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Control of Plant Cell Shape by the Irregular Trichome Branch Genes in Arabidopsis

Permanent Link: http://ufdc.ufl.edu/UFE0022058/00001

Material Information

Title: Control of Plant Cell Shape by the Irregular Trichome Branch Genes in Arabidopsis
Physical Description: 1 online resource (178 p.)
Language: english
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2008

Subjects

Subjects / Keywords: actin, arabidopsis, cell, cytoskeleton, plant, shape, trichome
Plant Molecular and Cellular Biology -- Dissertations, Academic -- UF
Genre: Plant Molecular and Cellular Biology thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: The control of plant cell shape is fundamentally important not only for the function of individual cells, but also for the morphogenesis of whole plants. The Arabidopsis leaf trichome is used as a cell model for genetic screens of mutations, called irregular trichome branch (itb) and disproportionate (dpp), which cause changes in trichome shape. Five genes (ITB1-ITB4 and DPP) were cloned through a positional cloning strategy and the functions of these genes were characterized in this study. ITB1 is a plant homolog of the actin-related protein2/3 complex activator Scar/WAVE, which regulates actin and microtubule organization. Disruption of ITB1 causes disorganization of actin filaments and microtubules, generating distorted trichomes. ITB2 is a putative member of the aminophospholipid translocase (ALA) family. Mutations in this gene result in defective trichomes with reduced branch length. ITB3 is a plant-specific protein that regulates actin organization through interaction with actin depolymerizing factor (ADF). The absence of ITB3 severely changes actin cytoskeleton organization by forming actin rings, but no change was observed in microtubule organization. The trichomes in itb3 mutants are reduced in size and branch length. ITB4 is the plant homolog of cleavage stimulation factor 64 that influences not only trichome morphogenesis, but also floral development. Compared to wild type, mutations in ITB4 reduce the trichome branch number and increase sepal and petal numbers. DPP is a keto acyl reductase and is involved in trichome cell expansion. Mutations homozygous for dpp are lethal. At the restrictive temperature (22 degrees C), heterozygous plants of the dpp mutants display trichomes with reduced branch length and increased stalk length. Although the five gene products described above have different functions in plant cells, their mutations all cause changes in Arabidopsis leaf trichome shape. These results indicate that plant cell shape can be controlled by different genes with a wide range of functions on multiple dynamic cell processes such as cytoskeleton dynamics and endomembrane dynamics.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Thesis: Thesis (Ph.D.)--University of Florida, 2008.
Local: Adviser: Oppenheimer, David.

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2008
System ID: UFE0022058:00001

Permanent Link: http://ufdc.ufl.edu/UFE0022058/00001

Material Information

Title: Control of Plant Cell Shape by the Irregular Trichome Branch Genes in Arabidopsis
Physical Description: 1 online resource (178 p.)
Language: english
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2008

Subjects

Subjects / Keywords: actin, arabidopsis, cell, cytoskeleton, plant, shape, trichome
Plant Molecular and Cellular Biology -- Dissertations, Academic -- UF
Genre: Plant Molecular and Cellular Biology thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: The control of plant cell shape is fundamentally important not only for the function of individual cells, but also for the morphogenesis of whole plants. The Arabidopsis leaf trichome is used as a cell model for genetic screens of mutations, called irregular trichome branch (itb) and disproportionate (dpp), which cause changes in trichome shape. Five genes (ITB1-ITB4 and DPP) were cloned through a positional cloning strategy and the functions of these genes were characterized in this study. ITB1 is a plant homolog of the actin-related protein2/3 complex activator Scar/WAVE, which regulates actin and microtubule organization. Disruption of ITB1 causes disorganization of actin filaments and microtubules, generating distorted trichomes. ITB2 is a putative member of the aminophospholipid translocase (ALA) family. Mutations in this gene result in defective trichomes with reduced branch length. ITB3 is a plant-specific protein that regulates actin organization through interaction with actin depolymerizing factor (ADF). The absence of ITB3 severely changes actin cytoskeleton organization by forming actin rings, but no change was observed in microtubule organization. The trichomes in itb3 mutants are reduced in size and branch length. ITB4 is the plant homolog of cleavage stimulation factor 64 that influences not only trichome morphogenesis, but also floral development. Compared to wild type, mutations in ITB4 reduce the trichome branch number and increase sepal and petal numbers. DPP is a keto acyl reductase and is involved in trichome cell expansion. Mutations homozygous for dpp are lethal. At the restrictive temperature (22 degrees C), heterozygous plants of the dpp mutants display trichomes with reduced branch length and increased stalk length. Although the five gene products described above have different functions in plant cells, their mutations all cause changes in Arabidopsis leaf trichome shape. These results indicate that plant cell shape can be controlled by different genes with a wide range of functions on multiple dynamic cell processes such as cytoskeleton dynamics and endomembrane dynamics.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Thesis: Thesis (Ph.D.)--University of Florida, 2008.
Local: Adviser: Oppenheimer, David.

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2008
System ID: UFE0022058:00001


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73642510e8992ffa17ac9b3cae484c1e
d78ad1f4e1ec36cf51981b13c209681923396f8e







CONTROL OF PLANT CELL SHAPE BY IRREGULAR TRICHOME BRANCH GENES IN
ARABIDOPSIS























By

XIAOGUO ZHANG


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

2008



























O 2008 Xiaoguo Zhang


































To my great Mom









ACKNOWLEDGMENTS

I offer my sincere gratitude to my advisor and committee chair, Dr. David G.

Oppenheimer, for his guidance and encouragement. I especially acknowledge him for improving

my knowledge in plant developmental genetics, expanding my thinking on scientific questions,

and teaching me English science writing during my doctoral education. I would like also express

my great appreciation to our research coordinator, Paris Grey, for her strong support of my

research.

I am truly grateful to my supervisory committee, Drs. Alice C. Harmon, Bernard A.

Hauser, and Daniel L. Purich, for their valuable advice and inspiring discussions throughout my

research. I also thank Drs. Wenyun Song and Xiaodong Ding for training me in yeast two-

hybridization in their laboratory. I would also like to thank Drs. Kevin M. Folta and Amit

Dhingra for the use of the Gene Gun and help with the transient assays. I also thank the graduate

students in the Oppenheimer laboratory, Stacey Jeffries and Meredith Sullivan, for their support

and friendship during my education. I want to take this opportunity to thank all the faculty

members who taught me classes and all the people in PMCB who helped me during the past

three years. Finally I give special thanks to my wife, Qingping Yang, my son, Yuxiang Zhang,

and my daughter, Aiwen Zhang, as well as my parents-in-law, Xueren Yang and Dongying

Xiong.












TABLE OF CONTENTS


page

ACKNOWLEDGMENT S .............. ...............4.....


LI ST OF T ABLE S ................. ...............9................


LIST OF FIGURES .............. ...............10....


AB S TRAC T ............._. .......... ..............._ 12...


CHAPTER


1 LITERATURE REVIEW: PLANT CELL EXPANSION .............. ...............14....


Introducti on .........._....._.... ....._ __ ....._ __ .............1

Significance of Cell Expansion .........._..._. ........ ...............14....
General Characterization of Cell Expansion ..........._.. ......... ........._.. .......1
Cell Wall Dynamics and Cell Expansion .............. ...............15....
Cell W all Components .............. ...............16....
Cell Wall Synthesis and Cell Expansion .........._....__......___ ...._.._ ...........1
Plasma Membrane Dynamics and Cell Expansion .........._..__ ......._.. ........_._........2
Components of the Plasma Membrane ..........._._ ...._.... ........._._ ....... 2
Asymmetry of Plant Plasma Membranes ..........._.__....._.._........._ ...........2
Plasma Membrane Assembly and Cell Expansion ........_.............._. .........._....__.25
Endomembrane Dynamics and Cell Expansion .............. ...............28....
Endomembrane Trafficking Pathways .............. ...............29....
Vesicle Dynamics ................. ...............30.................
Vesicle Fission M achinery .............. ...............31....
Vesicle Fusion M achinery ................. .......... ...............3.. 1....
Vesicle Trafficking and Cell Expansion ................. ...............32........... ...
Cytoskeleton Dynamics and Cell Expansion ................. ...............35........... ...
The Microtubule Cytoskeleton and Cell Expansion............... ...............3
Actin Cytoskeleton Dynamics and Cell Expansion............... ...............4


2 IRREGULAR TRICHOME BRANCH 3 (ITB3) IS A NOVEL REGULATOR OF
ACTINT ORGANIZATION............... ..............4


Introducti on ................. ...............46.................
Materials and Methods .............. .. ...... ...............4
Plant Materials and Growth Conditions .............. ...............49....
Positional Cloning oflITB3 ................. ...............49...___ .....
Plasmid Construction................ .............5
RNA Extraction and RT-PCR ........._.___..... .__. ...............51...
Plant Transformation ........._.___..... .___ ...............51.....
Yeast Two-hybrid Assays............... ...............52.
Protein Isolation............... ...............5












Pull-down Assay ................. ...............53.................
M orphological Analysis .............. ...................... ...............5
Immunostaining of the Actin and Microtubule Cytoskeletons. ................. .............. .54
M icroscopy ............... ....._ ...............54....
Double Mutant Construction .............. ...............54....
R e sults.................. ......... .. ......... .............5
Cloning of the ITB3 Gene ................. ...............54........... ...
ITB3 is a Plant-specific Gene .................. ... ...... ..........5
ITB3 Over-Expression Did Not Generate Novel Phenotypes .............. ....................5
ITB3 Has No Specific Subcellular Location ................. ...............56...............
ITB 3 Interacts With ADF3 in Yeast ................. ...............57........... ..
ITB3 Directly Binds with ADF3 in Vitro................. ........... ..... .. .............5
The Trichomes are Defective in the Mutants of adJ itb31-4 and Their Double
M utants .............. ...............57....
D discussion ........._...... .... ......._.......... .... .._ ... ... ......... .. .......5
Disruption of Actin Cytoskeleton Organization Leads to Misshapen Trichomes ...........58
The Precise Role of the Actin Cytoskeleton in Trichome Morphogenesis ................... ..59
Actin Filament Reorganization Is Required for Cell Expansion ................. ................ .60
ITB3 is a Plant-Specific Regulator of Actin Organization ................. ............. .......60
Future Perspectives............... ..............6


3 IRREGULAR TRICHOME BRANCH 2 (ITB2) IS A PUTATIVE
AMINOPHOSPHOLIPID TRANSLOCASE THAT REGULATES TRICHOME
BRANCH ELONGATION IN ARABIDOPSIS .............. ...............74....


Introducti on .................. ...............74........_ .....
Materials and Methods .............. .. ....... ...............7
Plant Materials and Growth Conditions .............. ...............76....
Positional Cloning of ITB2 ................. ...............77....... .....
Plasmid Construction................ .............7
RNA Extraction and RT-PCR .............. ...............78....
Plant Transformation ................. ...............79.____.......
Re sults.................. ..... ..__... .......__ ............7
Characterization of the itb2 Mutants ................. ...............79........... ...
Cloning of the ITB2 Gene. .................. .. .......... .......... .... ...... .......7
Complementation of the itb2 Mutant and Over-expression of the ITB2 Gene ................81
Discussion ................. ...............83._ ___.......
Future Perspectives............... ..............8


4 DISPROPORTIONATE (DPP) ENCODES A KETOACYL REDUCTASE INVOLVED
IN TRICHOME CELL EXPANSION............... ...............9


Introducti on ................. ...............92.................
Materials and Methods .............. .. ....... ...............9
Plant Materials and Growth Conditions .............. ...............95....
Positional Cloning .............. ...............95....












Plasmid Construction................ .............9
Plant Transformation ................. ...............97.................
R e sults............... ... ......... ..... .. ........... .............9
Characterization of dpp Mutants .............. ...............97....
Positional Cloning of DPP ................. ...............98___.....
Identification ofDPP .............. ...............99....
Discussion ............... ... .. .. ... .... ... ............10
DPP Encodes a P-ketoacyl Reductase .............. .... ......... ..............10
DPP Has Pleiotropic Functions in Cell Expansion and Wax Synthesis .......................101
DPP is Vital for Plant Viability ................. ...............103..............
DPP is Likely to be DEADHEAD ........._._.. ...._ ...............104
Future Perspectives............... .............10

5 IRREGULAR 7RICHOM~EBRANCH 4 INT ARABIDOPSIS ENCODES THE PLANT
HOMOLOG OF THE 64 KDA SUBUNIT OF CLEAVAGE STIMULATION
FACTOR AND REGULATES TRICHOME MORPHOGENESIS AND FLORAL
DEVELOP MENT ............ ..... ._ ...............122...


Introducti on ............ ..... .._ ...............122...
Materials and Methods .............. .. ....... ..............12
Plant Materials and Growth Conditions .............. ...............124....
The itb4-2 Mutant Isolation and ITB4 Cloning ................. .....____.............. ....125
Trans gene Construction................ ............12
RNA Extraction and RT-PCR ............_...... .__ ...............127..
Plant and Yeast Transformations................ ...........12

M orphological Analysis .............. ...............128....
M icroscopy ................. ...............128......... ......
in situ Hybridization ................. ...............129._._.. ......
Re sults........._...... ...... ..._.._ ... .... ._. ............12
ITB4 Encodes AtC stF-64 ............... .. ......_. .... ........ ...............12
Loss-of-function Mutations in ITB4 Cause Aberrant Development of Trichomes
and Flowers .........._.... .. .... .._ __ .. ......_._.. .... ...............13
ITB4 is Highly Expressed in Growing and Proliferating Cells ............... .....................134
Loss of ITB4 Function Alters the Expression Pattern of Perianth Organ Identity
G enes...................... ..... ... ... .....................13
ITB4 Localizes to Nuclei, but Does Not Functionally Complement Tts Homolog in
Y east .............. ...............135....
D discussion ............... ........ .. .. ........ ..... .. .... .. .. ......... 3
ITB4 Plays a Crucial Role in Trichome Morphogenesis and Floral Development.......1 38
Loss of AtCstF-64 Function Influences the Expression of Multiple Genes that
Control Floral Organ Development .............. ............__ .... ........ .............4
Differences Exist in the Mechanism of mRNA 3' End Formation among Plants,
Yeast and Mammals ................. ...............140...............
Future Perspectives............... .............14











LIST OF REFERENCES ................. ...............155................

BIOGRAPHICAL SKETCH ................. ...............178......... ......


































































8










LIST OF TABLES


Table page

2-1 Primers used in this study .............. ...............63....

3-1 Segregation of the mutant plants in F2 with different genetic background ................... ..877

3-2 Trichome shapes of the transgenic plants ................ ................ ...................878

4-1 Primers sequence used in this study............... ...............107.

4-2 Segregation of trichome phenotypes in F2 of the dpp mutant crossed to wild-type
plants ................. ...............108................

4-3 Segregation of trichome phenotypes in Fl of the dpp mutant reciprocally crossed to
wild-type plants............... ...............108

4-4 Single nucleotide polymorphism identified between the Ler and RLD ecotypes ...........108

4-5 Segregation of phenotypes in the Fl of the dpp mutant crossed to the Salk lines...........109

5-1 Primers used in this study ................. ...............144..............

5-2 Alteration of trichome cell shape in the itb4-2 mutant ................. .........................145










LIST OF FIGURES


Figure page

2-1 Positional cloning of ITB3. ............. ...............64.....

2-2 Actin cytoskeleton is disorganized in the itb3 mutant. ................... ...............6

2-3 Actin cable organization in the stalk of trichomes ......... ................. ........._.._. ...66

2-4 Actin rings in the itb3 mutant .............. ...............67....

2-5 Phylogenic tree of the Arabidopsis ITB3 family members ................. ............ .........68

2-6 Alignment of ITB3 protein sequence with its homologs in other plants .................. .........69

2-7 ITB3-GFP is not specifically localized to any subcellular structure in transformed
onion epidermal cells ................. ...............70.._. ......

2-8 Yeast two-hybrid screen for ITB3 interactors. ............. ...............71.....

2-9 ITB3 directly interacts with ADF. ............. ...............72.....

2-10 Trichome shapes are defective in adf3 and itb31-4 mutants .............. ....................7

3-1 Defects in leaf trichome and cotyledon shape of itb2 mutants............_._ .........._._ ....88

3-2 Positional cloning and gene structure of ITB2. .............. ...............89....

3-3 Mutations and corrections of ITB2 cDNA. .............. ...............90....

3-4 Transgenic plants with ITB2 cDNA ................. ...............91...............

4-1 The dpp mutant trichomes in the RLD genetic background............._._ ........_._......110

4-2 Positional cloning of DPP ........... ......__ ...............111.

4-3 Single nucleotide polymorphism between RLD and Ler wild types ........._..... .............112

4-4 Equivocal sequencing result using the DNA template from plants heretozygous for
the dpp mutation ................. ...............112...............

4-5 Sequencing result of Atlg67730 using the dpp mutant DNA as a template with the
forward prim er ................. ...............113......... ......

4-6 Sequencing result of Atlg67730 using the dpp mutant DNA as a template with the
reverse prim er ................. ...............113......... ......

4-7 Schematic explanation of DPP identification ....._ .....___ ............._........14











4-8 Schematic explanation of deletion identification in dpp mutants ................. ................1 15

4-9 Novel phenotypes in the Fl of dpp mutants and T-DNA insertion lines. ................... .....1 16

4-10 Equivocal sequencing result of Atlg67730 .......___ ............ ......_.._.........17

4-11 Unequivocal sequencing result of Atl1g67730O................. ...............117........... .

4-12 Identification of DPP ..........._...... ._ ...............118..

4-13 Unequivocal sequence result of Atlg67730 using the DNA from ded plants of Fl .......1 19

4-14 Unequivocal sequencing result of Atlg67730 using the DNA from the wild-type
plants of Fl ........._ ............ ...............119...

4-15 GC deletion in dpp cloned into pBluescript SK............... ...............120..

4-16 Wild-type DPP cloned into pBluescript SK............... ...............120..

4-17 Transgenic plants with the mutated DPP in distinct genetic backgrounds......................121

5-1 Positional cloning of ITB4 .........____...... ..... ...............146..

5-2 Rescue of the itb4-1 zwi-3 double mutant phenotype by Atlg71800 ................... ...........147

5-3 The itb4-2 mutants display the trichome shape defects ....._____ ... .....___ ..............148

5-4 Floral defects of itb4-2 mutants ................. ...............149____ ....

5-5 Increased number of floral organs itb4-2 mutants ................ .......___ .........__ ..150

5-6 Leaf shape and color defects in itb4-2 mutants .............. ...............151....

5-7 ITB2 expression pattern in Col wild type .............. ...............152....

5-8 Altered expression patterns of floral organ identity genes in itb4-2 mutants ..................1 53

5-9 ITB4 is localized to the nucleus ...._.. ................. ............... .....15









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

CONTROL OF PLANT CELL SHAPE BY IRREGULAR TRICHOME BRANCH GENES IN
ARABIDOPSIS

By

Xiaoguo Zhang

May 2008

Chair: David G. Oppenheimer
Major: Plant Molecular and Cellular Biology

The control of plant cell shape is fundamentally important not only for the function of

individual cells, but also for the morphogenesis of whole plants. The Arabidopsis leaf trichome is

used as a cell model for genetic screens of mutations, called irregular trichome branch (itb) and

disproportionate (dpp), which cause changes in trichome shape. Five genes (ITBl-ITB4 and

DPP) were cloned through a positional cloning strategy and the functions of these genes were

characterized in this study.

ITB 1 is a plant homolog of the actin-related protein2/3 complex activator Scar/WAVE,

which regulates actin and microtubule organization. Disruption of ITB 1 causes disorganization

of actin filaments and microtubules, generating distorted trichomes. ITB2 is a putative member

of the aminophospholipid translocase (ALA) family. Mutations in this gene result in defective

trichomes with reduced branch length. ITB3 is a plant-specific protein that regulates actin

organization through interaction with actin depolymerizing factor (ADF). The absence of ITB3

severely changes actin cytoskeleton organization by forming actin rings, but no change was

observed in microtubule organization. The trichomes in itb3 mutants are reduced in size and

branch length. ITB4 is the plant homolog of cleavage stimulation factor 64 that influences not

only trichome morphogenesis, but also floral development. Compared to wild type, mutations in









ITB4 reduce the trichome branch number and increase sepal and petal numbers. DPP is a keto

acyl reductase and is involved in trichome cell expansion. Mutations homozygous for dpp are

lethal. At the restrictive temperature (22o C), heterozygous plants of the dpp mutants display

trichomes with reduced branch length and increased stalk length. Although the five gene

products described above have different functions in plant cells, their mutations all cause

changes in Arabidopsis leaf trichome shape. These results indicate that plant cell shape can be

controlled by different genes with a wide range of functions on multiple dynamic cell processes

such as cytoskeleton dynamics and endomembrane dynamics.









CHAPTER 1
LITERATURE REVIEW: PLANT CELL EXPANSION

Introduction

Significance of Cell Expansion

The intricate coordination of cell division and expansion allows plants to achieve a unique

developmental plasticity that reduces the constraints of various adverse environments for plant

survival. Plants need continued availability of light, water, and nutrients throughout their life

cycle. However, plants are stationary and plant cells are surrounded by rigid cell walls, which

make plants unable to escape resource-depleted environments. Therefore, plants adjust the rate

and direction of cell division and expansion. Rigid cell walls offer structural and mechanical

support for plant bodies, and ultimately are responsible for the plant architectural design and

morphology. Thus, plant cell expansion is the basis for both the whole plant morphogenesis and

plant flexibility to adapt environmental conditions (Thompson, 1917).

General Characterization of Cell Expansion

An expanding plant cell is similar to an inflating balloon in direction determination, force

requirement, and wall alteration. Balloons can form diverse shapes, if specific locations of the

balloon surface are forced by counter-pressure to change conformation. So can plant cells, too,

expanding in directions that follow their functional requirement. The driving force of cell

expansion comes from the internal turgor pressure, generated by the water content of the

protoplast. During cell expansion, water needs to enter into the protoplast across the semi-

permeable plasma membrane to keep a constant turgor pressure (Steudle and Zimmermann,

1977). Second, the direction of cell expansion is spatially determined by the orientation of

cellulose microfibrils, which in turn is controlled by the plant cytoskeleton. The plant

cytoskeleton is composed of microtubules and microfilaments. It is generally believed that









microtubules orient the deposition of cellulose microfibrils in cell walls, and microfilaments

serve as tracks for transporting vesicles to specific sites of the wall that are loosened by lytic

enzymes or wall-remodeling proteins. Through exocytosis, dynamic vesicles, which contain

polysaccharides and proteins from endomembrane compartments, are fused with the plasma

membrane, and their contents are discharged into the extracellular matrix space for building cell

walls (Murphy et al., 2005; Samaj et al., 2005; Johansen et al., 2006). Even though the volume of

an expanding cell increases to a thousand times the original volume, the walls maintain a

relatively constant thickness. All cell expansion is mechanistically divided into two basic types

of growth: tip growth and diffuse growth. For tip growth, a spatially focused cell expansion, cells

grow exclusively at the extreme tip of the cell. Pollen tube and and root hairs are cell types that

expand using tip growth. For diffuse growth, expansion can occur at multiple locations and

encompass larger regions than occurs during tip growth (Baskin, 2005).

Cell Wall Dynamics and Cell Expansion

The plant cell wall may seem to be a paradoxical entity because it serves to both maintain

cell shape through constraint of cell expansion, yet it promotes development of cell morphology

by extension of itself. The wall provides rigidity to a plant cell, strength, and protection against

various mechanical stresses. It also limits the entry of large molecules and pathogens into cells.

The wall further creates a stable osmotic environment by preventing osmotic lysis and retaining

water. Finally, the wall determines plant cells myriad shapes. All these functions of the wall

underlie its action as a physical barrier, which constrains cell expansion. To solve the paradox,

plants have evolved a dynamic cell wall. Thereby, this wall is not only suited to flexible cell

expansion, but also participates in cell to cell and cell to nucleus communication through

signaling molecules and receptors in the wall (Darley et al., 2001).









Cell Wall Components

As mentioned above, cell walls allow plants to form various shapes. Thus, cell expansion

appears to be its wall extension. To understand cell expansion it is first necessary to know the

chemical composition of the wall. Because only primary walls can be remodeled for cell

expansion, my focus here is on those aspects that are germane to understanding cell expansion.

Secondary walls, which are formed after the cessation of cell expansion, are not covered in this

literature review.

Plant cell expansion requires coordination between maintenance of osmotic potential and

changes in wall properties, which are determined by wall components. Plant cell walls are highly

organized; the diverse components include cellulose, glycans, pectins, proteins, and aromatic

substances, as well as metal ions. Cellulose, a maj or component of plant cell walls, plays the

most important role in the wall architecture. Approximately 36 chains of 1,4 linked, P-D-glucose

associate through intermolecular hydrogen bonds to form a microfibril, which serves as a

structural scaffold to support other wall components, and forms a fundamental framework of the

wall architecture. The microfibrils in walls are paracrystalline and resistant to hydrolysis by

acids, bases and enzymes (Reiter, 2002; Somerville, 2006).

Glycans, a mixture of branched polysaccharides, form molecular backbones through the

linkage of P-D-hexosyl residues with the same bond as in cellulose. Distinct glycans interlock

microfibrils by hydrogen bonding. Glycans and pectins form the secondary network, which

strengthens the wall architecture. Xyloglucan (XyG), glucuronoarabinoxylans (GAX), mixed

linkage glucans (MLG), and (gluco) mannans are found to be the four main glycans in primary

cell walls. Because glycans have a property of random arrays and amorphous structures in the

wall they are readily hydrolyzed by dilute acids, bases, or myriad enzymes (Reiter, 2002).









Pectins are a family of heterogeneous polysaccharides that all contain 1,4-linked a-D-

galacturonic acid. Because of this residue, pectins provide cell walls a hydrated and charged

surface. Additionally, a pectin polymer has a multitude of branches, which makes cell walls

porous. These componential and structural features of pectins also allow cell walls to modulate

pH and ion balance. Homogalacturonan (HG) and rhamnogalacturonans (RG-I and RG-II) are

fundamental constituents of pectins in primary cell walls (Willats et al., 2001).

The maj ority of proteins in cell walls are glycoproteins that have oligosaccharide chains

covalently attach to particular moieties of polypeptides. Hydroxyproline-rich glycoproteins

(HRGPs, also called extensins), the arabinogalactan proteins (AGPs), and the proline-rich

proteins (PRPs) are the most abundant glycoproteins in plant cells. Although proteins are not a

maj or component of cell walls, they display a wide variety of structures and functions. These

proteins make the wall a dynamic entity throughout the cell's life. For example, expansion induces

the pH-dependent wall extension and stress relaxation in a characteristically unique manner

(Cosgrove et al., 2002). Wall-associated kinases (WAKs), which are covalently bound to pectin,

have the potential to provide a physical and signaling continuum between the cell wall and the

cytoplasm (Wagner and Kohorn, 2001). The absence of AGPs in cell walls causes aberrant cell

expansion, which forms numerous bulges in root epidermal cells (Willats and Knox, 1996; Ding

and Zhu, 1997).

Minerals appear in micro amounts in cell walls, but they also play a role in the wall

structure. If plants lack them, normal cell wall formation is disrupted. For example, Ca2+ links

distinct pectin polymers (RG-I) through an ionic bond (Catoire et al., 1998). Another example is

boron, which is predominantly associated with rhamnogalacturonan II (RG-II). Disruption of the









linkage between borate and RG-II affects plant growth, and borate deficiency compromises plant

cell expansion in growing tissues (O'Neill et al., 2004).

Cell Wall Synthesis and Cell Expansion

Biosynthesis of plant cell walls is intimately connected with cell expansion. While a cell is

increasing its volume, its wall becomes thinner and thinner. Finally, if the cell wall fails to

incorporate new materials, it bursts. Thus, wall biosynthesis is required for cell expansion. In

recent years, the study of the biosynthesis of wall components has made substantial progress

through biochemical, genetic, and genomic approaches. Here, the emphasis is on the genes that

are involved in primary wall synthesis and their mutations that cause disruption of cell expansion

and generate aberrant cell shape.

Cellulose is synthesized by cellulose synthase (CESA), which has been identified in most

plant species. In the Arabidopsis genome, ten genes code for CESA; in rice there are at least nine

genes (Keegstra and Walton, 2006), and poplar has 18 (Djerbi et al., 2005). All CESAs share

common structural features: eight transmembrane domains, two glycosyltransferase domains,

and several mi crotubul e-interacti on domains. Six hexamers of CESA form a symmetri c rosette

on the plasma membrane plane; rosette movement is guided by cortical microtubules. Each

subunit of CESA in the rosette synthesizes one 1,4 P-D-glucose chain (Somerville, 2006).

Glycan synthesis includes backbone synthesis and side chain addition. Compared to

CESA, glycan synthesis is not well understood, although there is a structural similarity between

cellulose and glycans. Basing on this similarity, the CSL hypothesis has been established. It was

hypothesized that CELLULOSE SYNTHESIS LIKE (CSL) genes encode Golgi-localized glycan

synthases. This hypothesis is supported by recent discoveries of several glycan synthase genes

(Lerouxel et al., 2006). For example, mannan synthase, MLG synthase, XyG glucan synthase,

and galactomannan galactosyltransferase are all encoded by members of the CSL gene family.









Additionally, these enzymes are all responsible for the backbone synthesis of glycans. XyG

fucosyltransferase and XyG xylosyltransferase (XT1) were found to add side chains to the

backbone of glycans (Reiter, 2002; Lerouxel et al., 2006).

Pectin biosynthesis is much more complicated than cellulose and glycan synthesis because

it is difficult to investigate pectin synthases using traditional biochemical purification techniques

and forward genetics. Through reverse genetics and genomics approaches, more than 50

glycosyltransferases (GTs) are predicted to be required for pectin synthesis (Ridley et al., 2001).

At present, only a few genes for pectin biosynthetic GTs have been identified, and for some of

their products, the activity of the pectin synthesis is not clear (Bacic, 2006). HG

galacturonosyltransferase (GAUT 1) is the first functional identification of GTs using

biochemical and functional genomic approaches in Arabidopsis (Sterling et al., 2006). Other

putative GTs include QUA1 (Bouton et al., 2002), NpGUT1 (Iwai et al., 2002), and

PAR VUS GLZl (Lao et al., 2003).

During cell expansion, polysaccharides are deposited into existing walls. If polysaccharide

synthesis is disrupted, old wall reinforcement and new wall assembly cannot take place. Thus,

normal cell expansion is disrupted, resulting in aberrant cell shapes. For example, mutations in

CESA cause alterations of cell shape in diverse cell types because cellulose forms a fundamental

framework of the wall architecture. RSW1 encodes CESAl. The rswl mutation causes

disassembly of CESA complexes on the apoplastic side plane of the plasma membrane and

reduction of cellulose accumulation in cell walls (Arioli et al., 1998). The rswl mutants display

shorter roots with radial swellings (rsw), smaller leaf blades with shorter petioles, and aberrant

trichomes at the 310C restriction temperature. All these defects are indicative of abnormal cell

expansion owing to the rswl mutation (Williamson et al., 2001). The root radial swellings of the









rswl mutants exactly mimic phenotypic responses of wild-type roots to cellulose synthesis

inhibitors such as dichlorobenzonitrile. This observation suggests that the abnormal cell

expansion is due to reduction of cellulose synthesis. Further evidence for this view comes from

down-expression of CESA1 and CESA3, using transformation with antisense constructs. The

antisense phenotypes of CESA1 or CESA3 display shorter inflorescent shoots and stamen

filaments, a result of reductions in cell length rather than cell number. In addition, the severity of

the manifestation of both genes of interest is closely similar and intimately correlated to their

reduced expression (Burn et al., 2002). PROCUSTE1 (PRC1) is another gene, coding for

CESA6. Mutations in this gene also exhibit similar defects as rswl, including decreased cell

elongation, especially in roots and dark-grown hypocotyls. The cell elongation reduction is

correlated to a cellulose deficiency (Fagard et al., 2000). These observations indicate that the

reduction of cellulose in walls generally causes suppression of cell expansion.

Glycans form the cell wall matrix, which enhances the wall strength. Probably because of

this role, mutations in glycan synthase genes often cause only a slight alteration of cell expansion

and shapes. In addition, a specific mutation in one glycan synthase gene fails to cause a visible

phenotype. For example, M~UR2 encodes XyG fucosyltransferase (AtFUT1). The cell walls of

mur2 contain less than 2% of the wild-type amount of fucosylated XyG. The mur2 plants show a

normal growth habit and wall strength. On the other hand, M~UR1 codes for a 4,6-dehydratase,

responsible for the de novo synthesis of 1-fucose. Mutations in M~UR1 cause structural changes in

several cell wall polysaccharides (Bonin et al., 1997). Thus, murl mutants exhibit a dwarfed

growth habit and decreased wall strength, probably indicating aberrant cell expansion (Reiter et

al., 1993; Bonin et al., 1997). M~UR3 encodes XyG galactosyltransferase, which specifically

catalyzes formation of the a-L-Fuc-P-D-Gal- side group. Although the XyG in the mur3 cell walls










completely loses the fucosylated disaccharide side chain, mur3 plants are visibly

indistinguishable from wild-type plants except for a collapse of trichome papillae (Madson et al.,

2003). However, later investigators found that the galactose residues of XyG are essential to

maintain wall mechanical strength during rapid cell expansion. The mur3 mutations result in

reduced wall strength. Through studies using a scanning electron microscope, the defects in the

mur3 hypocotyl cells were observed. In addition, these defects are similar to the phenotype of

mutations in rswl, generating swollen cells of larger size (Pena et al., 2004). These defects

indicate abnormal cell expansion can be caused by a reduction of cell wall strength.

Pectins combine with glycans to form the secondary network of cell wall architecture.

Thus, mutations in the pectin synthesis genes share similar phenotypes to the phenotypes seen in

plants with defective GT genes. QLMSIM~ODO1 (QLM1) codes for a putative membrane-bound

GT. When the qual mutants were grown in light, the plants showed reduced height because of

the pectin deficiency in cell walls; similarly, the qual seedlings grown in the dark had shorter

hypocotyls, compared with the wild type. These defects are likely due to suppressed cell

expansion (Bouton et al., 2002). The dwarfism phenotype was also observed in the parvus

mutants because of both reduction in RG-I branching and alterations in the abundance of

xyloglucan linkages. PARVUS encodes another putative GT (Lao et al., 2003). Pectins play a

crucial role in pollen tube elongation because they are the only kind of molecule that makes a

single layer of wall at the growing tip (Stepka et al., 2000). VANGEMRD1 (YUD1) encodes a

pectin methylesterase (PME), which, depending on ambient pH, by enzymatic activity can lead

either to stiffening or to loosening of cell walls (Catoire et al., 1998; Denes et al., 2000).

Mutations in vgd1 cause a rupture of elongating pollen tubes in vitro and retarded growth in vivo

(Jiang et al., 2005). Moreover, when PME is exogenously added to growing pollen tubes, the









apical wall is thickened, resulting in inhibition of pollen tube elongation (Bosch et al., 2005).

PME activity that promotes cell wall loosening will be described below.

After cell division, through anisotropic expansion plant cells generally reach their final

sizes. At this time, polysaccharides have been almost equally deposited into primary cell walls.

A maj ority of the cells further differentially form specific shapes through further anisotropic

expansion. Before this process occurs, the cell wall at specific sites is biochemically "loosened"

for turgor-driven cell expansion (Cosgrove, 2000a). Additionally, the wall is loosened without

compromising the tensile strength of the pliant wall (Cosgrove, 2000b). The reason plant cells

have the capacity for this complex event is that the walls contain such unique proteins as

expansins, PME, and XyG endotransglucosylase (XET). PME de-esterifies highly

methylesterified pectins, converting the methoxyl groups into carboxyl groups on the

polygalacturonic acid chain, and releasing both methanol and protons. This conversion promotes

pectin gelation accompanying wall stiffening, due to the formation of the cooperative Ca2+ CTOSS-

bridges between free carboxyl groups of adjacent pectin chains (Catoire et al., 1998). In addition,

the de-esterification reduces the local pH, which promotes activity of several other cell wall-

loosening hydrolases, such as expansins, polygalacturonases, and pectate lyases (Cosgrove et al.,

2002). Under-expression ofPM~E by antisense RNA in transgenic pea reduced root hair

elongation. Moreover, the root length reduction was correlated with an increase in extracellular

pH (Wen et al., 1999). The activation of wall hydrolases by acidification of cell walls was also

supported by the functional analysis of DET3, which is a vacuolar H+-ATPase (V-ATPase).

Mutations in DET3 lead to defects in hypocotyl cell elongation. It was suggested that V-ATPases

contribute to maintaining the internal turgor pressure of plant cells through modulation of solute

uptake to vacuole (Schumacher et al., 1999). Subsequently, another kind of vacuolar H+-pump









called H+-PPase AVPI (AVPl) was found to be implicated in this transport process. AVPI

adjusted the distribution and abundance of H+-ATPases in the plasma membrane by controlling

its trafficking through the endocytic secretary pathways. Over-expression of AVP I increased the

accumulation and polar distribution of the H+-ATPase in the plasma membrane.

Consequentially, more H+ was pumped to the apoplast and the cell wall was acidified. Thus, cell

elongation occurred. In the avpl-1 null mutant, root cell elongation was severely disrupted (Li et

al., 2005).

Expansin, another cell wall protein, induces pH-dependent wall extension and stress

relaxation in a characteristically unique manner (Cosgrove et al., 2002). Expansin has been

proven to cause isolated wall extension in vitro under constant mechanical stress (McQueen-

Mason et al., 1992). Application of exogenous expansion from cucumber in excised Arabidopsis

hypocotyls stimulates cell elongation. At a high concentration of applied expansion, the tips of

growing root hairs burst; at a lower level, exogenous expansion caused radial swelling at the tip

(Cosgrove et al., 2002). Over-expression of Arabidopsis EXP10 results in transgenic plants with

longer petioles and larger leaf blades because of increased cell size (Cho and Cosgrove, 2000).

Over-expression of EXP1 in tomato fruit enhances fruit softening and cell wall breakdown

(Brummell et al., 1999).

The enzymatic activity of XET breaks the existing linkages in the XyG-cellulose network

and rej oins the resultant ends with new partners at different positions. XET loosens the wall

during cell expansion through cooperation with expansion (Nishitani and Tominaga, 1992). XET

activity is intimately correlated to the cell growth rate and epidermal lengthening in the growing

zone of maize leaves (Rose et al., 2002). XET also is specifically localized at the site of









trichoblast walls, where the future bulge is formed during root hair initiation. A locally high level

of XET activity stimulates trichoblasts to initiate root hairs (Vissenberg et al., 2001).

Plasma Membrane Dynamics and Cell Expansion

The plasma membrane abuts cell walls and participates in cell wall synthesis. As

mentioned above, the plasma membrane protein CESA synthesizes cellulose for direct wall

synthesis. The plasma membrane also actively performs exocytosis for indirect wall synthesis

during cell expansion. In addition, the structural asymmetry of the plasma membrane provides

cell polarity for anisotropic expansion (Fischer et al., 2004). To understand the roles of the

plasma membrane in cell expansion, I briefly describe its components, highlighting its structural

asymmetry. After this, the assembly of the plasma membrane and its relation to cell expansion

are described.

