and Desjardins, 1994) and

abnormal chlorophyll fluorescence responses (Hdider and Desjardins, 1994).

In sweetgum, highbush blueberry (Vaccinium corymbosum L.) and tobacco

(Nicotiana tabacum L.), stomatal density decreased in newly developed leaves after ex









vitro transplant (Wetzstein and Sommer, 1983; Noe and Bonini, 1996; Ticha et al., 1999).

The main reason for this was the enormous enlargement in leaf area after transfer.

Conversely, tissue cultured plants exhibited greater stomatal densities than acclimated

plants. However, stomatal indices, which depict the ratio of stomata number to total

epidermal cells and stomata per unit area, were not calculated in these studies. Hence,

the influence of leaf expansion on resultant stomata density cannot be evaluated.

In vitro sweetgum leaves contained numerous superficial and circular stomata as

opposed to the ellipsoid, depressed and less numerous stomata observed on acclimated

plants (Wardle et al., 1983). The general consensus has been that stomata developed in

vitro are poorly functional with an inability to close in conditions otherwise leading to

closing (Brainerd and Fuchigami, 1981; Wetzstein and Sommer, 1983; Conner and

Conner, 1984; Ziv et al., 1987; Marin et al., 1988; Sutter, 1988; Preece and Sutter, 1991;

Majada et al., 2001). Afreen (2005) concluded that the absence or reduction in leaf

epicuticular and cuticular waxes, combined with non-functional stomata under conditions

of low relative humidity and high light intensity, leads to abnormally high transpiration

rates during acclimatization, which decreases plant survival. Additionally, plants may

also guttate copiously, demonstrating their inability to control water loss (Donnelly and

Tisdall, 1993). Apart from reduced or nonexistent stomatal control and poor epicuticular

and cuticular wax formation, poor control of water loss could result from reduced

trichome numbers (Donnelly and Vidaver, 1984a; Sutter, 1985).

Influence of Exogenous Sugars

Sucrose must be included in tissue culture media in order to promote adequate

shoot regeneration, growth and development under low light conditions. The supply of

sucrose in the media is also possibly the factor most significantly affecting net









photosynthetic rate (Pn) of plants in vitro. Studies with micropropagated rose (Rosa

multiflora L. 'Montse') showed that plantlet photosynthesis was influenced by the levels

of sucrose in the culture medium; the lowest levels of sucrose resulted in the highest

plantlet Pn values (Capellades et al., 1991).

Sucrose is the form of sugar that is most often transported within a plant in vivo.

Culture medium and sugar levels decrease during the culture period. Isoforms of the

enzyme invertase naturally occur in cell walls, vacuoles and cytoplasm. Microcuttings

secrete invertase into the culture media in response to wounding (De la Vifia et al., 1999;

Sturm, 1999). This extracellular invertase hydrolyzes sucrose in the media into glucose

and fructose, but the degree of hydrolysis depends on the amount of invertase secreted,

which differs with each plant. Partial hydrolysis also occurs during autoclaving of the

media, when 10-15% of sucrose is converted to glucose and fructose (George, 1993).

Accumulation of starch and soluble sugars following glucose uptake inhibited

photosynthesis in leaves of spinach (Spinacea oleracea L.) (Krapp et al., 1991) and rose

(Capellades et al., 1991). High sugar accumulation in leaves influences photosynthesis

by feedback inhibition of photosynthetic enzymes (Azc6n-Bieto, 1983; Schafer et al.,

1992), and by down-regulation of expression of genes encoding photosynthetic enzymes

(Sheen, 1994; Jones et al., 1996; Jang and Sheen, 1997; Sheen et al., 1999; Smeekens,

2000). The concept of sugars having an integrated role in the adjustment of cellular

activity of the entire plant system provides a new framework for analysis. According to

Koch's feast/famine hypothesis, the accumulation of carbohydrates results in decreased

photosynthesis and a long-term reduction in the expression of photosynthetic genes

(Koch, 1996).









Photoautotrophic Culture

Kozai et al. (1997) showed that most chlorophyllous plantlets/microcuttings in vitro

have the ability to grow photoautotrophically, in a medium without sugars, provided that

the environmental conditions are favorable for photosynthesis. Plantlets grown in

conventional culture vessels are characterized by low net photosynthetic rates caused by

low CO2 concentrations in the vessels during the photoperiod and low light intensities

typical of culture rooms (Heo and Kozai, 1999). Amdncio et al. (1999) found that higher

light intensities increased photosynthetic competence and subsequently improved

survival of grape (Vitis vinifera L.) plantlets during acclimatization. Similar results have

also been observed with elevated CO2 concentration (Seon et al., 2000). Additionally,

several studies report that addition of sucrose in the medium inhibits photosynthesis of in

vitro formed leaves (Serret and Trillas, 2000; Seon et al., 2000; Van Huylenbroeck and

Debergh, 1996; Lees et al., 1991).

Successful photoautotrophic systems have been developed for enhancing

photosynthesis and growth of the plantlets by increasing CO2 concentration and light

intensity in the culture vessel (Kozai, 1988). Furthermore, to enhance acclimatization

potential, Nguyen et al. (1999a) recommend the use of porous supporting material in

addition to liquid medium instead of conventionally-used gelling agents to allow the

formation of roots in vitro with higher vascular system development. Plantlets grown in

these systems have been well characterized for many growth and developmental

parameters including fresh and dry weight biomass accumulation (Cristea et al., 1999),

net photosynthetic rate (Heo et al., 2001; Valero-Aracama et al., 2001), cuticular

development and stomatal functioning (Zobayed et al., 1999), and improved carbohydrate









status (Wilson et al., 2001). A similar system could be used to enhance sea oats growth

during in vitro culture and ex vitro acclimatization.

Carbon Status during Acclimatization

As previously mentioned, it is known that higher sugar concentrations in the

medium suppress photosynthesis of in vitro plantlets. In vitro plantlets are mixotrophic

in their mode of nutrition; they alternate between carbohydrates used from the medium

and CO2 fixation. Mixotrophy contributes to the recycling of respiratory and

photosynthetic products and affects photosynthetic carbon metabolism. Kozai (1988)

suggests heterotrophic/photomixotrophic growth influenced by the tissue culture

environment contributes to low survivability of plantlets during acclimatization. Yet,

Wilson et al. (2000) assert that appropriate higher levels of carbohydrates can favor

plantlet survival upon transfer to ex vitro conditions, improve acclimatization and

facilitate physiological adaptations to ex vitro conditions.

After transfer to ex vitro conditions, micropropagated plantlets are very susceptible

to various stresses because their physiological and morphological status may restrict them

from allocating sufficient energy resources (Chaves, 1994). Moreover, plants respond

differently to the removal of exogenous sugars in the culture medium. Some plants have

no difficulty undergoing the transition from a heterotrophic or photomixotrophic to a

photoautotrophic condition. For example, broadleaf arrowhead (Sagittaria latifolia

Willd.) exhibited high survival rates when unrooted microcuttings, multiplied with low

levels ofN6-benzyladenine (BA) in Stage II, were transferred to the greenhouse (Lane,

1999). Peace lily (Spathiphyllumfloribundum Schott. 'Petite') leaves formed in vitro

also exhibited the capacity to photosynthesize and maintain a positive carbon balance ex

vitro (Van Huylenbroeck et al., 1998). Stage II unrooted microcuttings of other species









survive when transferred to the greenhouse, but the leaves formed in vitro die, giving rise

to new, more photosynthetically capable leaves. These leaves demonstrate a

cotyledonary effect, described by Kane (2000) as a "lifeboat" in which old leaves become

the source of carbohydrate reserves to shoot meristems. This phenomenon has been

observed in plants such as strawberry (Fragaria x aananssa Duch. 'Kent'), grape, and

Calathea loiusae Gagnep. 'Maui Queen' (Hdider and Desjardins, 1995; Amancio et al.,

1999; Van Huylenbroeck et al., 1998). Grout and Millam (1985) make a distinction

between these two groups of leaves formed in vitro: those that are photosynthetically

competent, and those that are photosynthetically non-competent.

Photosynthetic Rates during Acclimatization

Net photosynthetic rates of potato (Solanum tuberosum L.) and peace lily plants

decreased during the first week after transplanting to greenhouse conditions (Baroja

Fernandez, 1993; Baroja Fernmandez et al., 1995; Van Hyulenbroeck and Debergh, 1996).

While Calathea leaves formed in vitro were not able to photosynthesize during the first

days after transfer, and peace lily leaves formed in vitro were photosynthetically

competent ex vitro, both plant species exhibited substantial photosynthetic activity after

new leaves were fully developed (Van Huylenbroeck et al., 1998).

Cytokinin Carryover Effects on Plantlet Acclimatization

Plant growth regulators incorporated into shoot multiplication culture media,

especially cytokinins, can have deleterious carryover effects on in vitro and ex vitro

growth and development (PospiSilova et al., 1992; Werbrouck et al., 1995). N6-

benzyladenine (BA) is the most widely used cytokinin for shoot multiplication in tissue

culture (Werbrouck et al., 1996), and this cytokinin is required for sea oats

micropropagation (Philman and Kane, 1994). Experimental evidence indicates that









reduction in rooting and acclimatization, observed in some plants produced on BA-

supplemented medium, may result from production of an inhibitory BA metabolite,

[9G]BA. This metabolite accumulates at the base of plantlets in vitro and remains for

more than 6 weeks (Werbrouck et al., 1995; Werbrouck et al., 1996). Moncalean et al.

(2001) found that kiwi (Acitidinia deliciosa Chev. 'Hayward') explants cultured over 2

days in medium containing BA followed by BA-free medium performed better during

rooting and acclimatization stages than those cultured in BA-containing medium for

longer periods. In vitro culture duration in medium containing BA appears to have an

effect on ex vitro acclimatization. Strnad et al. (1997) reported that meta-topolin, a

naturally occurring BA analog, produces a deleterious metabolite with a shorter half-life

and is less inhibitory than BA. Werbrouck et al. (1996) found that meta-topolin

effectively multiplies peace lily shoots, with better rooting in vitro than equimolar

concentrations of BA. Conceivably, meta-topolin may be an acceptable BA substitute for

sea oats micropropagation, which may enhance rooting and Stage IV survival.

C4 Photosynthesis

Sea oats is a C4 plant and it exhibits Kranz anatomy (Brown and Gracen, 1972;

Brown and Smith, 1974). Kranz anatomy is considered an evolutionary advancement

because it facilitates two mechanisms of CO2 fixation (Keeley, 1998) and it allows

greater water-, carbon-, and nitrogen-use efficiencies (Zelitch, 1982; Robichaux and

Pearcy, 1984). In vitro culture conditions of low irradiance, high humidity, low CO2

concentration, and high exogenous sugar levels in the medium are an atypical

environment for a C4 plant. These conditions could lead to limitations in growth and

photosynthetic rates of plantlets in vitro. Changes in the activity of PEPC have been

observed under these conditions. Furbank et al. (1997) observed a substantial increase in









PEPC activity in smelter's bush (Flaveria bidentis [L.] Kuntze) seedlings, a C4 plant,

when grown in the presence of exogenous sucrose. In the same treatment, rubisco

activity was markedly decreased. Phosphoenolpyruvate carboxylase has a higher affinity

for C02, but requires twice the energy of ATP (Salisbury and Ross, 1992). Additionally,

C4 plants have adapted to high heat and light intensity, and a large investment in ATP

may render them disadvantaged in the tissue culture environment. Sugarcane

(Saccharum officinarum L.), also a C4 species, can be micropropagated in vitro through

various culture techniques (Sauvaire and Galzy, 1978; Barba et al., 1978; Ho and Vasil,

1983; Chengalrayan and Gallo-Meagher, 2001). Enhancement of growth and

photosynthetic rates of sugarcane in vitro can be achieved by manipulating the

environmental culture conditions (Erturk and Walker, 2000; Xiao et al., 2003). Rubisco

and PEPC activities are possibly also affected by the changes of the environmental

conditions.

CAM Photosynthesis

Abnormal physiological stomatal responses in plantlets may result from the tissue

culture environment or changes in the Kranz anatomy of sea oats in vitro. Malda et al.

(1999) observed similar adaptations in nellie cory cactus (Coryphantha minima Baird).

Under high humidity in vitro, this cactus performed crassulacean acid metabolism (CAM)

photosynthesis in the light. During acclimatization, carbon fixation via the CAM

pathway reverted back to operating only in the dark. In sea oats, high humidity

conditions could prevent stomatal closure throughout the day and night periods. This

would result in increased transpiration rates after ex vitro transfer and subsequent water

stress.









Research Objectives

It is likely that the in vitro environment is influencing the anatomy, morphology

and physiology of sea oats resulting in the low survival observed in some sea oats

genotypes during acclimatization. Modifications of the culture environment and

comparison between genotypes with differing responses during ex vitro acclimatization

provide the opportunity to better understand sea oats physiology in vitro. Consequently,

in the present study, anatomical and physiological basis for poor acclimatization of

selected sea oats genotypes were examined.

To examine the physiological and anatomical basis for sea oats genotypic

differences in acclimatization capacity, genotypes were selected from plants established

in vitro from Egmont Key, in the Gulf Coast: EK 16-3 and EK 11-1, an easy- and

difficult-to-acclimatize. The main objective of the present research was to correlate

morphological, anatomical and physiological characteristics between the two genotypes

with survival rates ex vitro.

This dissertation research has been organized in three chapters:

Comparative Morphology and Anatomy of In Vitro and Ex Vitro Cultured Sea Oats
Genotypes

Changes in the in vitro culture environment, including vessel headspace gas

composition, culture medium mineral salt composition, plant growth regulators (PGR)

levels and sugars, occur with time. Conceivably, these changes could influence growth

and development of in vitro cultures and affect the acclimatization capacity of shoots. In

vitro time course experiments were designed to examine this further. The morphological

and anatomical characterizations of both sea oats genotypes cultured in vitro and ex vitro

were characterized by evaluating in vitro multiplication, rooting and survival ex vitro as a









function of Stage II and Stage III culture duration. Furthermore, leaf samples were

collected at different periods during multiplication, rooting, and after acclimatization ex

vitro to determine anatomical and morphological changes. To accomplish this, samples

were processed, observed and photographed using optical light, scanning electron and

transmission electron microscopy. These techniques facilitated making anatomical

comparisons at tissue, cellular and organelle levels.

Photosynthetic and Carbohydrate Status of Sea Oats Genotypes during In Vitro and
Ex Vitro Culture Conditions

During the first days of acclimatization, photosynthetic capacity of leaves and

carbohydrate status of plants become critical for survival ex vitro. Therefore, the effects

of in vitro culture on photosynthetic capacity of sea oats genotypes were studied by

measuring photosynthetic rates during ex vitro acclimatization. Subsequently, a study

was conducted to determine the carbohydrate status of sea oats genotypes during in vitro

rooting and ex vitro acclimatization. This was correlated with the analyses of the

enzymatic activities of rubisco and PEPC during in vitro multiplication and rooting, ex

vitro acclimatization, and after establishment to greenhouse conditions.

Influence of In Vitro Growth Conditions on In Vitro and Ex Vitro Photosynthetic
Rates of Sea Oats Genotypes

In this section, the effects of environmental conditions during micropropagation of

sea oats genotypes on survival ex vitro were studied. A system that induced

photoautotrophy in vitro was designed for sea oats genotypes. Furthermore, effects of

CO2 enrichment during in vitro micropropagation on in vitro and ex vitro growth of

plantlets were evaluated. In this experiment the photosynthetic capacity of plantlets

during in vitro and ex vitro culture under different environmental culture conditions was

compared.














CHAPTER 2
COMPARATIVE GROWTH, MORPHOLOGY AND ANATOMY OF IN VITRO AND
EX VITRO CULTURED EASY- AND DIFFICULT-TO-ACCLIMATIZE SEA OATS
(Uniolapaniculata L.) GENOTYPES

Introduction

Sea oats (Uniolapaniculata L.), is a perennial dune grass native to the southeast

U.S. This species is commonly used for beach restoration and dune stabilization in

Florida after dune systems are damaged or destroyed by tropical storms, hurricanes or

human activity (Bachman and Whitwell, 1995). Sea oats has the ability to rapidly

colonize and establish, and exhibits high tolerance to heat, drought, and salinity (Wagner,

1964; Woodhouse, 1982). The root and rhizome systems hold sand particles together,

while the vegetation above ground retards wind and water driven erosion and promotes

sand deposition (Woodhouse, 1982).

Sea oats is propagated under nursery conditions from field-collected seed.

However, seed sources are limited due to increased beach erosion. Moreover, concerns

regarding the introduction of un-adapted ecotypes to revegetation sites have also limited

the use of seeds and or plant materials obtained from distant geographic sources.

Consequently, alternative propagation methods, including micropropagation, have been

developed for mass production of sea oats plants.

A micropropagation protocol for sea oats was developed by Philman and Kane

(1994). Additionally, 28 different sea oats genotypes were established, multiplied, and

rooted using this protocol. However, significant differences in survival and

acclimatization capacity were observed between genotypes when plants were transferred









to ex vitro conditions. Similarly, many other horticultural plants species are readily

micropropagated in vitro but exhibit poor acclimatization and subsequent survival ex

vitro (Debergh and Zimmerman, 1991). Alterations in the morphological and anatomical

characteristics of plants during and after in vitro culture appear to play a critical role in

successful acclimatization to greenhouse conditions (Donnelly and Vidaver; 1984a;

Serret and Trillas, 2000; Wetzstein and Sommer, 1982). In sea oats, a better

understanding of the morphological and anatomical differences between genotypes is

required to develop more efficient micropropagation procedures.

In vitro plantlets produced under high humidity conditions and transferred ex vitro

to the soil are usually very susceptible to desiccation (Wetzstein and Sommer, 1983).

Typically, low light intensities in vitro result in the production of leaves that resemble

shade or hydrophytic leaves ex vitro (Lee et al., 1988). These leaves often have little

epicuticular and cuticular wax formation (Grout, 1975) and malfunctioning stomata

(Brainerd and Fuchigami, 1981). Leaves produced in vitro also have low chlorophyll

content (Grout and Aston, 1977), restricted leaf blade expansion (Kozai et al., 1992), low

stomatal density (Ziv, 1995), poorly differentiated spongy and palisade tissues (Donnelly

et al., 1985), low percent dry matter, and/or hyperhydrated shoots (Ziv, 1991). All of

these characteristics negatively impact the potential for ex vitro acclimatization.

Some researchers have attributed hyperhydration to the effect of certain plant

growth regulators in the medium, such as N6-benzyladenine (BA) (Ziv, 1991; Khan et al.,

2002). Other medium components, such as sucrose, also affect the anatomy of plantlets.

Some in vitro plants contain chloroplastic starch granules while others have little or none









depending upon sucrose concentration (Dhawan and Bhojwani, 1987; Lee et al., 1985;

Queralt, 1989).

These morphological and anatomical characteristics result in insufficient

photosynthetic capacity to achieve a positive carbon balance (Grout and Aston, 1978).

The degree to which plants are affected by the in vitro environment depends on the plant

species. Furthermore, genotypic differences during in vitro propagation have also been

reported (Llorente and Ap6stolo, 1998). The extent of the differences between sea oats

genotypes needs further investigation.

The only effective cytokinin for shoot multiplication of sea oats is BA. Werbrouck

et al. (1995) identified a negative carryover effect of BA during ex vitro acclimatization.

The extent of this carryover effect appeared to depend upon the formation of BA

derivatives that accumulated in the base of plantlets and negatively affected rooting and

survival ex vitro. Furthermore, duration of in vitro incubation with BA appeared to affect

survival and acclimatization ex vitro (Moncalean et al., 2001). Preliminary studies with

sea oats genotypes indicated that there is similar negative BA carryover effect after ex

vitro transfer (Valero-Aracama et al, 2003). The extent of the effects of BA on

multiplication, rooting and survival ex vitro of sea oats genotypes needs further

investigation.

Additionally, other in vitro culture conditions, such as nutrient, sugar or plant

growth regulator concentration in the medium and gas composition in the vessel

headspace, change with culture duration. Understanding the effects of these changing

conditions with time on the anatomy and morphology of sea oats genotypes and









consequently on acclimatization and survival ex vitro will help improve the sea oats

micropropagation protocol.

In the present study, we compared the morphology of easy- and difficult-to-

acclimatize sea oats genotypes as affected by in vitro multiplication and rooting

conditions. Comparative analysis of leaf anatomy, stomatal formation in leaf surfaces

and chloroplast ultrastructure in mesophyll cells and bundle sheath cells of in vitro and ex

vitro leaf samples were conducted. Ultimately, we investigated the relationship between

morphological and anatomical development of sea oats genotypes and their differing

capacity for acclimatization.

Materials and Methods

Culture Conditions

Two established, stabilized and indexed sea oats genotypes (Uniolapaniculata L.),

genotyped using random amplified polymorphic DNA (RAPD) genetic analyses

(Ranamukhaarachchi, 2000), previously characterized as easy- and difficult-to-

acclimatize (EK 16-3 and EK 11-1, respectively) were used in this study. Five sea oats

shoot clusters (each consisting of three shoots, 25-mm long) of EK 16-3 and EK 11-1

genotypes were subcultured in 80 mL sterile multiplication medium (Stage II) into

separate Magenta GA7 vessels (Magenta Corp., Chicago, IL). Culture medium consisted

of Murashige and Skoog (MS) inorganic salts (Murashige and Skoog, 1962),

supplemented with 87.6 mM sucrose, 0.56 mM myo-inositol, 1.2 [tM thiamine-HC1, 2.2

[tM N6-benzyladenine (BA), and solidified with 8 g L-1 TCTM agar (PhytoTechnology

Laboratories, Shawnee Mission, KS). The medium was adjusted to pH 5.7 with 0.1 N

KOH prior to the addition of agar and autoclaving at 1.2 kg cm-2 and 121 C for 20 min.









Cultures were maintained for 8 weeks in a growth chamber at 24 1 C, 58 5%

relative humidity (RH), and 16-h photoperiod provided by cool-white fluorescent lamps

(General Electric F20WT12-CW), at a 40 5 [[mol m-2 s-1 photosynthetic photon flux

(PPF) as measured at culture level. Subsequently, 25-mm long single shoots from each

genotype were excised and transferred to rooting medium (Stage III).

Stage III rooting basal medium consisted of 80 mL sterile half-strength MS

medium, supplemented with 0.56 mM myo-inositol, 1.2 [LM thiamine-HC1, 87.6 mM

sucrose, and 10 [LM ca-naphthalene acetic acid (NAA), and adjusted to pH 5.7 with 0.1 N

KOH. Medium was solidified with 8 g L-1 TCTM agar and autoclaved at 1.2 kg cm-2 and

121 C for 20 min. Each GA7 culture vessel contained 8 single microcuttings, and were

maintained in a culture room at 22 2 C air temperature, 16-h photoperiod provided by

cool-white fluorescent lamps (General Electric F96T12-CW-WM), and 100 5 |tmol

m-2 S-1 PPF as measured at culture level.

Microcuttings were transferred to acclimatization conditions (Stage IV) into 48-cell

plug trays (8 six-celled blocks, each cell 4 x 6 x 5.5 cm; T.O. Plastics, Inc., Clearwater,

MN) containing coarse vermiculite as supporting material. Plug trays were placed in a

greenhouse under controlled environmental conditions. Plantlets were hand watered as

needed, and Peters 20N-20P-20K liquid fertilizer (150 mg N L-1; The Scotts Company,

Marysville, OH) was applied weekly.

Description of Treatments

Effect of Stage II duration on in vitro rooting and ex vitro survival of sea oats
genotypes

Plantlets of both genotypes were cultured for 4, 8 and 12 weeks in Stage II, and

subsequently, 48 unrooted microcuttings were directly transferred to Stage IV conditions.









Percent survival was determined after 4 weeks ex vitro. Another set of 64 microcuttings

per genotype, were concurrently transferred to Stage III rooting medium in vessels

arranged in a completely randomized design. Percent rooting was evaluated after 6

weeks in Stage III. Subsequently, 48 microcuttings were transferred to Stage IV

conditions and percent survival was determined after 4 weeks ex vitro culture. Arcsine

transformation was applied to the data where appropriate. Microcuttings were

maintained at 25/22.5 C day/night temperature from August 8 until December 12, 2002

in a greenhouse in Gainesville, FL with light intensities ranging 700-1200 [tmol m-2 s-1

This experiment was replicated once in time.