Components of the Plasma Membrane

The plasma membrane of plant cells is composed of lipids, proteins, and carbohydrates in a

molecular ratio of approximately 2 : 2 : 1. The membrane lipids include phospholipids,

sphingolipids, and sterols (Moreau et al., 1998; Jaillais and Gaude, 2008). Some phospholipids

such as phosphatidylcholine (PC) and phosphatidylethanolamine (PE) have head groups with

positive charges; whereas others are negative or neutral, depending on pH. More importantly, the

ratio of lipid classes in plant plasma membranes shows a wide range of variation among the

different organs in a given plant or identical organs in distinct plants (Jouhet et al., 2007). The

maj ority of carbohydrates in plant plasma membranes are present in the form of oligosaccharides

that are covalently linked to proteins to generate glycoproteins (Bacic et al., 1996; Classen et al.,

2005).









Asymmetry of Plant Plasma Membranes

Lipids, proteins, and carbohydrates of plant plasma membranes are regularly arranged in

an asymmetric bilayer structure. The lipid amphipathy allows for spontaneous assembly of

bilayers. The hydrophilic heads maximize their interactions with water, whereas the hydrophobic

tails interact with each other, minimizing their exposure in the exoplasmic leaflet (Holthuis and

Levine, 2005; Pomorski and Menon, 2006). Sphingolipids and sterols are abundant in

microdomains (lipid rafts), in which signaling proteins accumulate. Other proteins are also

localized at particular sites in the plasma membrane. For example, auxin carriers are positioned

at the apical or basal plasma membrane of root epidermal cells (Muller et al., 1998; Swarup et

al., 2001), whereas glycosylphosphatidylinositol (GPI)-anchored proteins (GAPs) such as

COBRA (COB) preferentially localizes to lateral membrane in root cells (Schindelman et al.,

2001). This asymmetric distribution establishes and maintains cell polarity, which is required for

longitudinal expansion at the elongation zone (Fischer et al., 2004; Kramer and Bennett, 2006).

Plasma Membrane Assembly and Cell Expansion

Plasma membranes of plant cells are assembled using various lipids and diverse proteins.

In addition, different lipids interact with distinct proteins, and specific proteins have particular

positions in lipid bilayers. During isotropic cell expansion, the plasma membrane evenly enlarges

its area, whereas in polar expansion it is assembled only at the growing sites. For any cell

expansion, the plasma membrane maintains the dynamic stability of chemical composition and

distribution of protein components. In addition, it has the capacity for flexible changes in these

features in response to intracellular and extracellular signalings for anisotropic expansion.

Perturbations of membrane stability cause abnormal cell expansion (Schrick et al., 2000; Souter

et al., 2002; Jaillais and Gaude, 2008).









Changes of lipid composition in the plasma membrane cause disruption of normal cell

expansion (Jaillais and Gaude, 2008). COTYLEDON VASCULAR PATTERN1~ (CVPl) and the

ORC gene encode a sterol methytransferase for sterol biosynthesis. Mutations in either CVP1 or

ORC alter the membrane sterol composition, which raises the level of cholesterol and

campesterol at the expense of sitosterol. The smtlore and smt2cvpl double mutants display defects

in polar cell expansion that result in the perturbed alignment of cells into parallel vascular cell

files in cotyledons because abnormal cell expansion leads to aberrant cell shapes (Carland et al.,

2002; Willemsen et al., 2003). Sterols are enriched in lipid crafts, which provide platforms that

anchor polar proteins to the plasma membrane. Thus, the influence of sterol composition on the

localization of polar proteins has been widely reported in yeasts and animals (Simons and

Ikonen, 1997; Bagnat et al., 2000; Bagnat and Simons, 2002; Danielsen and Hansen, 2006).

These events are also likely to occur in plants. For example, it has been found that, in

Arabidopsis, smtlore root cells mislocalized auxin carriers on the plasma membrane (Willemsen

et al., 2003).

Changes in anchored proteins of the plasma membrane also cause disruption of normal cell

expansion. In eukaryotic cells, glycosylphosphatidylinositol (GPI) anchored proteins (GAPs)

have been extensively reported to be plasma membrane bound proteins (Oxley and Bacic, 1999;

Sherrier et al., 1999; Zhao et al., 2002). Salt-overly-sensitive 5 (SOS5) encodes a plant GAP.

Mutations in SOS5 cause strong radial expansion of the cells in the elongation zone of roots,

instead of the longitudinal expansion seen in wild type. Thus, the epidermal, cortical, and

endodermal cells in sos5 roots display a swelling phenotype (Shi et al., 2003). COB, another

plant GAP, is anchored on the extracellular side of the plasma membrane and also released into

the wall. It is required for highly anisotropic expansion of all plant cells. The cob-4 null allele









shows greatly reduced growth of all organs in seedlings. However, because the cob-4 root cells

expanded radially in the elongation zone, the root diameter ultimately reaches nearly twice that

of the wild type. The root epidermal cells displayed a severe bulging phenotype. In addition,

their cell walls were occasionally broken. COB plays a crucial role in the oriented deposition of

cellulose microfibrils during rapid anisotropic expansion (Roudier et al., 2005). SKU5 has the

same cell localization as COB, which is in the cell wall and on the plasma membrane anchored

by GPI. SKES is expressed most strongly in expanding cells. Mutations in SKES cause skewed

roots and shortened hypocotyls because of abnormal cell expansion (Sedbrook et al., 2002).

Although PNT 1 is not a GAP, it encodes a mannosyltransferase for the GPI synthesis. All five

pnt mutants strongly reduce accumulation of GAPs. In the pnt1 mutants, the cell walls had

decreased crystalline cellulose; embryos showed delayed morphogenesis; apical meristems were

defective; and seedlings did not survive. All phenotypes were due to aberrant cell expansion

(Gillmor et al., 2005). ETH1 and SETH2 also are involved in GPI biosynthesis. The seth1 and

seth2 mutations specifically block or reduce pollen tube elongation because of abnormal callose

deposition. In addition, another 47 genes, which all encode potential GPI-anchored proteins, are

likely to play important roles in the establishment and maintenance of polarized pollen tube

expansion (Lalanne et al., 2004).

Normal cell expansion is disrupted by changes in the position of polar proteins.

Arabidopsis phospholipase D51 (AtPLDS1) is preferentially localized at the tip of growing root

hairs. Ectopic over-expression ofAtPLDS-1 disrupts its distribution in specific tissues and induces

non-root cells to form ectopic root hairs in cotyledons and hypocotyls. The raised level of

AtPLDS1 in root cells generates swollen or branched root hairs (Ohashi et al., 2003). It is likely

that the intrinsic polarized distribution of AtPLDS1 is perturbed, thus losing capacity for tip










growth. Recently, AtPLD52 was also found to be involved in cell expansion. Mutations in this

gene reduced the hypocotyls cell elongation (Li and Xue, 2007). More recently, it was found that

ROOT HAIR DEFECTIVE 2 (RHD2) is located at the growing tip of root hair cells. Mutations

in RHD2 causes defective root hairs (Takeda et al., 2008). AGC2 is a member of the

cAMP/cGMP-dependent kinase or protein kinase C family kinase (AGC kinase). It localizes to

the root hair tip. The agc2-1 mutants have short root hairs because of reduced cell elongation

(Anthony et al., 2004). Other polar proteins of plant plasma membranes that play a role in cell

expansion in response to phospholipid signaling are transported by oriented vesicle trafficking

involving such proteins as PIP5K3 (Kusano et al., 2008) and RHD4 (Thole et al., 2008). These

proteins are described below in the endomembrane dynamics and cell expansion section.

In recent years, numerous observations of transcytosis of the plasma membrane in plant

cells were reported as a result of the application of fluorescent styryl dyes, such as FM1-43 and

FM4-64, particularly in expanding root hairs and pollen tubes (Samaj et al., 2006). Through

exocytosis, secretary vesicles containing the cell wall cargo molecule fuse with the plasma

membrane, releasing their contents into the extracellular space for wall loosening, strengthening,

or assembling during cell expansion, whereas, through endocytosis, the extra plasma membrane

proteins and lipids are transferred into the cytosol for recycling at the growing sites (Samaj et al.,

2005; Johansen et al., 2006). Increasing evidence shows that blocking exocytosis with brefeldin

A (BFA) inhibits pollen germination and pollen tube elongation (Wang et al., 2005b). Because

transcytosis is tightly coupled with the endomembrane trafficking network, details of this topic

are described below.

Endomembrane Dynamics and Cell Expansion

Endomembrane trafficking is essential for cell expansion, especially for polar expansion

(Samaj et al., 2006). Except for cellulose, almost all cell wall components are synthesized in the










Golgi apparatus (GA) and endoplasmic reticulum (ER), further packaged into vesicles, and

finally transported to the wall space (Reiter, 2002; Dhugga, 2005; Lerouxel et al., 2006). The

oriented trafficking of vesicles serves to establish and maintain cell polarity, which initiates polar

expansion. In this part, I introduce general endomembrane pathways first, focusing on their

characteristics in plant cells. After that, I highlight the influence of the disruption of

endomembrane trafficking on plant cell expansion.

Endomembrane Trafficking Pathways

The endomembrane system is composed of all intracellular membranous compartments.

These compartments communicate with each other through exchanging molecules using

ubiquitous vesicles as carriers. The vesicles containing cargo molecules secrete from donor

compartments, and fuse with target compartments. During vesicle trafficking, the GA plays a

leading role. It lies at the heart of the membrane trafficking pathway, serving as the crossroad in

various trafficking events (Hawes and Satiat-Jeunemaitre, 2005a). At its cis-face, the GA

receives the vesicles from the ER (anterograde transport) and sends their vesicles back

(retrograde transport). At its trans-face, the GA sends its vesicles to endosomes, storage

vacuoles, lytic vacuoles, and the plasma membrane. The vesicles, from the plasma membrane

through endocytosis, are recycled in the GA. The vesicles exported from the ER can also bypass

the GA pathway and go directly to a vacuole or the plasma membrane (Hawes, 2005).

The GA is the sum of numerous polarized stacks of membrane-bounded cisternae. Within

the GA, cargo molecules are processed, concentrated, and packaged into vesicles (Hawes, 2005).

The mature cargo molecules in the vesicles are intracellularly routed to specific cellular

destinations within cells. For plant cells, the GA is an important organelle that specializes in the

synthesis and processing of complex components of cell walls, such as glycans and glycoproteins

(Hawes and Satiat-Jeunemaitre, 2005b). Probably to this end, the GA organization in plant cells









is different from its counterpart in animal cells, in which the GA is composed of many stacks that

are generally arranged side-by-side in a ribbon structure around the nucleus. In plants, the GA is

divided into individual Golgi stacks that are distributed through the cytoplasm (Latijnhouwers et

al., 2005). The number of Golgi stacks per cell and the number of cisternae per stack varies with

cell developmental stages and cell types (Neumann et al., 2003). Mammalian GA is a rather

static organelle, but plant GA is a highly mobile biosynthetic factory that moves over the ER on

an actin network at the speed of 2 Cpm per second (Boevink et al., 1998; Nebenfuhr et al., 1999).

This characteristic of the plant GA enables cargo molecules to be efficiently transported to the

extra-cellular matrix.

Vesicle Dynamics

In eukaryotic cells, the vesicle is a ubiquitous vector for endomembrane trafficking. The

highly dynamic vesicles are generated through membrane fission of donor compartments, and

disappear by membrane fusion at destination compartments. The processing of both the fission

and the fusion elements involves close contact between lipid bilayers and the final combination

of bilayer leaflets at specific sites (Markvoort et al., 2007). Fission begins with bending of

membranes at the export site. The bent membrane invaginates to an extreme curvature to form a

highly constricted neck. The neck further elongates and narrows until the two membranes merge,

which leads to the separation of the vesicles from the donor compartment. Membrane fusion

begins with docking of vesicles at acceptor compartments. The lipid bilayer of the docked

vesicle gradually unites with the membrane of the acceptor compartment (Sollner and Rothman,

1996; Atilgan and Sun, 2007). Membrane fission and fusion are completed by the vesicle

assembly machinery. In mammalian and yeast cells, the components of this machinery are well

known. Generally they include the coat proteins, adaptor proteins, cargo receptors, and small

GTPase proteins (Marks et al., 2001).









Vesicle Fission Machinery

During vesicle assembly, numerous proteins are recruited at the exit site. On the internal

surface of membranes, cargo receptors such as v-SNARES specifically bind outgoing molecules

from the lumen. On their external surface, the ADP-ribosylation factors (ARFs) are recruited

from the cytosol to bind tightly to the membrane. The ARFs further interact with their effectors

and regulators. A regulator such as GAP, contributes to hydrolysis of the active GTP-bound ARF

form to the inactive GDP-bound ARF form with opposite conversion catalyzed by GEF. The

change of the ARF conformation during GTP hydrolysis leads to structural alterations of both the

lipid and the effectors attaching with the membrane. This alteration promotes vesicle budding

and delivery from the donor compartment. The ARF effectors mainly include coat proteins

(Donaldson and Jackson, 2000; Donaldson et al., 2005). Three types of coat proteins have been

found: clathrin, COPI, and COPII. Clathrin coats the transport vesicle that shuttles between the

GA and the plasma membrane, the GA and endosomes, and the plasma membrane and

endosomes; COPs wrap the vesicles that shuttle between the GA and the ER. COPI coats

vesicles from the GA to the ER, while COPII coats the vesicles from the ER to the GA. Recent

research has shown that COPI interacts with Brefeldin-A ADP-ribosylated substrate (BARS)

(Yang et al., 2005; Yang et al., 2006) and the actin cytoskeleton provides force both for

membrane deformation during vesicle formation and for vesicle trafficking to the correct

destination (Goley and Welch, 2006; Kaksonen et al., 2006; Co et al., 2007).

Vesicle Fusion Machinery

After the vesicles detach from the donor membrane, the coat proteins release from the

vesicle surface into the cytosol for recycling. Thus, the vesicle surface marker v-SNAREs

(soluble N-ethylmaleimide-sensitive-factor attachment protein receptors) is exposed, as well as

their complementary SNAREs, termed t-SNAREs, on the target compartment. The interaction









between these two SNAREs causes fusion of the vesicle with the target membrane (Chen and

Scheller, 2001; Bonifacino and Glick, 2004). More than 30 members in the SNARE superfamily

were found in mammalian cells (Chen and Scheller, 2001). A large number of SNAREs were

also found in the Arabidopsis genome and other plants (Sanderfoot and Raikhel, 1999;

Sanderfoot et al., 2000; Sanderfoot, 2007).The regulatory protein Rab participates in specific

junctions of v-SNAREs with t-SNARSEs. Rab is a GTP-binding protein. It also provides energy

for driving membrane fusion through GTP hydrolysis, while binding with the t-SNAREs (Segev,

2001; Zerial and McBride, 2001)

Vesicle Trafficking and Cell Expansion

Identifying the molecular machinery of the plant membrane trafficking pathway reveals

significant homology with that of mammalian and yeast counterparts. Coat proteins (COPI,

COPII, and clathrin), small GTPases (Rabs, Arf, Sarl, and Rac), and fusion proteins (SNAREs)

appear to be well conserved throughout eukaryotic cells. In plant cells, the maj ority of the COPI

and COPII machinery and their associated effectors, such as Arf and Sarl, have been cloned, and

their functions are now being determined. More and more data have indicated that vesicle

trafficking contributes to establishment of the structural and molecular asymmetry at the cell

surface, which is the beginning of cell polar expansion (Xu and Scheres, 2005a; Friml et al.,

2006).

Brefeldin A (BFA) is a fungal toxin that has been widely used as a reversible inhibitor of

vesicle trafficking in yeast, mammalian, and plant cells. BFA blocks the detachment of vesicles

at the exit site of the GA membrane to form aggregates, which disrupts the normal flow of the

vesicles within cells. BFA has been shown to alter the distribution of such plasma membrane-

localized proteins as PIN1I, AUX1, PM-ATPase, and pectins in plant root cells. In addition, the

epidermal cells ofArabidopsis roots treated with BFA lost their polarity, displaying a decrease in









cell length and increase in the apical-basal initiation ratio, as well as formation of double root

hairs, in which two root hairs are derived from one trichoblast (Grebe et al., 2002). The

molecular mechanism of BFA inhibition of vesicle trafficking is due to the binding of BFA with

the ARFl1-GDP/ARF-GEF complex, which prevents ARF 1 activation necessary for vesicle

budding and cargo molecule selection (Robineau et al., 2000). In mammalian cells, ARF1 is a

core component of the vesicle assembly machinery, recruiting COPI and clathrin coat proteins to

membranes for vesicle assembly in mammalian cells (Boman, 2001; Rein et al., 2002; Spang,

2002; Song et al., 2006). In Arabidopsis, ARF 1 rescued the ARFl1/ARF2 lethal yeast double

mutant, which suggest that plant ARF 1 has similar functions to its yeast and mammalian

counterparts (Takeuchi et al., 2002). Moreover, Arabidopsis ARF 1 is also located to the GA and

endocytic organelles. The over-expression of the engineered ARF 1 with dominant activation and

inactivated formats that are targeted to interfere with the endogenous ARF1 function in

trichoblasts significantly inhibited polarized tip expansion, which produced shorter roots. After

strong heat shock induction, the trichoblasts and epidermal cells in the transgenic lines displayed

many more severe defects, such as apical-shifted root hairs, double root hairs, and bulged

epidermal cells. All these phenotypes show that the cells lost their polarity and capacity for polar

expansion (Xu and Scheres, 2005b). The ARF GTPase activating protein, ROOT AND POLLEN

ARFGAP (RPA), activates ARF 1 and plays a role in the elongation of root hairs and pollen tubes

in Arabidopsis. RPA is specifically expressed in roots and pollen; its product is located in the

GA. Additionally, RPA complements the glo3 gcs double mutants in yeast (Song et al., 2006).

GLO3 and GCS1 are two yeast ARFGAPs that function efficient retrograde trafficking of

vesicles from the GA to the ER (Poon et al., 1999; Robinson et al., 2006). ARFGAPI promotes

both vesicle formation and cargo sorting by functioning as a component of the COPI coat (Yang









et al., 2002; Lee et al., 2005). The loss-of-function mutant, rpa, causes root cells to isotropically

expand to generate short and branched root hairs, as well as a slowing of pollen tube elongation

(Song et al., 2006). Over-expression of the rice OsARFGAP also interfered with vesicle

trafficking, which influenced on the root hair formation and elongation. Transgenic plants of

Arabidopsis and rice had a reduced number of lateral roots and reduced root length. The ratio of

length and width of epidermal cells at the elongation region also decreased compared to wild-

type plants. Additionally, abnormal vesicle aggregates (the BFA compartment, a typical defect of

disrupted vesicle trafficking) were observed in the transgenic cells (Zhuang et al., 2005; Zhuang

et al., 2006). The ARF guanine-nucleotide exchange factor (ARF-GEF), GNOM, functions in the

establishment of apical-basal cell polarity by mediation of specific endosomal trafficking

pathway in Arabidopsis embryos and roots. GNOM localizes to endosomes. The loss-of-function

mutant of gnom lacked an apical-basal polar axis and embryonic root in early embryos (Geldner

et al., 2003). This phenotype is probably a result of defective cells that are unable to expand or

anisotropically expand.

Arabidopsis Rab GTPase RabA4b was found to function in cell directional expansion

through the regulation of vesicle trafficking involved in the polarized deposition of cell wall

components in tip-growing root hair cells (Preuss et al., 2004). Mammalian Rabl11 has a high

degree of homology to RabA4b, as does Arabidopsis Ara4, which was localized to Golgi-derived

vesicles, Golgi cisternae, and the trans-Golgi network (Ueda et al., 1996). Vesicles with the

EYFP-RabA4b labeling marker accumulated in the actively expanding zone in the growing root

hair. Such accumulation is necessary for root hair initiation and elongation. In the rhdl-1

mutant, and the rhd2-1 mutant, their defective cells did not show accumulation of the vesicles

with Rab4A (Preuss et al., 2004). Plant Rabl11 also plays a role in pollen tube tip growth.









Tobacco Rab 11b was localized to the pollen tube tip. Interference of endogenous Rab 11b

activity in its mutated variants gave rise to a reduction of pollen tube growth rate and change of

pollen tube morphology (de Graaf et al., 2005). Tobacco Rab2 also functions in cell polar

expansion of pollen tubes through a vesicle trafficking pathway at the GA. NtRab2 localizes to

the GA. The mutated NtRab2 blocks vesicle release from the GA so that the normal delivery of

Golgi cargo to their destinations, such as the cell surface, was disrupted, inhibiting pollen tube

expansion (Cheung et al., 2002).

Cytoskeleton Dynamics and Cell Expansion

The plant cytoskeleton includes microtubules and microfilaments that spatially control cell

expansion (Smith and Oppenheimer, 2005). It is generally believed that microtubules serve as a

scaffold for cells and are important for establishing and maintaining cell expansion direction,

whereas microfilaments function as a track for vesicles to specific sites to deliver cargo required

for expansion (Mathur and Hulskamp, 2002). Although many observations have shown that

cytoskeletal dynamics and proper organization are essential for cell expansion, much remains to

be learned, including the precise roles of microtubules and microfilaments in spatial control of

cell expansion. There is evidence, however, that shows that the direction of the wall's main

structural component, cellulose, is determined by microtubules. The arrangement of cellulose

microfibrils in the wall is a key determinant of the cell expansion pattern and is clearly related to

the arrangement of cortical microtubules in expanding cells (Smith and Oppenheimer, 2005). A

rapidly growing body of knowledge has accumulated about how the dynamics and organization

of both classes of filaments are controlled in expanding cells (Baskin, 2005). Here, however, we

emphasize recent work explaining regulation of the cytoskeleton and its contributions to

patterning of plant cell expansion.









The Microtubule Cytoskeleton and Cell Expansion

Microtubules orient the deposition of microfibrils in walls, which determines the cell

expansion direction. The direction of cell expansion is determined by the organizational pattern

of cortical microtubules because it was found that the latter normally mirrors the arrangement of

cellulose microfibrils, the key structural element of the wall, in growing cells. Based on this

observation, the co-alignment hypothesis was established. It was hypothesized that movement of

cellulose synthase enzyme complexes in the plasma membrane is constrained by interactions

with the cortical microtubules (Giddings and Stachelin, 1991). To accommodate later,

conflicting, observations, this hypothesis has further evolved into several distinct versions, such

as the template incorporation model (Baskin, 2001) and the microfibril length regulation

hypothesis (Wasteneys, 2004). Here I will describe the evidence that support these hypotheses,

with an emphasis on maj or advances in recent years.

Early observation showed that the deposition of cellulose microfibrils in elongating cells

was typically perpendicular to the axis of cellular expansion. In addition, disruption of these

fibrils with colchicine caused an isodiametric expansion. These characteristics led to the

prediction that cytoplasmic elements exist in the cell periphery, orient the deposition of cellulose

microfibrils, and constrain the pattern of cell expansion (Green, 1962). Only one year later,

electron microscopy showed slender tubules microtubuless) at the cell cortex. More importantly,

the orientation of these tubules mirrored that of the cellulose microfibrils in the adj acent cell

walls (Ledbetter and Porter, 1963). Thereafter, cortical microtubules were often observed to lie

parallel to the cellulose microfibrils (Hepler and Palevitz, 1974).

Microinj section of rhodamine-conjugated tubulin into the epidermal cells of pea internodes

showed the array shift of the cortical microtubules between the transverse organization pattern

and longitudinal one after application of gibberellic acid for induction of cell growth. This shift









is likely to be involved in a range of responses that alter the direction of cell expansion (Yuan et

al., 1994).

Disruption of the dynamics and organization of endogenous microtubules with

microtubule-modifying drugs gives rise to aberrant cell expansion. For instance, oryzalin is a

compound that causes microtubule depolymerization, whereas taxol has an opposite effect; it

promotes microtubule assembly. Arabidopsis seedlings treated with either oryzalin or taxol

display an identical defective phenotype, the radial expansion of root cells. This result indicates

the importance of microtubule dynamics in cell expansion. Additionally, the defective severity of

the cortical microtubules in the swelling root cells increases with drug concentration. At low

concentrations of oryzalin, microtubule arrays are disorganized; at medium concentrations they

are fragmented, and at high concentrations they are totally depleted. However, in the taxol-

treated root cells, the cortical microtubules at the elongation zone display disorganization in

directionality compared with the control cells. At 10 micromolar concentration, many stele cells

have more longitudinal microtubules, whereas many cortical cells appear to have more

transversely aligned microtubules (Baskin et al., 1994). These experiments were repeated later

(Sugimoto et al., 2003). The same results were obtained from an experiment with maize roots

treated with oryzalin or taxol (Hasenstein et al., 1999).

Microtubules are polymers of tubulin. Mutations in the genes for tubulin also cause

aberrant cell expansion, such as helical elongation. Dominant negative mutations in the a-tubulin

genes cause left-handed helical growth and clockwise twisting in elongating organs of

Arabidopsis because the mutant tubulins are incorporated into microtubules, producing right-

handed obliquely oriented cortical arrays in the root epidermal cells. Additionally, the cortical

microtubules in the mutants had increased sensitivity to microtubule-specific drugs, indicating









that the reduced microtubule stability can produce left-handed helical cell expansion

(Thitamadee et al., 2002). The same defective cell expansion was exhibited in transgenic plants

with the same mutated version of the a-tubulin gene (Abe and Hashimoto, 2005). This result

further confirms that disturbance of endogenous microtubules influences cell expansion.

During cell expansion, the dynamic and well organized microtubules are mediated by the

regulators, the maj ority of which are microtubule-associated proteins (MAPs). Thus, mutations

in the MAP genes suppress the dynamics of microtubules, block their reorganization, and affect

cell expansion. M~ICROTUBULE ORGANIZA TION1 (M~OR1) encodes a member of an ancient

family of MAPs. The amino acid sequence of MOR1 is similar to Xenopus MAP215. In

Arabidopsis MOR1 regulates cortical microtubule organization, likely through stabilization of

microtubules. Mutations in M~OR1 generate unstable microtubules. At the 290C restrictive

temperature, the cortical microtubules in leaf epidermal cells of mor1 mutants break into

fragments, but at the 210C permissive temperature, the microtubules revert to their normal

appearance. At the restrictive temperature, the mor1 plants are severely stunted, producing

radially swollen and short organs indicative of aberrant cell expansion (Whittington et al., 2001).

FRA2, another microtubule regulator, was found to have the activity of severing

microtubules in vitro. Through confocal microscopy and immunofluorescence, it was found that

the cortical microtubules are disorganized in fr~a2 mutants. Meanwhile, using field emission

scanning electron microscopy for studies on the walls, the fra2 mutation alters the normal

orientation of cellulose microfibrils in walls of expanding cells. The fra2 mutants show reduced

cell elongation. These findings strongly support the co-alignment hypothesis that microtubules

orient cell expansion through the control of directional deposition of cellulose microfibrils in the









wall (Burk and Ye, 2002). Using the same methods for the cob mutations, strong evidence

supporting this hypothesis was also obtained (Roudier et al., 2005).

COB is not a MAP, but an anchor to GAP, which is involved in regulation of cell polarity

(Fischer et al., 2004). As mentioned above, COB is polarly targeted to both the plasma

membrane and the longitudinal cell walls. Additionally it is distributed in a banding pattern

perpendicular to the longitudinal axis via a microtubule-dependent mechanism. The elongating

root cells in cob mutants lose capacity for anisotropic expansion and display a swelling

phenotype. The defective cells are accompanied by disorganization of the orientation of cellulose

microfibrils (Roudier et al., 2005).

The direct evidence supporting the microfibril and microtubule co-alignment hypothesis

was recently gained using spinning disk confocal microscopy. The process of cellulose

deposition was visualized in living cells by fluorescently-tagged CESA. The CESA complexes in

the plasma membrane moved at a constant rate in a linear track that was aligned and coincident

with cortical microtubules. Inhibition of microtubule polymerization changed the fine-scale

distribution and pattern of moving CESA complexes in the membrane, indicating a direct

mechanism for the guidance of cellulose deposition by microtubules (Paredez et al., 2006).

Signaling pathways of phospholipids and GTPases are involved in the regulation of the

microtubule organization for cell expansion. In recent years, rapid advances have been made on

understanding microtubule regulation by distinct signaling pathways, which are intimately

related to anisotropic cell expansion, particularly by phospholipids and Rho of plants (ROPs). A

wealth of mutations in the genes encoding components of these pathways causes aberrant cell

expansion.









Phospholipase D (PLD) is a key component of the phospholipid signaling pathway. It was

found that PLD decorates microtubules in plant cells (Marc et al., 1996) and is localized to the

plasma membrane (Marc et al., 1996; Gardiner et al., 2001). PLDs are enzymes that hydrolyze

structural phospholipids such as phosphatidylcholine to produce free choline and phosphatidic

acid (PA), which function as a second messenger in cell signaling. Biotic and abiotic stresses

such as wounding and pathogen infection rapidly stimulate PLD activity (Laxalt and Munnik,

2002; Wang et al., 2002; Meijer and Munnik, 2003). PLD activation triggers reorganization of

plant microtubules (Dhonukshe et al., 2003). Changes of PLD levels disrupt the phospholipid

signaling transmission, resulting in aberrant cell expansion likely because of the microtubule

disorganization. Thus, raised levels of AtPLDS1 generate either swollen or branched root hairs

(Ohashi et al., 2003). AtPLD52 absence reduced the hypocotyl cell elongation (Li and Xue,

2007).

ROP GTPases are plant-specific signaling molecules. They potentially interact with cell

surface-associated signal perception apparatus for such extracellular stimuli as hormones,

pathogen elicitors and abiotic stress. ROP GTPases mediate diverse cellular processes, including

microtubule dynamics and organization (Nibau et al., 2006). It was found that ROP2 inactivates

ROP-interactive CRIB motif-containing protein (RIC 1) in Arabidopsis epidermal leaf pavement

cells. RIC1 activity promotes well-ordered cortical microtubules. The RIC1i-dependent

microtubule organization not only locally inhibits outgrowth, but also in turn suppresses ROP2

activation in indentation zones. RIC1 over-expression suppresses lobe formation, and ric1

mutants exhibit wide neck regions (Fu et al., 2005).

Actin Cytoskeleton Dynamics and Cell Expansion

The mammalian actin cytoskeleton not only mechanically supports cells for formation of

various shapes, but also generates a driving force for such diverse cellular or intracellular events









as cell migration, vesicle trafficking, exocytosis, and endocytosis (Kaksonen et al., 2006). As

these events occur, actin cytoskeleton dynamics are essential for re-assembly of actin filaments

at distinct subcellular locations (Goley and Welch, 2006). The dendritic nucleation model of

actin polymerization is used for interpretation of the molecular mechanism of these events

(Mullins et al., 1998; Pollard and Borisy, 2003). A leading hypothesis for force generation is

through actoclampin, the ATP hydrolysis-dependent, affinity-modulated motor unit (Dickinson

and Purich, 2002; Dickinson et al., 2004; Zeile et al., 2005). However, the role of the plant actin

cytoskeleton is just coming of age. In recent years, increasing evidence has shown that the plant

actin cytoskeleton is important for cell expansion during cell morphogenesis, particularly for tip

growth and anisotropic expansion (Hussey et al., 2006). The role of the actin cytoskeleton is

generally believed to be the delivery of specific vesicles containing cell wall materials to

specified sites for local growth. The maj ority of investigations were done using cell-specific

models, such as pollen tubes, root hairs, trichomes, and leaf pavement cells (Mathur and

Hulskamp, 2002; Schellmann and Hulskamp, 2005; Smith and Oppenheimer, 2005; Cole and

Fowler, 2006).

Polarized organization of the actin cytoskeleton is required for tip growth. Both pollen

tubes and root hairs offer suitable models to study the roles of F-actin organization and dynamics

in tip growth. In these cells, it has been observed that at least two forms of F-actin exist. One is

the actin cables arranged along the elongation axis; and other is the dynamic Eine F-actin

localized to the tip (Hepler et al., 2001; Cole and Fowler, 2006). A wealth of observations shows

chemical and genetic disruption of F-actin dynamics and polarized organization in tip-growing

cells arrests tip growth.









For investigation of the roles of actin cytoskeleton in tip growth, LatB is the first chemical

agent used to inhibit actin polymerization. LatB-treated maize pollen tubes display a dose-

dependent depolymerization of F-actin. The elongation of the LatB-treated pollen tubes is

arrested because of F-actin depolymerization (Gibbon et al., 1999). The same result was obtained

in LatB-treated pollen tubes of Picea meyeri (Chen et al., 2007). The pollen tubes treated with

15 nM LatB for 20 hours show severe disruption of actin filaments. The polarized actin cables

become short fragments throughout the tubes. In addition, some actin fragments tend to

aggregate into clusters in the sub-apical region of the tube. The tip of LatB-treated pollen tubes

swelled because of its loss of polarity (Chen et al., 2007).

Genetic disruption of the actin cytoskeleton also causes aberrant tip growth. Formins are

actin-nucleating proteins that stimulate the de novo polymerization of actin filaments in

mammalian cells (Kovar, 2006). It has been found that plant formins appear to have the same

function as that in mammalian cells. Thus, changed levels of formin expression in pollen tubes

affect the dynamics of F-actin and disrupt tip growth. Over-expression of Arabidopsis formin

AFH1 in pollen tubes induces the formation of arrays of actin cables, resulting in depolarization

of tip growth and generation of a broadening tube. Moreover, severe membrane deformation was

observed in the apical region (Cheung and Wu, 2004).

Longitudinal actin cables serve as tracks for motor proteins that transport vesicles to the

tips of growing pollen tubes and root hairs. Active vesicle transport was observed in root hairs,

particularly at the growing tips. This cellular process is based on F-actin, which, when disrupted,

arrests vesicle trafficking (Voigt et al., 2005).

Precise organization of the actin cytoskeleton is important for cell morphogenesis. The

Arabidopsis trichome provides an excellent model for studies on cell morphogenesis. The









trichome is a large, single cell that develops on the epidermal surface. Its morphogenesis is a

complex process, in which an approximately round, epidermal cell develops into a stellate

symmetrical trichome (Schellmann and Hulskamp, 2005). Using this model cell, investigators

found indirect evidence supporting the importance of precise actin cytoskeleton organization for

trichome morphogenesis in pharmacological experiments with drugs that affect actin dynamics.

When developing trichomes were treated with microfilament destabilizing antagonists

(cytochalasin D and latrunculin B) or filamentous actin (F-actin) stabilizing inhibitors (phalloidin

and jasplakinolide), both observations shows stage-specific requirements for the actin

cytoskeletal array. Although the establishment of trichome cell polarity seems not to need precise

actin organization, the rapid expansion of trichome cells after branching is sensitive to the

inhibitors, causing an aborted, swollen stub or a highly elongated and distorted structure because

of their disorganized F-actin arrangement (Mathur et al., 1999; Szymanski et al., 1999).

The direct evidence supporting the necessity of a precise actin organization for trichome

morphogenesis comes from the discovery of the genes that encode the subunits of Arp2/3 and the

WAVE complexes in the Arabidopsis genome. The Arp2/3 complex by itself is inactive and

needs the WAVE complex to activate it. These two complexes coordinately regulate actin

polymerization, and influence F-actin reorganization in both mammalian (Goley and Welch,

2006) and plant cells (Schellmann and Hulskamp, 2005; Szymanski, 2005; Uhrig and Hulskamp,

2006). Recent work has indicated that mutations in the components of the Arp2/3 and WAVE

complexes cause a common trichome defect, resulting in distorted trichomes. In addition, the F-

actin is disorganized in the defective trichomes (Mathur et al., 2003; Basu et al., 2005; Zhang et

al., 2005b; Uhrig et al., 2007). The characterizations of the defective trichome phenotype and the

F-actin disorganization in the dis mutants are reminiscent of trichomes treated with anti actin










drugs. These uniform results strongly support a crucial role of a precise actin cytoskeleton in

trichome morphogenesis.

The Arabidopsis epidermal leaf pavement cells are another ideal model for cell

morphogenesis. These cells have a unique structure, which produces a jigsaw-like appearance.

They exhibit an interlocked arrangement that results from the interdigitation of adj acent cells

through the formation of complementary lobes and indentations. Working on these cells,

researchers discovered additional evidence supporting the critical role that the actin

reorganization plays in cell morphogenesis. ROP2, a small GTPase, is redundantly required for

normal pavement cell morphogenesis. Genetic disruption of ROP2 results in a severe decrease in

the lobe elongation of pavement cells. Additionally, Eine F-actin is also reduced at the lobes of

the defective cells (Fu et al., 2002). ROP2 promotes F-actin assembly through interaction with

RIC4, which is also expressed in leaves, and localizes preferentially at the cortical sites of

incipient lobe formation Moreover, RIC4 over-expression promotes the accumulation of Eine F-

actin and generates deep lobes. On the other hand, ric4 mutants display pavement cells with

shallow lobes. Thus, the Eine cortical F-actin at specific sites promotes outgrowth for lobe

formation (Fu et al., 2002; Fu et al., 2005).

Although the role of the actin cytoskeleton in the control of plant cell morphology is well

established, all these results do not yet offer an explanation of its molecular mechanism. Many

questions have yet to be answered. For example, why are abundant actin filaments observed in

defective trichomes of dis mutants? Why does the difference between distorted trichomes and

normal ones exist only at the late stage of trichome development? Also, tip-growing cells, such

as root hairs and pollen tubes, have a strict requirement for actin cytoskeleton. Why do they









display no defective phenotype in the dis mutants? More information is needed to put together

the elegant interdigitating mechanism of this jigsaw puzzle.









CHAPTER 2
IRREGULAR TRICHO1VE BRANCH 3 (ITB3) IS A NOVEL REGULATOR OF ACTINT
ORGANIZATION

Introduction

The actin cytoskeleton not only mechanically supports mammalian cells for the formation

of various shapes, but also generates a driving force for motility of diverse cellular or

intracellular events such as cell migration, vesicle trafficking, exocytosis, and endocytosis

(Kaksonen et al., 2006). As these events occur, actin cytoskeleton remodeling is active in the

assembly of actin filaments at specific subcellular locations (Goley and Welch, 2006). End

tracking motors (actoclampins) at the barbed end of growing actin filaments generate the

propulsive force for motile events (Dickinson and Purich, 2002; Dickinson et al., 2004).

The dendritic nucleation model of actin polymerization is well established for

interpretation of the molecular mechanisms of cell migration (Mullins et al., 1998). Human

epithelial fibroblasts and fish epithelial keratocytes are rapidly moving cells. They both form a

protrusion called a lamellipodium with a thin layer of cytoplasm containing a dense meshwork of

actin filaments. While the keratocyte migrates along the substrate surface, actin and its regulators

accumulate in lamellipodia. Actin depolymerizing factor (ADF) at the rear of the leading edge

severs and depolymerizes actin filaments and creates new plus ends for the growth of new actin

filaments at the front, where the actin nucleator ARP2/3 and its activator WASP/Scar/WAVE

polymerize actin filaments (Svitkina and Borisy, 1999). Actoclampin hydrolyzes ATP for free

energy and pushes the plasma membrane, propeling the cells forward (Dickinson and Purich,

2002; Dickinson et al., 2004). In mammalian cells, the actin tail, a comet-shaped structure of

actin filaments at the rear of rocketing cells, is responsible for driving the pathogenic bacteria

Listeria and .1/nlge// across the host cell cytoplasm using the same molecular mechanism as the

dendritic nucleation model (Cameron et al., 2000). Besides these moving cells, vesicle









trafficking also is dependent on the force generated from dynamic actin filaments. For example,

endocytic vesicles that budded from the yeast plasma membrane were observed to use a comet

tail for rapid trafficking deeper into the cytoplasm (Engqvist-Goldstein et al., 2004). Vesicle

formation also needs actin filaments, which participate in coated pit formation, vesicle

constriction, and vesicle scission (Yarar et al., 2005). During vesicle formation, the actin

filaments assembling at endocytic sites bind to dynamin through cortactin (Merrifield et al.,

2002; Merrifield et al., 2005). Dynamin and cortactin are important components of the vesicle

scission machinery. Cooperating with dynamin, dendritic actin filaments generate a strong

tension at the vesicle neck for vesicle budding (Roux et al., 2006). Cortactin may rearrange actin

filaments in specific directions (Kessels and Qualmann, 2005).