Comparative Stage II shoot multiplication and growth of sea oats genotypes

Plantlets from EK 11-1 and EK 16-3 consisting of 3 shoot clusters were cultured

for 4, 8, and 12 weeks under Stage II conditions. Number of shoots, number of leaves

per shoot, leaf length, and dry weight of plantlets from 5 replicate vessels per genotype

were measured after each culture period. Production of rootable microcuttings > 25 mm

long were considered as harvestable microcuttings. A completely randomized design was

used. This experiment was replicated once in time.

Effect of Stage III duration on in vitro rooting, growth and development and ex
vitro survival of sea oats genotypes

EK 11-1 and EK 16-3 plantlets were cultured for 8 weeks under Stage II

conditions. Subsequently, single shoots of each genotype were excised and transferred to

Stage III conditions for 3, 6 or 9 weeks in 8 replicate vessels per treatment and time

interval. After Stage III culture, 2 rooted plantlets per vessel were selected to measure

shoot number, leaf number per plant, leaf length, root number, root length, and dry

weight of shoots and roots per plantlet. The remaining 6 plantlets per vessel were









transferred to Stage IV conditions into the greenhouse. The same growth measurements

of 2 plants per six-celled pack were obtained after four weeks ex vitro culture.

Microcuttings were maintained at 25/22.5 C day/night temperature from June 12 until

July 24, 2003 in a greenhouse in Gainesville, FL with light intensities ranging 700-1000

[tmol m-2 s-1. This experiment was replicated once in time.

Comparative anatomy of sea oats genotypes during Stage II, Stage III and Stage IV
culture

Leaf histological cross sections collected from EK 11-1 and EK 16-3 microcuttings

cultured in vitro in Stage II and Stage III and from greenhouse-produced leaves were

made. For optical light microscopy (OLM) and transmission electron microscopy

(TEM), leaf sections approximately 2 mm from the center of the leaf blade were fixed in

Trumps fixative solution (McDowell and Trump, 1976). Fixative infiltration was

achieved under vacuum for 2 days. Leaf tissues were then rinsed 3 times in phosphate

buffer (pH 7.2), post-fixed in a 1% buffered osmium tetroxide solution and then rinsed in

phosphate buffer, 3 times in distilled water, and dehydrated in a five-step ascending ethyl

alcohol series (25, 50, 75, 95, 100%) followed by dehydration in 100% acetone. An

enbloc stain of 2% uranyl acetate was applied between the 75 and 95% steps of the ethyl

alcohol dehydration series. Leaf sections were then embedded in Spurr resin (Spurr,

1969). For OLM, thick leaf sections (500 nm) were obtained with a Leica Ultracut

ultramicrotome R (Leica Microscopy and Scientific Instruments, Deerfield, IL) and then

collected on glass slides. Sections were stained with 0.2% toludine blue and examined

using an Olympus BH-2 Epifluorescent Microscope (Olympus America Inc., Melville,

NY). Photographs were taken using a Pixera 120C digital camera attachment. For TEM,

ultrathin leaf sections (70 nm) were cut from the center part of the leaf blade with a Leica









Ultracut ultramicrotome R, collected on 0.35% form-var coated copper grids, stained

with methanolic uranyl acetate and lead citrate (Reynolds, 1963). Sections were viewed

on a Hitachi H7000 transmission electron microscope (Hitachi Scientific Instruments,

Danbury, CT) at 75 kV. Digital micrographs were taken on a BioScan/Digital

Micrograph 2.5 (Gatan Inc., Pleasanton, CA) at an exposure level optimized for viewing

the outer layer and processed with MEGA View III/AnalySIS 3.1 (Soft Imaging System

Corp., Lakewood, CO).

For scanning electron microscopy (SEM), leaf sections of approximately 5 mm

from the center of the leaf blade were immersed in 100% methanol. Leaf tissues were

collected from cultures 2 h after photoperiod started and from cultures 5 h after dark

period started. Sections were lyophilized using a Bal-Tec 030 critical point drier

(ICMAS Inc., Alcoa, TN) with liquid C02, sputter coated with gold-palladium using a

Denton Vacuum Desk II (Denton Vacuum, Moorestown, NJ) for approximately 50 s and

viewed with a Hitachi S-4000 FS scanning electron microscope (Hitachi Scientific

Instruments, Danbury, CT) operating at 6 kV. Digital images were processed using

SEMages 16 software (Advance Database Systems, Inc., Denver, CO).

Statistical Analyses

Percent data was transformed using the arc sine transformation, and the significant

differences among means were determined by two-way analysis of variance (ANOVA)

using the GLM procedure of SAS (SAS institute Inc., 1999). Interactions among

genotypes and time, where appropriate, are shown in the tables and graphs. Separate a

posteriori tests for significant differences among or between means were analyzed using

the Waller-Duncan procedure at P < 0.05.









Results

Effect of Stage II Duration on In Vitro Rooting and Ex Vitro Survival of Sea Oats
Genotypes

In vitro rooting

All Stage III EK 16-3 plantlets rooted regardless of Stage II duration. Rooting of

EK 16-3 plantlets was significantly greater (P = 0.0136) than EK 11-1 plantlets, with 93,

86 and 99% rooting at 4, 8 and 12 weeks Stage II, respectively. Significant differences in

root architecture and morphology were observed between the two genotypes after 4, 8

and 12 weeks Stage II culture following 6 weeks Stage III culture (Figure 2-1). The EK

11-1 root system consisted of a few thick elongated roots with small lateral branching

without visible root hairs after 8 and 12 weeks Stage II. The EK 16-3 root system

exhibited large root numbers that were shorter than EK 11-1 roots regardless of Stage II

duration. Visual assessment of shoot development during Stage III, revealed that EK 16-

3 plantlets had greater leaf length, leaf expansion and shoot thickness than EK 11-1

plantlets (Figure 2-1). However, increasing Stage II duration from 8 to 12 weeks resulted

in decreased shoot and root development in EK 16-3 plantlets after 6 weeks Stage III.

This decrease in plantlet growth was attributed to the quality of harvestable microcuttings

obtained after 12 weeks Stage II, since older shoots had died and the remaining

harvestable microcuttings were smaller than after 4 or 8 weeks culture. During Stage II,

visual assessments demonstrated that EK 16-3 produced longer leaves than EK 11-1.

Ex vitro survival

Survival of microcuttings transferred ex vitro directly from Stage II cultures was 0

(EK 11-1) and less than 17% (EK 16-3) (Figure 2-2A). There was no significant effect of

Stage II duration on survival of unrooted microcuttings within each genotype. EK 16-3





















































Figure 2-1. Comparative morphological differences in shoot and root development of EK
11-1 (left) and EK 16-3 (right) sea oats genotypes after 4, 8 and 12 weeks
Stage II culture (from top to bottom, respectively) followed by 6 weeks Stage
III culture. Scale = 1 cm.


Week4









Stage III microcuttings that rooted for 6 weeks Stage III, exhibited nearly 100% ex vitro

survival regardless of Stage II duration. In contrast, survivability of EK 11-1 rooted

microcuttings was 30, 54, and 64% respectively for 4, 8 and 12 weeks Stage II culture

(Figure 2-2B). Even though survival was higher in EK 11-1 plantlets cultured for 12

weeks rather than 8 weeks Stage II, we determined that the optimal Stage II culture

duration was 8 weeks, since the microcutting quality was higher at 8 weeks than at 12

weeks Stage II.

Comparative Shoot Multiplication and Growth of Sea Oats Genotypes during Stage
II Culture

Visual observations indicated that EK 11-1 and EK 16-3 sea oats genotypes

exhibited shoot multiplication and subsequent elongation of leaves after 4 weeks culture

(Figure 2-3; Table 2-1). However, unlike EK 11-1 leaves, EK 16-3 leaves continued

elongating after 8 weeks culture. At week 8 Stage II, both genotypes exhibited browning

of shoots and leaves, which continued until week 12 Stage II culture. At that time, most

of the longest shoots were brown and only the newly developed shoots remained green in

either genotype.

Shoot dry weights in both genotypes were greatest after four weeks culture (Figure

2-4). Browning and dying of tissue steadily increased with time (Figure 2-3). Shoot

number per plantlet (Table 2-1) of both genotypes increased from weeks 4 to 8 during

Stage II but decreased from weeks 8 to 12. Similarly, leaf number decreased during

Stage II culture in both genotypes due to leaf death (Table 2-1). Leaf length was

significantly higher for EK 16-3 than EK 11-1 plantlets throughout Stage II. After four

weeks culture, leaf length decreased in both genotypes (Table 2-1). The longest leaves,













.G- ** EK11-1 A
100 -Tx G:NS EK 16-3

3 80

E 60
o 40
g 20
40
0 20
t
T: *
G:- a a a B
100 T xG:
80 b
> b

0 I
2 40

M 20
0

4 8 12
Time in Stage II (weeks)

Figure 2-2. Comparative survival percentage after 4 weeks under Stage IV culture of A:
Stage II unrooted, and B: Stage III rooted sea oats microcuttings after 4, 8 or
12 weeks Stage II culture. Error bars indicate SE (n = 16). ANOVA is shown
on top left corner of each graph; T: Time, G: Genotype, NS, **: Non-
significantly or significantly different at P = 0.01, respectively. Different
letters on top of histobars are significantly different according to Waller-
Duncan test at P < 0.05.






















































Figure 2-3. Comparative morphological differences in shoot multiplication of EK 11-1
(left) and EK 16-3 (right) sea oats genotypes after 4, 8 and 12 weeks Stage II
culture (from top to bottom, respectively). Scale = 1 cm.

























350
350 T:**
G: NS EK 11-1
300 TxG:NS a EK 16-3
E ~EK 16-3
250 5
z 200 b
150
o 100
50
0
4 8 12
Time in Stage II (weeks)


Figure 2-4. Comparative Stage II shoot dry weights ofEK 11-1 and EK 16-3 sea oats
genotypes after 4, 8 and 12 weeks culture. Error bars indicate SE (n = 16).
ANOVA is shown on top left corer; T: Time, G: Genotype, NS, **: Non-
significantly or significantly different at P = 0.01, respectively. Different
letters on top of histobars are significantly different according to Waller-
Duncan test at P < 0.05.



















Table 2-1. Comparative shoot number, leaf number and leaf length of EK 11-1 and EK 16-3 sea oats genotypes after 4, 8 and 12
weeks Stage II culture.
Stage II Shoot Number Leaf Number Leaf Length (mm)
Duration
(weeks)z EK 11-1 EK 16-3 x EK 11-1 EK 16-3 x EK 11-1 EK 16-3
4 12.4 + 0.5 14.7 + 0.7 13.6 + 0.6b 102.2 + 3.8 106.8 + 4.5 104.5 + 4.2a 18.1 + 0.7b 25.5 0.9a
8 16.2 + 0.9 17.3 + 1.1 16.8 + 1.0a 92.4 + 5.2 81.7 + 9.4 87.1 + 7.3b 8.8 + 0.3d 16.5 0.7b
12 13.9 1.0 15.7 1.3 14.8 1.2b 80.7 5.7 69.7 6.0 75.2 5.9c 9.4 0.8d 11.4 0.8c
x 14.2 0.8B 15.9 + 1.0A
Analysis of variancex
Genotype (G) NS **
Time (T) ** ** **
G*T NS NS **
zMeans SE followed by different letters are significantly different according to Waller-Duncan test at P < 0.05.
YWhen no interaction was observed, means followed by lowercase letters indicate significant differences within Stage II duration.
Means followed by uppercase letters indicate significant differences between genotypes.
x **: Significant at P < 0.05 or 0.01, respectively (n = 16).











which were also the oldest, died and the newly formed leaves were considerably shorter.


Relative leaf length frequencies indicated a greater occurrence of longer leaves in EK 16-


3 than EK 11-1 plantlets throughout Stage II culture (Figure 2-5).


0,
0- o
C)


* 0

- a5 20





0

80

^, 60
60
0 0
4-zo


cr


Leaf length intervals (mm)


Figure 2-5. Comparative relative leaf length frequency per plantlet of sea oats genotypes
after A: week 4, B: week 8, and C: week 12 Stage II culture.