Unlike a mammalian cell, the plant cell is surrounded by a rigid cell wall, thus precluding

migration. The actin cytoskeleton is implicated in intracellular organelle motility and vesicle

trafficking, particular the Golgi apparatus (GA) movement. In mammalian cells, the GA is

located close to and around nuclei, but in plant cells, the Golgi carried by myosin, rapidly moves

along actin cable tracks throughout the whole cell (Brandizzi et al., 2003; Hawes and Satiat-

Jeunemaitre, 2005b; Latijnhouwers et al., 2005). In the tip growing cells of pollen tubes and root

hairs, a polarized actin cytoskeleton enables tip-directed organelle and vesicle trafficking.

Although pollen tubes and root hairs are two distinct cell types, they share a common

morphological form, consisting of a shank, a sub-apical zone, and an apical zone. The gradient of

actin filaments are regularly organized in these zones (Cole and Fowler, 2006; Samaj et al.,

2006). During pollen tube elongation, thick actin cables are arranged in parallel to the shank and

serve as tracks for myosin motors carrying organelles or vesicles to the growing site. In the sub-

apical zone, there is a dense fringe of actin filaments, which may promote vesicle formation from









endomembrane organelles (Cole and Fowler, 2006). In the apical zone, abundant vesicles are

embedded in the actin filament meshwork, which appears to propel vesicles to the plasma

membrane at the growing tip like the comet tail in animal cells (Hepler et al., 2001; Cole and

Fowler, 2006; Samaj et al., 2006). Root hairs demonstrate polarized characteristics similar to

pollen tubes. Abundant and highly motile endosomes were found in root hairs, and their

intracellular motility relied fully on the actin cytoskeleton. At the tip of root hairs, motile F-actin

patches have been presumed to propel endosomes to the plasma membrane (Voigt et al., 2005;

Samaj et al., 2006).

Although trichome cell expansion is not tip growth, it is a typical anisotropic diffuse

growth. It also was found that the actin cytoskeleton plays an important role in trichome

morphogenesis (Szymanski et al., 1999; Smith and Oppenheimer, 2005; Hussey et al., 2006).

Based on evidence from experiments with actin inhibitor-treated trichomes (Mathur et al., 1999;

Szymanski et al., 1999) and characterization of the actin cytoskeleton in distorted trichome

mutants (Mathur et al., 2003; Szymanski, 2005; Zhang et al., 2005b), the current hypothesis is

that the actin cytoskeleton maintains and coordinates the growth pattern established by

microtubules. MicroHilaments are expected to play the same role in trichome morphogenesis as in

tip growth, which is to deliver specific vesicles containing cargoes, such as cell wall materials to

specific sites for local growth (Mathur et al., 2002; Smith and Oppenheimer, 2005). However,

during trichome morphogenesis, a gradient of actin filaments was not found. Additionally,

microtubules were observed to act as tracks for GA transport. Mutations in KINESIN-13A cause

defects in the GA transport and result in misshaped trichomes (Lu et al., 2005). Therefore, the

precise role of the actin cytoskeleton in anisotropic cell expansion is still not known. In this

study, ITB3 was found to be a novel regulator of actin organization in Arabidopsis trichome










morphogenesis. Mutations of the ITB3 gene caused a change in trichome shape. The actin

cytoskeleton was aberrantly disorganized in itb3 mutants. Abundant rings formed by actin cables

were observed in the itb3 mutant, but never in the wild type. ITB3 was found to directly bind to

actin depolymerizing factor and inhibits it activity. These results indicated that the actin

cytoskeleton plays a crucial role trichome morphogenesis, which provides insight into the role of

the actin cytoskeleton in anisotropic cell expansion.

Materials and Methods

Plant Materials and Growth Conditions

The fast neutron induced mutant, itb3-2 7, was isolated in the Rschew (RLD) genetic

background (Zhang et al., 2005). The mutants, itb3-1 (SalK_073071) and itb3-2 (Salk_015997)

are T-DNA insertion mutant alleles in the Columbia (Col) genetic background from the SALK

T-DNA Insertion Database (http://signal. salk.edu/cgibin/tdnaexpress). The wild type used for

construction of the mapping population is the Landsberg erecta (Ler) ecotype.

Seeds were sown on a soilless potting medium, Fafard 2 Mix (Conrad Fafard, Inc.,

Agawam, MA). Seedlings were grown at 240C under constant light, provided by 40W cool white

fluorescent tubes. Plants were watered with PGP nutrient solution (Pollock and Oppenheimer,

1999) every two weeks.

Positional Cloning of ITB3

The mapping population for cloning the ITB3 gene was generated as described by Zhang et

al. (2005). Phenotypically itb3 mutant plants were selected from the F2 population. From each

selected plant, one of the cotyledons was removed for DNA extraction using the RED Extract-N-

Amp Plant PCR Kit (Sigma-Aldrich, St. Louis, MO). The isolated DNA was used to map the itb3

mutation relative to simple sequence length polymorphisms (SSLPs) (Bell and Ecker, 1994).









After the itb3 mutation was mapped to a narrow region, the expected large deletion was detected

by PCR.

Plasmid Construction

For expression of ITB3, the ITB3 gene, which covers the 1 13-bp 5' UTR, the 50 1-bp

coding sequence and the 277-bp 3' UTR was amplified using genomic DNA from RLD wild

type plants as the template using the appropriate primer pair (Table 2-1). The 891-bp PCR

product was cloned into pENTRIA (Invitrogen, Carlsbad, CA) using the BamIII and EcoRI sites,

and transferred into either pAM-PAT-GW (Bekir Ulker, Max Planck Institute for Plant Breeding,

Cologne, Germany) for expression from the 35S promoter or pCK86 (Arp Schnittger, Max

Planck Institute for Plant Breeding) for expression from the GL2 promoter through an LR

recombination. To localize ITB3, a 35S:ITB3-GFP construct was made. The 498-bp ITB3 open

reading frame was amplified using primer pairs that introduced Ncol sites at both ends of the

PCR product. The digested PCR product was cloned into the Ncol site of the GFP fusion vector,

pAVA319 (von Arnim et al., 1998). The resulting gene fusion was liberated by digestion with

EcoRI and Notl and transferred to pENTRIA through the same cut sites. Finally it was

transferred into the destination vectors, either pAM-PAT-GW or pCK86 as in the previous

constructs. Through the same strategy, for FRET assays, the 35S:ITB3CFP, 35S:ITB3L4CFP,

and 35S:ADF3YFP were constructed with pAVA574 containing CFP and pAVA 554 containing

YFP. To produce the ITB3 protein, the 501l-bp coding sequence of ITB3 was cloned into pET4 1

Ek/LIC, pET 15b through Ndel and BBBBBBBBBBBBBBBBBamII or pET41a through M~fel depending on the specific

tag needed (Novagen, Madison, WI). For pET41a-ITB3-GFP, the PCR products were amplified

using primers containing EcoRI sites at both ends of ITB3-GFP.

To construct pDBLeu-ITB3, used as bait for the yeast two-hybrid screen, the 501 bp

coding sequence of ITB3 was cloned into pDBLeu with kanamycin resistance (provided by Wen-









Yuan Song at the University of Florida) through NotI and Sall sites. All PCR products were

sequenced by the Interdisciplinary Center for Biotechnology Research (ICBR) at the University

of Florida to ensure no mutations were introduced.

RNA Extraction and RT-PCR

Total RNA was extracted from six-week-old Col wild-type plants using the RNeasy Plant

Mini Kit (Qiagen Inc. Valencia, CA) according to the manufacturer's instructions. The full

length ITB3 cDNA was amplified, following instructions for the cMaster RT plus PCR System

(Eppendorf AG, Hamburg, Germany). First-strand DNA synthesis was primed using oligo

(dT)20. The cDNA was amplified using the specific primers for ITB3 (see Table 1). The PCR

products were sequenced by ICBR at the University of Florida.

Plant Transformation

For ITB3 subcellular localization, the 35S:ITB3-GFP constructs were transferred into

onion epidermal cells by particle bombardment, using the Biolistic PDS-1000/He Particle

Delivery System (Bio-Rad, Richmond, CA), and the transformation protocol supplied by the

manufacturer was followed. A total of 5 CIL of DNA (1Cpg/CLL) was precipitated on 3 mg gold

microcarriers 0.6 Clm in diameter (Bio-Rad), by adding 50 CLL of 2.5 M CaCl2 and 20 CIL 0.1 M

of spermidine. After the precipitated DNA was washed once with 140 CLL 70% and once with

100% ethanol, it was resuspended in 50 CIL of 100% ethanol. 10 CLL of this solution was spread

on one rupture disk labeled with a burst pressure of 1,100 psi. Square tissue sections

approximately 2 x 2 cm were cut from onions and placed on Murashige and Skoog (MS) solid

medium for bombardment. Fluorescence was visualized after 36 hours incubation at room

temperature in darkness.

The constructs of 35S:ITB3, GL2:ITB3, 35S:ITB3-GFP, GL2:ITB3-GFP, and 35S:ITB3-

CFP were transferred into itb3-2 7 mutants by the floral dip method (Clough and Bent, 1998).









35S:ITB3L4CFP was transferred into the itb31-4 mutant, 35S:ADF3YFP was transferred into

adf3 mutants. The transgenic plants were selected using a 1000X dilution of Finale (Farnam

Companies Inc, Phoenix, AZ) with 5.78% glufosinate-ammonium.

Yeast Two-hybrid Assays

The protocol for the yeast two-hybrid assay was described by (Ding et al., 2004). The bait

construct pDBLeu-ITB3 was transferred into the yeast strain CGl945 through the Leu selection

marker using the Yeast Transformation Kit (Sigma, St. Louis, MO) according to the

manufacturer' s instructions. The prey, pPC86-cDNA, was a rice cDNA library, (provided by

Wen-Yuan Song at the University of Florida) with Trp as the selection marker in yeast strain

Y187. The mated cells of CGl945 and Y187 were spread on the YPD medium without Trp, Leu,

and His for positive selection. The plasmids were isolated from the grown yeast colonies with

Zymoprep Yeast Plasmid Miniprep (Zymoprep, Orange, CA 92867) and transferred into XL2-

Blue Ultracompetent Cells (Stratagene, LaJolla, CA). The genes of interest were sequenced using

plasmid DNA from individual bacterial colonies.

Protein Isolation

The constructs, pET41 EK/LIC-ITB3, pET15b-ITB3, and pET41a-ITB3, were transferred

into the host cell BL21 (DE3) (Novagen, Madison, WI) through chemical transformation for

ITB3 expression. The single colony with the target construct was inoculated into 500 ml

Overnight Express Instant TB Medium (Novagen, Madison, WI) for overnight culture. The

harvested host cells were lysed by lx FasBreak Cell Lysis Reagent (Promega, Madison, WI). For

pET15b-ITB3 with the 6xHis-tag, HisLink Protein Purification Resin (Promega) was used for

His-tagged ITB3 binding. For pGEX-profilin, pGEX-ADF 1 (provided by C. Staiger at Purdue

University) and pET41-ITB3, which the proteins of interest were tagged with GST, the GST

Binding Resin was used for the protein purification following the manufacturer' s instructions for









the BugBuster GST-Bind Purification Kit (Novagen). The 6xHis-tag was removed from the

fusion ITB3 with a Thrombin Cleavage Capture Kit (Novagen) and the GST tag was removed

using an Enterokinase Cleavage Capture Kit (Novagen) following the manufacturer' s

instructions. The concentration of the purified proteins was measured using a DC Protein Assay

Kit (Bio-Rad) following the manufacturer's instructions.

Pull-down Assay

The purified proteins in lx PBS were diluted to a 1lyg/C1l concentration with lx PBS. The

binding reaction was done in 100 Cll of binding buffer, which contained 5 mM Tris-HC1,

100 mM KC1, 1 mM MgC1, 1 mM CaCl2, 50 ng/CIl 6xHis-ITB3, and 100 ng/CIl ADF 1 or profilin

or BSA, pH 7.5. The reaction was allowed to proceed at room temperature for 80 minutes. After

the binding was completed, 12.0 Cll of 10x nickel resin binding buffer containing 1 M HEPES

and 100 mM imidazole (pH 7.5) and 8.0 Cll of 50% HisLink Protein Purification Resin

(Promega) were added to the binding buffer. The reaction tubes were rotated at 12 rpm at room

temperature for 60 min. The resin was spun down and washed three times for 10 minutes, with

washing buffer containing 100 mM HEPES, 10 mM imidazole, and 0. 1% NP-40. 3.0 Cll

NuPAGE LDS Sample Buffer (4x) (Invitrogen, Carlsbad, CA) was added to the pellet resin

suspension (~10.0 Cl). After a 5 minute heating at 95oC, the samples were loaded into NuPAGE

12% Bis-Tris Gel (Invitrogen, Carlsbad, CA) for protein separation.

Morphological Analysis

SALK lines (SALK_073071, 015997, 019320, 008148, 001114, 001117, and 019328),

which were ordered from Arabidopsis Biological Resource Center (The Ohio State University,

Columbus, OH) were examined under a dissecting microscope, and lines SALK_073071 and

015997 showed segregation of plants with the itb3 trichome phenotype.









Immunostaining of the Actin and Microtubule Cytoskeletons

The immunostaining protocol for actin filaments and microtubules, using specific

antibodies against tubulin or actin, were described in our previous studies (Zhang and

Oppenheimer, 2004; Zhang et al., 2005b)

Microscopy

Fluorescent images were collected with a Zeiss Axiocam HRm camera mounted on a Zeiss

Axioplan 2 Imaging microscope (Jena, Germany). The following filter sets were used to collect

fluorescent images: red fluorescence was obtained with Zeiss filter set 20 (excitation, 546/12;

dichroic, 560 LP; emission, 575 to 640), green fluorescence was obtained with Zeiss filter set 10

(excitation, 450 to 490; dichroic, 510 LP; emission, 515 to 565). Optical sections were collected

using the Zeiss Apotome and Axiovision 4. 1 software. Light micrographs were collected with a

Zeiss Axiocam MRc5 camera mounted on a Zeiss Stemi SV11 dissecting microscope. For

scanning electron microscopy (SEM), previously described methods were used (Luo and

Oppenheimer, 1999).

Double Mutant Construction

The double mutants were selected from individual F2 plants. The itb3-2 7 mutant was

crossed with the T-DNA insertion lines, itb31-4 and adf3. The selected putative double mutants

were self pollinated. These F3 individuals were crossed with their original parents to check for

mutations through exhibition of their specific mutant phenotypes.

Results

Cloning of the ITB3 Gene

The mutation in the itb3-2 7 mutant was mapped between the markers s2 (to BAC clone

T5Al4) and s3 (to BAC clone T30E16) on chromosome I (Zhang et al., 2005a). To clone the

ITB3 gene, we further mapped the itb3 mutation relative to simple sequence length









polymorphism (SSLP) markers (Bell and Ecker, 1994) to BAC clone F25Pl2 (Figure 2-1).

Because the itb3-2 7 allele was isolated from a fast neutron mutagenized population, we screened

BAC clone F25Pl2 for deletions by amplifying short regions spaced approximately every

1000 bp along F25Pl2. An approximately 42-kb deletion was found between the 92.4 and

134.4 kb positions in F25Pl2, but the deletion did not occur in wild type. In the deleted region,

there were 16 putative genes (Figure 2-1).

The SALK T-DNA insertion database was searched for insertions in these genes, and the

resultant insertion lines (Alonso and Stepanova, 2003) were screened for plants showing the itb3

trichome phenotype. Salk_073071 segregated plants that had strong itb3 phenotypes;

Salk_15997 line also segregated plants with a weak itb3 trichome phenotype. These three lines

all had a T-DNA insertion in Atlg56580. The itb3-2 7 mutant was crossed to these three lines for

complementation tests. The results indicated that the insertion mutations in the three lines were

allelic to itb3 (data not shown), which demonstrated that ITB3 is Atlg56580.

The trichomes on the leaves of all three itb3 mutant alleles fit the criterion set for irregular

trichome branching, where at least one branch that is shorter than the others in length, or at least

two branch points positioned separately on the stalk. Additionally, the trichome size and branch

number of itb3 mutants decreased when compared with wild type (Zhang et al., 2005a). To

determine why these defects occur in itb3 mutants, we examined the cytoskeletons in itb3 and

wild type trichomes by immunostaining with antibodies against actin and tubulin. Compared to

wild type, the microtubules in itb3 trichomes showed no apparent difference (data not shown),

but the actin filaments in itb3 trichomes were disorganized (Figure 2-2). In trichomes of

developmental stage 2-3, the actin cables in itb3 mutants accumulated near the bottom of the

stalk (Figure 2-2C), whereas in wild type more actin cables were distributed close to the top of









the trichome (Figure 2-2B). At stage 3, the disorganization of actin cables in itb3 mutants was

more pronounced. Most of the actin cables were parallel to the long axis of the stalk in the itb3

trichomes (Figure 2-2F), whereas in wild type actin cables were more abundant in the region

between branches (Figure 2-2E). At stage 4, more disorganized actin cables were arranged under

the trichome branch points in the itb3 mutant (Figures 2-21, 2-3A) compared with wild type

(Figures 2-2H, 2-3B). The greatest difference was the formation of actin rings in itb3 mutants

after stage 3, but these actin rings were rarely found in wild type (Figures 2-3I, 2-4).

ITB3 is a Plant-specific Gene

ITB3 encodes an unknown protein with a mass of about 18 KD and a pl of 6.64. No signal

peptide or other known motif was found in the ITB3 protein sequence. Using ITB3 to search the

complete Arabidopsis genome, we found a family consisting of 22 members. This family can be

grouped into two distinct clades: the ITB3-Like (AtlTB3L-02-05) clade in the green square and

ITB3 Related (AtlTB3R-01-13) clade in the yellow square, with 100% bootstrap support for their

occurrence (Figure 2-5). The search of all available genomes of other organisms indicated that

ITB3 is a plant-specific gene that is present in all land plants, including moss (Figure 2-6).

ITB3 Over-Expression Did Not Generate Novel Phenotypes

Over-expression oflTB3 using the 35S promoter completely complemented the defective

trichome phenotype and no additional phenotypes were found. Over-expression of ITB3 with the

same construct in wild type did not display any visible changes in the trichome shape or other

phenotypes (data not shown).

ITB3 Has No Specific Subcellular Location

When the ITB3-GFP fusion construct was transferred into onion epidermal cells through

particle bombardment, the GFP signal was located in the cytoplasm and nucleus of transformed

cells and was both indistinguishable from that of the control of GFP alone and remarkably









different from the control of the nucleus-localized GFP (Figure 2-7). The same gene fusion was

used to stably transform itb3 mutants. The signal distribution in the trichomes was the same as

that in the transgenic onion epidermal cells. Additionally, the itb3 mutant trichomes were

rescued into wild type trichomes (data not shown), which indicated that the ITB3-GFP carried

out normal functions in living cells.

ITB3 Interacts With ADF3 in Yeast

To search for ITB3 interactors, a yeast two-hybrid screen was performed using a rice

cDNA library as the prey and ITB3 as the bait. On the plate without Trp, the yeast containing the

bait plasmid, pDBLeu-ITB3, grew, but those containing the prey plasmids, pPC86-ADF3 and

DCD, did not. On the plate without Leu, the yeast containing the prey plasmids pPC86-ADF3

and DCD grew, but those containing the bait plasmid, pDBLeu-ITB3, did not. On the plate

lacking both Trp and Leu, the yeast with both bait and prey plasmids grew. Additionally, this

yeast also grew on the plate without Trp, Leu, and His (Figure 2-8). This result indicated that His

was produced by transcription activation through ADF3 or DCD bound to ITB3. Therefore, rice

ADF3 and DCD proteins are able to interact with Arabidopsis ITB3 in yeast cells.

ITB3 Directly Binds with ADF3 in Vitro

To further confirm the interaction between ITB3 and ADF3, we carried out a pull down

experiment. The purified His-tagged ITB3 pulled down GST-tagged AtADF 1 in vitro, but did

not pull down GST-tagged profilin (Figure 2-9). This data further supported the results from the

yeast two-hybrid assay.

The Trichomes are Defective in the Mutants of adf, itb31-4 and Their Double Mutants

To search for additional genetic evidence that ADF is involved with ITB3 in controlling

trichome development, the knockout lines of ADF and other ITB3 family members were

examined for trichome phenotypes. A T-DNA insertion in ITB3L-4 gave rise to trichomes with









fewer numbers of branches compared to wildtype. Plants homozygous for a T-DNA insertion in

ADF3 showed caused an increase in trichome branch number (Figure 2-10).

Double mutants of adf3 and itb3 displayed trichomes with more branches than itb3 mutants

and fewer branches than adf3 mutants. The mutation of ITB3 is additive to the mutation of ADF.

However, double mutants of adf3 and itb31-4 displayed trichomes with fewer branches than their

parents. Double mutants of itb3 and itb31-4 had much more severe decreases in branch number

compared to either parent. Therefore, these two mutations appear to be synergistic in controlling

trichome branching.

The transgenic plants of the itb3-2 7 mutants with 35S:ITB3GFP or 35S:ITB3CFP

displayed the wild-type trichome phenotype. However, the transgenic plants of either the itb31-4

mutants with 35S:ITB3L4CFP or the adf3 mutants with 35S:ADF3YFP exhibited mutant

trichome phenotypes. These results led us to devise the constructs that contain their own

endogenous promoters for driving the fusion gene expression in the future (data not shown).

Discussion

Disruption of Actin Cytoskeleton Organization Leads to Misshapen Trichomes

Pharmacological disruption of actin filaments with actin-specific drugs severely affects

trichome morphogenesis and changes trichome shapes (Mathur et al., 1999; Szymanski et al.,

1999). When the Arabidopsis trichomes were treated with the actin polymerization inhibitor

Latrunculin B, no effect was found at the initial developmental stages, but after the branching

events, the trichome cells rapidly expand. The inappropriately extended trichomes generate

distortion, shortened branches, and separate branch points. Genetic disruption of the actin

filament dynamics through loss-of-function mutations of the distorted class of Arabidopsis genes

gives rise to the aberrant trichome phenotypes, similar to those in the pharmacological

experiments. The distorted class genes encode subunits of the Arp2/3 complex and its activation










complex. Mutations in the distorted class genes disrupt actin filament dynamics, which causes an

altered distribution of cortical actin cables (Mathur et al., 2003). The dis mutants display changes

of overall trichome shape, a reduction of branch length, and an increase in the distance between

two branch points (Basu et al., 2005; Zhang et al., 2005b). These changes are similar to the

trichome defects seen in itb3 mutants. Although the distortion defect was considered as the

criterion for mutations in the DIS class genes (Hulskamp et al., 1994), it disappears in some

genetic backgrounds. For example, the itbl-1 allele, (dis3) does not display the distorted

trichomes in the Wassilewskij a (WS) genetic background. However, this mutant still exhibits the

itb trichomes (Zhang et al., 2005b). In itb3 mutant trichomes, the actin cytoskeleton is severely

disorganized. Therefore, our results strongly support the idea that the actin cytoskeleton has a

crucial role in the regulation of trichome morphogenesis.

The Precise Role of the Actin Cytoskeleton in Trichome Morphogenesis

A wealth of cellular observations indicates that a fine actin filament meshwork or diffuse

actin patches promote cell expansion, whereas dense actin bundles or actin cables serve as

structural scaffolds or tracks for myosin-based motors to transport organelles and vesicles. The

precise role of actin filaments in cell expansion is dependent on actin types and subcellular

locations. Our results show that the fine actin filament meshwork is abundant close to the rapidly

growing sites of normal trichomes. However, at rapid growth stages, the itb3 mutant trichomes

did not show this meshwork. We hypothesize that these fine actin meshworks promote cell

expansion. Similar actin patches have been observed in other cell types, such as root hairs

(Baluska et al., 2000) and elongating lobes of epidermal pavement cells (Fu et al., 2005).

Although dense actin cables were observed in both wild type and itb3 mutants, many of the actin

cables formed rings in itb3 mutants. These rings are unlikely to be normal tracks for vesicles. It

is conceivable that the rings caused reduced and misdirected delivery of vesicles to the cell









cortex, ultimately resulting reduced growth. This might be a reason why the trichome size of itb3

mutants is reduced when compared to normal trichomes.

Actin Filament Reorganization Is Required for Cell Expansion

In response to internal and external signals for cell expansion Actin filaments are rapidly

remodeled by multiple regulators. Some of the regulators modulate the size and activity of the

monomeric actin pool through interaction with actin monomers. Others change the disassembly

property of filamentous actin through binding to the filament sides (Staiger and Blanchoin, 2006).

Actin and its regulators establish a complex and adjustable system for plant cells in various

environments. When the delicate balance is impaired by loss of function of some regulators,

unusual consequences occur in actin assembly and cellular architecture. Additionally, recent

findings suggest that a class-specific interaction of actin with its regulators exist for proper

remolding of the actin filaments (Kandasamy et al., 2007). ADF is a key regulator of actin. It

binds with both actin monomers and filaments. ADF severs filaments, thereby generating new

barbed ends for subunit addition (Staiger and Blanchoin, 2006). ADF severing activity is

regulated by profilin (Didry et al., 1998), ACP1 (Bertling et al., 2004; Chaudhry et al., 2007),

AIP1 (Okada et al., 2002; Mohri et al., 2006), phosphorylation (Huang et al., 2006), and pH

(Gungabissoon et al., 2001; Chen et al., 2002). Our results show that plant ADF binds with ITB3.

Additionally, ADF activity is inhibited by ITB3 binding (Oppenheimer and Grey, unpublished

data). Our results provide new insight into differences in actin filament dynamics between plant

cells and animal cells, which in plant cells is the high ratio of monomeric to filamentous actin

(Snowman et al., 2002; Wang et al., 2005a).

ITB3 is a Plant-Specific Regulator of Actin Organization

A search of all known organisms shows ITB3 is present only in the plant kingdom.

Although actin and ADF both are conserved in all eukaryotic cells, only 3 1 of the 67 animal









actin-binding proteins appear to be conserved in plants (Hussey et al., 2002). In addition, plants

have a higher percentage of monomeric actin in the total actin pool (Gibbon et al., 1999;

Snowman et al., 2002; Wang et al., 2005a). Therefore, plant-specific actin regulators like ITB3

may provide the functions of the animal proteins that are missing from plants.

Future Perspectives

Plant cell shapes are controlled by the cytoskeleton. Our data show that Irregular Trichome

Branch 3 (ITB3) is a novel regulator of actin cytoskeleton organization. Mutations in ITB3

caused disorganization of the actin cytoskeleton in trichomes resulting in an altered trichome

shape. We showed that ITB3 interacts with and negatively regulates the function of actin

depolymerizing factor (ADF) in vitro. However, two important questions remain: First, how do

ITB3 and ADF work together to regulate actin organization, and second, how does this

interaction lead to site-specific cell expansion to generate normal trichome shape.

To answer the first question, we need additional information on the molecular mechanism

of ITB3 interaction with ADF. For example, does ITB3 regulate ADF binding to actin filaments?

Does ITB3 regulate ADF phosphorylation or dephosphorylation? To address these questions, we

can use actin depolymerization assays and actin filament-severing assays with purified ADF, in

the presence or absence oflITB3. In vitro polymerized actin filaments can be labeled by

including pyrene-labeled actin monomers in the assay, and fluorescence microscopy can be used

to visualize the actin filaments. The kinetics of F-actin depolymerization can be monitored by

continuous pyrene fluorescence measurements by using a Cary Eclipse fluorescence

spectrophotometer (Yokata et al., 2005). Although we have shown convincingly that ITB3

interacts with ADF in vitro, it is important to confirm this interaction in plan2ta. To this end,

Foirster resonance energy transfer" (FRET) can be used. An ITB3-CYP fusion can be used as the

donor; an ADF-YPF fusion can be used as the acceptor. The gene fusions are transferred into









Arabidopsis leaf epidermal cells through biolistic bombardment, and the FRET efficiency can be

determined. The higher the FRET efficiency, the higher the number of ITB3 and ADF

interactions.

Also, we know that both ADF and ITB3 are members of moderately sized families in

Arabidopsis. It is possible that there are family member specific interactions between ITB3 and

ADF proteins. Plant ADF family members are grouped into the vegetative and reproductive

ADFs based on expression pattern. There are sequence and activity differences between the two

groups of ADFs. The ITB3 family has been divided into two groups based on protein sequence

differences. Can ITB3 family members also be organized into vegetative and reproductive

groups? This can be done by examining the expression of the ITB3 family members in

microarray data sets. To address the binding of specific ITB3 members with specific ADF

members, two approaches can be applied. The first approach is to determine expression patterns

of ITB3 and ADF family members in trichomes using in situ hybridization. If two members are

co-expressed in specific tissues or organs, they may be binding partners. The second approach is

to use the yeast two-hybrid assay. Putative positives from this assay can be confirmed with pull

down assays in vitro, and in planta using FRET analysis.









Table 2-1. Primers used in this study


Primer name
F6D8-25F
F6D8-25R
Fl2Pl9-26F
Fl2Pl9-26R
F5Il4-55F
F5Il4-55R
T30E16-57F
T30E16-57R
F7F22-17F
F7F22-17R
F25Pl2-14F
F25Pl2-14R
F23H11-39F
F23H22-39R
T14Gll-12F
T14Gll-12R
T5Al4-14F
T5Al4-14R
T6H22-11F
T6H22-11R
ITB3 enF
ITB3enR
ITB3GSTpF
ITB3GSTpR
ITB3HISpF
ITB3HISpR


Sequence (5' 3')
CTACATTTGTTCATATACAGGGAGTTC
GCCGAGATATACTTGGATCATACTG
CTGGAAATATCTGCGAAGTGGAA
CATGAACTGTTTGTGCATCTCTG
CGGATGCGGTTATATAAATAGAGA
CCCTCCCTTTTCTTGCTACAAA
ACACTCTTTACTGGAAGATGCAA
AACACACCCATGCAAGTGAA
GCTCACACTTTCCAATGGTGT
CCTTGGAAGCGTAGACCCA
GCACGATCCTATGAGTTAGCA
TTACACGCGAGGAATGAAGA
TTTGATGGAGATTTTGCTGATT
ACCATTGACAGTGGAGCTACATT
TTTGGATGGATTTGTGCGTG
CGATGAGGTCAATCCTAAAGATCAG
GACCAATACAGAGATACAAAGCAA
TCCGCTAACTTATCCGACAA
GACAATTTTCTTC TATATAAGGATGTGG
GGTCATCCTTGCAAGATATCAA
GGGTGATTTCATACCACACCACC
TGGCTATGAAGTAACCGCTGAGAT
ATGGGTTTGG TTACAGATGAAGTG
AGCATCTGTGACTGCAACAGCTTC
ATGGGTTTGGTTACAGATGAAGTGAGAGC
CTAAGCATCTGTGACTGCAACAGC


Used for*
RLD 134/Ler 109

RLD 119/Ler 93

RLD 231/Ler 176

RLD 138/Ler 81

RLD 82/Ler 95

RLD 101/Ler 87

RLD 119/Ler 80

RLD 81/Ler 69

RLD 89/Ler 103

RLD 127/Ler 116

ITB3 over expression

GST-ITB3 protein

His-tagged ITB3


*Note: A maj ority of primers are used for ITB3 positional cloning, for example, RLD
134/Ler 109 is a SSLP marker that has PCR products of 134 bp in RLD wild type and 109
bp in Ler wild type.











Chromosome 1
about 42kb between 21187150 and 21228933


I I


nga


1248 nga240
I I


Recombinants and BACs~

1770 256 52 20 7 0 20 34 128 157
Total F7F22 F6D8 T5A14 T6H22 F25P12 T30E16 H23H11 F5114 F12P19
mmmm~ DM M


710 nga248

Genes in the
deleted region/


240 ngal11


a
r~~uo~


HlrlU~U(tll)ll
q~rrrr clrrll
~C


Rft~ll


Figure 2-1. Positional cloning of ITB3. The itb3 mutation was mapped near SSLP marker
ATPase on chromosome 1. Additional molecular markers were used to map the itb3
mutation to BAC clone F25Pl2. The numbers of recombinants (out of 1770
chromatids screened) are given above BAC clones. The locations of all putative genes
on BAC clone F25Pl2 are listed. The numbers inside the flags above specific genes
are numbers of recombinants for that marker. ITB3 is indicated by the vertical arrow.







































Figure 2-2. Actin cytoskeleton is disorganized in the itb3 mutant. (A), (D), and (G) are wild-
type trichomes showing different developmental stages through scanning electronic
microscope (SEM). (B), (E), and (H) are wild-type trichomes showing the normal
organization of actin cytoskeleton with the identical developmental stages with (A),
(D) and (G). (C), (F), and (I) are the itb3 mutant trichomes at the same developmental
stages as wild type showing disorganization of the actin cytoskeleton through
immunostaining and fluorescent microscopy. Bar = 100 Clm in all images.































Figure 2-3. Actin cable organization in the stalk of trichomes. (A) The itb3 mutant trichome
stalks showing clear differences in actin organization compared to the wild type in
(B). Bar = 10 lm.





























Figure 2-4. Actin rings in the itb3 mutant. Bar = 5 Clm









0.2


10 AtlTB3_At1g56580
70 Atl~ ITB3 L-01~_At 1 g 3 1 0

3 -AtlTB3L-02_A15g49600

100[ ~AI AtlT3 L-03 At4g241 30

SS- tlTB3L-043_A 1g30020]
AtllB13L-05 .AthodB230.
."lT3R04AT2GO3350

AitTB3R-03_AT5G37070D
100
AtT3-02 AT3008890

761 AtTB3R-01_t5 01610
At tTB8R-14_AT1 G55265

itTB3-09_AT5G1 9860

ft1 AtlTB3R-16_AT1G61667~

90 :1TB8-15 AT5G54530
lBR10_AT5G116380
71AtlTB8iR-11 AT3G074i60
100[
AtT3-08_AT3GO7470

itT3-12_AT5G19590

85 5 ,, tlTfB3R-13_AT1GO2813
97 AlTB3-0'7_AT4tGO2360

10At lTB3R~-06I_AT-IGO23 70
AtTB3~R-05_AT1GO2816




Figure 2-5. Phylogenic tree of the Arabidopsis ITB3 family members














~cl~lm~p ~ I8S$
Bime ITORI
Rice I~aki
mas! ITHR1
Con~sensu#75%.

.W18J~(LSb 1783
Pine i 1
Rica IT~Ki
m~a~s TBP1.1
Consenusurd7%

Atashiopsis~ IT83
Pine ITEL1
Rice ITB3L1
Woe~ ITB$1
CO~ea6sual?$%

Ateeidommis ITR
Prine ITOL1
RiPe FTBai
mass IT831.1
Conrsensual75%


. BL- ,Tp..? 2. -..LE 99it:' SE- ----- R: TLES. SM:-ol- Ir -~ r r. gP y ,pay:
s.:v., .2 Q 8;:..tS. .6 .Too 9------- !c ke~ iA es --. .IE rco vlr L.':Ae
I ..x TS ,g X;p ER, 1.TG ALE I :9ATRSD SP E C2FL DL ~~.~T- .1-F L, ~ -I.~ PI.C: J B
MAL. 1A. -.+9A11ia. .uaRe......... K .ALLEP .Lexe~ILLE 1.5.0..

:2r r.1.#!.9'Tl-Ax WA:. j t A1 5. r. b :Y


TOrywaX~KxR.E.xl.r..PGK.VSY.TEZITAall .Gkl LTV. l[ .51.e


D PS~ .ARI WTEESTOIESFTSAE25Wea RAC9 CT or :P V .J .

_ qansax-- ----------- ---~-------- r, .r ., E. ***. .TI .*P-.- Q--Q IB"
- ... ................ ............RT. + GiL S~rt SA~FLL.14 P;..





W............A. .v....


Figure 2-6. Alignment of ITB3 protein sequence with its homologs in other plants






























Figure 2-7. ITB3-GFP is not specifically localized to any subcellular structure in transformed
onion epidermal cells. (A): GFP alone; (B): GFP-NIa in nucleus; (C): ITB3-GFP.
Bar = 5pm
































Figure 2-8. Yeast two-hybrid screen for ITB3 interactors. (1): Bait ITB3; (2): bait ITB3 and
prey ADF; (3): prey ADF; (4): pre DCD; (5): bait ITB3 and prey DCD; (6): positive
control .


Trp -


Leu -


Trp Leu -


Trp Leu His -










M 1 2


M Mr
S52
--31


S1"1


GS T-AD F 1
GST-profi lin
6xH is- ITB3-+


."


""


Figure 2-9. ITB3 directly interacts with ADF in vitro. (1): GST-ADF; (2): the GST-ADF is
pulled down by His-ITB3; (3): the GST-profilin fails to be pulled down by His-ITB3;
(4) His-ITB3 alone; (5): GST-profilin alone.


m e n ~
























Figure 2-10. Trichome shapes are defective in adf3 and itb31-4 mutants. (A) and (D): Col wild
type; (B) and (E): adf3; (C) and (F): itb31-4. Bar = 100 Clm


rZ~
IE_~x~i~1 ,F









CHAPTER 3
IRREGULAR TRICHOME BRANCH 2 (ITB2) IS A PUTATIVE
AMINOPHOSPHOLIPID TRANSLOCASE THAT REGULATES TRICHOME
BRANCH ELONGATION INT ARABIDOPSIS

Introduction

Lipids are the maj or components of all eukaryotic membranes. They mainly include

phospholipids, sphingolipids, and sterols, which are distributed asymmetrically within

bilayer membranes. Phospholipids form the main homogenous planar architecture, but

sphingolipids and sterols are rich in microdomains called lipid shells/rafts, which

theoretically float freely in the more fluid surrounding membranes analogous to the so-

called "liquid disordered" phase. Distinct lipids also specifically localize the two leaflets

of membranes. In general, the aminophospholipid, phosphatidylserine (PS), and

phosphatidylethanolamine (PE) are concentrated in the cytosolic leaflet, whereas

phosphatidylcholine (PC) and sphingolipids are enriched in the exoplasmic leaflet

(Holthuis and Levine, 2005; Pomorski and Menon, 2006).