Effect of Stage III Duration on In Vitro Rooting and Growth and Ex Vitro Survival
and Acclimatization of Sea Oats Genotypes

In vitro rooting and growth

Shoot and root dry weights of both genotypes increased throughout Stage III

culture (Figure 2-6). At week 3, shoot dry weights of EK 16-3 plantlets were lower than

those ofEK 11-1 plantlets, but at weeks 6 and 9, EK 16-3 shoot dry weights were higher

than those ofEK 11-1 plantlets. Conversely, root dry weights were similar at week 3 but

higher for EK 11-1 than for EK 16-3 plantlets at weeks 6 and 9. Shoot to root dry weight

ratios were not significantly different between genotypes regardless of Stage III culture

duration (data not shown).

Shoot number increased with time in EK 11-1 plantlets and it was not significantly

different from weeks 6 to 9 in EK 16-3 plantlets (Figure 2-7; Table 2-2). Significantly

higher numbers of leaves were produced by EK 11-1 plantlets as the duration of Stage III

increased to nine weeks. However Stage III duration did not affect leaf number of EK

16-3 plantlets (Table 2-2). Conversely, by week 3, leaves were 2-fold longer in EK 16-3

than EK 11-1 plantlets and elongated rapidly, becoming 6.5-fold longer in EK 16-3 than

EK 11-1 plantlets by week 9 Stage III. Leaf length distributions (Figure 2-8) indicated

that leaf elongation was significantly inhibited in EK 11-1. Ninety-five percent of the EK

11-1 leaves were < 15 mm long by week 9, in contrast with EK 16-3, with 50% of the

leaves > 16 mm long.

Stage III root development differed significantly between genotypes. Root

production, at each culture interval, was higher in EK 16-3 than in EK 11-1 plantlets

(Table 2-3). Conversely, EK 11-1 roots were longer than EK 16-3 roots throughout Stage

III culture (Figure 2-7; Table 2-3). Root length distributions (Figure 2-9) indicated that










EK 11-1 plantlets produced a greater percentage of longer roots than EK 16-3 plantlets

from week 3, and continued elongating until 50% of the roots ranged from 243-550 mm

in length.


ba-r
4oa


o
o






a
B


^


180
160
140
120
100
80
60
40
20
0
160
140
120
100
80
60
40
20
0


3 6 9
Time in Stage III (weeks)


Figure 2-6. Comparative A: shoot dry weights, and B: root dry weights of sea oats
genotypes after weeks 3, 6 and 9 Stage III culture. Error bars indicate SE (n
32). ANOVA is shown on top left corner of each graph; T: Time, G:
Genotype, *, **: Significantly different at P 0.05 or 0.01, respectively.
Different letters on top of histobars are significantly different according to
Waller-Duncan test at P < 0.05.






















































Figure 2-7. Comparative morphological differences in rooting and shoot multiplication of
EK 11-1 (left) and EK 16-3 (right) sea oats genotypes after 8 weeks Stage II
followed by 3, 6 and 9 weeks Stage III culture (from top to bottom,
respectively). Scale = 1 cm.




















Table 2-2. Comparative shoot number, leaf number and leaf length of EK 11-1 and EK 16-3 sea oats genotypes after 3, 6 and 9 weeks
Stage III culture.
Stage III Shoot Number Leaf Number Leaf Length (mm)
Duration
Duration EK 11-1 EK 16-3 EK 11-1 EK 16-3 EK 11-1 EK 16-3
(weeks)
3 4.5 + 0.3cdz 4.3 0.3d 25.6 + 0.9c 24.3 1.2c 3.8 + 0.2c 7.8 0.4c
6 5.7 + 0.3b 5.3 + 0.3bc 39.2 + 1.8b 29.0 + 1.7c 5.6 + 0.5c 23.2 + 1.7b
9 7.4 + 0.5a 4.6 + 0.5cd 45.5 + 2.6a 24.0 1.5c 6.9 + 0.6c 44.9 3.3a
Analysis of variancey
Genotype (G) ** ** **
Time (T) ** ** **
G*T ** ** **
zMeans SE followed by different letters are significantly different according to Waller-Duncan test at P < 0.05.
Y **: Significantly different at P 0.01 (n = 32).

























Table 2-3. Comparative root number and root length of EK 11-1 and EK 16-3 sea oats
genotypes after 3, 6 and 9 weeks Stage III culture.
Stage III Root Number Root Length (mm)
Duration
Duration EK 11-1 EK 16-3 EK 11-1 EK 16-3
(weeks)
3 5.1 + 0.5dz 12.9 + 1.3c 11.4 0.1e 5.7 0.2e
6 6.5 + 0.5d 21.1 + 1.5b 121.6 + 5.8b 32.7 1.0d
9 6.8 0.7d 26.0 1.8a 251.1 6.1a 72.5 2.8c
Analysis of variancey
Genotype (G) ** **
Time (T) ** **
G*T ** **
zMeans SE followed by different letters are significantly different according to Waller-
Duncan test at P < 0.05.
Y **: Significantly different at P 0.05 (n = 32).







40











100 A
80 Week 3 EKll-1
S60 EK16-3

"> t 20-
20

r0




80 o Week 6 B

60
20

0






80 Week 9 C

60
o' 6' 20











20













Figure 2-8. Comparative relative leaf length frequency per plantlet of sea oats genotypes
after A: week 3, B week 6, and C: week 9 Stage III culture.
after A: week 3, B: week 6, and C: week 9 Stage III culture.







41











100
80 Week 3 EK 11-1 A
60 I i EK 16-3

40

5 30
20

10

0
0- ______-------------- _______
80 Week 6 B
60

a"- 40
o 40

30

20

10


80 Week 9 C
60

0 40

Sa 30

^ 20

0o -. on 0 I
10 HH.H,. I I I I I ,



0" n.H

Root length intervals (mm)

Figure 2-9. Comparative relative root length frequency per plantlet of sea oats genotypes
after A: week 3, B: week 6, and C: week 9 Stage III culture.









Ex vitro survival and acclimatization

Stage III duration had a significant effect on ex vitro survival of sea oats genotypes

especially in EK 11-1. After 4 weeks Stage IV, survivability was 79, 94 and 94% for EK

16-3, and 1, 20 and 26% for EK 11-1 at weeks 3, 6 and 9 Stage III culture, respectively.

Shoot and root dry weights after four weeks Stage IV, were significantly greater for EK

16-3 than EK 11-1 plantlets regardless of Stage III duration (Figure 2-10A, B). Shoot

and root dry weights increased in both genotypes with duration. Comparative ex vitro

survival and growth of rooted microcuttings of both genotypes previously cultured for

eight weeks in Stage II and six weeks in Stage III, are shown at weeks 0, 3 and 6 after

transfer to Stage IV in Figure 2-11.

Anatomical and Ultrastructural Comparisons

Optical light microscopy (OLM)

Leaf sections collected from greenhouse-produced leaves of plants established ex

vitro (Figure 2-12) served as reference for leaf development comparisons of plantlets in

vitro (Figures 2-13 and 2-14). The anatomy of ex vitro sea oats plants was typical of C4

plants, and was very similar between EK 11-1 and EK 16-3 genotypes (Figure 2-12).

Both genotypes possessed a compacted mesophyll composed of few large cells with most

visible chloroplasts present in cells surrounding the vascular bundle. Both genotypes

possessed extensive vascular bundles that were oval and occupied most of the cross

section of the leaves (Figure 2-12). Bundle sheath cells of greenhouse-produced leaves

were similar in shape between genotypes, contained larger chloroplasts than those of in

vitro plantlets (Figures 2-13 and 2-14) and were arranged centripetally in the bundle

sheath.



















500

400
+, ,

300

a200
o S
m 100

0

400

,300

S 200

100

0


Time in Stage III (weeks)

Figure 2-10. Comparative ex vitro A: shoot dry weights, and B: root dry weights of sea
oats genotypes after weeks 3, 6 and 9 Stage III culture followed by 4 weeks
Stage IV culture. Error bars indicate SE (n = 1, 14 and 16 for EK 11-1 at 3, 6
and 9 weeks, respectively and n = 32 for EK 16-3 during Stage III). ANOVA
is shown on top left corner of each graph; T: Time, G: Genotype, NS, **:
Non-significantly or significantly different at P < 0.01, respectively. Different
letters on top of histobars are significantly different with time according to
Waller-Duncan test at P < 0.05.


T:** EK 11-1 A
G: ** EK 16-3
Tx G: NS
a

b





T:** B
G: **
Tx G: NS







c n
_M 8 1


















































Figure 2-11. Comparative morphological differences in shoot multiplication of EK 11-1
(left) and EK 16-3 (right) sea oats genotypes previously cultured for 8 weeks
in Stage II and 6 weeks in Stage III, at weeks 0, 3, and 6 Stage IV culture
(from top to bottom, respectively).









Schlerenchyma tissue was clearly present between the epidermal layers and the

vascular bundles. A visible cuticle was present over the abaxial epidermis of both

genotypes. The adaxial epidermis of leaves contained bulliform cells. These cells were

responsible for the epidermal foldings which formed the characteristic adaxial ribs of the

leaf blades. Multicellular glands were observed on both abaxial and adaxial surfaces.

Early in Stage II (Figure 2-13A, D), large intercellular spaces in the mesophyll

tissue of both genotypes were present. Both genotypes comprised mesophyll cells of

varied sizes and shapes. At week 8 Stage II, these intercellular spaces were still present

in EK 11-1 plantlets whereas mesophyll of EK 16-3 leaves were more compact with

fewer intercellular spaces. A compact mesophyll was present in both genotypes by week

12 Stage II. Production of adaxial sinuses was only observed in EK 11-1 by week 12

Stage II. Adaxial sinuses were present in EK 16-3 plantlets by week 4 Stage II.

Conversely, EK 11-1 plantlets lacked adaxial sinuses during weeks 4 and 8, and

possessed round rather than oval vascular bundles. In greenhouse-produced leaves

(Figure 2-12) both genotypes exhibited a visible cuticle in the adaxial epidermis.

However, the abaxial epidermis of both genotypes lacked a visible cuticle during Stage II

(Figure 2-13). Throughout Stage II culture, schlerenchyma tissue was present between

the epidermal layers and the vascular bundles of EK 16-3 plantlets, whereas initial

development of schlerenchyma tissue was only present at week 12 culture in EK 11-1

plantlets. Based on visual observations, mesophyll and bundle sheath cells of either

genotype contained fewer numbers of chloroplasts than greenhouse-grown plants.

During Stage III culture, there were visible changes in leaf development in both

genotypes (Figure 2-14). At weeks 6 and 9 Stage III culture, EK 16-3 plantlets exhibited









anatomical features similar to greenhouse-grown plants, whereas differences in tissue

organization and anatomical features were observed between greenhouse-produced leaves

and leaves of Stage III EK 11-1 plantlets. Intercellular spaces appeared to be as frequent

during Stage III as during Stage II. EK 16-3 leaves were comprised of a more compacted

mesophyll than EK 11-1 plantlets, and larger intercellular spaces. Furthermore, the

mesophyll tissue in leaves of both genotypes was comprised of numerous small cells as

compared to that of greenhouse-produced leaves (Figure 2-12). Both genotypes lacked

adaxial sinuses at week 3 Stage III culture and exhibited large adaxial sinuses at weeks 6

and 9 (Figure 2-14B, C, E, F). Vascular bundles were round at week 3 (Figure 2-14A, D)

but became oval with time. At week 6, mesophyll and vascular bundles of EK 16-3

plantlets (Figure 2-14E) were highly structured and similar to those of greenhouse-

produced leaves (Figure 2-12). At that time, EK 16-3 vascular bundles were oval and

completely occupied the leaf cross section and filled the adaxial leaf sinuses. However,

mesophyll and vascular bundles were comparably disrupted and unorganized in EK 11-1

plantlets throughout Stage III (Figure 2-14A-C). EK 16-3 plantlets developed a visible

abaxial epidermal cuticle by week 6 (Figure 2-14E, F). In contrast, EK 11-1 plantlets

exhibited minimal cuticle development during Stage III (Figure 2-14A-C).

Schlerenchyma tissue between the epidermal layers and the vascular bundles was visible

in EK 16-3 plantlets throughout Stage III (Figure 2-14D-F), whereas it was less apparent

in EK 11-1 plantlets and only visible after six weeks Stage III (Figure 2-14B-C).



