The asymmetric distribution of lipids is generated by energy-dependent flippases that

hydrolyze ATPs for energy to translocate specific lipids across the lipid bilayer (Pomorski

and Menon, 2006). The P-type ATPase is such a flippase and its translocase activity was

found not only in the plasma membrane (PM) (Pomorski et al., 2004), but also in the

membranes of distinct vesicles (Zachowski and Gaudry-Talarmain, 1990; Alder-Baerens et

al., 2006) and the trans Golgi network (Nataraj an et al., 2004). In yeast, the DRS2 gene

codes for Drsp, a member of the P4-ATPase family in the P-type ATPase superfamily

(Ripmaster et al., 1993). Mutations in DRS2 caused an absence of low temperature uptake

of a labeled PS analog at the PM (Tang et al., 1996; Gomes et al., 2000). Additionally, loss

of the Golgi-associated P4-ATPases Drs2p and Dnf3p abolished the asymmetric









arrangement of endogenous PE in post-Golgi secretary vesicles (Alder-Baerens et al.,

2006). These findings indicated an essential role for P4-ATPases in generating and

maintaining lipid asymmetry during membrane flow through the Golgi. Additional P4-

ATPase family members also have been identified, and they are all associated with lipid

translocation in other species (Ujhazy et al., 2001; Perez-Victoria et al., 2003; Wang et al.,

2004), including plants (Gomes et al., 2000).

P4-ATPase is involved in vesicle formation. During the biogenesis of intracellular

transport vesicles, lipids need to translocate from the inner leaflet to the outer one by

flippases in membranes (Pomorski and Menon, 2006). In yeast, the absence of the two PM

associated P4-ATPases, Dnflp and Dnf~p, resulted in a cold-sensitive defect in the

biogenesis of endocytic vesicles (Pomorski et al., 2003) and inactivation of Drs2p caused a

decrease in clathrin-coated vesicle budding from the trans-Golgi (Gall et al., 2002;

Nataraj an et al., 2004). Stimulation of PS and PE inward translocation induced the

formation of endocytic vesicles in red blood cells (Birchmeier et al., 1979; Muller et al.,

1994). Conversely, enhancement of outward directed lipid translocation led to a defect in

endocytosis (Kean et al., 1997; Decottignies et al., 1998). The role of flippases in vesicle

biogenesis was considered a direct and mechanical action on vesicle budding. P4-ATPases

interacted with such cytosolic proteins as guanine nucleotide exchange factors (GEFs) and

small GTPases, which are crucial for the recruitment of such coat proteins as clathrin at

sites of the lipid translocation (Pomorski and Menon, 2006; Liu et al., 2007). The

membrane curvature, which is generated by lipid translocation that creates an area

difference between the two leaflets, promotes vesicle budding (Pomorski and Menon,

2006).









Vesicle-mediated membrane trafficking plays a crucial role in cell expansion. Polar

expansion such as elongation of pollen tubes and root hairs requires vesicles for

transporting materials to build cell walls (Samaj et al., 2006). Abundant vesicles are

transported and deposited cell wall molecules at growing sites. Normal vesicle trafficking

is also necessary for other cells to initiate or execute anisotropic expansion. Aberrant

vesicle trafficking caused lobe reduction ofArabidopsis epidermal pavement cells and

shape changes of trichomes (Zheng et al., 2005). In this study, we cloned the Arabidopsis

ITB2 gene through a map-based strategy. ITB2 is identical to ALA3, a member of the

putative aminophospholipid translocase (ALA) subfamily in the P4 ATPase family in the

P-type ATPase superfamily. Mutations in itb2 mutants reduced the trichome branch length.

We provide the evidence that plant P4 ATPases regulate cell expansion, likely through

contribution to the vesicle formation.

Materials and Methods

Plant Materials and Growth Conditions

The itb2-28, itb2-29, and itb2-12 (9412-12) mutants were isolated in the Rschew

(RLD) genetic background (Zhang et al., 2005). The itb2-4 (Salk_015929) mutant is a T-

DNA insertion line in the Columbia (Col) ecotype, based on the SALK T-DNA Expression

Database (http://signal.salk. edu/cgibin/tdnaexpress). The wild type used for construction of

the mapping population is the Landsberg erecta (Ler) ecotype. Seeds of the SALK lines,

SALK 015929, 006470, 067322, 139762, 129494, 133319, 082561, 066531, and 109350,

were ordered from the Arabidopsis Biological Resource Center (The Ohio State University,

Columbus, OH).

Seeds were sown on a soil-less potting medium, Fafard 2 Mix (Conrad Fafard, Inc.,

Agawam, MA). Seedlings were grown at 240C under constant light, provided by 40W cool









white fluorescent tubes. Plants were watered with PGP nutrient solution (Pollock and

Oppenheimer, 1999) every two weeks.

Positional Cloning of ITB2

The mapping population for cloning the ITB2 gene was generated as described by

Zhang et al., (2005). The phenotypically itb2 mutant plants were selected from the F2

generation of plants. From each selected plant, one of the cotyledons was removed for

DNA extraction using the RED Extract-N-Amp Plant PCR Kit (Sigma-Aldrich, St. Louis,

MO). The isolated DNA was used to map the itb2 mutation relative to simple sequence

length polymorphisms (SSLPs) (Bell and Ecker, 1994; Lukowitz et al., 2000). Because the

itb2 mutant was isolated from a fast neutron mutatgenized population, after the itb2

mutation was mapped to a relatively small region, the presence of a deletion was tested by

using itb2-28 DNA as the template to amplify about 500 bp target fragments. After the

deletion was found, all the genes in the deleted region were sequenced to check for

mutations using PCR products amplified from itb2-19 DNA.

Plasmid Construction

To complement the itb2 mutant, the 2146 bp element at the 5' end of ITB2 replaced

the 35S promoter on pAM-PAT-GW through XSbol and Asci, and the 1380 bp-element at

the 3' end of ITB2 was directionally cloned into the pAM-PAT-GW backbone using the

PstI restriction site. Because of the large size of the ITB2 genomic sequence, its cDNA was

used for construction. The full length coding sequence of ITB2 cDNA was amplified by

RT-PCR from total RNA of from Col wild type. The PCR fragment was cloned into the

pBluescript SK II (+) vector. The amplified PCR products were sequenced to identify any

mutations that were introduced during PCR. The mutations in the cloned cDNA of ITB2

were corrected in the Col wild-type version by multiple substitutions with the correct PCR









fragments. The resultant wild type ITB2 cDNA was cloned into pENRIA (Invitrogen,

Carlsbad, CA) using the BamBBBBBBBB~~~~~~~~~HI and EcoRI sites. Finally the ITB2 cDNA was transferred

into the modified pAM-PAT-GW vector through an LR reaction.

For overexpression of ITB2, the 35S:ITB2 and GL2:ITB2 constructs were made. The

ITB2 cDNA of the Col wild-type version was transferred into pAM-PAT-GW (Bekir Ulker,

Max Planck Institute for Plant Breeding, Cologne, Germany) for expression from the 35S

promoter, or pCK86 (Arp Schnittger, Max Planck Institute for Plant Breeding) for

expression from the GL2 promoter through an LR recombination.

To localize ITB2, a 35S:ITB2-CFP gene fusion was made. Cyan fluorescence protein

(CFP) was amplified by PCR using the pAVA574 (von Arnim et al., 1998) plasmid as a

DNA template. The stop codon was removed from pENRIA-ITB2 through sequence

substitution at the 3' end containing Ndel with the PCR product. The two PCR products

were ligated together through blunt ends. The fused fragment was cloned into pENRIA-

ITB2 using the Ndel and EcoRI sites. Finally the ITB2-CFP gene fusion was transferred

into pAM-PAT-GW for expression from the 35S promoter through an LR reaction.

RNA Extraction and RT-PCR

Total RNA was extracted from six-week-old Col wild-type plants using the RNeasy

Plant Mini Kit (Qiagen, Valencia, CA) according to the manufacturer's instructions. The

full length ITB2 cDNA was amplified, following the instructions for the cMaster RT plus

PCR System (Eppendorf AG, Hamburg, Germany). The first strand DNA synthesis was

primed using oligo (dT)20. The cDNA was amplified using specific primers oflTB2. The

PCR products were sequenced by the Interdisciplinary Center for Biotechnology Research

(ICBR) at the University of Florida.









Plant Transformation

The constructs, 5' untranslated region (UTR):ITB2:3 'UTR, 35S:ITB2, GL2:ITB2, and

35S:ITB2-CFP, were transferred into itb2-28 mutants by the floral dip method (Clough and

Bent, 1998). The transgenic plants were selected using a 1000X dilution of Finale (Farnam

Companies, Phoenix, AZ) with 5.78% glufosinate-ammonium 5.78%.

Results

Characterization of the itb2 Mutants

ITB2 controls the trichome shape mainly through regulation of branch expansion. All

the itb2 mutant alleles displayed defective trichomes in which one branch is longer than

others (Zhang et al., 2005a). The itb2-12, itb2-19, and itb2-28 mutant alleles are in the

RLD genetic background; and itb2-4 is in the Col genetic background. All mutants display

identical trichome shape that is a weak distortion and at leas one branch longer than others

(Figure 3-1G-L). In the RLD genetics background, cotyledon shape also was altered in itb2

mutants. A kidney- shape d cotyledons are shown in Figure 3-1A-F. In addition to this

effect, the genetic background also influences the frequency of the phenotypically itb2

mutant plants in F2. The segregation of the itb2 allele does not fit the monogenic model for

Mendelian segregation (Table 3-1). In the RLD and Col ecotypes, the ratio of wild-type

and mutant plants in F2 was approximately 15:1, which is closer to the segregation ratio

expected for two loci segregating independently. However, in the Ler genetic background,

the ratio was much lower, which also seems to indicate that the itb2 phenotype results from

multiple independent lesions (Table 3-1).

Cloning of the ITB2 Gene

ITB2 was mapped between ITB3 and ITB4 on chromosome I (Zhang et al., 2005a).

To clone the ITB2 gene, we executed its fine mapping relative to SSLP markers. Because









the itb2-28 mutant is in the RLD genetic background, additional SSLPs markers were

developed between the RLD and Ler ecotypes. In addition, we screened for DNA deletions

in the itb2-28 mutant. One set of PCR primers did not produce a product when itb2-28

DNA was used as the template, but produced products when wild-type DNA from RLD or

Ler was used. Additionally, the primers in this reaction are complementary to the

sequences that are located to the middle of the mapped region, the BAC clone F23H1 1

(Figure 3-2A). To determine the exact position of the detected deletion, we screened

F23H11 through amplification of short regions spaced approximately every 500 bp along

this BAC clone. Finally, an approximately 19-kb deletion was found between positions

62757 bp and 81830 bp on F23H11. The sequencing data showed that the breakpoint was

TTTAAGCCATGACGCTGAGCGAT//AAAAAGCTTTGATCGTCTGT The right

deletion breakpoint includes 779-bp cDNA of the Atlg59780 gene. The letf side of the

deletion covered a 1336 bp deletion of the Atlg59820 gene. Additionally, the deleted

region covered only five genes (Figure 3-2B), all of which were sequenced using the

genomic DNA from the itb2-19 allele as the template for PCR amplification. We identified

an approximately 800 bp deletion in the gene Atlg59820 in this mutant and further

sequenced the itb2-12 allele. The same deletion as the itb2-19 allele was found. These

results indicated that ITB2 was identical with Atlg59820, and that the itb2-19 and itb2-12

alleles were likely to derive from the same allele.

For an allelism tests, the itb2-28 mutant was crossed with itb2-19 and itb2-12

mutants. Their Fl generation all displayed the itb2 mutant trichome phenotype. To search

for more alleles of the itb2 mutant, the Salk T-DNA insertion lines were examined. The

Salk_015929 line segregated into two kinds of phenotypic plants, one of which displayed









the itb2 mutant trichomes. The PCR analysis indicated the two plants with the itb2 mutant

trichomes were homozygous for the T-DNA insertion in Atlg59820. The itb2-28 mutant

was crossed with the homozygous T-DNA lines of Salk_015929 as a complementation test.

The Fl plants all displayed the itb2 mutant trichome phenotype indicating that the plants

from SALK_015929 with the itb2 phenotype were itb2 alleles.

The ITB2 gene is relatively large, with an 8425-bp genomic coding sequence and a

3642-bp cDNA sequence. The coding sequence consists of 27 exons (Figure 3-2C).

Compared to the gene in the Col ecotype, the RLD ecotype has six single nucleotide-

alterations were identified. Among the SNPs detected, Hyve alterations were located in

introns and one alteration was located in the last exon, but this transition causes no change

in the protein sequence. Through RT-PCR, the same size the ITB2 cDNA was amplified

using the total RNA from Col wild type. The results from sequencing the ITB2 cDNA

indicated that the ITB2 structure is identical with the annotation of The Arabidopsis

Information Resource (TAIR) in Figure 3-2C.

Complementation of the itb2 Mutant and Over-expression of the ITB2 Gene

To complement itb2 mutants and over-express ITB2, we made constructs using the

ITB2 cDNA. One reason for using the ITB2 cDNA is that it is much smaller size than the

genomic DNA. The ITB2 cDNA that was cloned into the pBluescript SK II (+) vector (SK-

ITB2) had 9 base changes (Figure 3-3) compared to the Col wild type. Among these

mutations, fiye cause amino acid changes: at 878 (the position in the cDNA beginning with

I at A of ATG), A to G (the amino acid transition, K to R); at 2554, A to G (M to V); at

2690, T to C (L to P); at 3265, A to G (M to V); and at 3289, A to G (T to A). The

transition of L to P and T to A is likely to affect protein function and others are not,

because similarity of mutated amino acids with the wild type in structure and property.









The re-cloning of the full length ITB2 cDNA was unsuccessful and no full length

ITB2 cDNA clones were available from the stock center. To correct the mutations in the

resultant clone of the ITB2 cDNA, two EST clones from the ITB2 gene, G7B4 and

M65003, were ordered from the ABRC (The Ohio State University, Columbus, OH).

These two clones both contain about a 1 kb sequence at the 3' end region of the ITB2

cDNA, which can be used for substitution of the mutated part of SK-ITB2. The host cells

that contain G7B4 grew well and the plasmids could be isolated through mini-preparation.

However, the host cells that contain M650023 grew slowly and the plasmids could not be

isolated through standard metheds. The isolated plasmids were sequenced and a TC-

deletion was found in the G7B4 clone at the 2946-bp position of the ITB2 cDNA. The

target fragments were amplified through PCR using the M65003 DNA as the template and

cloned into pBluescript SK. The cells that hosted the resultant construct also grew slowly

and compared to the G7B4 clone, much less plasmid DNAs could be isolated using

standard plasmid isolation procedures. Therefore, in the later experiments, we added 0.5%

glucose to the medium to prevent expression of the genes of interest in the host cells.

Through multiple substitutions, all the changed bases in SK-ITB2 were corrected into the

Col wild-type version (Figure 3-3).

To complement the itb2 mutant, both the 2146-bp element of the ITB2 gene at the 5'

end of ITB2 and the 1380-bp element at the 3' end drove the ITB2 cDNA coding sequence

of the Col wild type version. The transgenic plants in the itb2-28 background displayed the

wild type trichome phenotype (Table 3-2 and Figure 3-4). This result provided further

evidence that the ITB2 gene is Atlg59820.









To over-express ITB2, the ITB2 cDNAs were cloned into either pAM-PAT-GW for

expression by the 35S promoter or pCK86 for expression by the GL2 promoter through a

homologous recombination. The transgenic plants containing the mutated ITB2 cDNA in

the itb2-28 mutant background displayed the itb2 mutant trichome phenotype (data not

shown). However, the transgenic plants of the same genetic background as the Col wild-

type ITB2 cDNA version exhibited wild-type trichomes (Table 3-2 and Figure 3-4). This

result indicated that the C-terminus is crucial for functionality. Additionally, no novel

visible phenotypes were observed for these transgenic plants.

To localize ITB2, we constructed the ITB2 gene fusion with CFP, using the Col wild

type ITB2 cDNA. The transgenic plants in the itb2 mutant background with the 35S: ITB2-

CFP construct displayed the itb2 trichome phenotype. This result further supports the

hypothesis that the C-terminus of ITB2 is crucial for functionality.

Discussion

Membrane trafficking is essential for establishment and maintenance of plant cell

polarity, especially for such tip growth as the elongation of pollen tubes and root hairs. At

the cellular level, tip growth is achieved through polar-specific and cell domain-specific

trafficking of vesicles (Hepler et al., 2001); at the molecular level, the tip growth

machinery is assembled mainly by small GTPases in the Rab, Arf, and Rop/Rac families

and their regulatory proteins such as Rho guanine nucleotide exchange factors (GEFs) and

GTPase activating proteins (GAPs) (Samaj et al., 2005; Yang and Fu, 2007). A trans-Golgi

network (TGN) was further proposed as a tip-localized vesicular compartment integrating

targeted secretion and endocytosis within the growing tip (Samaj et al., 2006).

When pollen tubes are growing, endomembrane trafficking activity transports

secretary vesicles along the flank of the tube to the tip; meanwhile, endocytic recycling









vesicles move back distally along the center of the tube, which forms a reverse-fountain

cytoplasmic streaming pattern (Hepler et al., 2001). The secretary vesicles accumulate

within the tip clear zone where they form clusters and fuse with the apical membrane,

depositing new membrane, proteins, and cell wall materials to support growth (Samaj et al.,

2006). The behavior of vesicles in the growing root hairs is reminiscent of that in growing

pollen tubes (Hepler et al., 2001; Voigt et al., 2005).

Although trichome branching is not considered to belong to the tip growth, its

process is a typical anisotropic cell expansion. Endomembrane trafficking also plays an

important role in plant epidermal cell expansion, such as Arabidopsis leaf pavement cells

and trichomes (Smith and Oppenheimer, 2005). The defective endocytic membrane traffic

can cause changes in the shapes of these cells. Using FM4-64 to track endocytic membrane

traffic, the larger vesicle clusters aggregated and were surrounded with an ring of Golgi

stacks in the leaf pavement cells of the cerl0 mutant (Zheng et al., 2005). These defects

resemble the compartments found in the brefeldin A (BFA)-treated cells. BFA is an

inhibitor that disrupts exocytosis (Baluska et al., 2002; Samaj et al., 2004). The cerl0

mutant leaf pavement cells were considerably smaller with less pronounced lobes, and the

cerl0 trichomes were smaller with short, crooked, and aberrant swollen stalks and branches

(Zheng et al., 2005). The alterations of these cell shapes indicated that their anisotropic

expansion was compromised, because of defects in endocytic membrane trafficking. A

further molecular model was proposed after analysis of Kinesin-13A, ZWI, and AN

functions (Lu et al., 2005; Smith and Oppenheimer, 2005). The plant Golgi apparatus and

secretary vesicles are transported by myosin(s) from the perinuclear region to the cell

cortex, dispersed to cell expanding sites by Kinesin-13A along the cortical microtubules.









Mutations of AtKinesin-13A increased the trichome branch number and a compromised

anisotropic trichome cell expansion. In the trichomes of the kinesin-13a-1 mutant, the

Golgi stacks aggregated (Lu et al., 2005). Further support for this modelcomes from the

aberrant trichome shapes found in myosin mutants (Oj angu et al., 2007).

ITB2 is putative aminophospholipid translocase 3 (ALA3) in the P4 ATPase family.

Although 12 members (ALAl-12) in this family were identified in Arabidopsis, their

biological functions are less known (Axelsen and Palmgren, 2001). Evidence suggests that

ALAl is involved in generating membrane lipid asymmetry. Down-regulation of ALAl

results in a significant reduction in plant size and an alteration of plant morphology at low

temperatures (Gomes et al., 2000). It is likely that the aberrant plants resulted from

defective cell expansion. ITB2 regulates anisotropic expansion in trichome morphogenesis

most likely through its role in generating membrane lipid asymmetry for vesicle formation

during endomembrane trafficking. To confirm our explanation, we will track membrane

trafficking in the itb2 mutants.

Future Perspectives

ITB2 is a putative flippase that translocates aminophospholipids from one leaflet to

another. It belongs to the P4 ATPase family. Our results show that mutations in ITB2 cause

a defective trichome shape phenotype and a slight change in cotyledon shape. These

mutants provide good experimental materials for further studies on function of ITB2 in

controlling plant cell shape. Recently published data showed that aminophospholipid

translocase 3 (ALA3) is identical to ITB2 (Poulsen et al., 2008). It was shown that ALA3

localizes to the Golgi. We predict that defective Golgi localization can also be observed in

itb2 mutant trichomes.









To observe Golgi localization in developing trichomes, we can cross the ST-YFP

construct to itb2 mutants. The ST-YFP construct is a fusion of the Golgi-localized

sialyltransferase enzyme to yellow fluorescent protein. This construct labels Golgi stacks in

transformed plant cells. Once this construct is introgressed into the itb2 mutant

background, localization of Golgi can be observed in living, developing trichomes using

confocal microscopy. By comparing the localization pattern and dynamics of Golgi stacks

in itb2 mutants with that of wild type trichomes, we can determine if Golgi dynamics and

localization is affected in itb2 mutants. If this is found, then it suggests that proper Golgi

localization is a key to directional cell expansion. Also this result would suggest that

flippase activity is important for Golgi function.

The key question that remains to be answered is whether or not ALA3/ITB2 is

actually a flippase. For this to be shown, flippase activity has to be reconstituted in vitro.

Many groups have attempted this, but have not been successful.












Table 3-1. Segregation of the mutant plants in F2 with different genetic background
Cross name Number of Number of X2 value* P value*


wt plants
424
867
1441


mutant plants
28
52
46


itb2-28/RLD wt
itb2-28/Col wt
itb2-28/Ler wt
* is for 15:1 ratio.


0.0022
0.5495
25.2859








% normal
89.3
91.8
94.6
96.5
10.7


>0.900
>0.500
<0.001








Total
799
972
815
713
651


Table 3-2. Trichome shapes of the transgenic plants
Constructname Promoter % irregular
XG61 ITB2 10.7
XG62 35S 8.2
XG63 GL2 5.4
RLD -3.5
itb2-28 -89.3














































Figure 3-1. Defects in leaf trichome and cotyledon shape of itb2 mutants. (A), (G), and
(J): RLD wild type; (D), Col wild type; (B), (H), and (K), itb2-28; (C), itb2-19;
(E), (I), and (L), itb2-4; (F), itb2-12. Bars = 1000 Clm in (A)-(F); 200 Clm in (G)-
(I); 100 Clm in (J)-(L)










A Chromosome 1
nga248 ngal11
I I


C Target gene
ITB2 : Atlg59820


3 1 1 0 4 8 12
T5A14 F25P12 T30E16 F23H11 F5114 F12P19 ngal11
"~~~ """


Recombinants and BACs


76
Total


25 9 5
nga248 F7F22 F6D8


deleted region
19074 bp-deletion


Atlg
79 bp 570


B Genes in the



Atlg 7
597807


Atlg
59800


Atlg
59810


Atl
1336 b
59820


__


mm m m m m


m1 m1 11 1 m a1 mmI m m m m mm mm M m m m


Figure 3-2. Positional cloning and gene structure of ITB2. The itb2 mutation was mapped
near SSLP markers between nga248 and ngall1 on chromosome 1. Additional
molecular markers were used to map the itb2 mutation to BAC clone F23H11.
The numbers of recombinants (out of 76 chromatids screened) are given above
BAC clones. All putative genes inside the identified deletion on BAC clone
F23H11 are listed. ITB2 structure is shown in (C). The thick bars in (C)
represent exons.













Point mutation


Amino acid change



SPCR product |


PCR
| product|
T AT


M65003


-


substitution 1


substitution 2


substitution 3


Figure 3-3. Mutations and corrections of ITB2 cDNA. All mutations of the cloned ITB2
cDNA are listed in the vectanglar boxes. The numbers on the above bases are
the positions of mutated bases. The respective amino acids encoded by the
change genetic codes are also listed below the thick line representing ITB2
cDNA. Approximate positions of the PCR primers used for substitution are
shown by the black arrows with the corrected base above them.


2554 2662 2690 3265 3289
A-,G T~C T~C A-G A-,G



































Figure 3-4. Transgenic plants with ITB2 cDNA. (A) and (C), RLD wild type; (B) and (D)
the transgenic plant. Bars = 500 Clm in (A) and (B); 100 Clm in (C) and (D)









CHAPTER 4
DISPROPORTIONA TE (DPP) ENCODES A KETOACYL REDUCTASE INVOLVED IN
TRICHOME CELL EXPANSION

Introduction

The control of plant cell shape has been suggested to occur in three sequential steps:

First, cell polarity is established by intracellular mechanisms and/or extracellular cues.

Second, using the established polarity, cytoskeletal rearrangements take place. Third, the

cytoskeletal changes enable polarized cell expansion, which includes the incorporation of

membrane and cell wall material at defined areas of the cell periphery (Hulskamp et al.,

1998; Smith and Oppenheimer, 2005).

Arabidopsis trichomes are and excellent model for studies on the control of plant cell

shape. Through a forward genetic approach, more than 20 genes that regulate trichome

development have been cloned (Marks, 1997; Schellmann and Hulskamp, 2005). The

products of these trichome genes are diverse; their functions include transcription initiation,

cytoskeletal organization and vesicle trafficking. For example, GL1, GL2, GL3, EGL3,

TTG1, TRY, ETC, MYB23 and CPC all function as transcriptional regulators (Schellmann

and Hulskamp, 2005). DIS1, DIS2, DIS3/ITB1, WOR I, CRK, GRL, A TRK1 and PIR/KLK

belong to the "distorted" group of trichome genes that encode regulators of the actin

cytoskeleton (Schellmann and Hulskamp, 2005; Szymanski, 2005). AN, ZWI, MYA2 and

KINESIN-13A are known or predicted to be involved in vesicle trafficking (Smith and

Oppenheimer, 2005). Recently, a group of genes identified by wax and cuticle phenotypes

whose products are involved in wax synthesis and transport (Kunst and Samuels, 2003)

have been shown to be involved in trichome morphogenesis (see below).

Wax is derived mainly from very long chain fatty acids (VLCFAs), which are

required for sphingolipid synthesis. VLCFA moieties in sphingolipids are essential for









determining the physical properties and characteristics of membranes. Sphingolipids are an

important class of lipids in the plasma membrane and the endomembrane system (Simons

and Toomre, 2000). In yeast and mammalian cells, sphingolipids are concentrated in lipid

rafts, which are involved not only in cellular trafficking of certain plasma membrane

proteins, but also play important roles in signal transduction and generation or maintenance

of cell polarity (Raj endran and Simons, 2005). Plant lipid rafts are also enriched in

sphingolipids, but their role in generation or maintenance of cell polarity has rarely been

reported (Bhat and Panstruga, 2005; Grennan, 2007).

VLCFA synthesis is a complex process including two stages in different cellular

compartments (Kunst and Samuels, 2003). The de novo fatty acid synthesis of C16 and

C18 acyl chains occurs in the stroma of plastids by the soluble enzyme complex called the

fatty acid synthase (FAS). The synthesized fatty acyl precursors are further extended to

C34 VLCFA chains through the same reactions as the de novo fatty acid synthesis, but

these reactions are catalyzed by membrane-bound enzyme complexes called fatty acid

elongases (FAE) located in the endoplasm reticulum. FAE is composed of four enzymes

that catalyze four sequential reactions: These are 1) condensation of malonyl-CoA to

acetyl-CoA by 3-ketoacyl-CoA synthase (KCS), 2) reduction of 3-ketoacyl-CoA by 3-

ketoacyl-CoA reductase (KCR), 3) dehydration of 3-hydroxyacyl-CoA by 3-hydroxyacyl-

CoA dehydrase (DCH), and 4) reduction of trans-2-enoyl-CoA by enoyl-CoA reductase

(ECR). The resultant VLCFAs are finally modified into different kinds of waxes (Kunst

and Samuels, 2003). Wax monomers are exported to the cell surface by the ABC

transporters such as ABCGl2/CER5 (Pighin et al., 2004) and ABCG11/WBC11 (Bird et

al., 2007).









LACERATA (LCR) encodes a monooxygenase, which catalyzes co-hydroxylation of

fatty acids ranging from C12 to C18. The trichomes on Icr mutant leaves exhibited under-

development with a variety of aberrant shapes (Wellesen et al., 2001). The FIDDLEHEAD

(FDH) gene codes for a KC S. Mutations in FDH have a deleterious effect on trichome

differentiation because leaf trichome number was reduced 2-fold in fdh mutants

(Yephremov et al., 1999). ECERIFERUM1 (CER10) encodes an ECR; mutations in this

gene caused defective leaf trichomes, which had short, crooked, and abnormally swollen

stalks and branches (Zheng et al., 2005). The BODYGUARD (BDG) gene encodes a

putative extracellular synthase responsible for the formation of the cuticle. The bdg mutants

displayed many misshapen leaf trichomes including ones with flat, bent, and collapsed

shapes (Kurdyukov et al., 2006). The maize (Zea mays) GLOSSY1 (GL1) gene codes for a

component in the pathway leading to cuticular wax biosynthesis in seedling leaves. The gl1

mutation results in leaf trichomes that are smaller than normal (Sturaro et al., 2005).

Mutations in YORE-YORE (YRE), a putative GL1 homolog in Arabidopsis, also led to small

trichomes. In addition, the trichome shape of yre cerl double mutants was greatly

deformed (Kurata et al., 2003). Genetic lesions in (DESPERADO) DSO/AtWBC11, an ATP

binding cassette (ABC) transporter for wax export, led to a dramatic alteration in wax load

and trichome development. The dso atwbc11 mutants had waxless stems and collapsed and

underdeveloped trichomes (Bird et al., 2007; Luo et al., 2007; Panikashvili et al., 2007).

Here, we report the cloning of the DPP gene through a novel strategy amenable to the map-

based cloning of other dominant, homozygous lethal mutations. We show that DPP codes

for a KCR, and dpp mutations lead to a waxless phenotype in addition to dramatically

altered trichome shape.









Materials and Methods

Plant Materials and Growth Conditions

The dpp mutant was isolated in a genetic screen of fast-neutron mutagenized seeds,

in the Rschew (RLD) genetic background, which were purchased from Lehle Seeds (Round

Rock, TX). The Landsberg erect (Ler) ecotype was used as wild type for construction of

the mapping population. Wild-type plants of the Columbia (Col) ecotype were used for

transformation. Seeds were sown on a soil-less potting medium, Fafard 2 Mix (Conrad

Fafard, Inc., Agawam, MA). Seedlings were grown at different temperatures (180 C, 200 C,

220 C and 240 C) in growth chambers under 16 hr light and 8 hr darkness. Light was

provided by 40W, cool white fluorescent tubes. Plants were watered with PGP nutrient

solution (Pollock and Oppenheimer, 1999) every two weeks.

Positional Cloning

For cloning DPP, a mapping population was generated from a cross between the dpp

mutant and Ler. Because the dpp mutant was dominant and homozygous lethal, 646

phenotypically wild type plants were selected from the F2 population for mapping. The first

leaf pair from each individual was used for DNA extraction, following a standard protocol

(Edwards et al., 1991). The isolated DNA was used as a template for PCR to map the dpp

mutation relative to simple sequence length polymorphisms (SSLPs) (Bell and Ecker 1994,

Lukowitz et al. 2000) and cleaved amplified polymorphic sequences (CAPS) (Konieczny

and Ausubel, 1993). After the dpp mutation was mapped into a narrow region, all the

candidate genes in the mapped region were amplified by PCR of genomic DNA from the

dpp mutant to check for deletions, as expected in a mutant isolated from fast neutron

mutagenesis. Primers sequences are listed in Table 4-1.









To determine the lesion in the dpp mutant, we crossed dpp plants to SALK lines that

were homozygous for T-DNA insertions in the candidate genes located within the mapped

region. The SALK lines were obtained from the Arabidopsis Biological Resource Center

(The Ohio State University, Columbus, OH). Plants homozygous for the T-DNA insertions

were identified through PCR with T-DNA and gene-specific primers designed using the

SALK T-DNA insertion primer design website (http://signal_ salk.edu/tdnaprimers.2.html).

The homozygous T-DNA insertion lines were used as the female and the dpp mutant was

used as the male for the crosses. The Fl progeny were grown at temperatures below 200C

to ensure that the dpp trichome phenotype was visible. The segregation of the trichome

phenotype in the Fl plants was recorded. DNA was extracted from the Fl plants using the

DNeasy Plant Mini Kit (Qiagen, Valencia, CA), following the manufacturer's instructions.

The candidate genes were amplified by PCR, and the products were sequenced to check for

mutations. The sequences were compared to that of RLD wild type.

Plasmid Construction

Genomic DNA from dpp mutants was used as the template for amplification of

Atlg67730 using primers DEd7 and DEd8. PCR reaction conditions were as follows: 94o

C, 3 minutes; 25 cycles of 94o C, 30 seconds; 64o C, 30 s; 680 C, 3 minutes. KOD XL DNA

polymerase (Novagen, Madison, WI) was used to amplify the entire Atlg67730 coding

sequence, which comprised 4967 bp. The PCR products were cloned into pBluescript SK

using the PstI and EcoRV sites. Positive clones were sequenced to check for mutations in

the cloned gene. The GC-deleted dpp allele was also cloned into pAM-PAT-GW (Bekir

Ulker, Max Planck Institute for Plant Breeding, Cologne, Germany) using the AscI and PstI

sites. This clone was used to transform Col and RLD wild type plants to regenerate the

dominant dpp trichome phenotype.









Plant Transformation

Plants were transformed by the floral dip method (Clough and Bent, 1998).

Transgenic plants were grown at 160 C, and sprayed with a 1000X dilution of Finale

(5.78% glufosinate-ammonium) (Farnam Companies, Inc., Phoenix, AZ).

Results

Characterization of dpp Mutants

To gain additional insight into trichome morphogenesis, we screened fast neutron-

mutagenized Arabidopsis populations for plants that displayed defects in trichome shape.

One of the mutants isolated in this screen showed shorter branches and longer stalks

(Figures 4-1B, 4-1C) than wild type (Figures 4-1E, 4-1F). This mutant was named

disproportionate (dpp). During routine plant growth, we noticed that the dpp phenotype

was temperature sensitive; at temperatures below 220C, plants displayed the dpp

phenotype, while at temperatures above 240C, the trichomes developed normally. At

temperatures between 220C and 240C, most of the trichomes on the first leaf pair had a dpp

phenotype, but later leaves had a higher proportion of normal trichomes. In addition, dpp

leaf blades adhered to each other at an early developmental stage (Figure 4-1A), whereas

the wild type leaves are open at the same stage (Figure 4-1D).

When dpp plants were crossed to other wild type plants, the resulting Fl plants

segregated for the dpp phenotype. This result showed that the dpp mutation was dominant.

The resulting F2 progeny also displayed segregation of both dpp and wild type plants in a

ratio of 2:1, which fits a monogenic model for dpp (Table 4-2), assuming that the dpp

homozygotes are lethal. We also noticed that all dpp plants segregated both dpp and wild

type plants in the next generation. This phenomenon was examined in more detail by

scoring the phenotypes of at least 50 progeny from 108 individual dpp plants. The plants in









all the observed populations segregated for dpp and normal trichomes. This result indicated

that the populations were derived from the homozygous plants. Therefore, dpp plants are

homozygous embryo lethal because no homozygous plants were found.

To test transmission of the dpp allele, the dpp mutant was reciprocally crossed with

RLD wild type or Salk_143503. When the dpp mutant was used as the female parent, a

higher frequency of Fl plants with the dpp phenotype occurred than when the dpp mutant

was used as the male parent (Table 4-3). Because dpp mutants are heterozygous, when dpp

is used as the male parent, both dpp and wild type pollen are transferred to the stigma. This

result indicated that the dpp pollen was less competitive than wild type pollen during

pollination and/or fertilization.

Positional Cloning of DPP

To clone the DPP gene, we generated an F2 mapping population consisting of 646

wild type plants. Wild type plants were used because the dpp homozygotes do not survive,

and known homozygous plants need to be genotyped to identify recombinants. The DPP

locus was mapped relative to SSLP markers to two BAC clones, TIFl5 and Fl12A21. It

was further mapped relative to CAPS markers to an 82 Kb region on Fl2A21. According to

the TAIR database, this region covers 34 putative genes (Figure 4-2).

Because the dpp mutant was isolated from fast neutron-mutagenized seeds, we

expected this mutation was caused by a deletion. We designed primers to amplify 0.5 Kb to

3 Kb fragments that cover the region of Fl2A21 that contained dpp. PCR products were

amplified from both wild type and dpp mutants, and the sizes of the PCR products were

compared with each other to determine the deletion position. Results showed that PCR

products that differed in size from those predicted for wild type were generated by non-

specific binding of the primers, and not due to a deletion in the dpp mutant. The PCR









products were further sequenced through amplification of the coding sequences of the

putative genes in this region using the genomic DNA from the dpp mutant plants. This

strategy identified single nucleotide polymorphisms (SNPs) between the of RLD and Ler

ecotypes (Table 4-4 and Figure 4-3), and these SNPs were used as the markers for further

mapping the dpp mutation. Many sequencing results were equivocal because the genomic

DNA used for sequencing was extracted from the heterozygous dpp plants. For example,

see sequencing results of Atlg67760 (Figure 4-4) and Atlg67730 (Figure 4-5, 4-6).

Atlg67760 encodes a chaperonin and Atlg67730 encodes a ketoacyl reductase. There were

no clear connections between these two gene products and the regulation of trichome

shapes. Therefore, they were not immediately pursued further.

Identification ofDPP

The homozygous lethality of the dpp mutation presented a challenge to identify the

molecular lesion in the dpp allele because all dpp plants were heterozygous. Therefore, to

identify the lesion in the dpp allele, we developed a strategy whereby we could specifically

amplify the dpp allele from the heterozygous dpp plants, which was accomplished by

crossing the dpp mutant to a homozygous SALK lines (Figure 4-7).

The dpp mutation is dominant, the heterozygous plants grow normally except for the

dpp trichome phenotype, and the homozygous dpp plants are lethal. If the T-DNA insertion

in the Salk lines causes a decrease in DPP gene expression, the Fl plants may display a

novel phenotype. The result of phenotypic segregation in the Fl plants from crosses of dpp

to homozygous SALK lines confirmed this prediction. Among the 10 homozygous SALK

lines (with T-DNA insertions in 7 distinct genes) two Fl families showed lower

transmission of the dpp mutation (Table 4-5). These two SALK lines had a T-DNA inserted

in the 5' UTR of Atlg67730. The Fl plants from one of these combinations segregated for









a novel phenotype: crinkled leaves, collapsed trichomes, waxless stems, and fused floral

organs (Figure 4-9). Some of these phenotypes are similar to those displayed by deadhead

(ded) mutants (Lolle et al., 1998).

The crinkled leaves and severe defective trichomes led us to focus our attention on

Atlg67730. We re-sequenced the PCR products using the DNA from the dpp plants as the

template. The result remained equivocal (Figure 4-10). However, the forward reaction

worked well with the same DNA sample (Figure 4-11), suggesting that a deletion or

insertion likely occurred in this gene. When the Fl plants that had the novel phenotypes

grew larger, DNA was isolated from their leaves for further analysis. By designing primers

that flank the T-DNA insertion, such as primers 720S1 and 720S2 or 730S1 and 730S2 (see

Table 4-1 and Figure 4-12), we could specifically amplify the dpp allele, using the isolated

DNA from the Fl plants. Sequencing of the PCR product revealed a 2 bp deletion (GC) at

position 1595 and 1596 of the coding sequence of gene Atlg67730 (Figure 4-13). This

deletion was not observed in the sequence of PCR products from DNA extracted from Fl

plants that had a wild type trichome phenotype (Figure 4-14). To confirm that the GC-

deletion also exsits in the dpp mutant and that DPP is Atlg67730, the entire coding

sequence of Atlg67730 including upstream and downstream intergenic regions was

amplified from DNA isolated from dpp plants using primers DEd7 and DEd8 (Figure 4-

12). The PCR products were cloned into the pBluescriptSK vector. The plasmid DNAs

from two individual colonies were sequenced. One of them showed the GC-deletion

(Figure 4-15) (and other was wild type (Figure 4-16). The GC-deleted DPP was used to

transform wild type plants. Plants transformed with this GC-deleted transgene showed the

same trichome defect as the original dpp mutant (Figure 4-17). These results further









confirm that DPP is Atlg67730, which encodes a ketoacyl reductase, basing on the gene

annotation (Arabidopsis Information Resource).