EK 11-1










EK 16-3 .



bc


Figure 2-12. Comparative histological leaf sections of A: EK 11-1, and B: EK 16-3 sea
oats genotypes in greenhouse-produced leaves. (b: bundle sheath cell, c:
cuticle, m: mesophyll cell, s: stoma). Scale = 100 |tm.
















EK 11-1

B









IllI Ill

EK 16-3







n,





4 weeks 8 weeks 12 weeks

Figure 2-13. Comparative histological leaf sections of EK 11-1 (A, B, C) and EK 16-3
(D, E, F) genotypes at week 4 (A, D), week 8 (B, E) and week 12 (C, F) Stage
II culture. (b: bundle sheath cell, i: intercellular space, m: mesophyll cell, s:
stoma). Scale = 100 |tm.















EK 11-1









EK 16-3______ __
S,,,'






Iml b Dt M E









3 weeks 6 weeks 9 weeks

Figure 2-14. Comparative histological leaf sections of EK 111 (A B, C) andEK 16-3
(D, E, F) genotypes at week 3 (A, D), week 6 (B, E) and week 9 (C, F) Stage
III culture. (b: bundle sheath cell, c: cuticle, i: intercellular space, m:
mesophyll cell, s: stoma). Scale = 100 |tm.









Scanning electron microscopy (SEM)

Leaf sections collected from greenhouse-grown plants (Figures 2-15 and 2-16)

served as a reference for leaf surface and stomata comparisons with in vitro produced

plantlets (Figures 2-17 and 2-18). Adaxial epidermal surface features and stomate

morphology of greenhouse-produced leaves were similar between both genotypes

(Figures 2-15 and 2-17). Both genotypes possessed glands on the adaxial ribs (Figure 2-

15). Stomata were found arranged in rows inside the invaginations of the adaxial leaf

surfaces. Each row had 1 or 2 epidermal cells between each stomate in either genotype.

Epidermal cells were raised, convex and rectangular. A closer image of the stomata

(Figure 2-16) of both genotypes revealed significant epicuticular wax layer arranged in a

homogeneous layer and bearing a crystalline structure. Stomata were characteristic of

grass species, with elongated guard cells surrounded by two large subsidiary cells, and

very small stomatal apertures. Leaves were amphistomatous (with stomata in both

epidermal surfaces), and based on visual assessments, stomatal density was greater on the

adaxial surface. Yet, because stomata were present in the invaginations of the adaxial

epidermises, stomatal density could not be accurately quantified. Stomata were arranged

in longitudinal rows of cells throughout the leaf blade length.

Similarly to that observed in histological sections, leaves produced in Stage II

exhibited no or relatively low wax deposition in both genotypes (Figures 2-17A-D). Wax

clusters were observed randomly over the leaf surfaces, instead of in a homogeneous

layer as in greenhouse-grown plants. Stomata apertures were fused or blocked with wax

depositions regardless of being collected during light or dark period (images not shown)

during Stage II culture. Both genotypes exhibited similar stomatal structure and

epidermal surface features regardless of Stage II duration.









During Stage III culture (Figures 2-18A-D), increased wax deposition was

observed on the leaf surfaces of both sea oats genotypes, especially on EK 16-3 leaves

after nine weeks culture. Abaxial cuticle development was minimal at the OLM level in

EK 11-1 plantlets (Figure 2-18C). Wax deposition was more uniform in EK 16-3 than in

EK 11-1 leaf surfaces (Figure 2-18F). While EK 11-1 stomatal apertures appeared

blocked in most cases, clearly defined stomatal apertures without blockage were observed

in EK 16-3 leaves. The stomatal structure was similar between genotypes regardless of

Stage III duration.







EK 11-1











EK 16-3







Figure 2-15. Comparative SEM of adaxial epidermis of A: EK 11-1, and B: EK 16-3
genotypes in greenhouse-produced leaves. (g: gland, s: stomate). Scale = 75
[tm.





























EK 11-1










EK 16-3







Figure 2-16. Comparative SEM of stomata on adaxial epidermis of A: EK 11-1, B: EK
16-3 genotypes in greenhouse-produced leaves. Scale = 10 atm.











































4 weeks 8 weeks 12 weeks

Figure 2-17. Comparative SEM of stomata on adaxial epidermis of A-C: EK 11-1, and
D-F: EK 16-3 genotypes at week 4 (A, D), week 8 (B, E) and week 12 (C, F)
Stage II culture. Scale = 10 m.










































3 weeks 6 weeks 9 weeks
Figure 2-18. Comparative SEM of stomata on adaxial epidermis of A-C: EK 11-1, and D-
F: EK 16-3 genotypes at week 3 (A, D), week 6 (B, E), and week 9 (C, F)
Stage III culture. Scale = 10 tm.









Transmission electron microscopy (TEM)

Ultra-thin section examination of samples collected from EK 11-1 and EK 16-3

plants revealed additional details, such as differences in chloroplast ultrastructure in both

mesophyll and bundle sheath cells. Chloroplast ultrastructure in acclimatized plants was

very different in mesophyll cells compared to bundle sheath cells (Figure 2-19).

Ultrastructural characteristics of ex vitro leaves were similar between genotypes.

Mesophyll cell chloroplasts (Figures 2-19A, C) consisted of a highly compacted

thylakoid membrane system with minimal stroma. A compact thylakoid membrane

system was similarly observed in bundle sheath chloroplasts (Figures 2-19B, D) and

included large starch granules. Mesophyll cell chloroplasts were smaller and randomly

arranged within the cells, whereas those in bundle sheath cells were larger and arranged

centripetally within the bundle sheath.

Mesophyll cells of EK 11-1 Stage II plantlets contained chloroplasts with similar

thylakoid membrane distribution at weeks 4 and 8 (Figures 2-20A, B), however signs of

senescence, including thylakoid disruption, were observed by week 12 Stage II (Figure 2-

20C). Similar signs of senescence were present at week 12 Stage II culture in EK 16-3

chloroplasts (Figure 2-201). Bundle sheath cells of either genotype contained

chloroplasts that exhibited compacted thylakoid membrane throughout Stage II culture.

While no visible starch granules were observed in chloroplasts of EK 11-1 plantlets,

regardless culture duration, EK 16-3 chloroplasts of either cell type contained starch

granules at week 8 Stage II.

During Stage III culture, EK 11-1 mesophyll cell chloroplasts contained swollen

thylakoids at weeks 6 and 9 (Figure 2-21B, C), whereas typical compacted thylakoid

membranes were observed in EK 16-3 chloroplasts (Figure 2-21H, I). While large starch









granules were only observed in the early weeks of Stage III in either genotype, smaller

starch granules were observed at week 6 Stage III culture (Figure 2-21D, E, J).



M.-








EK 11-1



















EK 16-3











Figure 2-19. Comparative TEM of chloroplasts of A-B: EK 11-1, and C-D: EK 16-3
genotypes after Stage IV acclimatization in the greenhouse. Mesophyll (A, C)
and bundle sheath cell (B, D) chloroplasts are shown. Scale = 2 jtm. (ch:
chloroplast, s: starch granule, t: thylakoid membranes)






















































4 weeks


8 weeks 12 weeks


Figure 2-20. Comparative TEM of chloroplasts of EK 11-1 (A-F) and EK 16-3 (G-L)
genotypes at week 4 (A, D, G, J), week 8 (B, E, H, K), and week 12 (C, F, I,
L) Stage II culture. Mesophyll (A-C and G-I) and bundle sheath cell (D-F and
J-L) chloroplasts are shown. Scale = 2 |tm. (s: starch granule, t: thylakoid
membranes)


















































3 weeks 6 weeks 9 weeks


Figure 2-21. Comparative TEM of chloroplasts of EK 11-1 (A-F) and EK 16-3 (G-L)
genotypes at week 3 (A, D, G, J), week 6 (B, E, H, K), and week 9 (C, F, I, L)
Stage II culture. Mesophyll (A-C and G-I) and bundle sheath cell (D-F and J-
L) chloroplasts are shown. Scale = 2 |tm. (s: starch granule, t: thylakoid
membranes)









Discussion

We observed differing responses between the two genotypes tested in vitro, and

especially during ex vitro acclimatization. Anatomical and morphological differences

among plants are largely attributed to genotypic differences and phenotypical plasticity,

but also to different responses to these conditions (Majada et al., 2000). The culture

duration during multiplication and rooting stages also had differing effects on

acclimatization and survival of both genotypes to ex vitro conditions.

During in vitro multiplication and rooting conditions, the rate of leaf differentiation

and development differed between genotypes. In EK 11-1 development of grass-like

leaves with expanded blades was significantly suppressed. Therefore, the developmental

stages of leaves of both genotypes differed when the anatomical leaf comparisons were

made. However, our main interest was to evaluate the anatomical state of leaves at time

of transfer to rooting conditions and to ex vitro conditions. This information is helpful to

understand the differences in growth and survival ex vitro between both genotypes.

Histological observations indicated that unorganized tissues and abnormal

anatomical features, such as lack of cuticle, intercellular spaces in the mesophyll, blocked

stomata, or disrupted chloroplast ultrastructure, were common during Stage II for both

genotypes, especially in the difficult-to-acclimatize genotype, EK 11-1. Additionally,

SEM micrographs showed that both genotypes similarly lacked a homogeneous

epicuticular wax layer and exhibited fused or blocked stomata during Stage II.

Transmission electron micrographs showed chloroplast disruption in both

genotypes throughout Stage II. Several authors have described similar chloroplast

disruption as a characteristic of hyperhydric leaves (Wetzstein and Sommer, 1982; Ziv et

al., 1983; Lee et al., 1985; Capellades et al., 1991) and leaves produced under low light









intensities (Lee et al., 1988). Furthermore, like hyperhydric leaves, Stage II sea oats

leaves had fewer chloroplasts with reduced thylakoid stacking compared to greenhouse-

produced leaves (Drennan and van Staden, 1986; Jones et al., 1993).

Low light levels during culture result in swollen thylakoids in chloroplasts of

certain plant species (Queralt, 1989). Similarly, under the low light intensity

characteristic of Stage II, swollen thylakoids appeared more frequently in sea oats

genotypes than when they were cultured under higher light intensity during Stage III.

Frequently, swollen thylakoids occur after starch granules have accumulated within the

chloroplast (Queralt, 1989). When plants utilize their stored reserves, starch is used but

the thylakoid membranes remain swollen. Anatomical modifications caused by the tissue

culture environment have direct impact on diffusion of CO2 inside the in vitro produced

leaves and thus in photosynthesis in vitro (Desjardins, 1995). Alterations of the

chloroplast ultrastructure are associated with disorganization of the light harvesting

pigments. As a result, the photosynthetic capacity of developing leaves in vitro is low

(Lee et al, 1985), causing poor acclimatization and survival ex vitro of Stage II plantlets.

This state may contribute to the low ex vitro survival of sea oats Stage II microcuttings.

Another possible cause of mortality during acclimatization is poor control of water

loss (Brainerd and Fuchigami, 1982). In angiosperms, this phenomenon has been related

to poor stomatal functioning (Brainerd and Fuchigami, 1982) and reduced or abnormal

structure of epicuticular wax (Grout and Aston, 1977, Dhawan and Bhojwani, 1987,

Sutter, 1988). These characteristics were common in Stage II sea oats leaves. Therefore,

Stage II unrooted sea oats microcuttings of either genotype transferred directly ex vitro

probably exhibited poor control of water loss during acclimatization. Additionally, Stage









II unrooted microcuttings did not root after ex vitro transfer, possibly because the energy

resources of shoots were not sufficient to initiate root development or to continue shoot

growth. Unrooted microcuttings subsequently had minimal capacity to uptake water

from the substrate. Water uptake by roots, together with reduced water loss from shoots,

is critical for maintenance of water balance during acclimatization of in vitro produced

plants (Fila et al., 1998).

For Stage II sea oats shoots multiplication, only the cytokinin BA is effective

(Philman and Kane, 1994). The negative BA carry over effect of Stage II on ex vitro

survival of the difficult-to-acclimatize genotype was observed. However, EK 11-1

survival increased by increasing Stage II culture duration. This may, in part, be due to a

detrimental effect of using BA for Stage II multiplication. Moncalean et al. (2001)

observed decreased ex vitro survival of kiwi (Actidinia deliciosa Chev. 'Hayward')

explants with increasing Stage II incubation with BA. However, these authors used 35-

day subculture periods consisting of shorter BA incubation periods (from 30 min to 2

days) followed by incubation in plant growth regulator-free medium. Werbrouck et al.