Discussion

DPP Encodes a P-ketoacyl Reductase

We isolated the dpp mutant during a screen for mutants with altered trichome shape.

We found that the dpp mutation was dominant and homozygous lethal. In addition, the dpp

mutants had a temperature sensitive phenotype; at 240C and above, the trichomes appeared

perfectly normal, whereas at 220C and below, the trichomes displayed an elongated stalk

and short branches. To clone DPP, we combined microarray information from laser-

capture microdissected trichome cells with traditional positional cloning methods to

identify the most likely candidate gene, Atlg67730, for further analysis. Atlg67730

encodes a P-ketoacyl reductase (KCR) that is involved in cuticular wax biosynthesis. To

overcome the problems associated with identifying the dpp mutation from DNA isolated

from heterozygous plants, we devised a novel strategy using primers flanking a T-DNA

insertion in Atlg67730. Using these primers and DNA from Fl plants from a cross of dpp

with Atlg67730 T-DNA insertion, only the dpp allele was amplified by PCR and the dpp

mutation unequivocally identified. Recapitulation of the dpp phenotype in wild type plants

containing the mutant dpp transgene confirmed that Atlg67730 is DPP. The

straightforward cloning of dpp demonstrates the efficacy of our approach for the positional

cloning of other dominant, homozygous lethal alleles.

DPP Has Pleiotropic Functions in Cell Expansion and Wax Synthesis

The mutation in the dpp mutant caused a 31amino acid-deletion at the C-terminus of

KCR. The mutated KCR resulted in plant lethality, but heterozygous plants in the RLD

ecotype background, at the restrictive temperature, exhibited a trichome phenotype.









Otherwise the plants grew normally. Expression of the truncated KCR with the endogenous

promoter in Col wild type plants exhibited novel phenotypes such as waxless stems,

crinkled leaves, fused organs and severe aberrant trichome shapes. These phenotypes are

similar to those seen in the Fl plants from the cross of the dpp mutant with the T-DNA

insertion line, except that the phenotype of the transgenic plants is less severe. This may be

due to the fact that only one mutant copy of dpp is present along with two- copies of wild

type DPP in the transgenic plants. This suggests that the severity of the phenotype is

dependent on the dosage of the DPP gene relative to the mutant allele.

The truncated DPP protein encoded by the dpp allele is likely to have less function

compared to wild type. The more dpp protein that gets assembled into FAE, the less

VLCFAs are synthesized. The dpp-homozygous plants are lethal because of the VLCFA

deleption. Therefore, these long-chain fatty acids are essential for plant viability. The

possibility that the dpp protein is a gain-of-function allele cannot be excluded. For

example, the dpp protein might interfere with wax synthesis or transport.

Plants with the dpp allele in the Col genetic background lacked cuticular wax and had

fused leaves and floral organs. These phenotypes are typical of the defects resulting from

mutations in genes that are involved in wax synthesis.

The GL8 gene in maize (Zea mays L.) codes for a KCR. gl8 mutants reduced the

amount and altered the composition of seedling cuticular waxes (Xu et al., 1997). The

disruption of CER6/CUT1 coding for a KCS reduced wax accumulation on stems (Millar et

al., 1999; Fiebig et al., 2000). The FIDDLEHEAD (FDH) gene encodes another KCS

(Fiebig et al., 2000; Pruitt et al., 2000). This mutant displayed defects in organ fusion

(Lolle et al., 1992). Mutation of the CER10 gene, which encodes an ECR, caused a









reduction of wax abundance (Zheng et al., 2005). However, the dpp mutation in the RLD

background did not cause a reduction in wax accumulation or organ fusion. The reason for

this difference is not known, but this difference was also observed in other studies. For

example, the appearance of crinkled and fused leaves in the abnormal leaf shapel (alel)

mutant depended on the genetic background, and the mutant phenotypes could be observed

in the Landsberg erecta background but not in the Columbia and Wassilewskij a genetic

backgrounds (Watanabe et al., 2004).

Mutations in CER10/ECR caused a change in the VLCFA content of sphingolipids

(Zheng et al., 2005). Sphingolipids play a role in generation and maintenance of cell

polarity. Genetic or pharmacological inactivation of sphingolipid synthases not only

prevents polarized hyphal growth, but it also abolishes cell polarity establishment (Cheng

et al., 2001). This result was further confirmed by distinct sphingolipid synthases with a

chemical genetic approach (Li et al., 2006). The epidermal leaf pavement cells in the cerl0

mutants displayed a three-fold reduction in size and less pronounced lobing, when

compared to these cells in wild-type leaves. Aberrant cell shapes were caused by a

disruption of trafficking since the Golgi stacks aggregated, forming ring-like structures in

the cerl0 mutants (Zheng et al., 2005). The dpp mutant in the RLD background also

exhibited a change in the trichome shape at the restrictive temperature, but the wax and

other phenotypes were normal.

DPP is Vital for Plant Viability

DPP codes for a KCR, a subunit of FAE for VLCFA synthesis. In distinct FAEs,

specificity of each elongation reaction on different chain length substrates is determined by

the selectivity of a KCS (Millar and Kunst, 1997). A large family of 21 KCS-like

sequences in the Arabidopsis genome contributes to wax biosynthesis, which takes place









in several different tissues at different stages of plant development (Kunst and Samuels,

2003; Costaglioli et al., 2005; Suh et al., 2005). In contrast, two KCRs apparently have no

particular acyl chain length specificity and are shared by distinct FAEs (Kunst and

Samuels, 2003). In the yeast genome, because only one gene was found to code for each

KCR, TSC13 coding for ECR is essential for yeast viability (Kohlwein et al., 2001), but

YBR159w coding for a KCR is not essential (Beaudoin et al., 2002). In the Arabidopsis

genome, five genes code for ECRs (Costaglioli et al., 2005). Mutations in CER10/ECR

disrupted normal shoot development and cell expansion, but plants were viable (Zheng et

al., 2005). The genes coding for KCR are Atlg67730/DPP and Atlg24470. These two

proteins are 44% identical and 68% similar to each other (Kunst and Samuels, 2003).

Based on microarray analysis, Atlg67730 was expressed to significant levels, but

Atlg24470 expression is low (Costaglioli et al., 2005). Therefore, it is likely that DPP is

the maj or component of FAE. The Salk-096487 line having a T-DNA inserted in an exon

of Atlg24470 displayed no visible phenotype (Xiaoguo Zhang, Unpublished data).

DPP is Likely to be DEADHEAD

The deadhead mutation was mapped to chromosome I. It maps to the same location

as the dpp mutation. More importantly, among deadhead, bulkhead and hothead, only the

deadhead mutant displayed no wax on stems (Lolle et al., 1998). The Fl from the

combination of the dpp mutant and Salk linel43505 also displayed this waxless phenotype

on stems. Therefore, it is possible that the DPP gene is the DEADHEAD gene.

Unfortunately, a complementation test between dpp and ded is not possible, because dpp is

dominant.









Future Perspectives

DPP encodes a ketoacyl reductase that is one maj or component of the fatty acid

synthesis complex. The dpp mutation causes disproportionate trichomes, which have longer

stalks and shorter branches as compared to wild type trichomes. Interestingly, the dpp

trichome phenotype is temperature sensitive: below 220C, the trichomes showed the dpp

phenotype, whereas at temperatures above 240C, the trichomes appeared normal.

Additionally, the dpp mutation was monogenic, dominant, and homozygous lethal. A two-

base (GC) deletion in the dpp gene causes a 30 amino acid-truncation from the C terminus

of encoded ketoacyl reductase. Based on these results, several interesting questions come to

mind. For example, why does this truncated ketoacyl reductase generate a dominant

trichome phenotype? Is it a gain-of-function mutation or a loss-of-function mutation? Why

and how is this truncated protein so sensitive to temperature? To answer these questions,

biochemical assays in vitro need to be performed. For example, activity of the wild type

and truncated DPP can be compared in vitro, at different temperatures.

Plants heterozygous for dpp have only defective trchomes. However, our results

shows the Fl plants from the cross of the dpp mutant with the T-DNA insertion line

containing a T-DNA in 5' UTR of DPP display pleitropic phenotypes including collapsed

trichomes, crinkled leaves, fused floral organs, and waxless stems. Some of these

phenotypes are similar to the phenotypes seen in deadhead (ded) mutants. In addition, the

ded mutation has been roughly mapped to the south end of chromosome 1, which is the

same location of DPP. Currently it is not known if DPP and DED are same gene. A simple

complementation test between dpp and ded cannot be done, because dpp is a dominant

mutation. To address this question, the DPP genes from multiple alleles of ded mutants









need to be sequenced. If all the ded alleles have mutations, then it is likely that ded and dpp

are the same gene.

Published data show that mutations in the gene encoding an enoyl reductase, which is

another maj or subunit of the fatty acid synthesis complex, caused defects in the Golgi

apparatus. Leaf epidermal cells of the mutant display many clustered Golgi stacks. It would

be interesting to examine the organization of the Golgi stacks in the dpp mutant. If the dpp

mutant shows altered Golgi organization, then this result would support the idea that the

Golgi apparatus plays an important role in cell expansion, and that fatty acid synthesis is

crucial for proper membrane trafficking. To visualize the Golgi apparatus in live plant

cells, the Golgi-labeled markers such as ST-YFP can be used for this study.









Table 4-1. Primers sequence used in this study


* The primers here are used for the DPP mapping and identification. For mapping, dCAPS
and SNP markers are listed, for example, two primers, dCAPS3F and dCAPS3R, are a
primer pair for a dCAPS marker which is located at the gene Atlg68060. When the PCR
products from this primer pair are digested by Ndel a difference in the size of the digested
DNAs between the ecotypes of RLD and Ler will be observed. The degenerated bases are
bold and underlined. Similarly, T30E16-57F and T30E16-57R is a primer pair for a SNP
marker. The PCR products from them display a difference in size. For the RLD ecotype, a
138-bp DNA is seen, whereas for the Ler ecotype the size is 81 bp.


Primer name
dCAPS3F
dCAPS3R
dCAPS6F
dCAPS6R
dCAPS7F
dCAPS7R
dCAPS9F
dCAPS9R
dCAPS10F
dCAPS10R
T30E16-57F
T30E16-57R
Fl2Pl9-26F
Fl2Pl9-26 R
TIFl5-42F
TIFl5-42R
F5A8-15F
F5A8-15R
T22E19-6F
T22E19-6R
T26J14-42F
T26J14-42R
T6C23-20F
T6C23-20R
720S1
720S2
730S1
730S2
730codl
730cod2
DEd7
DEd8


Sequence(5' 3')
GTAGTCGCCTTGAGAAAATCTTCA
CCATTGCCTTTGTTAAAGTTTCA
GGTCGCTTCGAGAACAACATTA
GGTGTGGTCAGGAGTCCTTTA
GAGAGAATC ACAC GAAT TCAAAAGAAAC C
GGTGATAGCAGAAAGGCCAAAA
GGGGTTCTGTCTACTGTGGTAACTCCAT
GGTATTGGATCTTATTTAGAAGCCTC
CCACTCTTTAAATGGAAAATCTGGTCATCATCTA
TGCTTGCAATTGTGATCATCTTG
ACACTCTTTACTGGAAGATGCAA
AACACACCCATGCAAGTGAA
CTGGAAATATCTGCGAAGTGGAA
CATGAACTGTTTGTGCATCTCTG
GCTGATAAGCGTATCATCACACA
GGTGCGCCATCAAATAATGT
TGGAGTTAACATATTTTTAATTTATCC
GTGGTCAACATCACATTAAAAACA
CCCAATCTAACGGATTTGAAT
GGGCTTTGTTTCTTGTGAAAT
GTCTTTCAACTGGTTTCAAATTTGT
GTTCCATTTTGGTACTTAGTAATGGAC
CGCTACTAAATTTGGTGGGGGTT
TGAGCCTAAAACTTTAACTTCTGC
GCGACCTATAGAGGAGGCATTATTGCG
CCTTTTGTTCTGTCTCAAGTTACAGG
GGCAACAGCAACCAAGTGCATGTCTC
CCTTGCTTACTAGCTTCCTCGAGC
CCTTGAAGAGACGCAAACCAT
TTCTATCCACCTTCGTCCCTT
CGTCTTCTCTTCCCTCAGCTA
CACTAGACTGGCTAACTCGGC


Use for*
Ndel at Atlg68060

M~sel at Atlg67865

M~nlI Atlg68140

aTagI at ATIG67850

Xbal at BAC Fl2A21

RLD 138/Ler 81

RLD 119/Ler 93

RLD 153/Ler 111

RLD 143/Ler 128

RLD 92/Ler 86

RLD 102/Ler 102

RLD 118/Ler 98

Identification ofDPP

Identification ofDPP

Identification ofDPP

Identification ofDPP












Genotype dpp Wt Total X2 Value* P value*
dpp/+ RLD wild type 403 148 551 10.3662 >0.001
dpp/+ Col wild type 178 65 243 4.7407 >0.01
dpp/+ Ler wild type 322 135 457 2.8981 >0.05
* is the ratio of 2:1.


Genotype dpp Wt Total X2 Value (1:1) P value (1:1)
LD wild type/dpp 72 79 151 0.3245 >0.50
dpp/RLD wild type 75 106 181 5.3094 >0.01
Salk_143503/dpp 23 35 58 2.4828 >0.20
dpp/Salk_143503 44 114 158 31.0127 <0.001


Table 4-4. Single nucleotide polymorphism identified between the Ler and RLD ecotypes
Gene name Position* Ler RLD
Atlg67790 ATT C CGTAT GACGATAC ATA/T GAC CGATT C TTT A T
Atlg67760 ATTACTCAGC TCCATTAATT/GTTCAACTTCATC T G
Atlg67760 ATCAACCTTTTGTCTTTTAG/ACTGTCTTCACCT G A
Atlg67670 ATAGGATT TGT CGAGAC TTG/TT TT TTGTT TAT G T
* Red bold letters represent single nucleotide polymorphisms between the Ler and RLD
ecotypes


Table 4-2. Segregation of trichome phenotypes in F2
type plants


of the dpp mutant crossed to wild-


Table 4-3. Segregation of trichome phenotypes in Fl
crossed to wild type plants


of the dpp mutant reciprocally










Table 4-5. Segregation of phenotypes in the Fl of the dpp mutant crossed to the Salk lines
Gene name Salk line Position dpp Wt Total % dpp oC
Atlg67730 Salk_143503 300-UTR5 11 41 52 0.21 20
Atlg67730 Salk_143503 300-UTR5 9 22 31 0.29 24
Atlg67730 Salk_039982 300-UTR5 14 41 45 0.31 20
Atlg67730 Salk_039982 300-UTR5 10 40 50 0.20 24
Atlg67750 Salk_095735c 300-UTR5 6 13 19 0.32 20
Atlg67750 Salk_017335c 300-UTR5 34 28 62 0.55 24
Atlg67750 Salk_017336 300-UTR5 36 50 86 0.42 24
Atlg67680 Salk_025786 Exon 11 11 22 0.50 24
Atlg67770 Salk_129146 Exon 34 47 81 0.42 24
Atlg67620 Salk_100543 Exon 26 32 58 0.49 24
Atlg67630 Salk_017965 300-UTR3 20 40 60 0.33* 24
*The hybrid seeds were collected 9 days after pollination, whereas others were 15 days
after pollination.




















































Figure 4-1. The dpp mutant trichomes in the RLD genetic background. (A), (B), and (C):
dpp mutants; (D), (E), and (F): RLD wild type. Bar = 1000 Clm in (A) and (D),
100Cl in others.








110














ATPase

I


Chromosome 1


Recombinants and BACs


1292 107
Total T30E16


31
F12P19


5
F5A8


4/0
T1F15


0/1
F12A21
1- I


8
T22E19


10 38 123
T26J14 T6C23 ATPase


The mutated gene


P
~~Cnlr qilcna
em" r~
~UII~1~


T~""
qilcmr
~f"' T"~""
y~rrrl r!~nasr~rsrl


~"rm'
nlC'tl


~nll
~"


Rl~n*ll
TW"'
*l~rus
r


Figure 4-2. Positional cloning of DPP. The dpp mutation was mapped near SSLP marker
ATPase on chromosome 1. Additional molecular markers were used to map the
dpp mutation to BAC clone F l2A21. The numbers of recombinants (out of 1292
chromatids screened) are given above BAC clones. The locations of all putative
genes on BAC clone Fl12A21 are listed. The numbers inside the flags above
specific genes are recombinant event numbers at that the specific location. DPP
is indicated by the vertical arrow.


1L~MIICIL114(
o~n,,

~'~C" ~'"U"'





A TC CT T A


T in RLD; A in Ler


QA.TA


CAT
470


Nd G A C C O A T
475 480r


Figure 4-3. Single nucleotide polymorphism between RLD and Ler wild types. N is
indicating a single nucleotide polymorphism in the gene Atlg67790 between
RLD and Ler wild types. N = T in the RLD wild type; N = A in the Ler wild
type


TOOT
630


T O C A Gi
535


Figure 4-4. Equivocal sequencing result using the DNA template from plants heretozygous
for the dpp mutation. The overlapping peaks at an exon in Atlg67760 are
showing in the rectangle, using DNA from dpp mutant plants.



















TOG
271


AC


C A AC AG
255 250


G O A A T
285


Figure 4-5. Sequencing result of Atlg67730 using the dpp mutant DNA as a template.
The indistinguishable bases are showed inside the rectangles, but they are
distinguishable in the repeated experiment in Figure 4-11.







?A G?


G: OG G OA A T G C A G A C AC C
320 325 h 330


Figure 4-6. Sequencing result of Atlg67730 using the dpp mutant DNA as a template.
The bases of interest are showing indistinguishable inside the rectangles.
















Salk lines
Homozygous for
T-DNA insertion


Pollen
The dpp mutant


~


IF11


The dpp mutant plant cells


Figure 4-7. Schematic explanation ofDPP identification. The cross ofdpp mutants with
T-DNA insertion lines for DPP identification. The Fl is generated by a cross
with dpp mutants as the male parent and Salk lines homozygous for T-DNA
insertion as the female parent. DNA is amplified by PCR with primers flanking
the inserted T-DNA. No PCR product is amplified because of a large size of T-
DNA between the regions, parts of which end sequences are identical with the
designated primer pairs.























-
- ,:---. -
-
-
-
-

-


-
- -
-
-
-
- -

-


F1 plants





SLarger deletion I

dpp wt


SSmaller deletion

dpp wt


5000 bp
3000 bp
2000 bp
800 bp
400 bp

100 bp

50 bp


5000 bp
3000 bp
2000 bp
800 bp
400 bp

100 bp

50 bp


Figure 4-8. Schematic explanation of deletion identification in dpp mutants. Fl seedlings
are segregated into the dpp and wild-type (wt) plants. DNA is extracted from
these two kinds of plants, using for PCR as the template with primers indicated
in Figure 4-7. The size of PCR products from dpp mutants and wt DNA are
identified by running gels. The deletion ofdpp mutations is roughly determined
by band positions.

































"Ji~


Figure 4-9. Novel phenotypes in the Fl of dpp mutants and T-DNA insertion lines. (A),
(B), (C), and (D): RLD wild type; (E), (F), and (G): dpp mutants heterozygous
for the dpp mutation and wild type. I, J, and K: Fl heterozygous for the dpp
mutation and T-DNA insertion. (H) Showing the wax stem of the Col wild type
(left) and the waxless stem of the Fl plant (right). (L) Showing fused organs of
floral meristems. Bar = 1000 Clm in (A), (E), and (I); 100 Clm in others.

















G O O O C:
a~i


AAT
3510


Figure 4-10. Equivocal sequencing result of Atlg67730, using the DNA from dpp mutants
as the template by the reward primer in the repeated experiment. The bases of
interest are showing indistinguishable in the rectangle.


C:A A


C A
275


rj A C G Ij
200


O O


Figure 4-11. Unequivocal sequencing result of Atlg67730, using the same DNA as in
Figure 4-10 in the repeated experiment by the forward primer. The bases of
interest, indicating two indistinguishable bases in Figure 4-5, are showing
unequivocal .


G, IGIAC/A G A G A


Yes C


AATOr
285



















~lllbil0
Rtlqr7130


720S2 7095 720S1


p ~Y


T-DNA
insertion


730S2 4202 bp 730S1


2055 bp


720cd2


730cd1


4967 bp


DEd8 DEd7


Figure 4-12. Identification of DPP. Gene structure of DPP and its neighboring genes are
showing exons in green segmental lines. The T-DNA insertion is showing
between the coding sequences ofDPP and its neighbor genes. The specific
primers used in the DPP identification are showing their positions.



































118


















aGocO
("1 nos


A A TA 0
son


;A A CA A
seeb


Figure 4-13. Unequivocal sequence result of Atlg67730, using the DNA from ded plants
of Fl in the combination of dpp mutants and Salk_143503 as the template. The
deleted bases of interest are showing inside the rectangle.


G G G O C A
305


G A G A C A
315


Figure 4-14. Unequivocal sequencing result of Atlg67730, using the DNA from the wild-
type plants of Fl in the combination of dpp mutants and Salk_143503 as the
template. The bases of interest are showing inside the rectangle.


ATOCA
310





























Figure 4-15. GC deletion in dpp cloned into pBluescript SK





SWild type


\i'


GC deletion


SC A A TA a O
3P05


60
300


SA O A C A A
310


GGCAA
300


TOCA
305


e AG ACA A
310 315


Figure 4-16 Wild-type DPP cloned into pBluescript SK






































Figure 4-17. Transgenic plants with the mutated DPP in distinct genetic
backgrounds. (A), (E), and (I): RLD wild type; (B), (F) (J), and (M):
transgenic plants with the mutated DPP in the RLD wild-type ecotype;
(C) (G) (K), and (0): transgenic plants with the mutated DPP in the Col
wild-type ecotype; (D), (H) (L), and (P): Col wild type. (N): Fl between
dpp mutants and Salk_143503. Bar = 500 Clm in (A), (B), (C) and (D);
200 Clm (E), (F) (G), and (H); 100 Clm in (I), (J), (K), and (L); 1000 Clm in
(M), (N), (0), and (P).









CHAPTER 5
IRREGULAR TRICHOM~E BRANCH 4 INT ARABIDOPSIS ENCODES THE PLANT
HOMOLOG OF THE 64 KDA SUBUNIT OF CLEAVAGE STIMULATION FACTOR AND
REGULATES TRICHOME MORPHOGENESIS AND FLORAL DEVELOPMENT

Introduction

Polyadenylation includes two sequential reactions-endonucleolytic cleavage and

adenylate-residue addition at specific poly(A) sites in 3' non-coding sequences of pre-mRNAs.

During eukaryotic development, polyadenylation plays an important role in the regulation of

gene expression through contributions to transcription (Zorio and Bentley, 2004), mRNA export

to the cytoplasm (Vinciguerra and Stutz, 2004), mRNA stability, and translation efficiency

(Wilusz et al., 2001). The specific cleavage at the 3'-end of pre-mRNAs is performed mainly by

two complexes: cleavage and polyadenylation specificity factor (CPSF) and cleavage stimulation

factor (CstF) (Zhao et al., 1999). In mammalian cells, the former is a heterotetramer consisting of

the CPSF-160, CPSF-100, CPSF-73, and CPSF-30 subunits, which are necessary for both

cleavage and polyadenylation (Murthy and Manley, 1992), while the latter is a heterotrimer that

is composed of CstF-77, CstF-64, and CstF-50 subunits (Takagaki et al., 1990). A a recent model

hypothesized that each component in the CstF complex is a homodimer (Bai et al., 2007). The

CstF complex is required for cleavage, but is dispensable for the synthesis of the poly(A) tail.

CPSF-73 is an endoribonuclease that cleaves pre-mRNAs at poly(A) sites (Mandel et al., 2006).

CstF-77 functions as a bridge between CstF-64 and CstF-50 (Takagaki and Manley, 1994), and

also interacts with the CPSF complex through CPSF-160 (Murthy and Manley, 1995). CPSF-160

specifically binds to the canonical AAUAAA element (Murthy and Manley, 1992). CstF-64

recognizes and binds to a conserved U- or G/U-rich downstream sequence element (DSE) in pre-

mRNAs (Takagaki and Manley, 1997). The CstF complex cooperates with the CPSF complex to









facilitate stable binding of both complexes to a pre-mRNA, and enhances polyadenylation

efficiency (Murthy and Manley, 1992).

CstF-64 is the limiting component for assembly of the active CstF complex (Takagaki et

al., 1996). There is increasing evidence that shows that altered levels of CstF-64 expression has a

significant impact on 3' mRNA processing, and thus regulates specific gene expression, which

modulates eukaryotic development. For example, decreases in CstF-64 expression had a variety

of effects on B-lymphoma cells (Takagaki and Manley, 1998). Decreased CstF-64 expression in

chicken B cells caused lower IgM heavy chain mRNA accumulation as well as an isoform

change due to the utilization of an alternative poly(A) site. Also the cell cycle prolonged, cells

arrested in GO/Gl, and eventually entered apoptosis. In contrast, increased CstF levels in

differentiating mouse and human B-cells had the opposite effect; the cells transitioned from GO

to S phase and were induced to proliferate (Takagaki et al., 1996; Martincic et al., 1998). In

mouse macrophages following lipopolysacchride stimulation, a 10-fold increase in CstF-64

expression significantly altered the expression of 51 genes (Shell et al., 2005). The change in

gene expression was due to alternative polyadenylation through the choice of either a strong (or

weak) poly(A) site, which removed (or retained) instability elements in the mature transcripts

(Shell et al., 2005).

Five poly(A) sites, L1 to LS, in the adenovirus maj or late transcription unit (MLTU) are

used for generation of distinct mRNAs through alternative polyadenylation (Larsson et al., 1992;

Mann et al., 1993). During the course of adenovirus infection, the activity of CstF in HeLa cells

varies (Mann et al., 1993). In the later phases of infection, the activity of CstF was substantially

decreased. Additionally, the interaction of CstF with the L3 poly(A) site of the MLTU was found

to be more stable than the interaction of CstF with the L1 poly(A) site and the L3 site was used









three times more frequently than the L1 site. These events are essentially a reverse of the events

observed during the early phases of infection (Mann et al., 1993). CstF-64 expression was also

reported to vary in other cell types. Mouse testicular cells contain at least 250-fold more CstF-64

mRNA than liver cells (Dass et al., 2001). Male germ line cells in meiosis, have no detectable

CstF-64 mRNA (Wallace et al., 1999).

The Arabidopsis homologs of mammalian CstF have been cloned, and were named

AtCstF-77, AtCstF-64 and AtCstF-50. Additionally, biochemical assays in vitro show that

AtCstF-64 binds to mRNA 3' non-coding regions, and interacts with AtCstF-77 similarly to the

mammalian CstF-64 (Yao et al., 2002). However, the role of AtCstF in the control of plant

development has not been demonstrated. Here, we report the identification of a developmental

role for AtCstF-64 in Arabidopsis. Our results show that AtCstF-64, encoded by the ITB4 gene,

is highly expressed in growing and proliferating cells, and it is required for normal trichome

morphogenesis and floral development. Although the basic mechanism is widely conserved in all

eukaryotic cells, results of our functional analysis of CstF-64 suggest that key differences exist in

the specific mechanisms of mRNA 3'-end processing among yeast, plants and mammals,

Materials and Methods

Plant Materials and Growth Conditions

The itb4-1 and zvi-3 single mutants and the itb4-1 zvi-3 double mutant are in the

Columbia (Col) genetic background (Zhang et al., 2005b). A novel itb4 mutant allele (itb4-2) is

also in the Col genetic background. Seeds were sown on a soil-less potting medium, Fafard 2

Mix (Conrad Fafard Inc. Agawam, MA 01001, USA). Seedlings were grown at 240C under

constant light, which was provided by 40W cool white fluorescent tubes. Plants were watered

with PGP nutrient solution (Pollock and Oppenheimer, 1999) every two weeks.









The itb4-2 Mutant Isolation and ITB4 Cloning

The F2 mapping population for cloning ITB4 was generated as described by Zhang et al.,

(2005). A total of 23 10 phenotypically itb4 zwi-3 double mutant plants were selected from the F2

population. One of the cotyledons was removed from each selected plant for DNA extraction

with the RED Extract-N-Amp plant PCR kit (Sigma-Aldrich, St. Louis, MO). The isolated DNA

was used to map the itb4-1 mutation relative to simple sequence length polymorphisms (SSLPs)

(Bell and Ecker, 1994; Lukowitz et al., 2000). After the itb4-1 mutation was mapped to a narrow

region where the SSLP markers were not available, cleaved amplified polymorphic sequence

(CAPS) markers were used (Konieczny and Ausubel, 1993). Landsberg er genomic information

provided by Monsanto's Cereon SNP database (http://www. arabidopsis. 0rg/Cereon/index~ijsp),

was used to design the SSLP and CAPS primers for fine scale mapping. Finally the itb4 mutation

was narrowed down to an approximately 41-kb region, where no additional markers were

available. All of the putative genes in this region were examined for mutations by sequencing

PCR products amplified from itb4-1 mutant DNA. Putative mutations were re-checked through

sequencing or enzymatic digestion of the PCR products from different template DNAs isolated

from 20 individual double mutant plants from the F2 mapping population, the zwi-3 single

mutant and Col wild type. The primers used for these gene amplifications are listed Table 1.

To confirm that we had identified ITB4, the SALK T-DNA insertion line, SALK 131655,

was ordered from Arabidopsis Biological Resource Center (The Ohio State University,

Columbus, OH). The seeds were sown on plates containing MS medium with 1% sucrose and 50

Clg/ml kanamycin. Kanamycin resistant seedlings were transferred into pots, and seeds of

individual plants were collected separately and sown into different pots. The phenotypes of the

resulting plants were characterized. The T-DNA insertion in SALK_131655 was confirmed by









PCR and sequencing using the ITB4-specific primers. Loss of detectable full-length ITB4

transcript in this allele was verified by RT-PCR.

Transgene Construction

For ITB4 overexpression, the ITB4 genomic region was expressed from either the

constitutive 35S promoter, or the trichome and root-hair specific GL2 promoter (Szymanski et

al., 1998). The resulting constructs were named 35S:ITB4 and GL2:ITB4, respectively. The

genomic sequence was amplified by PCR from Col wild type genomic DNA. The PCR fragment

was cloned into pENTRIA (Invitrogen, Carlsbad, CA) and transferred into either pAM-PAT-

GW (a gift from B. Ulker, Max Planck Institute for Plant Breeding, Cologne, Germany) for

expression from the 35S promoter or pCK86 (a gift from A. Schnittger, Max Planck Institute for

Plant Breeding) for expression from the GL2 promoter through an LR recombination. To localize

ITB4, 35S:ITB4-GFP and GL2:ITB4-GFP were constructed. The PCR fragment was amplified

using primer pairs that introduced an Ncol site at both ends of the PCR products. The digested

PCR products were cloned into the Ncol site on the GFP fusion vector, pAVA319 (von Arnim et

al., 1998). The resulting gene fusions were liberated by digestion with Smal and Xhol, and

transferred to either pAM-PAT-GW or pCK86, which were digested by Pstl, filled in using

Klenow, and then digested with XSbol. To produce Gall:ITB4, the full-length cDNA of ITB4 was

amplified by RT-PCR using total RNA from six-week-old Col wild-type plants. The PCR

product was cloned into pYES-DEST52 (Invitrogen, Carlsbad, CA) through an LR

recombination. To produce 35S:hCstF-64, 35S:h zstF-64,GL2:hCstF-64, and GL2:h zstF-64,

the same 5'-leader sequence (TL) and 3'-terminator (Ter) as in the construct of 35S:ITB4-GFP

were used. The coding sequences of hCstF-64 and h zOstF-64 were amplified by PCR from the

plasmids containing hCsF-64 or h OstF-64 (gifts from C. C. Macdonald, Texas Tech University









Health Sciences Center, Lubbock, TX). The PCR products were cloned into the modified

pNENRIA with TL and Ter elements using KpnI and BglII. The resulting TL-hCstF-64-Ter and

TL-h zCstF-64-Ter fusions were transferred into either pAM-PAT-GW or pCK86 through an LR

recombination. All primers used for the constructs are listed in Table 1, and all PCR products

were sequenced to confirm that no mutations were introduced. Sequencing was performed by the

Interdisciplinary Center for Biotechnology Research (ICBR) at the University of Florida.

RNA Extraction and RT-PCR

Total RNA was extracted from six-week-old Col wild-type plants using the RNeasy Plant

Mini Kit (Qiagen Inc. Valencia, CA) according to the manufacture's instructions. The full length

ITB4 cDNA was amplified using the cMaster RT plus PCR System (Eppendorf AG, Hamburg,

Germany). First strand cDNA synthesis was primed using oligo (dT)20. The cDNA was amplified

using the ITB4-specific primers (see Table 5-1). The PCR products were sequenced by ICBR at

the University of Florida.

Plant and Yeast Transformations

For ITB4 subcellular localization, the 35S:ITB4-GFP construct was used to transiently

transform onion epidermal cells by particle bombardment using the Biolistic PDS-1000/He

Particle Delivery System (Bio-Rad, Richmond, CA). The transformation was carried out using

the manufacturer's instructions. Briefly, 5 CIL DNA (1 Clg/CIL) was precipitated onto 3 mg gold

microcarriers (Bio-Rad) of 0.6 Clm in diameter, by adding 50 CIL 2.5 M CaCl2 and 20 CL 0. 1 M

spermidine. After the precipitated DNA was washed once each with 140 CIL 70% and 100%

ethanol, it was resuspended in 50 CIL 100% ethanol. Ten microliters of this DNA suspension

solution was spread on one 1,100 psi rupture disk. The onion epidermal tissue was placed on

solid MS medium for bombardment. Fluorescence was visualized after a 36-hour incubation at

room temperature in darkness.









The 35S:ITB4, GL2:ITB4, 35S:ITB4-GFP and GL2:ITB4-GFP constructs were used to

transform itb4-1 zwi-3 double mutants and Col wild type plants. The 35S:hCstF-64, 35S: h TCstF-

64, GL2:hCstF-64 and GL2:h TCstF-64 constructs were used to transform itb4-1 zwi-3 double

mutants. Transformation was accomplished using the floral dip method (Clough and Bent, 1998).

The transgenic plants were selected using a 1000X dilution of Finale (5.78% glufosinate-

ammonium) (Famnam Companies, Inc., Phoenix, AZ).

The Gall:ITB4 construct was used to transform yeast using a Yeast Transformation Kit

(Sigma-Aldrich, St. Louis, MO) according to the manufacture's instructions. The yeast strains

transformed were mal4-1, rnal5-1, and w303 wild type. These yeast strains were a generous gift

from Frangois Lacroute, Centre de Genetique Moleculaire, Yvette, France. The mal4-1 strain

contains a mutation in RNAl4 gene, a homolog of CstF-77 in mammals and AtCstF-77 in

Arabidopsis; the mal5-1 strain contains the mutation in RNAl5 gene, a homolog of CstF-64 in

mammals and AtCstF-64 in Arabidopsis (http://www.uky .edu/~aghunt00/polyA20 10.html).

Morphological Analysis

Arabidopsis trichomes were isolated from leaves and stained with Toluidine Blue as

previously described (Zhang and Oppenheimer, 2004). About 50 plants of itb4-2 and Col wild

type were grown in constant light. After one week when the first flower opened on each plant, 50

flowers for each genotype were selected from different plants, and dissected under the

microscope to determine the number of sepals, petals and stamens.

Microscopy

GFP images were obtained with a Zeiss Axiocam HRm camera mounted on a Zeiss

Axioplan 2 Imaging microscope using Zeiss Filter set 10 (excitation: 450-490, dichroic: 510 LP,

emission: 515-565). Zeiss Filter Set 02 (excitation: 365, dichroic: 395 LP, emission: 420 LP) was









used to collect fluorescent signal from DAPI stained tissue. A Zeiss Axiocam MRc5 camera

mounted on a Zeiss Stemi SV11 dissecting microscope was used to obtain light micrographs.

Environmental Scanning electron microscopy was carried out at the National High Magnetic

Field Laboratory (Tallahassee, FL), in an Electroscan Model E-3 environmental scanning

electron microscope. Tissue samples were mounted on moist paper towels and scanned at 20 kV

under 1-2 torr pressure.

in situ Hybridization

Fixation, dehydration, and embedding of Arabidopsis inflorescences and young siliques

were performed as previously described (Zhang et al., 2005a). RNA probes were made using the

DIG RNA Labeling Kit (Roche Diagnostics, Indianapolis, IN) following the manufacturer's

instructions. The gene-specific primer pairs and their antisense primer pairs were designed such

that a T7 promoter was introduced (Table 5-1) and the transcription templates were prepared by

PCR. The steps of in situ hybridization were essentially those described by the Meyerowitz's

laboratory at http://www.its. caltech. edu/~plantlab/html/protocols.html, except for the following

changes: the 50% Denhardt's Solution (in the hybridization solution) was replaced with 10%

Blocking Reagent (Roche Diagnostics, GmbH, Mannheim, Germany), the hybridization

temperature was 450 C, and the washing temperature was 500 C.

Results

ITB4 Encodes AtCstF-64

To further understand the role of ITB4 in trichome development, we cloned the ITB4 gene

using positional cloning methods. We generated an F2 mapping population through a cross of the

itb4-1 zwi-3 double mutant with Ler wild type. The itb4-1 zwi-3 double mutant was used as a

parent for the following reasons: first, the itb4-1 mutant trichomes have three branches with a

subtle change only in trichome branch length; second, the zwi-3 mutant trichomes have two









branches and its mutation was known to be located on chromosome V, unlinked to itb4-1; finally,

the itb4-1 zwi-3 double mutant displayed an unequivocal trichome phenotype, i.e., unbranched

trichomes. Therefore, we were able to unambiguously identify itb4 plants in the F2 mapping

population. A total of 23 10 plants with unbranched trichomes were selected from the F2

population. Through the use of SSLP markers, we mapped the itb4-1 mutation to a region

between BAC clones F23N20 and F28P5 BAC. Using CAPS markers, itb4-1 was further

narrowed down to an approximately 41-kb region on the BAC clone F l4023 according to the

TAIR database (Figure 5-1A). This region contains 10 putative genes. All open reading frames in

this region were amplified by PCR using genomic DNA from itb4-1 seedlings. The PCR

products were sequenced, and a C to T transition at base 1896 (starting at the A in the ATG start

codon) was found in gene Atg71800. The mutation, which was located in the last second exon of

Atlg71800, created a new TAA stop codon, causing an 88 amino acid truncation from the C-

terminus of the putative ITB4 protein (Figure 5-1A).

To confirm that this mutation also exists in the itb4-1 zwi-3 double mutants, we randomly

selected 20 DNA samples from the F2 mapping population for amplification of the target

sequence that contains the mutated base. Because the mutation creates an M~se I cut site in the

amplified target sequence, the PCR products were digest with M~sel. Our results showed all the

itb4-1 zwi-3 double mutants indeed had the mutated base in the ITB4 gene (Figure 5-1B).