(1995) observed a rapid accumulation of BA derivatives one day after incubation of

peace lily (Spathiphyllumfloribundum Schott 'Petite') plantlets on BA containing

medium. One of these derivatives ([9G]BA) accumulated at the base of the plantlets, did

not seem to be transported, and had longer half-life than BA. It was concluded that

[9G]BA accumulation was detrimental to root formation and to ex vitro survival of

plantlets. The increased survival in EK 11-1 with increasing Stage II duration was

possibly caused by the degradation of [9G]BA with time. These results are in agreement

with those reported by Moncalean et al. (2001), because longer incubation periods with









BA would result in higher accumulation of [9G]BA at the base of the plantlets,

consequently resulting in lower survival ex vitro.

To evaluate multiplication rates, we considered microcutting yield along with

quality of shoots, including shoot biomass, leaf number, leaf length and visual quality

assessments. Our observations indicated that eight weeks Stage II culture yielded the

highest multiplication rates of both genotypes. At week 8, multiplication rates decreased

mainly because leaf senescence was extensive, possibly a consequence of the

translocation of nutrients and energy reserves to newly developing tissues (Thomas and

Sadras, 2001). During senescence, at the molecular level chloroplasts and chlorophyll

are degraded. Proteins and lipids are also degraded in the form of amino acids and

sugars. The various culture conditions that change with time, such as BA concentration,

secondary metabolite production, sugar content, and gas composition inside the vessel

headspace, could cause the senescence of older leaves and the translocation of resources

to new shoots or storage areas (Buchanan-Wollaston et al., 2003). Therefore, in a

multiplication stage longer than eight weeks, medium nutrient and sugar depletion could

be limiting to the growth and multiplication of sea oats genotypes and induce leaf

senescence.

During Stage III culture, shoot and root growth increased in both genotypes, but

clear differences in plantlet morphology were observed between genotypes. At week 6

Stage III, EK 16-3 plantlets had greater shoot but lower root biomass than EK 11-1.

While energy sources were directed mainly to root growth in EK 11-1, EK 16-3 plantlets

predominantly utilized their energy sources towards growth of elongated leaves with

expanded leaf blades. These differences in developmental patterns are important during









the transition from in vitro photomixotrophic or heterotrophic conditions to the ex vitro

photoautotrophic conditions. During this transition, plantlets must adjust from using

medium sucrose or their stored reserves to producing their own photoassimilates to

continue growth and development ex vitro (Piqueras et al., 1998). Although the

importance of photosynthetic capacity during acclimatization has been emphasized in

many studies, Van Huylenbroeck and Debergh (1996) concluded that the photosynthetic

ability at time of ex vitro transfer is of secondary importance; the primary requirement

are carbohydrate reserves large enough to overcome the transition to ex vitro conditions.

However, in sea oats, photosynthetic capacity is also critical during acclimatization since

in previous studies we observed low ex vitro survival of EK 11-1 plants that exhibited

high starch reserves at time of ex vitro transfer (Valero-Aracama et al., 2004b). These

starch reserves are rapidly depleted during the first 2 weeks of ex vitro acclimatization

(Chapter 3). Leaves in these plants were also short, thick, and without expanded blades,

and plants also exhibited large root systems.

One significant difference between the easy- and difficult-to-acclimatize genotype

is the ability of the former to produce leaves during Stage III of similar morphology and

anatomical features as ex vitro produced leaves (Valero-Aracama et al., 2003). These

features may be reflected in normal photosynthetic capacity when transferred ex vitro.

Optical light micrographs of leaves from EK 16-3 during Stage III revealed similar leaf

tissue organization between plantlets cultured for six weeks in Stage III and after being

acclimatized to greenhouse conditions. At weeks 6 and 9 Stage III, tissues were highly

organized in the easy-to-acclimatize genotype compared to the difficult-to-acclimatize

genotype. EK 11-1 leaf sections exhibited disorganized mesophyll tissue throughout









Stage III. This has also been reported by other authors (Brainerd et al., 1981; Donnelly

and Vidaver, 1984a; Johansson et al., 1992, Ticha and Kutik, 1992; Dami and Hughes,

1995; Noe and Bonini, 1996). Likewise, whole leaf morphology, especially expanded

blades, was similar between acclimatized plantlets and EK 16-3 plantlets cultured for six

or nine weeks in Stage III. Conversely, EK 11-1 plantlets produced small thin leaves

throughout Stage III culture. Several studies have indicated that in vitro produced leaves

of some species were both functionally and structurally anomalous (Sutter, 1981).

Leaves of cauliflower (Brassica oleracea L.) (Wardle et al., 1979), and apple (Malus

pumila Mill.) (Brainerd and Fuchigami, 1981) had reduced stomatal functioning as well

as reduced photosynthetic capacity (Grout and Aston, 1977) during in vitro culture.

These leaves would be more likely injured under the stress conditions characteristic

during the change from in vitro to ex vitro culture.

A decreased biomass would imply a loss of photosynthetic capacity during

acclimatization. Loss in shoot dry weight (from 81 mg to 71 mg) was observed when EK

11-1 plantlets were compared from week 6 Stage III (time 0 Stage IV) to week 4 Stage

IV. In contrast, EK 16-3 plantlets exhibited a 2.7-fold increase in shoot biomass (from 94

mg to 261 mg) for the same culture periods. After transfer ex vitro, a positive carbon

balance is required for plantlet acclimatization and continued growth (Grout and Ashton,

1978). A positive carbon balance was not attained in EK 11-1 plantlets. The anomalous

anatomy in EK 11-1 limited photosynthesis ex vitro resulting in decreased survival and

initially limiting growth of those that survived. Additionally, stored carbohydrate

reserves were likely depleted before plants could produce sufficient photoassimilates to

overcome the energy demands for growth (Chapter 3).









Although adventitious root formation was necessary to increase ex vitro survival of

both sea oats genotypes, excessive root biomass may have rendered EK 11-1 plants

disadvantaged during acclimatization ex vitro. In sea oats, large root systems are sources

of starch reserves and carbohydrates accumulated in vitro (Valero-Aracama et al.,

2004b). However, being composed of heterotrophic tissue, the root often has high energy

demands.

In vitro produced roots of sea oats genotypes differed significantly in architecture

and morphology. Several investigations have indicated that adventitious roots of various

plant species formed in vitro display particular anatomical and morphological features

induced by the physical characteristics of the gelled culture medium (Mohammed and

Vidaver, 1988; McClelland and Smith, 1988; McClelland et al., 1990). Therefore, the

roots produced in vitro may promote growth in vitro but may not be functional after ex

vitro transfer (McClelland et al., 1990; Bonal and Monteuuis, 1997).

During the initial acclimatization period, the fraction of dry matter allocated to

roots has been reported to be small or unchanged (Fila et al., 1998). When comparing in

vitro and ex vitro root dry weights of plants cultured for six weeks in Stage III and four

weeks in Stage IV, we observed a 38% decrease in root biomass of EK 11-1 whereas

there was a 57% increase in that of EK 16-3 plantlets. The decrease ofEK 11-1 root

biomass indicated that those roots partially or totally died ex vitro and either a portion of

the roots were still functional or newly developed roots were formed. Conversely, the

increase of EK 16-3 root biomass ex vitro indicated that those roots formed in vitro were

possibly functional and continued growing after ex vitro transfer. Possibly, high energy









demands by roots, limited root functionality, and limited photosynthetic capacity of

shoots in EK 11-1 plants, contributed to death of roots.

These morphological and anatomical differences between sea oats genotypes may

influence photosynthetic capacity of plants. This should be verified by comparative

determination of photosynthetic capacity in vitro and ex vitro in both genotypes.

Additionally, the carbohydrate status of both genotypes during the same culture periods

should be investigated.

Conclusions

The abnormal anatomy and morphology of the difficult-to-acclimatize sea oats

genotype in vitro correlated with poor survival and acclimatization ex vitro. In vitro

produced sea oats plantlets require the formation of elongated leaves with expanded

lamina, and highly organized leaf tissues for successful acclimatization. These

anatomical and morphological characteristics possibly facilitate the transition from

photomixotrophic or heterotrophic conditions to photoautotrophic conditions by

improving the control of water loss and the photosynthetic capacity ex vitro.

Additionally, formation of roots in vitro appeared critical for survival ex vitro due to: 1)

increased water demands ex vitro, and 2) reduced energy demands to produce roots ex

vitro because this energy is supplied in vitro (Stage III). Furthermore, the Stage II

duration should be long enough to overcome the negative ex vitro carryover effect of BA.

Yet, to increase production efficiency, it would be beneficial to culture sea oats

genotypes for eight weeks rather than twelve weeks Stage II to limit leaf senescence and

to obtain higher shoot multiplication rate and quality of microcuttings in vitro.

During Stage III, sea oats plantlets require at least six weeks culture to produce

elongated shoots and roots. This stage is critical for the development of photosynthetic









competence and to accumulate carbohydrate reserves. The high survival observed in the

easy-to-acclimatize genotype indicated that its carbon balance was positive after

acclimatization. Conversely, the difficult-to-acclimatize genotype exhibited poor

development of shoots and an extensive heterotrophic root system that likely resulted in a

negative carbon balance ex vitro, thus leading to low survival.

Acknowledgements

This project was developed under the guidance of the Florida Sea Grant College

Program and funded from NOAA, Department of Commerce, Grant No. NA16RG-2195.

The authors would like to thank Ramon Littell for his statistical expertise and Karen

Kelley and Fred Bennett in the Electron Microscopy Core Laboratory at the UF, for their

guidance and assistance in histological studies.














CHAPTER 3
PHOTOSYNTHETIC AND CARBOHYDRATE STATUS OF EASY- AND
DIFFICULT-TO-ACCLIMATIZE SEA OATS (Uniolapaniculata L.) GENOTYPES
DURING IN VITRO CULTURE AND EX VITRO ACCLIMATIZATION

Introduction

Micropropagation has been extensively used for the rapid production of many plant

species and cultivars (Debergh and Zimmerman, 1991; Jeong et al., 1995; Hartmann et

al., 2002). However, despite its extraordinary potential, this technology is still

confronted with many problems. Among these, one of the most important is the poor

survival of plantlets following ex vitro transfer, during acclimatization to greenhouse or

field conditions (Pospisilova et al., 1999). This problem originates from poor

development of photosynthetic capacity in vitro, which has been attributed to the

presence of sugar in the medium (Kozai, 1991a; Pospisilova et al., 1992), low light and

inadequate CO2 supply (Kozai and Iwanami, 1988; De et al., 1993), and poor control of

water loss caused by high relative humidity within the vessel (Desjardins, 1995; Estrada-

Luna et al., 2001). These conditions can ultimately influence plant development and

photosynthetic performance (Kozai, 1991b; Preece and Sutter, 1991).

Kozai et al. (1997) demonstrated that most chlorophyllous plantlets/microcuttings

in vitro have photosynthetic ability provided that the environmental conditions are

favorable for photosynthesis. However, low ventilation rates, characteristic of

conventional culture vessels, limit CO2 availability during almost the entire photoperiod

(Kadle6ek et al., 2001). Consequently, plantlets commonly exhibit low net









photosynthetic rates caused by low CO2 concentrations in the vessels during the

photoperiod and low light intensities typical of culture rooms (Heo and Kozai, 1999).

In vitro culture conditions frequently result in alterations in mesophyll development

as well as chloroplast structure, namely grana development (Wetzstein and Sommer,

1982). At the biochemical level, low ribulose 1,5-bisphosphate carboxylase /oxygenase

(rubisco) activity (Grout, 1988) and high phosphoenolpyruvate carboxylase (PEPC)

activity (Triques et al, 1997) is often encountered in C3 species. These conditions also

contribute to low photosynthetic activity.