To confirm that the phenotype observed in the itb4-1 mutant is caused by the mutation in

Atlg71800, the wild type Atlg71800 gene was expressed in itb4-1 zwi-3 double mutant plants.

The construct used for transformation was a 2800-bp genomic fragment, which contained the

5'UTR (250 bp), the Atlg71800 coding sequence (2245 bp) and the 3' UTR (300 bp), and was

expressed from either the 35S promoter or the GL2 promoter. Over 30 independent transgenic










plants for each construct displayed trichomes with two branches identical to the trichomes on

zwi-3 mutants (Figures 5-2C, 5-2D). Occasionally, three-branched trichomes were observed on

some transgenic plants (Figure 5-2E). The rescue of the itb4-1 mutant phenotype by the

Atlg71800 coding sequence demonstrates that ITB4 is Atlg71800.

Loss-of-function Mutations in ITB4 Cause Aberrant Development of Trichomes and
Flowers

ITB4 contains three functional domains: the RNA recognition motif (RRM) that directly

binds to mRNA; the Hinge domain that interacts with CstF-77 in mammals and Arabidopsis; and

the PC4/subl/res1 domain that interacts with polymerase processivity factors in mammals and

yeast (Herr et al., 2006). The mutation in itb4-1 causes a truncation of 88 amino acids at the C-

terminus of ITB4. The truncated part includes the KIWI/KELP domain (Cormack et al., 1998),

which is homologous to PC4 in mammals and Sub1 in yeast (Herr et al., 2006). The deletion of

the KIWI/KELP domain in ITB4 causes only a slight alteration in trichome branch length and

branch position. To further analyze the function of ITB4 in trichome morphogenesis, we used a

reverse genetic approach to seek a complete loss-of-function mutation in the ITB4 gene. By

searching the SALK T-DNA Express Database (http://signal.salk. edu/cgi-bin/tdnaexpress), we

identified eight lines that contained insertions in the Atlg71800 gene: Salk_131655,

Salk 088885, Salk 088876, Salk 088877 Salk 150929, Salk 133589, Salk 038729, and

Salk_150929. After selection for kanamycin resistance, the progeny from line Salk_131655

segregated plants showing a defective trichome phenotype. Additionally, the segregation ratio of

wild type to mutant was nearly 3:1. ITB4 in Salk_131655 is interrupted by a T-DNA insertion in

its last exon (Figure 5-1A). Plants homozygous for the T-DNA insertion in this line were

identified by PCR-based screening (Figure 5-1C). This mutant allele was named itb4-2. ITB4










expression was not detectable in the itb4-2 mutant by RT-PCR (Figure 5-1D), indicating that

itb4-2 is likely to be a complete loss-of-function mutant allele of ITB4.

The mutant phenotype caused by the T-DNA insertion mutation in the itb4-2 allele is

considerably more severe than the itb4-1mutant. Because the latter exhibits shorter trichome

branches and separate branch positions, it probably represents a partial loss-of-function mutation;

however, the former shows dramatic defects in trichome morphogenesis and floral development.

The itb4-2 plants displayed changes in both trichome shape and trichome cell fate. The

branch length and branch positions of the itb4-2 trichomes were clearly distinguished from wild

type. In Col wild type trichomes, 96% trichomes have three branches (Table 5-2) and the lengths

of the branches are almost equal. Additionally, the positions of the primary and secondary branch

are adj acent or close to each other (Figure 5-3A). However, the branch number of the itb4-2

trichomes covered a wide range from one branch (Figures 5-3D, 5-3F, Table 5-2) to five

branches (Figure 5-3J, Table 5-2) and the percentage of three-branched trichomes is notably

decreased. In contrast, the percentage of two-branched trichomes is prominently increased (Table

5-2). The branch lengths and positions of the trichomes on the itb4-2 mutant show the same

characteristics as the trichomes on the itb4-1 mutant, i.e., unequal branch lengths and separated

branch positions (Figures 5-3B, 5-3G-J) (Zhang et al., 2005b).

Interestingly, the itb4-2 mutation produced trichome clusters, which appeared as "twins"

on itb4-2 plants. The twin frequency was 2.52% in the itb4-2 mutant, but never observed in Col

wild type (Table 5-2). The branch numbers of the twin trichomes varied (Figures 5-3C, 5-3F, 5-

3K-N), and they all seem to be conj oined at the bottom of their stalks (Figures 5-3C, 5-3E, 5-3F,

5-3K-J). Separation of the twin trichomes was resistant to treatments with both EGTA and










pectinase (Figure 5-3E) (Zhang and Oppenheimer, 2004), which suggests that the trichomes

share part of a cell wall.

The other maj or defect of the ibt4-2 mutant is abnormal development of flowers.

Beginning with stage 3 during floral development on wild type plants, the abaxial sepal

primordia arise first, followed by the adaxial primordia. Entering stage 4, they elongate, curve

inward and cover the dome-shaped meristem before petals arise (Figure 5-4A). The developing

primordia rapidly enlarge from 30 Clm at stage 3 to 70 Clm at stage 4 (Smyth et al., 1990).

Compared with wild type, the floral primordia on the itb4-2 mutant displayed aberrant

development. First, the sepals elongate slowly and do not cover the floral organ primordia after

the petals arise (Figure 5-4B, large arrow). Second, the abaxial sepals preferentially grow

(Figures 5-4B, 5-4F) and often fuse with adjacent sepals (Figure 4B, small arrow). Additionally,

the numbers of sepals and petals is significantly increased (Figures 5-4B, 5-4C, 5-4F, 5-5); floral

buds with seven sepals were often observed (Figure 5-4C). Throughout flower development, the

defective sepals were unable to completely enclose the stamens and carpels as in wildtype

(Figures 5-4D, 5- 4E). Third, the stigmatic papillae were aberrant (Figures 5-4H, 5-41). On wild-

type stigmas, the papillae stand straight and are regularly arranged, but the papillae on the itb4-2

mutant displayed irregular shapes and clusters (Figures 5-4H, 5-41). The anthers of the itb4-2

mutant contained few pollen grains and the ones that did not form, were mostly unviable

(Figures 5-4K, 5-4L). In spite of the extra floral organs, the overall floral bud size was smaller

than wild type (Figures 5-4B, 5-4E).

The itb4-2 mutant plants also showed a relatively minor alteration of rosette leaf color and

shape. Col wild-type plants produce green rosette leaves with smooth edges (Figures 5-5A, 5-5C,

5-5E). However, itb4-2 plants produced yellow-and-green mosaic rosette leaves with serrated









edges (Figures 5-5B, 5-5D). These defects are most prominent in the first leaf pair (Figure 5-6B),

while subsequent leaves display a gradual transition toward normal leaf morphology during

vegetative growth (Figures 5-6D, 5-6F).

ITB4 is Highly Expressed in Growing and Proliferating Cells

To gain insight into why itb4 mutations caused the defects in specific cell types and

organs, we examined the ITB4 expression pattern by in situ hybridization. We found that ITB4 is

highly transcribed in developing trichome cells, embryos, meristems and floral primordia during

plant vegetative growth and reproductive development (Figure 5-7). At the globular stage of

embryo development in Col wild type, ITB4 was highly expressed in the developing embryo, but

not in cells of the suspensor (Figure 5-7A). At the heart stage, ITB4 expression became stronger

(Figure 5-7B), but in mature embryos, ITB4 expression decreased (data not shown). During

germination, ITB4 was expressed in the apical meristem and provascular cells. The actively

dividing cells in the meristem showed higher ITB4 expression than the growing provascular cells

(Figure 5-7C). Developing trichome cells displayed a higher ITB4 expression level than other

epidermal cells (Figure 5-7E). At stage 1 of trichome development, ITB4 is strongly expressed.

The nuclei of trichomes at this stage undergo endoreduplication from 2C to an average of about

8C with a concomitant increase in nuclear volume. This increase in nuclear size distinguishes

cells committed to the trichome fate from other epidermal cells (Hulskamp et al., 1994). The

strong expression of ITB4 remains until stage 3 or 4, at which time the trichome size increases

rapidly from about 20 Clm to 400 lm. During flower development, ITB4 is strongly expressed in

floral meristems (Figure 5-7F). At later developmental stages, strong expression oflTB4 can be

observed in developing stamens and carpels, but weak expression is observed in developed

sepals (Figure 5-7G).









Loss of ITB4 Function Alters the Expression Pattern of Perianth Organ Identity Genes

Complete loss-of-function of ITB4 caused an increase in the number of sepals and petals,

but no difference for stamen and carpel numbers. To understand how ITB4 influences perianth

development, we examined the expression of the floral organ identity genes, AP1, AP3 and Pl in

the itb4-2 mutant and Col wild type flowers through in situ hybridization. At stage 3 of floral

development, API expression domain is restricted to the outer two whorls because AG represses

API expression in the inner two whorls of wild type flowers (Figure 5-8A) (Gustafson-Brown et

al., 1994). However, in the itb4-2 mutant, in addition to its strong expression in the outer two

whorls at stage 3, API is ectopically expressed in the inner two whorls at stage 3 and even in

stamens and carpels at stage 7 (Figure 5-8B). In wildtype, AP3 is expressed from stage 3 in the

presumptive second and third whorls. After floral stages 5 and 6, AP3 is expressed throughout

the developing petals and stamens, and at the adaxial base of sepals (Figure 5-8C) (Jack et al.,

1992). In itb4-2 mutant flowers, AP3 expression appears precociously and ectopically in the

inner two whorls at the stage 3 (Figure 5-8D). The PI expression pattern in ibt4-2 mutants is

indistinguishable from that in the wild type (Figures 5-8E, 5-8F).

ITB4 Localizes to Nuclei, but Does Not Functionally Complement Tts Homolog in Yeast

ITB4 was annotated as a cleavage stimulation factor by both the Arabidopsis Genome

Initiative (http://arabidopsis .org/info/agi.html) and GSF-MIPS

(http://mipsggsf de/proj /plant/j sf/athal/searchj sp/index.j sp). The full length cDNA of ITB4 was

amplified by RT-PCR using Col wild type RNA as a template. The PCR product is identical with

the computer-based prediction of both of the databases mentioned above. In silico translation of

the ITB4 coding sequence predicts a protein of 461 amino acids with an isoelectric point of 9.32.

This protein sequence is identical to that reported by Yao et al. (2002) for AtCstF-64. Therefore,

we renamed the ITB4/Atlg71800 protein to AtCstF-64.









AtCstF-64 functions in the cleavage reaction for mRNA 3' end formation. It contains the

same three functional domains as its homologs in yeast and mammals. It is generally believed

that the CstF complex in plants functions in a similar way to the animal counterparts for mRNA

3'-end processing in nuclei (Yao et al., 2002; Herr et al., 2006). To support this idea, a GFP-

tagged version of AtCstF-64, 35S:ITB4-GFP, was transiently expressed in onion epidermal cells

through biolistic transformation. The ITB4-GFP signal was found in nuclei (Fig. 5-9). This result

is consistent with the nuclear location of RNAl5 in yeast (Bonneaud et al., 1994) and CstF-64 in

mammals (Schul et al., 1996). To test the functionality of ITB4-GFP in vivo, 35S:ITB4-GFP and

GL2:ITB4-GFP were used to transform itb4-1 zwi-3 double mutant plants through

Agrobacterium-mediated stable transformation. Both constructs rescued the unbranched

trichome phenotype of the itb4-1 zwi-3 double mutant, and transformants showed the two-

branched trichome phenotype of the zwi-3 mutant. However, the nuclear localized ITB4-GFP

signal was weaker than that observed in onion cells (data not shown). The nuclear localization of

the fusion protein supports a function for AtCstF-64 in pre-mRNA processing in Arabidopsis.

To determine if AtCstF-64 function is conserved, we examined the ability of AtCstF-64 to

complement yeast CstF subunit mutants. GAL1:ITB4 was used to transform three different yeast

strains: rnal51-, rnal4-1, and w303 (wild type). The rnal5-1 strain contains a mutation in

RNAl5, the counterpart of CstF-64 in mammals and AtCstF-64 in Arabidopsis; the rnal4-1

strain contains a mutation in RNAl4, the counterpart of CstF-77 in mammals and AtCstF-77 in

Arabidopsis. The mutations in both strains caused a temperature-sensitive phenotype. At the

permissive temperature, 280 C, the rnal5-1 and rnal4-1 strains grow. However, at the restrictive

temperature, 33o C, their growth stops. The transformed rnal5-1 or rnal4-1 strains that

contained the GAL1:ITB4 construct could not grow at 33o C, but the transformed wild type strain










grew normally (data not shown). These results indicate that AtCstF-64 does not functionally

complement its counterpart in yeast.

To further examine the functional conservation of AtCstF-64, we attempted to rescue the

itb4 phenotype using CstF-64 homologs from mammals. 35S:hCstF-64, 35S:h zCstF-64,

GL2:hCstF-64 and GL2:h zCstF-64 were used to stably transform itb4-1 zwi-3 double mutant

plants. The human counterpart of AtCstF-64, hCstF-64, exhibits 32% identity and 44% similarity

to the Arabidopsis protein sequence. The human hzCstF-64 protein is a paralog of hCstF-64 that

exhibits 74% protein sequence identity with hCstF-64 and is expressed in male germ cells to

maintain normal spermatogenesis (Dass et al., 2001; Dass et al., 2002). None of the transgenic

plants showed rescue or any other change in the itb4-1 zwi-3 double mutant trichome phenotype

(data not shown). These results indicate that the mammalian CstF-64 does not functionally

complement its counterpart in Arabidopsis.

Discussion

Polyadenylation is a common event that occurs in the nuclei of all eukaryotic cells, during

which a maj ority of mRNAs receive a string of A residues. Additionally, 25% mRNAs in

Arabidopsis use alternative polyadenylation sites (Meyers et al., 2004). Alternative

polyadenylation of a number of mRNAs has been shown to affect plant development (Cheng et

al., 2003; Quesada et al., 2005). Although there is increasing data to suggest that alternative

polyadenylation contributes to the control of gene expression in animals, its role in the regulation

of development in plants is not well understood. The results presented in this paper are the first

report that a plant homolog of a subunit of the CstF complex influences specific events during

plant development.









ITB4 Plays a Crucial Role in Trichome Morphogenesis and Floral Development

The itb4-1 mutation leads to a truncated AtCstF-64 protein that lacks the KIWI/KELP

domain, and itb4-1 plants display only a relatively mild trichome shape defect. The truncated

AtCstF-64 protein retains the other important functional domains such as the RRM and Hinge.

The function of the KIWI/KELP domain is not well understood in Arabidopsis. It has been

proposed that the KIWI/KELP domain interacts with general transcription factors for activation

of gene transcription (Cormack et al., 1998). Because AtCstF-64 is single copy gene in

Arabidopsis and lack of the KIWI/KELP domain produces few phenotypic effects compared

with the likely null itb4-2 mutation, it is likely that the KIWI/KELP domain is nonessential for

AtCstF-64 function. The relatively mild phenotype produced by the itb4-1 mutation greatly

contrasts with the phenotype of the itb4-2 mutation, which led to profound changes in trichome,

leaf, and flower development. These phenotypic defects suggest an important developmental role

for AtCstF-64.

The occurrence of twin trichomes in the itb4-2 mutant suggests that proper trichome cell

fate specification requires AtCstF-64 function. Normally, once an epidermal cell acquires the

trichome fate, division of that cell ceases. The twin trichomes seen in itb4-2 mutants are

reminiscent of the trichome clusters seen in siamnese (sim) mutants, where incipient trichomes

still divide due to a failure to properly enter the endoreduplication cycle (Walker et al., 2000).

This phenotype suggests that in itb4-2, mRNAs encoding proteins involved in the control of

endoreduplication may have altered in expression levels, and hence be direct or indirect targets

of AtCstF-64.

Plants homozygous for itb4-2 also displayed a significant increase in the number of sepals

and petals, compared to wild type, although the organs were relatively normal in appearance.

These results suggest that early developmental events are more sensitive to loss of At CstF-64









function than later differentiation events. This idea is supported by our finding that ITB4 is most

highly expressed in actively proliferating tissue and organ primordia. A similar developmental

role for CstF-64 is seen during mammalian cell differentiation. High expression of CstF-64 is

required for normal development of B-lymphocytes. Reduced expression of CstF-64 gave rise to

aberrant differentiation and apoptotic cell death (Takagaki and Manley, 1998) Our results

support the finding in mammals that the slowly growing or inactively dividing cells may be able

to tolerate lower levels of CstF than the rapidly growing or actively dividing cells (Takagaki et

al., 1996).

In contrast to the CstF-64 mutations in other organisms, the itb4-2 homozygous plants are

viable. Depletion of CstF-64 in chicken and mouse B lymphocytes caused apoptotic cell death

(Takagaki and Manley, 1998). RNAl5, the homolog of CstF-64 in yeast, is also essential for cell

viability (Minvielle-Sebastia et al., 1991). These differences between plant and animal CstF-64

mutants suggest that plants are more tolerant of loss of CstF-64 function than other eukaryotes.

The recent identification of ESP1 (Atlg73 840) in Arabidopsis suggests that there are at

least two complexes that contain CstF-64 homologs in Arabidopsis (Herr et al., 2006). The

enhanced silencing phenotype (esp) mutants affect gene silencing and are involved in RNA

metabolism. ESP1 encodes an AtCstF-64 like protein that lacks the RNA-binding RRM domain.

It has been postulated that the standard CstF complex contains AtCstF-64 that uses the RRM

domain to bind pre-mRNAs. The other putative complex contains ESP1 and uses a separate

RNA-binding protein to recognize pre-mRNAs (Herr et al., 2006). The two complexes function

redundantly in mRNA 3' end formation, which is likely to be a reason why the itb4-2 mutation is

not lethal.









Loss of AtCstF-64 Function Influences the Expression of Multiple Genes that Control
Floral Organ Development

In mammalian cells, altered levels of CstF-64 expression has been shown to influence the

expression of at least 51 genes and induce alternative poly(A) site selection (Shell et al., 2005).

This is also likely to also occur in plant cells. Our results showed that loss-of-function mutations

in ITB4 caused changes in trichome fate, shape, and floral structure. It is likely that these

phenotypic defects are due to changes in the expression of the genes involved in control of

trichome morphogenesis and floral development. There are at least 30 genes known to regulate

trichome morphogenesis (Schellmann and Hulskamp, 2005). For example, STI, AN, ZWI, FRC1-

4 and GL3 function as positive regulators that promote trichome branching (Hulskamp et al.,

1994; Oppenheimer et al., 1997; Luo and Oppenheimer, 1999); loss of function mutations in

these genes cause a decrease in the trichome branch number. Conversely, TFCA, RFI, KAK,

PYM, and SUZ4 function as a negative regulators that suppress trichome branching; mutations in

these genes result in an increase in trichome branch number (Krishnakumar and Oppenheimer,

1999; Perazza et al., 1999; Kirik et al., 2002). Mutations in ITB2 and ITB3 affect trichome

branch length (Zhang et al., 2005b), and mutations in TRY cause trichome clusters (Hulskamp et

al., 1994). It is likely that the pre-mRNAs from the above-mentioned genes may be substrates of

ITB4. Without ITB4 function, may not be correctly polyadenylated, and therefore, their

expression levels are likely to be altered. Similar events may occur in the genes that regulate

floral development. Regulators of floral organ identity genes may have altered expression

leading to observed changes in floral architecture.

Differences Exist in the Mechanism of mRNA 3' End Formation among Plants, Yeast and
Mammals

Formation of mRNA 3' ends is carried out by multiple trans-factors including CstF, CPSF,

cleavage factors (CF) and the poly(A) polymerase (PAP). Homologs of subunits for each of









these factors have been identified in yeast, plants and mammals (Zhao et al., 1999; Yao et al.,

2002; Elliott et al., 2003; Herr et al., 2006). It has been widely accepted that mRNA 3' end

formation is similar in all eukaryotic cells, based on the high level of protein sequence identity of

these factors, and the cooperative RNA-protein and protein-protein interactions within the

processing machinery (Yao et al., 2002; Elliott et al., 2003; Delaney et al., 2006; Herr et al.,

2006; Xu et al., 2006).

The results of our in vivo functional assays indicated that the homologs of CstF-64 in

yeast, plants and mammals are not functionally equivalent. It is possible that this is due to the

differences in the sequences of the poly(A) signals in pre-mRNAs in yeasts, plants and mammals

(Mogen et al., 1990; Mogen et al., 1992; Li and Hunt, 1997; Zhao et al., 1999; Loke et al., 2005;

Herr et al., 2006; Ji et al., 2007). The common minimal poly(A) signal is composed of an A-rich

sequence, a U-rich element, and a PyA cleavage site in all eukaryotes. Nonetheless, the

requirement for specific sequence elements differs greatly between yeast, plants, and animals

(Zhao et al., 1999).

In mammals, a single copy of the AAUAAA element is highly conserved and absolutely

necessary for precise 3' end formation. The AAUAAA element is located about 15-30

nucleotides upstream of the poly(A) site; and the DSE is located within 50 nucleotides (Zhao et

al., 1999). CPSF-160 and CstF-64 specifically recognize and bind to the AAUAAA element and

DSE in pre-mRNAs, respectively (Murthy and Manley, 1992; Takagaki and Manley, 1997), and

cleavage occurs preferentially at CA (Zhao et al., 1999). However, the plant poly(A) signal lacks

a consensus element as in mammals. It has a wide distribution of multiple AAUAAA-like

sequences, and the GU-rich elements are located upstream of the poly(A) site, not downstream as

in mammals, and cleavage occurs preferentially at PyA(Wu et al., 1995; Li and Hunt, 1997;









Zhao et al., 1999). Previous work has shown that plant cells do not properly recognize animal

polyadenylation signals (Hunt, 1987). It is therefore likely, that human CstF-64 does not

recognize plant polyadenylation signals. This may explain why the human CstF-64 was not able

to functionally complement the itb4-2 mutant in Arabidopsis. The yeast poly(A) signal seems to

be more similar to that in plants. In yeast, usually redundant A-rich and U-rich elements are

located upstream of the cleavage site, and no unambiguous DSEs have been identified. As in

plants, the preferential cleavage site is PyA. However, yeast and plants do not share a consensus

element in their poly(A) signal sequences (Zhao et al., 1999). This may explain our result that

ITB4 could not complement the yeast CstF-64 mutant.

In summary, we have shown that ITB4 encodes AtCstF-64, which is highly expressed in

growing and proliferating cells, and is required for normal trichome morphogenesis and floral

development. Loss of ITB4 results in aberrant floral architecture due, in part, to the altered

expression of floral organ identity genes. The finding that ITB4 is not functionally equivalent to

its yeast counterpart, and that the mammalian CstF-64 could not functionally complement the

itb4-2 mutant, supports the idea that there exists key differences between polyadenylation in

yeast, plants, and animals even though the basic mechanism is conserved in all eukaryotic cells

(Zhao et al., 1999; Yao et al., 2002; Elliott et al., 2003; Delaney et al., 2006; Herr et al., 2006; Ji

et al., 2007). Given the importance of 3' end processing for a host of cellular processes including

gene regulation and cell proliferation (Danckwardt et al., 2008), understanding these differences

is needed to unravel the role of CstF in development.

Future Perspectives

Polyadenylation is a common event that occurs in all eukaryotic nuclei, but alternative

polyadenylation events that affect plant development have rarely been reported. Our results show

that absence of the plant homolog of CstF-64 encoded by ITB4 causes pleiotropic effects in plant









development such as severe defects in trichomes and floral organs. ITB4 is highly expressed in

these rapidly expanding and proliferating cells. ITB4 protein was localized to nuclei. The itb4

mutants had altered gene expression of important floral developmental regulators. These results

provide a link between Cst-64 and plant development. However, the function of ITB4 is still far

away from being fully understood. Multiple questions need to be answered. The most important

is which genes are affected by either the aberrant alternative polyadenylation or the failure of

poly(A) addition in the itb4 mutants. To answer these questions, the following approach can be

used. Through microarray experiments using mRNA isolated from itb4-2 mutants and wild type,

genes whose expression is altered in the mutant can be identified. The poly(A) site of these genes

can be examined by using 3' rapid amplification of cDNA ends (3' RACE).

Although polyadenylation has been considered as a conserved mechanism in all eukaryotic

cells, and the 3' end formation machinery is found in mammalian, yeast and plant cells, our

results suggest that there may be important differences in 3' end processing between these

groups. We were unable to rescue a yeast CstF-64 mutant using the Arabidopsis coding

sequence, and the mammalian CstF-64 could not functionally complement itb4 mutants.

However, alternative explanations exist. First, the mammalian CstF-64 gene may not be properly

expressed in plants. To check this, we could use western blotting of proteins extracted from

transformants using an antibody to the mammalian CstF-64 protein. Likewise, we have to rule

out lack of expression before we conclude that ITB4 cannot rescue the yeast mutant.









Table 5-1. Primers used in this study


Primer name
itb4m F
itb4m R
ITB4g F
ITB4g R
ITB4cDNA F
ITB4cDNA R
ITB4GFP F
ITB4GFP R
CstF64 F
CstF64 R
zCstF64 F
zCstF64 R
ITB4il 1F
ITB4i 1R *
ITB4ic F *
ITB4ic R
ITB4i2 F
ITB4i2 R
ITB4i3 F
ITB4i3 R
APi F
APi R
APlic F
APlic R
AP3i F
AP3i R
Pi F
Pi R


Sequence (5' 3')
TGGCAAAAGAATAAACGAGGG
ATTCAGGGCATTCTAAGCGA
CTCCTATCGACGACGAATACGAAAG
AGGGGCCACAGGATTAAAACCA
CTCCTATCGACGACGAATACGAAAG
CTATGAAGGCTGCATCATGTGGTCTTGC
ATGGCTTCAT CATCATCCCA ACGACGC
TGAAGGCTGCATCATGTGGTCCTTGCTTG
ATGGCGGGTTTGACTGTGAGAGACCC
TACAGGTGCTCCAGTGGATTTCTGTATTTGTTCC
ATGTCGAGTTTGGCGGTGAGAGACCC
GGAGGAGGGAAACCCTAATCCAAGTGTGGG
ATGGCTTCATCATCATCCCAAC
GTGCCTTTGTCATTCTCAGCAA
ATGGCTTCATCATCATCCCAAC
GTGCCTTTGTCATTCTCAGCAA
CGCCAAATATTGTTCAGGCCC
TTGGTAATGCTTGGTGGGG
AAGCAGATTGGAGGGCCAGTAGATT
TTTGCGTAAACTGCGAACCGA
GGGAAGGGGTAGGGTTCAATTGAAGA
GAC AACAAGAGCAAC TTC AGCATCAC
GGGAAGGGGTAGGGTTCAATTGAAGA
GAC AACAAGAGCAAC TTC AGC ATCAC
CGAGAGGGAAGATCCAGATCAAGA
GCTAGAGAACATGATAATCGAAACCC
GGAGGAATGGATTGGTGAAGAAGGCT
GCCAGATAACTTCTGGTATTGGTCCA


Used for
itb4-1 mutation
identification
ITB4 expression
construct
ITB4 cDNA
amplification
ITB4-GFP
expression construct
CstF64 expression
construct
TCstF64 expression
construct
ITB4 in situ
hybridization
ITB4 in situ
hybridization
ITB4 in situ
hybridization
ITB4 in situ
hybridization
API in situ
hybridization
API in situ
hybridization
AP3 in situ
hybridization
PI in situ
hybridization


* Primers used for in situ hybridization included the T7 promoter sequence,
5' taatacgactcactataggg3 at the 5' end, for example: ITB4il1 R,
5' taatacgactcactatagggGTGCCTTTGTCATTCTCAGCAA' for the antisense RNA probe, and
ITB4ic F, 5 'taatacgactcactatagggATGGCTTCATCATCATCCCAAC' for the control sense
RNA probe.










Table 5-2. Alteration of trichome cell shape in the itb4-2 mutant
Trichome branches % a Total %Twin
Strain nmeb clstr
0 2 3 4 5 nme lse
Col wt 0 0.1 96.0 3.9 0 1120 0
itb4-2 3.4 18.0 70.3 7.8 0.5 1540 2.52
a Numbers represent percentages of the total number of trichomes with the indicated number of
branches.
b Total number of trichomes counted on at least ten leaves.
" Numbers represent percentages of the total number of trichomes that were present as twins.













nga111on theBAC F28P12


F14023


2 2 0/OI 1 1
0 28 35 48 76 89 98



the target gene


i1b4-1, C A; Gin stop


it~b4-2, Salk -131655


Chromosome 1





BAC~s





Recombilnants
Position (kb~)
Fl4023

Candidate
genes



Atlg71800


F413


F5A18 F23N20


F17M819 F8P5


F28P22


1TB2 cDNA~


Figure 5-1. Positional cloning of ITB4. (A) Positional cloning strategy to identify ITB4 gene.
(B) Confirmation of the C to T transition in the itb4-1 allele by digestion of PCR
products with M~sel. The C to T transition creates an M~sel site in the itb4-1 allele.
PCR products amplified using ITB4 specific primers (see Table 1) were digested with
M~sel and subj ected to electrophoresis through an agarose gel. (C) Identification of the
itb4-2 mutant homozygous for the T-DNA insertion. Genomic DNA from wildtype,
heterozygous or homozygous itb4-2 mutants was amplified using ITB4 and T-DNA
specific primers (see Table 1), and the products were subj ected to electrophoresis
through an agarose gel. (D) Results of RT-PCR using ITB4 specific primers showing
similar levels of ITB4 transcript in itb4-1 and wild type plants, and no detectable
transcript in itb4-2 mutants.


B
DNA Col double
Ladder wt routert rd





ol



4111115"


4 8b3 t-2


wmm1 *-I- ACTINh2





































Figure 5-2. Rescue of the itb4-1 zwi-3 double mutant phenotype by Atlg71800. (A) Light
micrograph showing unbranched trichomes on the itb4-1 zwi-3 double mutant. (B)
Light micrograph showing the two-branched trichomes on the zwi-3 single mutant.
(C) Light micrograph of an itb4-1 zwi-3 double mutant transformed with the
GL2:ITB4 construct. The transgenic double mutant shows the same phenotype as the
zwi-3 single mutant, demonstrating rescue of the unbranched trichome phenotype. (D)
Light micrograph of an itb4-1 zwi-3 double mutant transformed with the 35S:ITB4
showing two-branched trichomes. (E) Magnified image from panel D showing a
three-branched trichome. Scale bar = 200 lm.

























I I
F


I I


G HI I J






K L M~lN





Figure 5-3. The itb4-2 mutants display the trichome shape defects. (A) (D) Scanning electron
micrographs. (E) (H) Light micrographs of isolated trichomes. (A) Symmetrical Col
wild type trichomes with equal branch length. (B) Irregular trichomes on an itb4-2
mutant showing unequal branch length and multiple branch points. (C) Developing
twin trichome on an itb4-2 mutant showing a trichome cluster. (D) Developing
unbranched trichome on an itb4-2.mutant. (E) Twin trichome still attached following
treatment with EGTA and pectinase. (F) Twin unbranched trichome on an itb4-2
mutant. (G) (J) Irregular trichomes on itb4-2 mutants showing different numbers of
branches and separated branch positions. (K) (N) Twin trichomes with different
numbers of branches.Scale bar = 100 Clm in (A) (F), and 50 Clm in (G) (N).

















































Figure 5-4. Floral defects of itb4-2 mutants. (A), (D), (G), and (J) SEMs of Col wild type
flowers. (B), (C), (E), (F), (H), (I), (K), and (L) SEMs of flowers from itb4-2 mutants.
(A) Wildtype developing flower buds showing four sepal primordia. (D) Older
wildtype buds showing the sepal completely enclosing the floral organs. (G) Wildtype
stigma with normal papillae. (J) Wildtype anther containing normal pollen. (B)
Developing itb4-2 flower buds showing the developing petal primordia (large arrow).
Small Arrow indicates fused sepal primordia. (C) itb4-2 flower bud with extra sepals.
(E) Developing itb4-2 flower buds showing the exposed stigmas. (F) Preferentially
growing sepal on an itb4-2 flower bud (H) and (I) Abnormal itb4-2 stigma showing
malformed papillae. (K) Abnormal itb4-2 anther with abortive pollen. (L) Abnormal
itb4-2 anther lacking pollen. Scale bar = 100 lm.










A 6.5i 0 Col wt
6 itM-12









Sepal Petal Stamen
Floral organ number

B








Figure 5-5. Increased number of floral organs itb4-2 mutants. (A) Floral organ numbers for Col
wild type and itb4-2 mutant flowers. (B) Representative flowers from wild type, left,
and an itb4-2 mutant, right. Scale bar = 1 mm.

































Figure 5-6. Leaf shape and color defects in itb4-2 mutants. (A), (C), and (E) Col wild type
plants. (B), (D), and (F) itb4-2 mutant plants. (A) Wild type plant showing normal
first leaf pair. (B) itb4-2 mutant showing yellow first leaf pair. (C) Wild type seedling
showing smooth leaf edge. (D) itb4-2 mutant seedling showing serrated leaf edge and
less yellow leaf color. (E) Wild type mature plant. (F) itb4-2 mutant plant showing
normal color leaves. Scale bar in (A) and (B) = 0.1 mm; Scale bar in (C) and (D) = 5
mm; Scale bar in (E) and (F) = 10 mm;









AIP:- P& B C D









Fiue -. T2 xresonpttr i olwldtpe A)Goblr tgeebroshwn
stogexrsion. B) eart staeeby so ingsrn xrsin C
Gemnaig ed hwigstog xresoni terotca ndsoo pia
meisem D)Gemiain sedhbrdiedwthth ngtie onrl ene N
prbe (E) Logiudna seto fasx-ekodseligsoigsrogepeso
in~ ~~~~~~~~~~~~~ theI deeoiglavsadtihoe.()Ln ituia ecinotefoa
meriste shoin stogepeso ntefoa mrse n eeoigfoe
buds (G lrlognpiori hwn togepreso i h evlpn
stmn n apl n ekepeso in the deelpe seas (H )Foaorn
primordiaa hyrdzdwt h eatv oto es N poe cl a 0p
in () an (B, 50pm () ad (C, 10 pm in (E) ? H)




























rl


.5~


Scale~~i bar= 5 pm



























Figure 5-9. ITB4 is localized to the nucleus. Onion epidermal cells were transiently transformed
with either 35S:ITB4-GFP (A) (C), or 35S:GFP. (D) -(F). (A) GFP localization
confined to the nucleus. (B) Same cell as in (A), but stained with DAPI to show
position of the nucleus. (C) DIC image of the same cell as in (A) showing the position
of the nucleus (arrow). (D) GFP localization in both the cytoplasm and the nucleus.
(E) The same cell as in (D) stained with DAPI to show the position of the nucleus. (F)
DIC image of the same cell as in (D) showing the position of the nucleus (arrow).
Scale bar = 100 Clm










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BIOGRAPHICAL SKETCH

Xiaoguo Zhang was born in Hubei province, China. After completing his college education

at Huazhong Agricultural University, he attended China Agricultural University for his master's

degree in crop genetics and breeding. After his graduation, he worked at Wuhan University. He

started his Ph.D. program in plant molecular and cellular biology in 2005 at University of

Florida.





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1 CONTROL OF PLANT CELL SHAPE BY IRREGULAR TRICHOME BRANCH GENES IN ARABIDOPSIS By XIAOGUO ZHANG A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2008

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2 2008 Xiaoguo Zhang

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3 To my great Mom

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4 ACKNOWLEDGMENTS I offer m y sincere gratitude to my advisor and committee chair, Dr. David G. Oppenheimer, for his guidance and encouragemen t. I especially acknowledge him for improving my knowledge in plant developmental genetics, expanding my thinking on scientific questions, and teaching me English science writing during my doctoral education. I w ould like also express my great appreciation to our re search coordinator, Paris Grey for her strong support of my research. I am truly grateful to my supervisory co mmittee, Drs. Alice C. Harmon, Bernard A. Hauser, and Daniel L. Purich, for their valuab le advice and inspiring discussions throughout my research. I also thank Drs. Wenyun Song and Xiaodong Ding for training me in yeast twohybridization in their laboratory. I would also like to thank Drs. Kevin M. Folta and Amit Dhingra for the use of the Gene Gun and help with the transient assays. I also thank the graduate students in the Oppenheimer laboratory, Stacey Je ffries and Meredith Sullivan, for their support and friendship during my education. I want to take this opportunity to thank all the faculty members who taught me classes and all the peop le in PMCB who helped me during the past three years. Finally I give sp ecial thanks to my wife, Qingping Yang, my son, Yuxiang Zhang, and my daughter, Aiwen Zhang, as well as my parents-in-law, Xueren Yang and Dongying Xiong.