Acclimatization, the transition period from in vitro to ex vitro conditions, is critical

because this is when developmental and physiological abnormalities of plants produced

in vitro need to be corrected to ensure survival and continued plant growth (Debergh and

Zimmerman, 1991; Preece and Sutter, 1991). During acclimatization, in vitro cultured

plants must go through a transition from a heterotrophic or photomixotrophic mode to a

fully photoautotrophic mode in the greenhouse. It has been frequently reported that high

sugar concentration in the culture medium results in feedback inhibition of

photosynthesis in C3 plants (Hdider and Desjardins, 1994). Also, several investigations

indicate that high sugar concentration decreases rubisco activity (Hdider, 1994; Hdider

and Desjardins, 1994). The activity of this enzyme and the presence of starch granules in

chloroplasts, which is common in in-vitro cultured plantlets, are known to affect the

regeneration of ribulose 1,5-bisphosphate (RuBP), the substrate for rubisco (Desjardins,

1995). It has also been reported that low light levels, typical of in vitro culture, inhibit

rubisco activity by reduced activation of rubisco activase (Potris, 1992). During the









initial days of acclimatization, typically low rubisco activity steadily increases to levels

common in greenhouse-grown plants (Desjardins, 1990).

During initial stages of acclimatization, in vitro leaves of certain plant species serve

as carbohydrate storage organs to cover metabolic demands of growing tissues (Van

Huylenbroeck et al., 1998; Piqueras et al., 1998). However, the function of in vitro

produced leaves during acclimatization varies depending upon plant species. Van

Huylenbroeck et al. (1998) concluded that in some plant species such as Calathea

(Calathea loiusae Gagnep. 'Maui Queen'), in vitro formed leaves can function as storage

organs, whose energy reserves are consumed during the first days of acclimatization.

These types of leaves have limited photosynthetic ability. However, in other plant

species such as peace lily (Spathiphyllumfloribundum Schott. 'Petite'), in vitro leaves are

photosynthetically competent and function similarly to greenhouse-produced leaves.

Furthermore, cauliflower (Brassica oleracea L.) or strawberry (Fragaria x aananssa

Duch. 'Kent') leaves are net respirers, and their in vitro produced leaves senesce rapidly

after transplantation ex vitro (Grout and Aston, 1978; Grout and Millam, 1985).

Sea oats (Uniolapaniculata L.) is a perennial C4 grass, native to the southeastern

U.S., and commonly used for beach and dune restoration and stabilization (Wagner,

1964; Brown and Smith, 1974). This species is usually propagated by seed. However,

alternative vegetative propagation methods are necessary because of the limitation on the

natural sources of plants and seeds (Hester and Mendelssohn, 1987; Bachman and

Whitwell, 1995; Burgess et al., 2002; Burgess et al., 2005). Furthermore, concerns

regarding the use of unadapted ecotypes collected from distant locations have also limited

the collection of plant materials. Consequently, a micropropagation protocol was









developed to mass produce sea oats genotypes from localized sites (Philman and Kane,

1994). This protocol was defined using a single genotype. However, for the purpose of

maintaining genetic diversity, it is critical to be able to micropropagate a wide range of

sea oats genotypes. When this protocol was applied to multiple sea oats genotypes,

microcuttings of different genotypes transferred ex vitro displayed differing

acclimatization capacities.

Understanding the reasons for low acclimatization capacity of some sea oats

genotypes is needed to efficiently produce a wide range of sea oats genotypes using

micropropagation. The difference in acclimatization capacity between sea oats genotypes

could be the result of differing photosynthetic and carbohydrate status during in vitro

culture. Furthermore, the physiological changes occurring during the initial period of

acclimatization appear critical during ex vitro establishment. The objective of the present

study was to compare the photosynthetic characteristics, photosynthetic enzymatic

activity and carbohydrate status of in vitro and ex vitro sea oats genotypes to correlate

this with their differing capacity for acclimatization.

Materials and Methods

Culture Conditions

Established and indexed in vitro shoot cultures of two sea oats (Uniola paniculata

L.) genotypes, collected from Egmont Key, on the Florida Gulf coast, genotyped using

random amplified polymorphic DNA (RAPD) genetic analyses, and previously

characterized as easy- and difficult-to-acclimatize (EK 16-3 and EK 11-1, respectively)

were used. Five sea oats shoot clusters each consisting of three shoots, 25 mm long of

EK 16-3 and EK 11-1 genotypes, were subcultured into separate Magenta GA7 vessels

(Magenta Corp., Chicago, IL) containing 80 mL sterile Stage II medium. Stage II









medium consisted of Murashige and Skoog (MS) inorganic salts (Murashige and Skoog,

1962), supplemented with 87.6 mM sucrose, 0.56 mM myo-inositol, 1.2 ptM thiamine-

HC1, 2.2 [tM N6-benzyladenine (BA), and solidified with 8 g L-1 TCTM agar

(PhytoTechnology Laboratories, Shawnee Mission, KS). All media were adjusted to pH

5.7 with 0.1 N KOH prior to the addition of agar and autoclaving at 1.2 kg cm-2 and 121

C for 20 min.

Cultures were maintained for eight weeks in a growth chamber at 24 1 C, 58 +

5% relative humidity (RH), 16-h photoperiod provided by cool-white fluorescent lamps

(General Electric F20WT12-CW), and at 40 5 [tmol m-2 s-1 photosynthetic photon flux

(PPF) as measured at culture level. Subsequently, the shoot clusters of each genotype

were subdivided into single shoots and transferred to Stage III rooting medium.

Stage III rooting medium consisted of 80 mL sterile half-strength MS medium,

supplemented with 0.56 mM myo-inositol, 1.2 [tM thiamine-HC1, 87.6 mM sucrose, and

10 ptM ca-naphthalene acetic acid (NAA), contained in GA7 vessels. Culture vessels

contained eight single microcuttings each, and were maintained in a culture room at 22 +

2 C, under a 16-h photoperiod provided by cool-white fluorescent lamps (General

Electric F96T12-CW-WM), at 100 5 [[mol m-2 s-1 PPF, as measured at culture level.

After six weeks, rooted microcuttings were placed in 48-cell plug trays (8 six-

celled blocks, each cell 4 x 6 x 5.5 cm; T.O. Plastics, Inc., Clearwater, MN) containing

coarse vermiculite as supporting material and transferred to Stage IV conditions.

Plantlets were watered as needed, and Peters 20N-20P-20K liquid fertilizer (150 mg N

L-1; The Scotts Company, Marysville, OH) was applied weekly. Greenhouse set points









for cooling and heating were 24 and 22C, respectively, and natural solar PPF ranged

900-1200 [[mol m-2 s-1 at noon.

Photosynthesis Studies

Shoot clusters of the EK 11-1 and EK 16-3 sea oats genotypes were cultured for

eight weeks under Stage II conditions as previously described. Subsequently, single

microcuttings were transferred to Stage III conditions for six weeks, with 10 replicate

GA7 vessels per genotype. All rooted microcuttings were transferred in 48-cell plug

trays (6 eight-celled blocks, each cell 4 x 6 x 5.5 cm; Summit Plastics Inc., Clearwater,

OH) to a greenhouse under natural solar PPF ranging 900-1200 [tmol m-2 s-1 during the

measurements and day/night temperatures of 25/22 C, respectively.

Net photosynthetic rates per leaf area (Pni) were determined with a PP System

Model Ciras-1 (PP System Co., Ltd., UK) without a supplemental light source and inlet

CO2 concentration fixed at 400 + 10 [tmol mol-P. Measurements were taken in full sun

near midday (10 am to 12 pm) on newly formed with fully expanded leaves every week

beginning the day after establishment ex vitro. Percent survival was scored every week

during Stage IV. Two plants per plug tray were measured, compiling data from 20 plants

per genotype. Data were collected weekly for 7 consecutive weeks.

Photosynthesis Enzymatic Studies

Shoot clusters from EK 11-1 and EK 16-3 were cultured for 8 weeks in GA7

vessels under Stage II conditions as previously described. There were 4 replicate vessels

per genotype, each vessel containing 5 plants. Concurrently, another set of sea oats shoot

clusters were cultured for 8 weeks under the same conditions for shoot multiplication









prior to transfer to Stage III conditions for 3, 6 or 9 weeks. Each treatment in Stage III

consisted of 4 replicate vessels per genotype each containing 8 microcuttings.

Shoots of EK 11-1 and EK 16-3 cultures from Stage II and Stage III in vitro and

Stage IV ex vitro conditions were harvested 3 h after the beginning of photoperiod for

rubisco and PEPC analyses. Each replicate collected from in vitro conditions consisted

of all 5 shoot clusters per vessel from Stage II or all 8 clusters per vessel from Stage III,

which immediately were placed in liquid N2. During ex vitro Stage IV, each replicate

contained 3 plants, which were placed in liquid N2 in the greenhouse. Subsequently, all

replicates were ground to a fine powder in a mortar cooled with liquid N2 and placed in

plastic vials in a -80 C freezer before analyses.

For enzymatic extraction, powdered shoot tissue (-225 mg per sample) was placed

in a glass mortar with 1.8 mL extraction buffer containing 100 mM bicine (pH 8.0 at 25

C), 10 mM MgCl2, 0.1 mM EDTA, 5 mM dithiothreitol (DTT), 10 mM isoascorbate, 2%

PVP-40 (w/v) and 0.1% TX-100 (w/v). Leaf tissue was ground with extraction buffer on

ice for up to 2 min, transferred into 2 microfuge tubes, and centrifuged for 45 s. A 0.2

mL aliquot of the supernatant was incubated (on ice for a minimum of 5 min) with 0.01

mL 500 mM MgCl2 plus 0.011 mL 200 mM NaHCO3 to obtain fully-carbamylated

rubisco ("activated extract") prior to assay of rubisco activity. A separate aliquot of the

supernatant was held at room temperature for immediate assay for PEPC. Both enzymes

were assayed in total volumes of 0.5 mL at 25 C, in triplicate, and assays were

completed within 30 min from start of extractions procedure with a Hitachi Model U-

2000 Double-Beam UV/VIS Spectrophotometer (Hitachi Instruments, Inc., Danbury,

CT).









Phosphoenolpyruvate carboxylase was assayed spectrophotometrically at 340 nm

by following the reduction of OAA by NADH in the presence of excess malate

dehydrogenase [MDH] (Ashton et al., 1990). The reaction mixture contained 100 mM

bicine (pH 8.0), 10 mM MgCl2, 0.1 mM EDTA, 10 mM NaHCO3, 5 mM DTT, 2.5 units

MDH, and 0.2 mM NADH. After addition of 0.01 mL extract, a steady baseline was

established and the reaction was initiated by addition of PEP to a final concentration of 5

mM. The linear decrease in absorbance was recorded over a period of 150 s.

Total rubisco activity (fully activated rubisco) was assayed spectrophotometrically,

based on the method by Lilley and Walker (1974). This enzyme-linked assay couples the

activity of rubisco with the oxidation of NADH using 3-phosphoglyceric phosphokinase

and glyceraldehyde 3-phosphate dehydrogenase extracted from rabbit muscle (linking

enzymes). The linking enzymes were purchased as ammonium sulphate precipitates.

Sulphate was removed prior to use either by de-salting solubilized precipitates or by

dissolving the precipitates in 20% glycerol solution in buffer (Sharkey et al., 1991). The

reaction mixture for measuring rubisco activity contained 100 mM bicine (pH 8.0), 20

mM MgCl2, 1 mM EDTA, 20 mM NaC1, 10 mM NaHCO3, 5 mM DTT, 2.5 mM ATP, 5

mM phosphocreatine, 5 units creatine phosphokinase, 5 units each of the linking

enzymes, 0.2 mM NADH and 0.6 mM ribulose 1,5-bisphosphate. After a steady baseline

absorbance at 340 nm was established, the reaction was initiated with 0.011 mL of

activated extract. The linear decrease in absorbance resulting from oxidation of NADH

was recorded over a period of 150 s.

Chlorophyll content was determined using the method described by Arnon (1949).

From each tube containing the crude extract after grinding, an aliquot of 0.1 mL was









transferred to 2 microfuge tubes, each containing 0.1 mL water. Subsequently, 0.8 mL

100% acetone were added in each tube, and incubated in the dark for at least 30 min in

ice. After 3 min centrifuging, the supernatant was collected and absorbance was

measured at 645 and 663 nm for total chlorophyll determination. Total soluble protein

(TSP) in the extracts was quantified by the dye-binding assay of Bradford (1976) using

bovine serum albumin as standard.

Transmission Electron Micrograph Studies

Leaf histological cross sections from EK 11-1 and EK 16-3 microcuttings cultured

in vitro in Stage III were obtained to compare bundle sheath chloroplast ultrastructure.