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5 TABLE OF CONTENTS page ACKNOWLEDGMENTS...............................................................................................................4LIST OF TABLES................................................................................................................. ..........9LIST OF FIGURES.......................................................................................................................10ABSTRACT...................................................................................................................................12CHAPTER 1 LITERATURE REVIEW: PL ANT CELL EXPANSION ..................................................... 14Introduction................................................................................................................... ..........14Significance of Cell Expansion....................................................................................... 14General Characterization of Cell Expansion...................................................................14Cell Wall Dynamics and Cell Expansion............................................................................... 15Cell Wall Components.................................................................................................... 16Cell Wall Synthesis and Cell Expansion......................................................................... 18Plasma Membrane Dynamics and Cell Expansion................................................................. 24Components of the Plasma Membrane............................................................................ 24Asymmetry of Plant Plasma Membranes........................................................................ 25Plasma Membrane Assembly and Cell Expansion.......................................................... 25Endomembrane Dynamics and Cell Expansion..................................................................... 28Endomembrane Trafficking Pathways............................................................................29Vesicle Dynamics............................................................................................................30Vesicle Fission Machinery..............................................................................................31Vesicle Fusion Machinery...............................................................................................31Vesicle Trafficking and Cell Expansion.......................................................................... 32Cytoskeleton Dynamics and Cell Expansion..........................................................................35The Microtubule Cytoskel eton and Cell Expansion........................................................36Actin Cytoskeleton Dynamics and Cell Expansion......................................................... 402 IRREGULAR TRICHOME BRANCH 3 (ITB3) IS A NOVEL REGULATOR OF ACTIN ORGANIZATION ..................................................................................................... 46Introduction................................................................................................................... ..........46Materials and Methods...........................................................................................................49Plant Materials and Growth Conditions..........................................................................49Positional Cloning of ITB3 ..............................................................................................49Plasmid Construction....................................................................................................... 50RNA Extraction and RT-PCR......................................................................................... 51Plant Transformation....................................................................................................... 51Yeast Two-hybrid Assays................................................................................................52Protein Isolation.............................................................................................................. .52

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6 Pull-down Assay.............................................................................................................. 53Morphological Analysis.................................................................................................. 53Immunostaining of the Actin and Microtubule Cytoskeletons........................................ 54Microscopy......................................................................................................................54Double Mutant Construction...........................................................................................54Results.....................................................................................................................................54Cloning of the ITB3 Gene................................................................................................54ITB3 is a Plant-specific Gene.......................................................................................... 56ITB3 Over-Expression Did Not Generate Novel Phenotypes......................................... 56ITB3 Has No Specific Subcellular Location................................................................... 56ITB3 Interacts With ADF3 in Yeast................................................................................ 57ITB3 Directly Binds with ADF3 in Vitro........................................................................ 57The Trichomes are Defective in the Mutants of adf itb3l-4 and Their Double Mutants........................................................................................................................ 57Discussion...............................................................................................................................58Disruption of Actin Cytoskeleton Organization Leads to Misshapen Trichomes........... 58The Precise Role of the Actin Cytoskeleton in Trichome Morphogenesis..................... 59Actin Filament Reorganization Is Required for Cell Expansion.....................................60ITB3 is a Plant-Specific Regul ator of Actin Organization..............................................60Future Perspectives.................................................................................................................613 IRREGULAR TRICHOME BRANCH 2 (ITB2) IS A PUTATIVE AMINOPHOSPHOLIPID TRANSL OCASE THAT REGULATES TRICHOME BRANCH ELONGATION IN ARABIDOPSIS.................................................................... 74Introduction................................................................................................................... ..........74Materials and Methods...........................................................................................................76Plant Materials and Growth Conditions..........................................................................76Positional Cloning of ITB2 ..............................................................................................77Plasmid Construction....................................................................................................... 77RNA Extraction and RT-PCR......................................................................................... 78Plant Transformation....................................................................................................... 79Results.....................................................................................................................................79Characterization of the itb2 Mutants............................................................................... 79Cloning of the ITB2 Gene................................................................................................79Complementation of the itb2 Mutant and Over-expression of the ITB2 Gene................81Discussion...............................................................................................................................83Future Perspectives.................................................................................................................854 DISPROPORTIONATE (DPP) E NCODES A KETOACYL REDUCTASE INVOLVED IN TRICHOME CELL EXPANSION.................................................................................... 92Introduction................................................................................................................... ..........92Materials and Methods...........................................................................................................95Plant Materials and Growth Conditions..........................................................................95Positional Cloning........................................................................................................... 95

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7 Plasmid Construction....................................................................................................... 96Plant Transformation....................................................................................................... 97Results.....................................................................................................................................97Characterization of dpp Mutants..................................................................................... 97Positional Cloning of DPP ..............................................................................................98Identification of DPP ......................................................................................................99Discussion.............................................................................................................................101DPP Encodes a -ketoacyl Reductase..........................................................................101DPP Has Pleiotropic Functions in Ce ll Expansion and Wax Synthesis....................... 101DPP is Vital for Plant Viability.....................................................................................103DPP is Likely to be DEADHEAD .................................................................................104Future Perspectives...............................................................................................................1055 IRREGULAR TRICHOME BRANCH 4 IN ARABIDOPSIS ENCODE S THE PLANT HOMOLOG OF THE 64 KDA SUBUNI T OF CLEAVAGE STIMULATION FACTOR AND REGULATES TRICHOM E MORPHOGENESIS AND FLORAL DEVELOPMENT.................................................................................................................122Introduction................................................................................................................... ........122Materials and Methods.........................................................................................................124Plant Materials and Growth Conditions........................................................................124The itb4-2 Mutant Isolation and ITB4 Cloning.............................................................125Transgene Construction................................................................................................. 126RNA Extraction and RT-PCR....................................................................................... 127Plant and Yeast Transformations................................................................................... 127Morphological Analysis................................................................................................ 128Microscopy....................................................................................................................128in situ Hybridization......................................................................................................129Results...................................................................................................................................129ITB4 Encodes AtCstF-64...............................................................................................129Loss-of-function Mutations in ITB4 Cause Aberrant Development of Trichomes and Flowers................................................................................................................131ITB4 is Highly Expressed in Growing and Proliferating Cells..................................... 134Loss of ITB4 Function Alters the Expressi on Pattern of Perianth Organ Identity Genes..........................................................................................................................135ITB4 Localizes to Nuclei, but Does Not Functionally Complement Tts Homolog in Yeast..........................................................................................................................135Discussion.............................................................................................................................137ITB4 Plays a Crucial Role in Trichome Morphogenesis and Floral Development....... 138Loss of AtCstF-64 Function Influences th e Expression of Multiple Genes that Control Floral Organ Development...........................................................................140Differences Exist in the Mechanism of mRNA 3 End Formation among Plants, Yeast and Mammals...................................................................................................140Future Perspectives...............................................................................................................142

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8 LIST OF REFERENCES.............................................................................................................155BIOGRAPHICAL SKETCH.......................................................................................................178

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9 LIST OF TABLES Table page 2-1 Primers used in this study................................................................................................. .633-1 Segregation of the mutant plants in F2 with different genetic background.....................8773-2 Trichome shapes of the transgenic plants........................................................................ 8784-1 Primers sequence used in this study................................................................................. 1074-2 Segregation of trichome phenotypes in F2 of the dpp mutant crossed to wild-type plants......................................................................................................................... .......1084-3 Segregation of trichome phenotypes in F1 of the dpp mutant reciprocally crossed to wild-type plants............................................................................................................... .1084-4 Single nucleotide polymorphism identif ied between the Ler and RLD ecotypes........... 1084-5 Segregation of phenotypes in the F1 of the dpp mutant crossed to the Salk lines........... 1095-1 Primers used in this study................................................................................................ 1445-2 Alteration of trichome cell shape in the itb4-2 mutant.................................................... 145

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10 LIST OF FIGURES Figure page 2-1 Positional cloning of ITB3 ................................................................................................642-2 Actin cytoskeleton is disorganized in the itb3 mutant.......................................................652-3 Actin cable organization in the stalk of trichomes............................................................. 662-4 Actin rings in the itb3 mutant............................................................................................672-5 Phylogenic tree of the Arabi dopsis ITB3 family members............................................... 682-6 Alignment of ITB3 protein sequence with its homologs in other plants........................... 692-7 ITB3-GFP is not specifically localized to any subcellular stru cture in transformed onion epidermal cells......................................................................................................... 702-8 Yeast two-hybrid screen for ITB3 interactors................................................................... 712-9 ITB3 directly interacts with ADF......................................................................................722-10 Trichome shapes are defective in adf3 and itb3l-4 mutants.............................................. 733-1 Defects in leaf trichome and cotyledon shape of itb2 mutants..........................................883-2 Positional cloning and gene structure of ITB2 ...................................................................893-3 Mutations and corrections of ITB2 cDNA......................................................................... 903-4 Transgenic plants with ITB2 cDNA................................................................................... 914-1 The dpp mutant trichomes in the RLD genetic background............................................ 1104-2 Positional cloning of DPP ...............................................................................................1114-3 Single nucleotide polymorphism between RLD and Ler wild types............................... 1124-4 Equivocal sequencing result using the DNA template from plants heretozygous for the dpp mutation.............................................................................................................. 1124-5 Sequencing result of At1g67730 using the dpp mutant DNA as a template with the forward primer.................................................................................................................1134-6 Sequencing result of At1g67730 using the dpp mutant DNA as a template with the reverse primer..................................................................................................................1134-7 Schematic explanation of DPP identification.................................................................. 114

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11 4-8 Schematic explanation of deletion id entification in dpp mutants.................................... 1154-9 Novel phenotypes in the F1 of dpp mutants and T-DNA insertion lines......................... 1164-10 Equivocal sequencing result of At1g67730..................................................................... 1174-11 Unequivocal sequencing result of At1g67730................................................................. 1174-12 Identification of DPP .......................................................................................................1184-13 Unequivocal sequence result of At1g67730 using the DNA from ded plants of F1.......1194-14 Unequivocal sequencing result of At1g67730 using the DNA from the wild-type plants of F1......................................................................................................................1194-15 GC deletion in dpp cloned into pBluescript SK...............................................................1204-16 Wild-type DPP cloned into pBluescript SK....................................................................... 1204-17 Transgenic plants with the mutated DPP in distinct genetic backgrounds...................... 1215-1 Positional cloning of ITB4 ...............................................................................................1465-2 Rescue of the itb4-1 zwi-3 double mutant phenotype by At1g71800.............................. 1475-3 The itb4-2 mutants display the tr ichome shape defects................................................... 1485-4 Floral defects of itb4-2 mutants....................................................................................... 1495-5 Increased number of floral organs itb4-2 mutants........................................................... 1505-6 Leaf shape and color defects in itb4-2 mutants............................................................... 1515-7 ITB2 expression pattern in Col wild type........................................................................ 1525-8 Altered expression patterns of floral organ identity genes in itb4-2 mutants.................. 1535-9 ITB4 is localized to the nucleus....................................................................................... 154

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12 Abstract of Dissertation Pres ented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy CONTROL OF PLANT CELL SHAPE BY IRREGULAR TRICHOME BRANCH GENES IN ARABIDOPSIS By Xiaoguo Zhang May 2008 Chair: David G. Oppenheimer Major: Plant Molecular and Cellular Biology The control of plant cell shape is fundamen tally important not onl y for the function of individual cells, but also for the morphogenesis of whole plants. The Arabidopsis leaf trichome is used as a cell model for gene tic screens of mutations, called irregular trichome branch ( itb ) and disproportionate ( dpp) which cause changes in trichome shape. Five genes ( ITB1 ITB4 and DPP) were cloned through a positio nal cloning strategy and the f unctions of these genes were characterized in this study. ITB1 is a plant homolog of the actin-related protein2/3 complex activator Scar/WAVE, which regulates actin and microt ubule organization. Disruption of ITB1 causes disorganization of actin filaments and microtubules, generating distorted trichomes. ITB2 is a putative member of the aminophospholipid translocase (ALA) famil y. Mutations in this ge ne result in defective trichomes with reduced branch length. ITB3 is a plant-specific protein that regulates actin organization through interaction w ith actin depolymerizing factor (ADF). The absence of ITB3 severely changes actin cytoskeleton organization by forming actin rings, but no change was observed in microtubule organi zation. The trichomes in itb3 mutants are reduced in size and branch length. ITB4 is the plant homolog of cleava ge stimulation factor 64 that influences not only trichome morphogenesis, but al so floral development. Compared to wild type, mutations in

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13 ITB4 reduce the trichome branch number and increase sepal and petal numbers. DPP is a keto acyl reductase and is involved in tricho me cell expansion. Mutations homozygous for dpp are lethal. At the restrictive temperature (22 C), heterozygous plants of the dpp mutants display trichomes with reduced branch length and incr eased stalk length. Although the five gene products described above have di fferent functions in plant cell s, their mutations all cause changes in Arabidopsis leaf trichome shape. These results indicate that plant cell shape can be controlled by different genes with a wide range of functions on multiple dynamic cell processes such as cytoskeleton dynamics and endomembrane dynamics.

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14 CHAPTER 1 LITERATURE REVIEW: PL ANT CELL EXPANSION Introduction Significance of Cell Expansion The intricate coordination of cell division and ex pansion allow s plants to achieve a unique developmental plasticity that reduces the constr aints of various adverse environments for plant survival. Plants need continued availability of light, water, and nutrients throughout their life cycle. However, plants are stationary and plan t cells are surrounded by ri gid cell walls, which make plants unable to escape resource-depleted e nvironments. Therefore, plants adjust the rate and direction of cell division and expansion. Rigi d cell walls offer structural and mechanical support for plant bodies, and ulti mately are responsible for the plant architectural design and morphology. Thus, plant cell expansion is the ba sis for both the whole plant morphogenesis and plant flexibility to adapt environmental conditions (Thompson, 1917). General Characterization of Cell Expansion An expanding plant cell is sim ilar to an inflating balloon in directi on determination, force requirement, and wall alteration. Balloons can form diverse shapes, if spec ific locations of the balloon surface are forced by counter-pressure to change conformation. So can plant cells, too, expanding in directions that follow their functi onal requirement. The driving force of cell expansion comes from the internal turgor pr essure, generated by the water content of the protoplast. During cell expansion, water needs to enter into the protoplast across the semipermeable plasma membrane to keep a consta nt turgor pressure (Steudle and Zimmermann, 1977). Second, the direction of cel l expansion is spatially dete rmined by the orientation of cellulose microfibrils, which in turn is c ontrolled by the plant cytoskeleton. The plant cytoskeleton is composed of microtubules and microfilaments. It is generally believed that

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15 microtubules orient the depositi on of cellulose microfibrils in cell walls, and microfilaments serve as tracks for transporting vesicles to specific sites of the wall that are loosened by lytic enzymes or wall-remodeling proteins. Through e xocytosis, dynamic vesicles, which contain polysaccharides and proteins from endomembrane compartments, are fused with the plasma membrane, and their contents are discharged in to the extracellular matrix space for building cell walls (Murphy et al., 2005; Samaj et al., 2005; Johansen et al., 2006). Ev en though the volume of an expanding cell increases to a thousand times the original volume, the walls maintain a relatively constant thickness. A ll cell expansion is mechanistica lly divided into two basic types of growth: tip growth and diffuse growth. For tip growth, a spatially focu sed cell expansion, cells grow exclusively at the extreme tip of the cell. Pollen tube and and root hairs are cell types that expand using tip growth. For diffuse growth, expansion can occur at multiple locations and encompass larger regions than occu rs during tip growth (Baskin, 2005). Cell Wall Dynamics and Cell Expansion The plant cell wall m ay seem to be a paradoxical entity because it serves to both maintain cell shape through constraint of cell expansion, yet it promotes development of cell morphology by extension of itself. The wall provides rigidity to a plant cell, strengt h, and protection against various mechanical stresses. It also limits the entry of large molecules and pathogens into cells. The wall further creates a stable osmotic enviro nment by preventing osmotic lysis and retaining water. Finally, the wall determines plant cells myriad shapes. All thes e functions of the wall underlie its action as a physical barrier, which constrains ce ll expansion. To solve the paradox, plants have evolved a dynamic cell wall. Thereby, this wall is not only suited to flexible cell expansion, but also participates in cell to cell and cell to nucleus communication through signaling molecules and receptors in the wall (Darley et al., 2001).

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16 Cell Wall Components As m entioned above, cell walls allow plants to form various shapes. Thus, cell expansion appears to be its wall extension. To understand cell expansion it is first necessary to know the chemical composition of the wall. Because only primary walls can be remodeled for cell expansion, my focus here is on those aspects that are germane to understanding cell expansion. Secondary walls, which are formed after the cessation of cell expa nsion, are not covered in this literature review. Plant cell expansion requires coordination between maintenance of osmotic potential and changes in wall properties, which are determined by wall components. Plant cell walls are highly organized; the diverse components include cellulo se, glycans, pectins, proteins, and aromatic substances, as well as metal ions. Cellulose, a major component of plant cell walls, plays the most important role in the wall architect ure. Approximately 36 chains of 1,4 linked, -D-glucose associate through intermolecular hydrogen bonds to form a micro fibril, which serves as a structural scaffold to support other wall compone nts, and forms a fundamental framework of the wall architecture. The microfibr ils in walls are paracrystalline and resistant to hydrolysis by acids, bases and enzymes (Reiter, 2002; Somerville, 2006). Glycans, a mixture of branched polysaccharides, form molecular backbones through the linkage of -D-hexosyl residues with the same bond as in cellulose. Distinct glycans interlock microfibrils by hydrogen bonding. Glycans and p ectins form the secondary network, which strengthens the wall architecture. Xyloglucan (XyG), glucuronoarabinoxylans (GAX), mixed linkage glucans (MLG), and (gluco) mannans are found to be the four main glycans in primary cell walls. Because glycans have a property of ra ndom arrays and amorphous structures in the wall they are readily hydrolyzed by dilute acids bases, or myriad enzymes (Reiter, 2002).

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17 Pectins are a family of heterogeneous pol ysaccharides that all contain 1,4-linked -Dgalacturonic acid. Because of this residue, pec tins provide cell walls a hydrated and charged surface. Additionally, a pectin polymer has a multitude of branches, which makes cell walls porous. These componential and structural features of pectins also allo w cell walls to modulate pH and ion balance. Homogalacturonan (HG) and rhamnogalacturonans (RG-I and RG-II) are fundamental constituents of pectins in primary cell walls (Willats et al., 2001). The majority of proteins in cell walls are glycoproteins that have oligosaccharide chains covalently attach to particular moieties of polypeptides. Hydroxyprolin e-rich glycoproteins (HRGPs, also called extensins), the arabinogala ctan proteins (AGPs), and the proline-rich proteins (PRPs) are the most a bundant glycoproteins in plant cells. Although proteins are not a major component of cell walls, th ey display a wide variety of structures and functions. These proteins make the wall a dynamic entity throughout the cells life. For example, expansin induces the pH-dependent wall extension and stress re laxation in a characteri stically unique manner (Cosgrove et al., 2002). Wall associated kinases (WAKs), which are covalently bound to pectin, have the potential to provide a physical and signaling continuum between the cell wall and the cytoplasm (Wagner and Kohorn, 2001). The absence of AGPs in cell walls causes aberrant cell expansion, which forms numerous bulges in r oot epidermal cells (Willats and Knox, 1996; Ding and Zhu, 1997). Minerals appear in micro amounts in cell walls but they also play a role in the wall structure. If plants lack them, normal cell wall formation is disrupted. For example, Ca2+ links distinct pectin polymers (RG-I) through an ioni c bond (Catoire et al., 1998). Another example is boron, which is predominantly associated with rhamnogalacturonan II (RG-II). Disruption of the

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18 linkage between borate and RG-II affects plant grow th, and borate deficiency compromises plant cell expansion in growing tissues (O'Neill et al., 2004). Cell Wall Synthesis and Cell Expansion Biosynthesis of plant cell walls is intimately connected with cell expansion. While a cell is increasing its volume, its wall becomes thinner an d thinner. Finally, if the cell wall fails to incorporate new materials, it bursts. Thus, wall biosynthesis is required for cell expansion. In recent years, the study of the biosynthesis of wall components has made substantial progress through biochemical, genetic, and genomic approaches. Here, the emphasis is on the genes that are involved in primary wall synt hesis and their mutations that cau se disruption of cell expansion and generate aberrant cell shape. Cellulose is synthesized by cellulose synthase (CESA), which has been identified in most plant species. In the Arabidopsis genome, ten genes code for CESA; in rice there are at least nine genes (Keegstra and Walton, 2006), and poplar ha s 18 (Djerbi et al., 2005 ). All CESAs share common structural features: eight transmembrane domains, two glycosyltransferase domains, and several microtubule-interacti on domains. Six hexamers of CESA form a symmetric rosette on the plasma membrane plane; rosette movement is guided by cortical microtubules. Each subunit of CESA in the rosette synthesizes one 1,4 -D-glucose chain (Somerville, 2006). Glycan synthesis includes backbone synthesis and side chain addition. Compared to CESA, glycan synthesis is not we ll understood, although there is a structural similarity between cellulose and glycans. Basing on th is similarity, the CSL hypothesi s has been established. It was hypothesized that CELLULOSE SYNTHESIS LIKE (CSL) genes encode Golgi-localized glycan synthases. This hypothesis is supported by recent discoveries of several glycan synthase genes (Lerouxel et al., 2006). For example, mannan synt hase, MLG synthase, XyG glucan synthase, and galactomannan galactosyltransferas e are all encoded by members of the CSL gene family.

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19 Additionally, these en zymes are all responsible for the ba ckbone synthesis of glycans. XyG fucosyltransferase and XyG xylosyltransferase (X T1) were found to add side chains to the backbone of glycans (Reiter, 2002; Lerouxel et al., 2006). Pectin biosynthesis is much more complicated than cellulose and glycan synthesis because it is difficult to investigate pectin synthases us ing traditional biochemical purification techniques and forward genetics. Through reverse genetics and genomics approaches, more than 50 glycosyltransferases (GTs) are pr edicted to be required for pec tin synthesis (Ridley et al., 2001). At present, only a few genes for pectin biosynthe tic GTs have been iden tified, and for some of their products, the activity of the pectin synthesis is not clear (Bacic, 2006). HG g alacturonosyltransferase (GAUT1) is the first functional identifi cation of GTs using biochemical and functional genomic approaches in Arabidopsis (Sterling et al., 2006). Other putative GTs include QUA1 (Bouton et al., 2002), NpGUT1 (Iwai et al., 2002), and PARVUS/GLZ1 (Lao et al., 2003). During cell expansion, polysaccharides are depos ited into existing walls. If polysaccharide synthesis is disrupted, old wall reinforcement and new wall assembly cannot take place. Thus, normal cell expansion is disrupted, resulting in ab errant cell shapes. For example, mutations in CESA cause alterations of cell shape in diverse cel l types because cellulose forms a fundamental framework of the wall architecture. RSW1 encodes CESA1. The rsw1 mutation causes disassembly of CESA complexes on the apoplasti c side plane of the plasma membrane and reduction of cellulose accumulation in cell walls (Arioli et al., 1998). The rsw1 mutants display shorter roots with radial swellings ( rsw), smaller leaf blades with shorter petioles, and aberrant trichomes at the 31 C restriction temperature. All these defects are indicative of abnormal cell expansion owing to the rsw1 mutation (Williamson et al., 2001). The root radial swellings of the

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20 rsw1 mutants exactly mimic phenotypic responses of wild-type roots to cellulose synthesis inhibitors such as dichlorobenzonitrile. This observation suggests that the abnormal cell expansion is due to reduction of cellulose synthesis. Further evidence for this view comes from down-expression of CESA1 and CESA3, using transformation with antisense constructs. The antisense phenotypes of CESA1 or CESA3 display shorter inflorescent shoots and stamen filaments, a result of reductions in cell length ra ther than cell number. In addition, the severity of the manifestation of both genes of interest is closely similar and intimately correlated to their reduced expression (Burn et al., 2002). PROCUSTE1 ( PRC1 ) is another gene, coding for CESA6. Mutations in this gene al so exhibit similar defects as rsw1, including decreased cell elongation, especially in roots and dark-grown hypocotyls. The cell elongation reduction is correlated to a cellulose deficien cy (Fagard et al., 2000). These observations indicate that the reduction of cellulose in walls genera lly causes suppression of cell expansion. Glycans form the cell wall matrix, which enhances the wall strength. Probably because of this role, mutations in glycan synthase genes ofte n cause only a slight alteration of cell expansion and shapes. In addition, a specific mutation in one glycan synthase gene fa ils to cause a visible phenotype. For example, MUR2 encodes XyG fucosyltransferase (AtFUT1). The cell walls of mur2 contain less than 2% of the wild-type amount of fucosylated XyG. The mur2 plants show a normal growth habit and wall strength. On the other hand, MUR1 codes for a 4,6-dehydratase, responsible for the de novo synthe sis of l-fucose. Mutations in MUR1 cause structural changes in several cell wall polysaccharides (Bonin et al., 1997). Thus, mur1 mutants exhibit a dwarfed growth habit and decreased wall st rength, probably indicating aberra nt cell expansion (Reiter et al., 1993; Bonin et al., 1997). MUR3 encodes XyG galactosyltransferase, which specifically catalyzes formation of the -L-Fuc-D-Galside group. Although the XyG in the mur3 cell walls

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21 completely loses the fucosylated disaccharide side chain, mur3 plants are visibly indistinguishable from wild-type plants except for a collapse of trichome papillae (Madson et al., 2003). However, later investigator s found that the galactose resi dues of XyG are essential to maintain wall mechanical strength during rapid cell expansion. The mur3 mutations result in reduced wall strength. Through studies using a sc anning electron microscope, the defects in the mur3 hypocotyl cells were observed. In addition, these defects are similar to the phenotype of mutations in rsw1, generating swollen cells of larger si ze (Pena et al., 2004). These defects indicate abnormal cell expansion can be caused by a reduction of cell wall strength. Pectins combine with glycans to form the s econdary network of ce ll wall architecture. Thus, mutations in the pectin synthesis genes sh are similar phenotypes to the phenotypes seen in plants with defective GT genes. QUASIMODO1 ( QUA1 ) codes for a putative membrane-bound GT. When the qua1 mutants were grown in light, the plants showed reduced height because of the pectin deficiency in cell walls; similarly, the qua1 seedlings grown in the dark had shorter hypocotyls, compared with the wild type. Thes e defects are likely due to suppressed cell expansion (Bouton et al., 2002). The dwarfi sm phenotype was also observed in the parvus mutants because of both reduction in RG-I bran ching and alterations in the abundance of xyloglucan linkages. PARVUS enco des another putative GT (Lao et al., 2003). Pectins play a crucial role in pollen tube elongation because th ey are the only kind of molecule that makes a single layer of wall at the growing tip (Stepka et al., 2000). VANGUARD1 (VGD1) encodes a pectin methylesterase (PME), which, dependi ng on ambient pH, by enzymatic activity can lead either to stiffening or to loosening of cell walls (Catoire et al., 1998; Denes et al., 2000). Mutations in vgd1 cause a rupture of elongating pollen tubes in vitro and retarded growth in vivo (Jiang et al., 2005). Moreover, when PME is exogenously added to grow ing pollen tubes, the

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22 apical wall is thickened, resul ting in inhibition of pollen tube elongation (Bosch et al., 2005). PME activity that promotes cell wall loosening will be described below. After cell division, through anis otropic expansion plant cells generally reach their final sizes. At this time, polysaccharides have been a lmost equally deposited into primary cell walls. A majority of the cells further differentially form specific shapes th rough further anisotropic expansion. Before this process o ccurs, the cell wall at specific s ites is biochemically loosened for turgor-driven cell expansion (Cosgrove, 2000a). Additi onally, the wall is loosened without compromising the tensile strength of the plia nt wall (Cosgrove, 2000b). The reason plant cells have the capacity for this complex event is th at the walls contain such unique proteins as expansins, PME, and XyG endotransgluc osylase (XET). PME de-esterifies highly methylesterified pectins, converting the methoxyl groups into carboxyl groups on the polygalacturonic acid chain, and releasing both methanol and prot ons. This conversion promotes pectin gelation accompanying wall stiffening, due to the formation of the cooperative Ca2+ crossbridges between free carboxyl groups of adjacent pectin chains (Catoire et al., 1998). In addition, the de-esterification reduces th e local pH, which promotes activ ity of several other cell wallloosening hydrolases, such as expansins, polygalacturonases, a nd pectate lyases (Cosgrove et al., 2002). Under-expression of PME by antisense RNA in transgenic pea reduced root hair elongation. Moreover, the root lengt h reduction was correlated with an increase in extracellular pH (Wen et al., 1999). The activation of wall hydrol ases by acidification of cell walls was also supported by the functional analysis of DET3, which is a vacuolar H+-ATPase (V-ATPase). Mutations in DET3 lead to def ects in hypocotyl cell elongation. It was suggested that V-ATPases contribute to maintaining the internal turgor pressure of plan t cells through modulation of solute uptake to vacuole (Schumacher et al., 1999). Subs equently, another kind of vacuolar H+-pump

PAGE 23

23 called H+-PPase AVP1 (AVP1) was found to be im plicated in this transport process. AVP1 adjusted the distribution and a bundance of H+-ATPases in the pl asma membrane by controlling its trafficking through the endocyt ic secretory pathways. Over-expr ession of AVP1 increased the accumulation and polar distribution of th e H+-ATPase in the plasma membrane. Consequentially, more H+ was pumped to the ap oplast and the cell wall was acidified. Thus, cell elongation occurred. In the avp1-1 null mutant, root cell elongation was severely disrupted (Li et al., 2005). Expansin, another cell wall protein, induces pH-dependent wall extension and stress relaxation in a characte ristically unique manner (Cosgrove et al., 2002). Expansin has been proven to cause isolated wall extension in vitr o under constant mechanical stress (McQueenMason et al., 1992). Application of exogenous expansin from cucumber in excised Arabidopsis hypocotyls stimulates cell elongation. At a high concentration of applied expansin, the tips of growing root hairs burst; at a lower level, e xogenous expansin caused radial swelling at the tip (Cosgrove et al., 2002). Over-expression of Arabidopsis EXP10 results in transgenic plants with longer petioles and larger leaf blades because of increased cel l size (Cho and Cosgrove, 2000). Over-expression of EXP1 in tomato fruit enhances fruit softening and cell wall breakdown (Brummell et al., 1999). The enzymatic activity of XET breaks the existing linkages in the XyG-cellulose network and rejoins the resultant ends with new partne rs at different positions. XET loosens the wall during cell expansion through cooperation with e xpansin (Nishitani and Tominaga, 1992). XET activity is intimately correlated to the cell growth rate and epidermal lengthening in the growing zone of maize leaves (Rose et al., 2002). XET also is specifica lly localized at the site of

PAGE 24

24 trichoblast walls, where the future bulge is formed during root hair initiation. A locally high level of XET activity stimulates tr ichoblasts to initiate root hairs (Vissenberg et al., 2001). Plasma Membrane Dynamics and Cell Expansion The plasm a membrane abuts cell walls and participates in cell wall synthesis. As mentioned above, the plasma membrane protein CESA synthesizes cellulose for direct wall synthesis. The plasma membrane also actively pe rforms exocytosis for indirect wall synthesis during cell expansion. In addition, the structural asymmetry of the plasma membrane provides cell polarity for anisotropic e xpansion (Fischer et al., 2004). To understand the roles of the plasma membrane in cell expansion, I briefly describe its components, highlig hting its structural asymmetry. After this, the assembly of the plasma membrane and its relation to cell expansion are described. Components of the Plasma Membrane The plasm a membrane of plant cells is composed of lipids, proteins, and carbohydrates in a molecular ratio of approximately 2 : 2 : 1. The membrane lipids include phospholipids, sphingolipids, and sterols (Moreau et al., 1998; Jaillais and Gaude, 2008). Some phospholipids such as phosphatidylcholine (PC) and phosphatidylethanolamine (PE) have head groups with positive charges; whereas others are negative or neutral, depending on pH. More importantly, the ratio of lipid classes in plan t plasma membranes shows a wide range of variation among the different organs in a given plant or identical organs in distinct plants (Jouhet et al., 2007). The majority of carbohydrates in plant plasma membranes are present in the form of oligosaccharides that are covalently linked to proteins to generate glycoproteins (Bacic et al., 1996; Classen et al., 2005).

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25 Asymmetry of Plant Plasma Membranes Lipids, proteins, and carbohydrates of plant plasma membranes are regularly arranged in an asymmetric bilayer structur e. The lipid amphipathy allows for spontaneous assembly of bilayers. The hydrophilic heads ma ximize their interactions with water, whereas the hydrophobic tails interact with each other, minimizing their exposure in the exoplasmic leaflet (Holthuis and Levine, 2005; Pomorski and Menon, 2006). S phingolipids and sterols are abundant in microdomains (lipid rafts), in which signaling proteins accumulate. Other proteins are also localized at particular sites in the plasma me mbrane. For example, auxin carriers are positioned at the apical or basal plasma membrane of r oot epidermal cells (Muller et al., 1998; Swarup et al., 2001), whereas glycosylphosphatidylinositol (GPI)-anchored proteins (GAPs) such as COBRA (COB) preferentially localizes to lateral membrane in root cells (Schindelman et al., 2001). This asymmetric distributi on establishes and main tains cell polarity, which is required for longitudinal expansion at the elongation zone (F ischer et al., 2004; Kramer and Bennett, 2006). Plasma Membrane Assembly and Cell Expansion Plasm a membranes of plant cells are assemble d using various lipids and diverse proteins. In addition, different lipids interact with distinct proteins, and sp ecific proteins have particular positions in lipid bilayers. During isotropic cell e xpansion, the plasma membrane evenly enlarges its area, whereas in polar expansion it is a ssembled only at the growing sites. For any cell expansion, the plasma membrane maintains the d ynamic stability of chemical composition and distribution of protein components. In addition, it has the capacity for flexible changes in these features in response to intracellular and ex tracellular signalings for anisotropic expansion. Perturbations of membrane stabili ty cause abnormal cell expansion (Schrick et al., 2000; Souter et al., 2002; Jaillais and Gaude, 2008).

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26 Changes of lipid composition in the plasma membrane cause disruption of normal cell expansion (Jaillais and Gaude, 2008). COTYLEDON VASCULAR PATTERN 1 ( CVP1 ) and the ORC gene encode a sterol methytransferase for sterol biosynthesis. Mutations in either CVP1 or ORC alter the membrane sterol composition, wh ich raises the level of cholesterol and campesterol at the expense of sitosterol. The smt1orc and smt2cvp1 double mutants display defects in polar cell expansion th at result in the perturbed alignment of cells into parallel vascular cell files in cotyledons because abnormal cell expansion leads to aberrant cell shapes (Carland et al., 2002; Willemsen et al., 2003). Sterols are enriched in lipid crafts, which provide platforms that anchor polar proteins to the plasma membrane. Thus, the influence of sterol composition on the localization of polar proteins has been widely reported in yeasts and animals (Simons and Ikonen, 1997; Bagnat et al., 2000; Bagnat and Simons, 2002; Danielsen and Hansen, 2006). These events are also likely to occur in pl ants. For example, it has been found that, in Arabidopsis, smt1orc root cells mislocalized auxin carri ers on the plasma membrane (Willemsen et al., 2003). Changes in anchored proteins of the plasma membrane also cause disruption of normal cell expansion. In eukaryotic cells, glycosylphosphati dylinositol (GPI) anchor ed proteins (GAPs) have been extensively reported to be plasma membrane bound proteins (Oxley and Bacic, 1999; Sherrier et al., 1999; Zhao et al., 2002). Salt-overly-sensitive 5 ( SOS5) encodes a plant GAP. Mutations in SOS5 cause strong radial expansion of the cells in the elongation zone of roots, instead of the longitudinal expansion seen in wild type. Thus, the ep idermal, cortical, and endodermal cells in sos5 roots display a swelling phenotype (Shi et al., 2003). COB, another plant GAP, is anchored on the extracellular side of the plasma membrane and also released into the wall. It is required for highly anisot ropic expansion of a ll plant cells. The cob-4 null allele

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27 shows greatly reduced growth of all organs in seedlings. However, because the cob-4 root cells expanded radially in the elongation zone, the root diameter ultimately reaches nearly twice that of the wild type. The root epidermal cells displayed a severe bulgi ng phenotype. In addition, their cell walls were occasionally broken. COB plays a crucial role in the oriented deposition of cellulose microfibrils during ra pid anisotropic expansion (Roudi er et al., 2005). SKU5 has the same cell localization as COB, which is in the cell wall and on the plasma membrane anchored by GPI. SKU5 is expressed most strongly in expanding cells. Mutations in SKU5 cause skewed roots and shortened hypocotyl s because of abnormal cell ex pansion (Sedbrook et al., 2002). Although PNT1 is not a GAP, it encodes a mannosy ltransferase for the GPI synthesis. All five pnt mutants strongly reduce accumulation of GAPs. In the pnt1 mutants, the cell walls had decreased crystalline cellulose; embryos showed delayed morphogenesis; apical meristems were defective; and seedlings did not survive. All phenotypes were due to aberrant cell expansion (Gillmor et al., 2005). ETH1 and SETH2 also are involved in GPI biosynthesis. The seth1 and seth2 mutations specifically block or reduce pollen tube elongation because of abnormal callose deposition. In addition, another 47 genes, which a ll encode potential GPIanchored proteins, are likely to play important roles in the establis hment and maintenance of polarized pollen tube expansion (Lalanne et al., 2004). Normal cell expansion is disrupted by ch anges in the position of polar proteins. Arabidopsis phospholipase D 1 (AtPLD 1) is preferentially localized at the tip of growing root hairs. Ectopic over-expression of AtPLD 1 disrupts its distribution in specific tissues and induces non-root cells to form ectopic root hairs in cotyledons and hypocotyls. The raised level of AtPLD 1 in root cells generates swollen or branched root hairs (Ohashi et al., 2003). It is likely that the intrinsic polarized distribution of AtPLD 1 is perturbed, thus losing capacity for tip

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28 growth. Recently, AtPLD 2 was also found to be involved in cell expansion. Mutations in this gene reduced the hypocotyls cell elongation (Li and Xue, 2007). More recently, it was found that ROOT HAIR DEFECTIVE 2 (RHD2) is located at the gr owing tip of root hair cells. Mutations in RHD2 causes defective root hairs (Takeda et al., 2008). AGC2 is a member of the cAMP/cGMP-dependent kinase or pr otein kinase C family kinase ( AGC kinase). It localizes to the root hair tip. The agc2-1 mutants have short root hairs because of reduced cell elongation (Anthony et al., 2004). Other polar pr oteins of plant plasma membranes that play a role in cell expansion in response to phospholip id signaling are transported by oriented vesicle trafficking involving such proteins as PIP5K3 (Kusano et al., 2008) and R HD4 (Thole et al., 2008). These proteins are described below in the endomembrane dynamics and cell expansion section. In recent years, numerous observations of tran scytosis of the plasma membrane in plant cells were reported as a result of the application of fluorescent styryl dyes, such as FM1-43 and FM4-64, particularly in expanding root hairs and pollen tubes (Samaj et al., 2006). Through exocytosis, secretory vesicles containing the cell wall cargo molecule fuse with the plasma membrane, releasing their contents into the extracellular space for wall loosening, strengthening, or assembling during cell expansion, whereas, through endocytosis, the extra plasma membrane proteins and lipids are transferred into the cytosol for recycling at the growing sites (Samaj et al., 2005; Johansen et al., 2006). Incr easing evidence shows that blocking exocytosis with brefeldin A (BFA) inhibits pollen germination and po llen tube elongation (Wa ng et al., 2005b). Because transcytosis is tightly coupled with the endomembrane trafficking network, details of this topic are described below. Endomembrane Dynamics and Cell Expansion Endom embrane trafficking is essential for cel l expansion, especially for polar expansion (Samaj et al., 2006). Except for cellulose, almost all cell wall components are synthesized in the

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29 Golgi apparatus (GA) and endoplasmic reticulum (ER), further packaged into vesicles, and finally transported to the wall space (Reiter, 2002; Dhugga, 2005; Leroux el et al., 2006). The oriented trafficking of vesicles serves to establ ish and maintain cell polari ty, which initiates polar expansion. In this part, I in troduce general endomembrane path ways first, focusing on their characteristics in plant cells. After that, I highlight the influence of the disruption of endomembrane trafficking on plant cell expansion. Endomembrane Trafficking Pathways The endom embrane system is composed of all intracellular membranous compartments. These compartments communicate with each other through exchanging molecules using ubiquitous vesicles as carriers. The vesicles containing cargo molecules secrete from donor compartments, and fuse with target compartm ents. During vesicle trafficking, the GA plays a leading role. It lies at the hear t of the membrane trafficking path way, serving as the crossroad in various trafficking events (Hawes an d Satiat-Jeunemaitre, 2005a). At its cis -face, the GA receives the vesicles from the ER (anterograd e transport) and sends their vesicles back (retrograde transport). At its trans-face, the GA sends its vesi cles to endosomes, storage vacuoles, lytic vacuoles, and the plasma membra ne. The vesicles, from the plasma membrane through endocytosis, are recycled in the GA. The vesicles exported from the ER can also bypass the GA pathway and go directly to a vacuol e or the plasma membrane (Hawes, 2005). The GA is the sum of numerous polarized stacks of membrane-bounded cisternae. Within the GA, cargo molecules are proces sed, concentrated, and packaged into vesicles (Hawes, 2005). The mature cargo molecules in the vesicles ar e intracellularly routed to specific cellular destinations within cells. For plant cells, the GA is an important organelle that specializes in the synthesis and processing of complex components of cell walls, such as glycans and glycoproteins (Hawes and Satiat-Jeunemaitre, 2005b). Probably to this end, the GA organization in plant cells

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30 is different from its counterpart in animal cells in which the GA is composed of many stacks that are generally arranged side-by-si de in a ribbon structure around th e nucleus. In plants, the GA is divided into individual Golgi stacks that are di stributed through the cytoplasm (Latijnhouwers et al., 2005). The number of Golgi stacks per cell and the number of cisternae per stack varies with cell developmental stages and cell types (Neu mann et al., 2003). Mammalian GA is a rather static organelle, but plant GA is a highly mobile biosynthetic factory that moves over the ER on an actin network at the speed of 2 m per second (Boevink et al., 1998; Nebenfuhr et al., 1999). This characteristic of the plant GA enables cargo molecules to be efficiently transported to the extra-cellular matrix. Vesicle Dynamics In eukaryotic cells, the vesicle is a ubiquit ous vector for endomemb rane trafficking. The highly dynamic vesicles are generated through membrane fission of donor compartments, and disappear by membrane fusion at destination co mpartments. The processing of both the fission and the fusion elements involves close contact be tween lipid bilayers and the final combination of bilayer leaflets at specif ic sites (Markvoort et al., 2007). Fission begins with bending of membranes at the export site. The bent membrane invaginates to an extreme curvature to form a highly constricted neck. The neck further elonga tes and narrows until the two membranes merge, which leads to the separation of the vesicles from the donor compartment. Membrane fusion begins with docking of vesicles at acceptor compartments. The lipid bilayer of the docked vesicle gradually unites with the membrane of the acceptor compartment (Sollner and Rothman, 1996; Atilgan and Sun, 2007). Membrane fission and fusion are completed by the vesicle assembly machinery. In mammalian and yeast cells the components of this machinery are well known. Generally they include the coat proteins adaptor proteins, cargo receptors, and small GTPase proteins (M arks et al., 2001).