Transmission electron microscopy (TEM) was used, for which leaf sections

approximately 2 mm from the center of the leaf blade were fixed in Trumps fixative

solution (McDowell and Trump, 1976). Fixative infiltration was achieved under vacuum

for 2 days. Leaf tissues were then rinsed 3 times in phosphate buffer (pH 7.2), post-fixed

in a 1% buffered osmium tetroxide solution and then rinsed in phosphate buffer, 3 times

in distilled water, and dehydrated in a five-step ascending ethyl alcohol series (25, 50, 75,

95, 100%) followed by dehydration in 100% acetone. An enbloc stain of 2% uranyl

acetate was applied between the 75 and 95% steps of the ethyl alcohol dehydration series.

Leaf sections were then embedded in Spurr resin (Spurr, 1969). Ultrathin leaf sections

(70 nm) were cut from the center part of the leaf blade with a Leica Ultracut

ultramicrotome R (Leica Microscopy and Scientific Instruments, Deerfield, IL), collected

on 0.35% form-var coated copper grids, stained with methanolic uranyl acetate and lead

citrate (Reynolds, 1963). Sections were viewed on a Hitachi H7000 transmission

electron microscope (Hitachi Scientific Instruments, Danbury, CT) at 75 kV. Digital

micrographs were taken on a BioScan/Digital Micrograph 2.5 (Gatan Inc., Pleasanton,









CA) at an exposure level optimized for viewing the outer layer and processed with

MEGA View III/AnalySIS 3.1 (Soft Imaging System Corp., Lakewood, CO).

Carbohydrate Studies

Plantlets from EK 11-1 and EK 16-3 sea oats genotypes were cultured for 8 weeks

under Stage II conditions, with 20 replicate vessels per genotype, each containing 5 shoot

clusters. Subsequently, single shoots from each genotype were cultured under Stage III

conditions for 3, 6 or 9 weeks. Each Stage-III duration treatment consisted of 5 replicate

vessels per genotype, each containing 8 single shoot microcuttings.

At week 8 Stage II conditions, clusters of plantlets obtained from 5 vessels were

taken out of the medium, washed, and packaged in aluminum foil envelopes. Every

package contained 3 plantlets (sub-samples), which were collected from the same vessel.

Samples were frozen in liquid N2 and stored at -800C prior to drying in a 10-MR-TR

freeze drier (The Virtis Company, Gardiner, NY) for 7 days. Dry weights of each sub-

sample were then recorded. At weeks 3, 6 and 9 of Stage III conditions, all plantlets from

5 vessels were collected and processed in the same way, this time separating roots from

shoots.

A parallel study was completed to analyze the carbohydrate status of both

genotypes during Stage IV duration. Plantlets form EK 11-1 and EK 16-3 sea oats

genotypes were cultured for 8 weeks under Stage II conditions, followed by 6 weeks

Stage III conditions and then transferred to Stage IV conditions for 4 weeks. Five

replicate samples consisting of 3 shoots or roots per genotype were collected at weeks 2

and 4 Stage IV and processed as previously described.

Procedures for total soluble sugar extraction were modified following the

description by Boersig and Negm (1985) and Miller and Langhans (1989). Glass Pasteur









pipettes with glass wool plugs were loaded with 50 mg of each sample. Soluble sugars

were extracted with 1.5 mL of methanol:chloroform:water (MCW) (12:5:3 v:v:v)

overnight. One hundred ptL mannitol (10 mg mL-1) was added as an internal standard to

each sample. One hour extractions were repeated twice with 1.5 mL MCW followed by

two additional MCW (1.5 mL) rinses. Nanopure water (3.5 mL) was added to extract

prior to 20-min centrifugation at 4,000 rpm. The aqueous phase was removed and

applied to polyethylene columns containing 3 mL 1 methanol: water (v:v, MW) and

cation and anion exchange resin (1 mL Amberlite IRA-67 layered with 1 mL Dowex 50-

W, Sigma-Aldrich Co., St. Louis, MO). Soluble sugars were eluted and rinsed twice with

methanol: water (MW) (1:1 v:v) prior to complete evaporation using a RapidVap vacuum

evaporator system (Labconco Corp., Kansas City, MO). The dry residue was re-

suspended in 1 mL HPLC-grade water and filtered through a 0.45 [t membrane prior to

HPLC injection. Total soluble sugars (TSS; sucrose, glucose and fructose) were

analyzed using a Waters 2695 High Pressure Liquid Chromatograph (Waters

Technological Corporation, Milford, MA) equipped with a Waters 2414 refractive index

detector (Waters Technological Corporation, Milford, MA) and two connected BioRad

Aminex HPX-87C columns (BioRad Laboratories, Hercules, CA). Column and detector

temperatures were maintained at 80 and 50 C, respectively. High pressure liquid

chromatography grade water was used as the mobile phase, at a flow rate of 0.6 mL

min

For starch analysis, procedures were modified as described by Haissig and Dickson

(1979) and Miller and Langhans (1989). The tissue residue (remaining in pipets after

soluble sugar extraction) was oven dried overnight at 50 oC, suspended in 4 mL Na-









acetate buffer (100 mM, pH 4.5), and placed in a boiling water bath (90 C) for 30 min.

After cooling to room temperature, 1.0 mL amyloglucosidase solution (from Rhizopus

mold, Sigma-Aldrich Co., St. Louis, MO) (50 units/assay in 0.1 M pH 4.5 Na-acetate

buffer) was added to each test tube to hydrolyze starch to glucose. Samples were

incubated for 48 h at 55 C with occasional agitation. An aliquot (100-[tL) of each

sample (glucose hydrolyzate) was transferred to a clean test tube and subjected to an

enzyme assay containing glucose oxidase (5 units mL-1) and peroxidase (200 units mL-1).

After the addition of 2.2 N HC1 (1.0 mL), absorbance at 450 nm was determined using a

Bechman DU-64 spectrophotometer (Beckman Coulter Inc., Fullerton, CA) and starch

content was calculated based on the regression equation of the glucose calibration line

(0.0 to 1.0 [tmol).

For ex vitro analyses, two replicate sub-samples were taken from each tissue

sample per genotype. These sub-samples were averaged to estimate starch and total

soluble sugar contents.

Experimental Designs and Statistical Analyses

All experiments were arranged in completely randomized designs. For

photosynthesis measurements, each plant per genotype was considered a replication. For

carbohydrate and enzymatic analyses all shoots or roots obtained from each vessel were

considered a replication. Main treatment effects and interactions were evaluated using

the general linear model (GLM) procedures developed by Statistical Analysis System

(SAS Institute Inc., 1999) and mean separation was evaluated using Waller-Duncan at P

< 0.05.









Results

Photosynthetic and Transpiration Status Ex Vitro

The ability of plantlets to acclimatize to ex vitro conditions differed significantly

between genotypes. EK 16-3 plants exhibited nearly 100% survival after 6 weeks,

whereas EK 11-1 plants started senescing after 2 weeks acclimatization, resulting in 29%

survival after 6 weeks ex vitro culture (Figure 3-1A). Significant differences in Pni were

also observed during initial plant growth ex vitro, with Pn1 values being greater for EK

16-3 than for EK 11-1 plants (Figure 3-1B). At week 0, ex vitro Pni for EK 11-1 was 1.9

[tmol m-2 s-1, which was significantly lower than Pni of EK 16-3 plants. Surviving EK 11-

1 plants exhibited a significant increase ofPni over time that was not significantly

different to the Pni of EK 16-3 plants after 5 weeks ex vitro culture. Transpiration rates of

plants at initial ex vitro transfer were significantly higher for EK 11-1 than EK 16-3

plantlets. After one week ex vitro culture, transpiration rates in both genotypes increased

until after week 3 when they decreased (Figure 3-1C).

Carbohydrate Status in Vitro and Ex Vitro

During weeks 3 to 9 of Stage III, starch content in shoots was greater in EK 11-1

than EK 16-3 plantlets and both genotypes exhibited a steady decrease of shoot starch

with time (Figure 3-2A). After 6 weeks in Stage III, both genotypes exhibited a

remarkable decrease in starch content when transferred ex vitro to the greenhouse. By

week 4 Stage IV, shoot starch content of EK 11-1 plantlets was 3.8 times lower than that

of EK 16-3 plantlets. Transmission electron micrographs (Figure 3-3) of bundle sheath

cell chloroplasts indicated differences in the size and distribution of starch grain

chloroplasts with time and between genotypes. While large starch granules were found

after 3 weeks in Stage III, smaller starch grains were observed after 6 and 9 weeks







81


100 -
A
-0- EK 11-1
80 -0- EK16-3


60


40


20 -


20
B
25

20




10 -

5

0




8 8









2


0
0 1 2 3 4 5 6
Time in Stage IV (weeks)



Figure 3-1. Effect of in vitro culture conditions on A: ex vitro survival B: ex vitro
transpiration rate per leaf area, and C: ex vitro net photosynthetic rate per leaf
area (Pni) of EK 11-1 and EK 16-3 genotypes during Stage IV culture. Means
+ SE are shown (n = 10).









Stage III. Chloroplasts in EK 11-1 plantlets after 3 weeks Stage III (Figure 3-3A)

exhibited larger numbers of plastoglobuli than in EK 16-3 (Figure 3-3D). Furthermore,

at the same time, thylakoid membranes ofEK 11-1 chloroplasts were separated and

appeared disrupted (Figure 3-3A).

At 6 weeks Stage III, root starch content (Figure 3-2B) was lower in both

genotypes compared to shoot starch content (Figure 3-2A), and it was not significantly

different between genotypes at 6 and 9 weeks Stage III culture. Root starch content also

decreased from 6 to 9 weeks under Stage III culture conditions. Throughout Stage IV

acclimatization, root starch content was similar among genotypes. At 2 weeks Stage IV,

root starch content was nearly depleted (13.2 and 6.5 times lower than that of EK 11-1

and EK 16-3, respectively, at week 0).

During Stage III, shoot soluble sugars (sucrose and hexose) increased from week 0

to 3 and then gradually decreased from weeks 3 to 9 (Figure 3-4 A-C). At 0 weeks Stage

III (after 8 weeks in Stage II), there was significantly higher shoot sucrose content per dry

weight in EK 16-3 than in EK 11-1 plantlets, whereas hexose and TSS were non

significantly different between genotypes. Although shoot sucrose, hexose and TSS were

similar among genotypes at 3 and 9 weeks Stage III, at 6 weeks, sucrose content was 21%

greater in EK 11-1 shoots than in EK 16-3 shoots. During Stage IV (after 6 weeks in

Stage III), shoot soluble sugars decreased from week 0 to 2, and then remained steady

from weeks 2 to 4 (Figure 3-4 A-C). Furthermore, EK 16-3 plants exhibited significantly

higher hexose and TSS contents than EK 11-1 plants after 2 and 4 weeks ex vitro.

During Stage III, root soluble sugars (sucrose and hexose) decreased similarly

among genotypes between weeks 6 and 9 (Figure 3-5 A-C). Sucrose, hexose and TSS










contents in roots were significantly higher for EK 11-1 than EK16-3 plantlets, and

decreased after 9 weeks Stage III in both genotypes (Figure 3-5A-C). During Stage IV

(after 6 weeks in Stage III) root sucrose and hexose of both genotypes dramatically

decreased within two weeks, being nearly depleted during acclimatization.


~aoT


0 3 6 9 0 2 4
Time in Stage III (weeks) Time in Stage IV (weeks)


Figure 3-2. Comparative A: shoot starch content and B: root starch content of EK 11-1
and EK 16-3 genotypes during in vitro Stage III (left) and after microcuttings
were rooted for 6 weeks Stage III and transferred to Stage IV (right). Error
bars indicate SE (n = 10). ANOVA analysis is shown on top left corner of
each graph; T: Time, G: Genotype, NS, **: Non-significant or significant at P
< 0.01, respectively. Different letters on top of histobars within each culture
stage are significantly different according to Waller-Duncan test at P < 0.05.










































3 weeks 6 weeks 9 weeks


Figure 3-3. Comparative TEM of chloroplasts of A-C: EK 11-1, and D-F EK 16-3
genotypes after 3 weeks (A, D), 6 weeks (B, E) and 9 weeks (C, F) Stage III
culture conditions in bundle sheath cells. Scale = 2 |tm. (p: plastoglobuli, s:
starch granule, t: thylakoid membranes).