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31 Vesicle Fission Machinery During vesicle assem bly, numerous proteins are recruited at th e exit site. On the internal surface of membranes, cargo receptors such as v-SNARES specifically bind outgoing molecules from the lumen. On their external surface, th e ADP-ribosylation factors (ARFs) are recruited from the cytosol to bind tightly to the membrane. The ARFs further interact with their effectors and regulators. A regulator such as GAP, cont ributes to hydrolysis of the active GTP-bound ARF form to the inactive GDP-bound ARF form with opposite conversion catalyzed by GEF. The change of the ARF conformation during GTP hydrolysis leads to structural alterations of both the lipid and the effectors attaching with the me mbrane. This alteration promotes vesicle budding and delivery from the donor compartment. The AR F effectors mainly in clude coat proteins (Donaldson and Jackson, 2000; Donaldson et al., 2005) Three types of coat proteins have been found: clathrin, COPI, and COPII. Clathrin coats the transport vesi cle that shuttles between the GA and the plasma membrane, the GA and e ndosomes, and the plasma membrane and endosomes; COPs wrap the vesicles that shuttle between the GA and the ER. COPI coats vesicles from the GA to the ER, while COPII coat s the vesicles from the ER to the GA. Recent research has shown that COPI interacts with Brefeldin-A ADP-ribosylated substrate (BARS) (Yang et al., 2005; Yang et al., 2006) and th e actin cytoskeleton provides force both for membrane deformation during vesicle formation and for vesicle trafficking to the correct destination (Goley and Welch, 2006; Ka ksonen et al., 2006; Co et al., 2007). Vesicle Fusion Machinery After the vesicles detach fr om the donor membrane, the coat proteins release from the vesicle surface into the cyto sol for recycling. Thus, the vesicle surface marker v-SNAREs (soluble N-ethylmaleimide-sensitive-factor attach ment protein receptors) is exposed, as well as their complementary SNAREs, termed t-SNAREs, on the target compartment. The interaction

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32 between these two SNAREs causes fusion of the vesicle with the target membrane (Chen and Scheller, 2001; Bonifacino and Glick, 2004). More than 30 members in the SNARE superfamily were found in mammalian cells (Chen and Sche ller, 2001). A large number of SNAREs were also found in the Arabidopsis genome and other plants (S anderfoot and Raikhel, 1999; Sanderfoot et al., 2000; Sanderfoot 2007).The regulatory protein Rab participates in specific junctions of v-SNAREs with t-SNARSEs. Rab is a GTP-binding protein. It also provides energy for driving membrane fusion through GTP hydrolys is, while binding with the t-SNAREs (Segev, 2001; Zerial and McBride, 2001) Vesicle Trafficking and Cell Expansion Identifying the m olecular machinery of the plant membrane trafficking pathway reveals significant homology with that of mammalian and yeast counterparts. Coat proteins (COPI, COPII, and clathrin), small GTPases (Rabs, Ar f, Sar1, and Rac), and fusion proteins (SNAREs) appear to be well conserved throughout eukaryotic cells. In plant cells, th e majority of the COPI and COPII machinery and their associated effector s, such as Arf and Sar1, have been cloned, and their functions are now being determined. More and more data have in dicated that vesicle trafficking contributes to establishment of the structural and molecular asymmetry at the cell surface, which is the beginning of cell polar e xpansion (Xu and Scheres, 2005a; Friml et al., 2006). Brefeldin A (BFA) is a fungal toxin that has been widely used as a reversible inhibitor of vesicle trafficking in yeast, mammalian, and plant cells. BFA blocks the detachment of vesicles at the exit site of the GA membrane to form aggregates, which disrupts the normal flow of the vesicles within cells. BFA has been shown to a lter the distribution of such plasma membranelocalized proteins as PIN1, AUX 1, PM-ATPase, and pectins in pl ant root cells. In addition, the epidermal cells of Arabidopsis roots treated with BFA lost their polarity, displaying a decrease in

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33 cell length and increase in the ap ical-basal initiation ratio, as we ll as formation of double root hairs, in which two root ha irs are derived from one tric hoblast (Grebe et al., 2002). The molecular mechanism of BFA inhibition of vesicle trafficking is due to the binding of BFA with the ARF1-GDP/ARF-GEF complex, which preven ts ARF1 activation necessary for vesicle budding and cargo molecule selectio n (Robineau et al., 2000). In mammalian cells, ARF1 is a core component of the vesicle assembly machinery, recruiting COPI and clathrin coat proteins to membranes for vesicle assembly in mammalian cells (Boman, 2001; Rein et al., 2002; Spang, 2002; Song et al., 2006). In Arabidopsis ARF1 rescued the ARF1/ARF2 lethal yeast double mutant, which suggest that plant ARF1 has si milar functions to its yeast and mammalian counterparts (Takeuchi et al., 2002). Moreover, Arabidopsis ARF1 is also located to the GA and endocytic organelles. The over-e xpression of the engineered ARF1 with dominant activation and inactivated formats that are targeted to in terfere with the endoge nous ARF1 function in trichoblasts significantly inhibited polarized tip expansion, which produced shorter roots. After strong heat shock induction, the tric hoblasts and epidermal cells in the transgenic lines displayed many more severe defects, such as apical-shift ed root hairs, double root hairs, and bulged epidermal cells. All these phenotypes show that the cells lost their polarity and capacity for polar expansion (Xu and Scheres, 2005b). The ARF GTPase activating protein, ROOT AND POLLEN ARFGAP (RPA), activates ARF1 and plays a role in the elongation of root hairs and pollen tubes in Arabidopsis RPA is specifically expressed in roots a nd pollen; its product is located in the GA. Additionally, RPA complements the glo3 gcs double mutants in yeast (Song et al., 2006). GLO3 and GCS1 are two yeast ARFGAPs that f unction efficient retrograde trafficking of vesicles from the GA to the ER (Poon et al., 1999; Robinson et al., 2006). ARFGAP1 promotes both vesicle formation and cargo sorting by functi oning as a component of the COPI coat (Yang

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34 et al., 2002; Lee et al., 2005) The loss-of-f unction mutant, rpa causes root cells to isotropically expand to generate short and branched root hair s, as well as a slowing of pollen tube elongation (Song et al., 2006). Over-e xpression of the rice OsARFGAP also interfered with vesicle trafficking, which influenced on the root hair formation and el ongation. Transgenic plants of Arabidopsis and rice had a reduced number of lateral roots and reduced root length. The ratio of length and width of epidermal ce lls at the elongation region also decreased compared to wildtype plants. Additionally, abnormal vesicle aggregates (the BFA compartment, a typical defect of disrupted vesicle trafficking) we re observed in the transgenic cells (Zhuang et al., 2005; Zhuang et al., 2006). The ARF guanine-nu cleotide exchange factor (ARF -GEF), GNOM, functions in the establishment of apical-basal cell polarity by mediation of specific endosomal trafficking pathway in Arabidopsis embryos and roots. GNOM localizes to endosomes. The loss-of-function mutant of gnom lacked an apical-basal polar axis and embryonic root in early embryos (Geldner et al., 2003). This phenotype is pr obably a result of def ective cells that are unable to expand or anisotropically expand. Arabidopsis Rab GTPase RabA4b was found to func tion in cell directional expansion through the regulation of vesicle trafficking invo lved in the polarized deposition of cell wall components in tip-growing root hair cells (Preuss et al., 2004). Mammalian Rab11 has a high degree of homology to RabA4b, as does Arabidopsis Ara4, which was localized to Golgi-derived vesicles, Golgi cisternae, and the trans-Golgi network (Ueda et al., 1996 ). Vesicles with the EYFP-RabA4b labeling marker accumulated in the actively expanding zone in the growing root hair. Such accumulation is necessary for root hair initiation and elongation. In the rhd1-1 mutant, and the rhd2-1 mutant, their defective cells did no t show accumulation of the vesicles with Rab4A (Preuss et al., 2004). Plant Rab11 also plays a role in pollen tube tip growth.

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35 Tobacco Rab11b was localized to the pollen tube tip. Interference of endogenous Rab11b activity in its mutated variants gave rise to a re duction of pollen tube growth rate and change of pollen tube morphology (de Graaf et al., 2005). Tobacco Rab2 also functions in cell polar expansion of pollen tubes through a vesicle trafficking pathway at the GA. NtRab2 localizes to the GA. The mutated NtRab2 blocks vesicle releas e from the GA so that the normal delivery of Golgi cargo to their destinations, such as the cell surface, was disrupted, inhibiting pollen tube expansion (Cheung et al., 2002). Cytoskeleton Dynamics and Cell Expansion The plant cytoskeleton includes m icrotubules an d microfilaments that spatially control cell expansion (Smith and Oppenheimer 2005). It is generally believed that microtubules serve as a scaffold for cells and are important for establ ishing and maintaining cel l expansion direction, whereas microfilaments function as a track for vesicl es to specific sites to deliver cargo required for expansion (Mathur and Hulskamp, 2002). Although many observations have shown that cytoskeletal dynamics and proper organization are essential for cel l expansion, much remains to be learned, including the precise ro les of microtubules and microf ilaments in spatial control of cell expansion. There is evidence, however, that shows that the direc tion of the walls main structural component, cellulose, is determined by microtubules. The arrangement of cellulose microfibrils in the wall is a key determinant of the cell expansion pattern a nd is clearly related to the arrangement of cortical microtubules in expanding cells (Smith and Oppenheimer, 2005). A rapidly growing body of knowledge has accumulate d about how the dynamics and organization of both classes of filaments ar e controlled in expanding cells (B askin, 2005). Here, however, we emphasize recent work explaining regulation of the cytoskeleton and its contributions to patterning of plant cell expansion.

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36 The Microtubule Cytoskeleton and Cell Expansion Microtubules orient the deposit ion of m icrofibrils in walls, which determines the cell expansion direction. The directi on of cell expansion is determin ed by the organizational pattern of cortical microtubules because it was found that the latter normally mirrors the arrangement of cellulose microfibrils, the key structural elemen t of the wall, in growing cells. Based on this observation, the co-alignment hypothesis was estab lished. It was hypothesized that movement of cellulose synthase enzyme comp lexes in the plasma membrane is constrained by interactions with the cortical microtubules (Giddings and Staehelin, 1991). To accommodate later, conflicting, observations, this hypothesis has further evolved into several distinct versions, such as the template incorporation model (Baskin, 2001) and the microfibr il length regulation hypothesis (Wasteneys, 2004). Here I will describe the evidence that support these hypotheses, with an emphasis on major advances in recent years. Early observation showed that the deposition of cellulose microfibrils in elongating cells was typically perpendicular to th e axis of cellular expansion. In addition, disruption of these fibrils with colchicine caused an isodiametric expansion. These char acteristics led to the prediction that cytoplasmic elements exist in the cell periphery, orient the deposition of cellulose microfibrils, and constrain the pattern of cell expansion (Green, 1962) Only one year later, electron microscopy showed slender tubules (micro tubules) at the cell cort ex. More importantly, the orientation of these tubules mirrored that of the cellulose microfibrils in the adjacent cell walls (Ledbetter and Porter, 1963). Thereafter, cortical microtubules were often observed to lie parallel to the cellulose microfib rils (Hepler and Palevitz, 1974). Microinjection of rhodamine-conj ugated tubulin into the epidermal cells of pea internodes showed the array shift of the co rtical microtubules be tween the transverse organization pattern and longitudinal one after applicat ion of gibberellic acid for induc tion of cell growth. This shift

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37 is likely to be involved in a range of responses that alter the dire ction of cell expansion (Yuan et al., 1994). Disruption of the dynamics and organi zation of endogenous microtubules with microtubule-modifying drugs gives rise to aberrant cell expansion. For instance, oryzalin is a compound that causes microtubule depolymerizati on, whereas taxol has an opposite effect; it promotes microtubule assembly. Arabidopsis seedlings treated with ei ther oryzalin or taxol display an identical defective phenotype, the radial expa nsion of root cells. This result indicates the importance of microtubule dynamics in cell expa nsion. Additionally, the defective severity of the cortical microtubules in the swelling root cells increases with drug concentration. At low concentrations of oryzalin, micr otubule arrays are disorganized; at medium concentrations they are fragmented, and at high concentrations they are totally depleted. However, in the taxoltreated root cells, the cortical microtubules at the elongation z one display disorganization in directionality compared with th e control cells. At 10 micromolar concentration, many stele cells have more longitudinal microtubules, whereas many cortical cells appear to have more transversely aligned microtubules (Baskin et al., 1994). These e xperiments were repeated later (Sugimoto et al., 2003). The same results were ob tained from an experiment with maize roots treated with oryzalin or ta xol (Hasenstein et al., 1999). Microtubules are polymers of tubulin. Mutations in the genes for tubulin also cause aberrant cell expansion, such as helical elongation. Dominant negative mutations in the -tubulin genes cause left-handed helical growth and clockwise twisting in elongating organs of Arabidopsis because the mutant tubulins are incor porated into microtubules, producing righthanded obliquely oriented cortical arrays in th e root epidermal cells. Additionally, the cortical microtubules in the mutants had increased sensi tivity to microtubule-spec ific drugs, indicating

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38 that the reduced microtubule stability can produce left-han ded helical cell expansion (Thitamadee et al., 2002). The same defective cell expansion was exhibited in transgenic plants with the same mutated version of the -tubulin gene (Abe and Hash imoto, 2005). This result further confirms that disturbance of endoge nous microtubules influences cell expansion. During cell expansion, the dynamic and well or ganized microtubules are mediated by the regulators, the majority of wh ich are microtubule-associated pr oteins (MAPs). Thus, mutations in the MAP genes suppress the dynamics of micr otubules, block their reor ganization, and affect cell expansion. MICROTUBULE ORGANIZATION 1 (MOR1) encodes a member of an ancient family of MAPs. The amino acid sequence of MOR1 is similar to Xenopus MAP215. In Arabidopsis MOR1 regulates cortical mi crotubule organization, likel y through stabilization of microtubules. Mutations in MOR1 generate unstable microtubul es. At the 29C restrictive temperature, the cortical microtubul es in leaf epidermal cells of mor1 mutants break into fragments, but at the 21C perm issive temperature, the microtubules revert to their normal appearance. At the restrictive temperature, the mor1 plants are severely stunted, producing radially swollen and short organs in dicative of aberrant cell expansion (Whittington et al., 2001). FRA2, another microtubule regulator, wa s found to have the activity of severing microtubules in vitro. Through confocal microscopy and immunofluorescence, it was found that the cortical microtubules are disorganized in fra2 mutants. Meanwhile, using field emission scanning electron microscopy for studies on the walls, the fra2 mutation alters the normal orientation of cellulose microfibr ils in walls of expanding cells. The fra2 mutants show reduced cell elongation. These findings strongly support the co-alignment hypothesis that microtubules orient cell expansion through the control of directional deposition of cellulose microfibrils in the

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39 wall (Burk and Ye, 2002). Using the same methods for the cob mutations, strong evidence supporting this hypothesis was also obtained (Roudier et al., 2005) COB is not a MAP, but an anchor to GAP, whic h is involved in regula tion of cell polarity (Fischer et al., 2004). As mentioned above, COB is polarly ta rgeted to both the plasma membrane and the longitudinal cell walls. Additi onally it is distributed in a banding pattern perpendicular to the longitudinal axis via a microtubule-dependent mechanism. The elongating root cells in cob mutants lose capacity for anisotropic expansion and display a swelling phenotype. The defective cells are accompanied by disorganization of the orientation of cellulose microfibrils (Roudier et al., 2005). The direct evidence supporting the microfib ril and microtubule co-alignment hypothesis was recently gained using spinning disk conf ocal microscopy. The process of cellulose deposition was visualized in living cells by fluorescently-tagged CESA. The CESA complexes in the plasma membrane moved at a c onstant rate in a linear track that was aligned and coincident with cortical microtubules. I nhibition of microtubule polymeri zation changed the fine-scale distribution and pattern of m oving CESA complexes in the membrane, indicating a direct mechanism for the guidance of cellulose depos ition by microtubules (P aredez et al., 2006). Signaling pathways of phospholipids and GTPase s are involved in the regulation of the microtubule organization for cell ex pansion. In recent years, rapid advances have been made on understanding microtubul e regulation by distinct signali ng pathways, which are intimately related to anisotropic cell e xpansion, particularly by phospholipid s and Rho of plants (ROPs). A wealth of mutations in the genes encoding comp onents of these pathways causes aberrant cell expansion.

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40 Phospholipase D (PLD) is a key component of the phospholipid signaling pathway. It was found that PLD decorates microtubules in plant cells (Marc et al., 1996) and is localized to the plasma membrane (Marc et al., 1996; Gardiner et al., 2001). PLDs are en zymes that hydrolyze structural phospholipids such as phosphatidylc holine to produce free choline and phosphatidic acid (PA), which function as a second messenger in cell signaling. Biotic and abiotic stresses such as wounding and pathogen infection rapidl y stimulate PLD activity (Laxalt and Munnik, 2002; Wang et al., 2002; Meijer and Munnik, 2003). PLD activation triggers reorganization of plant microtubules (Dhonukshe et al., 2003). Changes of PLD levels disrupt the phospholipid signaling transmission, resulting in aberrant ce ll expansion likely because of the microtubule disorganization. Thus, raised levels of AtPLD 1 generate either swollen or branched root hairs (Ohashi et al., 2003). AtPLD 2 absence reduced the hypocotyl cell elongation (Li and Xue, 2007). ROP GTPases are plant-specific signaling molecules. They potentially interact with cell surface-associated signal perception apparatus fo r such extracellular stimuli as hormones, pathogen elicitors and abiotic st ress. ROP GTPases mediate divers e cellular proce sses, including microtubule dynamics and organization (Nibau et al., 2006). It was found that ROP2 inactivates ROP-interactive CRIB motif-cont aining protein1 (RIC1) in Arabidopsis epidermal leaf pavement cells. RIC1 activity promotes well-ordered co rtical microtubules. The RIC1-dependent microtubule organization not only lo cally inhibits outgrowth, but al so in turn suppresses ROP2 activation in indentation z ones. RIC1 over-expression s uppresses lobe formation, and ric1 mutants exhibit wide neck regions (Fu et al., 2005). Actin Cytoskeleton Dynamics and Cell Expansion The m ammalian actin cytoskeleton not only mechanically supports cell s for formation of various shapes, but also generates a driving force for such diverse cellular or intracellular events

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41 as cell migration, vesicle trafficking, exocytosis, and endocytosis (Kaksonen et al., 2006). As these events occur, actin cytosk eleton dynamics are essential for re-assembly of actin filaments at distinct subcellular locations (Goley and Welch, 2006). The dendritic nucleation model of actin polymerization is used for interpretation of the molecular mechanism of these events (Mullins et al., 1998; Pollard and Borisy, 2003). A leading hypot hesis for force generation is through actoclampin, the ATP hydrolysis-depende nt, affinity-modulated motor unit (Dickinson and Purich, 2002; Dickinson et al., 2004; Zeile et al., 2005). However, the role of the plant actin cytoskeleton is just coming of age. In recent y ears, increasing evidence has shown that the plant actin cytoskeleton is important for cell expansion during cell morphogenesis, particularly for tip growth and anisotropic expansion (Hussey et al ., 2006). The role of the actin cytoskeleton is generally believed to be the de livery of specific vesicles containing cell wall materials to specified sites for local growth. The majority of investigations were done using cell-specific models, such as pollen tubes, root hairs, tr ichomes, and leaf pavement cells (Mathur and Hulskamp, 2002; Schellmann and Hulskamp, 20 05; Smith and Oppenheimer, 2005; Cole and Fowler, 2006). Polarized organization of the actin cytoskeleton is required for tip growth. Both pollen tubes and root hairs offer suitable models to st udy the roles of F-actin organization and dynamics in tip growth. In these cells, it ha s been observed that at least two forms of F-actin exist. One is the actin cables arranged along th e elongation axis; and other is the dynamic fine F-actin localized to the tip (Hepler et al., 2001; Cole and Fowler, 2006). A wealth of observations shows chemical and genetic disruption of F-actin dyn amics and polarized organization in tip-growing cells arrests tip growth.

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42 For investigation of the roles of actin cytoskeleton in tip growth, LatB is the first chemical agent used to inhibit actin polymerization. LatB-treated mai ze pollen tubes display a dosedependent depolymerization of F-actin. The elon gation of the LatB-treated pollen tubes is arrested because of F-actin depolymerization (Gibbon et al., 1999). The same result was obtained in LatB-treated pollen tubes of Picea meyeri (Chen et al., 2007). The pollen tubes treated with 15 nM LatB for 20 hours show severe disruption of actin filaments. The polarized actin cables become short fragments throughout the tubes. In addition, some actin fragments tend to aggregate into clusters in the s ub-apical region of the tube. The tip of LatB-treated pollen tubes swelled because of its loss of polarity (Chen et al., 2007). Genetic disruption of the actin cytoskeleton al so causes aberrant tip growth. Formins are actin-nucleating proteins that stimulate the de novo polymerization of actin filaments in mammalian cells (Kovar, 2006). It has been found that plant formins appear to have the same function as that in mammalian cells. Thus, change d levels of formin expression in pollen tubes affect the dynamics of F-actin and disrupt tip growth. Over-expression of Arabidopsis formin AFH1 in pollen tubes induces the formation of a rrays of actin cables, re sulting in depolarization of tip growth and generation of a broadening tube. Moreover, severe membrane deformation was observed in the apical re gion (Cheung and Wu, 2004). Longitudinal actin cables serve as tracks for moto r proteins that trans port vesicles to the tips of growing pollen tubes and root hairs. Active vesicle transpor t was observed in root hairs, particularly at the growing tips. This cellular process is based on F-acti n, which, when disrupted, arrests vesicle traffick ing (Voigt et al., 2005). Precise organization of the actin cytoskelet on is important for cell morphogenesis. The Arabidopsis trichome provides an excellent model for studies on cell morphogenesis. The

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43 trichome is a large, single cell that develops on the epidermal surface. Its morphogenesis is a complex process, in which an approximately ro und, epidermal cell devel ops into a stellate symmetrical trichome (Schellmann and Hulskam p, 2005). Using this model cell, investigators found indirect evidence supporting the importance of precise actin cytoskeleton organization for trichome morphogenesis in pharmacological experime nts with drugs that affect actin dynamics. When developing trichomes were treated w ith microfilament dest abilizing antagonists (cytochalasin D and latrunculin B) or filamentous actin (F-actin) stabilizing inhibitors (phalloidin and jasplakinolide), both observations shows stage-specific requirements for the actin cytoskeletal array. Although the es tablishment of trichome cell polar ity seems not to need precise actin organization, the rapid expansion of tricho me cells after branching is sensitive to the inhibitors, causing an aborted, swollen stub or a highly elongated and distorted structure because of their disorganized F-actin arrangement (Mathur et al., 1999; Szymanski et al., 1999). The direct evidence supporting the necessity of a precise actin organization for trichome morphogenesis comes from the discovery of the gene s that encode the subunits of Arp2/3 and the WAVE complexes in the Arabidopsis genome. The Arp2/3 complex by itself is inactive and needs the WAVE complex to activate it. These two complexes coordinately regulate actin polymerization, and influence F-actin reorga nization in both mammalian (Goley and Welch, 2006) and plant cells (Schellmann and Hulskamp 2005; Szymanski, 2005; Uhrig and Hulskamp, 2006). Recent work has indicate d that mutations in the com ponents of the Arp2/3 and WAVE complexes cause a common trichome defect, result ing in distorted tricho mes. In addition, the Factin is disorganized in the defective trichomes (Mathur et al., 2003; Basu et al., 2005; Zhang et al., 2005b; Uhrig et al., 2007). The ch aracterizations of the defec tive trichome phenotype and the F-actin disorganization in the dis mutants are reminiscent of tr ichomes treated with anti actin

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44 drugs. These uniform results strongly support a crucial role of a precise actin cytoskeleton in trichome morphogenesis. The Arabidopsis epidermal leaf pavement cells are another ideal model for cell morphogenesis. These cells have a unique struct ure, which produces a jigsaw-like appearance. They exhibit an interlocked arrangement that resu lts from the interdigitation of adjacent cells through the formation of complementary lobes and indentations. Working on these cells, researchers discovered additional evidence suppo rting the critical ro le that the actin reorganization plays in cell morphogenesis. ROP 2, a small GTPase, is redundantly required for normal pavement cell morphogenesis. Genetic disrupti on of ROP2 results in a severe decrease in the lobe elongation of pavement cells. Additionally, fine F-actin is also reduced at the lobes of the defective cells (Fu et al., 2002). ROP2 promotes F-actin a ssembly through interaction with RIC4, which is also expressed in leaves, and lo calizes preferentially at the cortical sites of incipient lobe formation Moreover, RIC4 over-expression promotes the accumulation of fine Factin and generates deep l obes. On the other hand, ric4 mutants display pavement cells with shallow lobes. Thus, the fine cortical F-actin at specific sites promotes outgrowth for lobe formation (Fu et al., 2002; Fu et al., 2005). Although the role of the actin cytoskeleton in the control of plant cell morphology is well established, all these results do not yet offer an explanation of its molecular mechanism. Many questions have yet to be answered. For example, why are abundant actin filaments observed in defective trichomes of dis mutants? Why does the difference between distorted trichomes and normal ones exist only at the late stage of tric home development? Also, tip-growing cells, such as root hairs and pollen tubes, have a strict requirement for actin cytoskeleton. Why do they

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45 display no defective phenotype in the dis mutants? More information is needed to put together the elegant interdigitating mechanism of this jigsaw puzzle.

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46 CHAPTER 2 IRREGULAR TRICHOME BRANCH 3 (ITB3) IS A NOVEL REGUL ATOR OF ACTIN ORGANIZATION Introduction The actin cytoskeleton not only m echanically supports mammalian cells for the formation of various shapes, but also generates a driving force for motility of diverse cellular or intracellular events such as cel l migration, vesicle trafficki ng, exocytosis, and endocytosis (Kaksonen et al., 2006). As these events occur, actin cytoskeleton rem odeling is active in the assembly of actin filaments at specific subcellular locati ons (Goley and Welch, 2006). End tracking motors (actoclampins) at the barbed end of growing actin filaments generate the propulsive force for motile events (Dickinson and Purich, 2002; Dickinson et al., 2004). The dendritic nucleation model of actin polymerization is well established for interpretation of the molecular mechanisms of cell migration (Mullins et al., 1998). Human epithelial fibroblasts and fish epithelial keratocy tes are rapidly moving cells. They both form a protrusion called a lamellipodium with a thin laye r of cytoplasm containing a dense meshwork of actin filaments. While the keratocyte migrates along the substrate surface, actin and its regulators accumulate in lamellipodia. Actin depolymerizing factor (ADF) at the rear of the leading edge severs and depolymerizes actin filaments and crea tes new plus ends for the growth of new actin filaments at the front, where the actin nucleator ARP2/3 and its activator WASP/Scar/WAVE polymerize actin filaments (Svitkina and Bori sy, 1999). Actoclampin hydrolyzes ATP for free energy and pushes the plasma membrane, propel ing the cells forward (Dickinson and Purich, 2002; Dickinson et al., 2004). In mammalian cells, the actin tail, a cometshaped structure of actin filaments at the rear of rocketing cells, is responsible for driving the pathogenic bacteria Listeria and Shigella across the host cell cytopl asm using the same molecular mechanism as the dendritic nucleation model (Cameron et al., 2000). Besides these moving cells, vesicle

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47 trafficking also is dependent on the force genera ted from dynamic actin filaments. For example, endocytic vesicles that budded fr om the yeast plasma membrane were observed to use a comet tail for rapid trafficking deep er into the cytoplasm (Engqvist-Goldstein et al., 2004). Vesicle formation also needs actin filaments, which participate in coated pit formation, vesicle constriction, and vesicle scis sion (Yarar et al., 2005). Duri ng vesicle formation, the actin filaments assembling at endocytic sites bind to dynamin through cortactin (Merrifield et al., 2002; Merrifield et al., 2005). Dynamin and cort actin are important com ponents of the vesicle scission machinery. Cooperating with dynamin, dendritic actin filaments generate a strong tension at the vesicle neck for vesicle budding (R oux et al., 2006). Cortactin may rearrange actin filaments in specific directions (Kessels and Qualmann, 2005). Unlike a mammalian cell, the plant cell is surr ounded by a rigid cell wa ll, thus precluding migration. The actin cytoskeleton is implicated in intracellular organelle motility and vesicle trafficking, particular the Golg i apparatus (GA) movement. In mammalian cells, the GA is located close to and around nuclei, but in plant cells, the Golgi ca rried by myosin, rapidly moves along actin cable tracks throughout the whole cel l (Brandizzi et al., 2003; Hawes and SatiatJeunemaitre, 2005b; Latijnhouwers et al., 2005). In the tip growing cel ls of pollen tubes and root hairs, a polarized actin cytoskeleton enables tip-directed orga nelle and vesicle trafficking. Although pollen tubes and root hairs are two distinct cell types, they share a common morphological form, consisting of a shank, a sub-apic al zone, and an apical zone. The gradient of actin filaments are regularly organized in th ese zones (Cole and Fowler, 2006; Samaj et al., 2006). During pollen tube elongation, thick actin cables are arranged in parall el to the shank and serve as tracks for myosin motors carrying organelles or vesicles to the growing site. In the subapical zone, there is a dense fringe of actin fi laments, which may promote vesicle formation from

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48 endomembrane organelles (Cole and Fowler, 2006). In the apical zone, abundant vesicles are embedded in the actin filament meshwork, whic h appears to propel vesicles to the plasma membrane at the growing tip like the comet tail in animal cells (Hepler et al., 2001; Cole and Fowler, 2006; Samaj et al., 2006). Root hairs dem onstrate polarized characteristics similar to pollen tubes. Abundant and highly motile endosom es were found in root hairs, and their intracellular motility re lied fully on the actin cytoskeleton. At the tip of root hairs, motile F-actin patches have been presumed to propel endosomes to the plasma membrane (Voigt et al., 2005; Samaj et al., 2006). Although trichome cell expansion is not tip gr owth, it is a typical anisotropic diffuse growth. It also was found that the actin cytoskeleton plays an important role in trichome morphogenesis (Szymanski et al., 1999; Smith and Oppenheimer, 2005; Hussey et al., 2006). Based on evidence from experiments with actin i nhibitor-treated trichomes (Mathur et al., 1999; Szymanski et al., 1999) and charac terization of the actin cytosk eleton in distorted trichome mutants (Mathur et al., 2003; Szymanski, 2005; Zhang et al., 2005b), the current hypothesis is that the actin cytoskeleton maintains and c oordinates the growth pattern established by microtubules. Microfilaments are e xpected to play the same role in trichome morphogenesis as in tip growth, which is to deliver sp ecific vesicles contai ning cargoes, such as cell wall materials to specific sites for local growth (Mathur et al ., 2002; Smith and Oppenheimer, 2005). However, during trichome morphogenesis, a gradient of actin filaments was not found. Additionally, microtubules were observed to act as tr acks for GA transport. Mutations in KINESIN-13A cause defects in the GA transport and result in missh aped trichomes (Lu et al., 2005). Therefore, the precise role of the actin cytosk eleton in anisotropic cell expansion is still not known. In this study, ITB3 was found to be a novel re gulator of actin organization in Arabidopsis trichome

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49 morphogenesis. Mutations of the ITB3 gene caused a change in trichome shape. The actin cytoskeleton was aberrantly disorganized in itb3 mutants. Abundant rings formed by actin cables were observed in the itb3 mutant, but never in th e wild type. ITB3 was found to directly bind to actin depolymerizing factor and inhibits it activity. These resu lts indicated that the actin cytoskeleton plays a crucial role trichome morphogenesis, which pr ovides insight into the role of the actin cytoskeleton in anisotropic cell expansion. Materials and Methods Plant Materials and Growth Conditions The fast neutron induced m utant, itb3-27, was isolated in the Rschew (RLD) genetic background (Zhang et al., 2005). The mutants, itb3-1 (SalK_073071) and itb3-2 (Salk_015997) are T-DNA insertion mutant alleles in the Colu mbia (Col) genetic background from the SALK T-DNA Insertion Database (http://s ignal.salk.edu/cgibin /tdnaexpress). The wild type used for construction of the mapping population is the Landsberg erecta (Ler) ecotype. Seeds were sown on a soilless potting medi um, Fafard 2 Mix (Conrad Fafard, Inc., Agawam, MA). Seedlings were grown at 24C unde r constant light, provi ded by 40W cool white fluorescent tubes. Plants were watered with PGP nutrient solution (Pol lock and Oppenheimer, 1999) every two weeks. Positional Cloning of ITB3 The m apping population for cloning the ITB3 gene was generated as described by Zhang et al. (2005). Phenotypically itb3 mutant plants were selected from the F2 population. From each selected plant, one of the cotyledons wa s removed for DNA extraction using the RED Extract-NAmp Plant PCR Kit (Sigma-Aldrich, St. Louis, MO). The isolated DNA was used to map the itb3 mutation relative to simple sequence length polymorphism s ( SSLPs) (Bell and Ecker, 1994).

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50 After the itb3 mutation was mapped to a narrow region, th e expected large de letion was detected by PCR. Plasmid Construction For expression of ITB3 the ITB3 gene, which covers the 113-bp 5 UTR, the 501-bp coding sequence and the 277-bp 3 UTR was a m plified using genomic DNA from RLD wild type plants as the template using the appropr iate primer pair (Table 2-1). The 891-bp PCR product was cloned into pENTR1A (I nvitrogen, Carlsbad, CA) using the BamHI and EcoR I sites, and transferred into either pAM-PAT-GW (Bekir Ulker, Max Planck Institute for Plant Breeding, Cologne, Germany) for expression from the 35S promoter or pCK86 (Arp Schnittger, Max Planck Institute for Plant Breeding) for expression from the GL2 promoter through an LR recombination. To localize ITB3, a 35S:ITB3-GFP construct was made. The 498-bp ITB3 open reading frame was amplified usi ng primer pairs that introduced NcoI sites at both ends of the PCR product. The digested PCR product was cloned into the NcoI site of the GFP fusion vector, pAVA319 (von Arnim et al., 1998). The resulting gene fusion was liberated by digestion with EcoR I and Not I and transferred to pENTR1A through the same cut sites. Finally it was transferred into the destination vectors, eith er pAM-PAT-GW or pCK86 as in the previous constructs. Through the same st rategy, for FRET assays, the 35S:ITB3CFP, 35S:ITB3L4CFP and 35S:ADF3YFP were constructed with pAVA574 c ontaining CFP and pAVA 554 containing YFP. To produce the ITB3 protein, the 501-bp coding sequence of ITB3 was cloned into pET41 Ek/LIC, pET15b through Nde I and BamHI or pET41a through Mfe I depending on the specific tag needed (Novagen, Madison, WI). For pET41a -ITB3-GFP, the PCR products were amplified using primers containing EcoR I sites at both ends of ITB3-GFP. To construct pDBLeu-ITB3, used as bait for the yeast two-hybr id screen, the 501 bp coding sequence of ITB3 was cloned into pDBLeu with ka namycin resistance (provided by Wen-

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51 Yuan Song at the University of Florida) through Not I and Sal I sites. All PCR products were sequenced by the Interdisciplinary Center for Biotechnology Research (ICBR) at the University of Florida to ensure no mutations were introduced. RNA Extraction and RT-PCR Total RNA was extra cted from six-week-old Co l wild-type plants using the RNeasy Plant Mini Kit (Qiagen Inc. Valencia, CA) according to the manufacturers instructions. The full length ITB3 cDNA was amplified, following instructions for the cMaster RT plus PCR System (Eppendorf AG, Hamburg, Germany). First-stra nd DNA synthesis was primed using oligo (dT)20. The cDNA was amplified usi ng the specific primers for ITB3 (see Table 1). The PCR products were sequenced by ICBR at the University of Florida. Plant Transformation For ITB3 subcellula r localization, the 35S:ITB3-GFP constructs were tra nsferred into onion epidermal cells by particle bombardment using the Biolistic PDS-1000/He Particle Delivery System (Bio-Rad, Richmond, CA), and the transformation protocol supplied by the manufacturer was followed. A total of 5 L of DNA (1 g/ L) was precipitated on 3 mg gold microcarriers 0.6 m in diameter (Bio-Rad), by adding 50 L of 2.5 M CaCl2 and 20 L 0.1 M of spermidine. After the precip itated DNA was washed once with 140 L 70% and once with 100% ethanol, it was resuspended in 50 L of 100% ethanol. 10 L of this solution was spread on one rupture disk labeled with a burst pr essure of 1,100 psi. Square tissue sections approximately 2 x 2 cm were cut from onions and placed on Murashige and Skoog (MS) solid medium for bombardment. Fluorescence was vi sualized after 36 hours incubation at room temperature in darkness. The constructs of 35S:ITB3 GL2:ITB3 35S:ITB3-GFP, GL2:ITB3-GFP, and 35S:ITB3CFP were transferred into itb3-27 mutants by the floral dip method (Clough and Bent, 1998).

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52 35S:ITB3L4CFP was transferred into the itb3l-4 mutant, 35S:ADF3YFP was transferred into adf3 mutants. The transgenic plants were select ed using a 1000X dilution of Finale (Farnam Companies Inc, Phoenix, AZ) with 5.78% glufosinate-ammonium. Yeast Two-hybrid Assays The protocol for the yeast two-hybrid assay wa s described by (Ding et al., 2004). The bait construct pDBLeu-ITB3 was transf erred into th e yeast strain CG1945 through the Leu selection marker using the Yeast Transformation Kit (Sigma, St. Louis, MO) according to the manufacturers instru ctions. The prey, pPC86-cDNA, wa s a rice cDNA library, (provided by Wen-Yuan Song at the University of Florida) with Trp as the selection marker in yeast strain Y187. The mated cells of CG1945 and Y187 were sp read on the YPD medium without Trp, Leu, and His for positive selection. The plasmids were isolated from the grown yeast colonies with Zymoprep Yeast Plasmid Miniprep (Zymoprep, Orange, CA 92867) and transferred into XL2Blue Ultracompetent Cells (Stratagene, LaJolla, CA). The genes of interest were sequenced using plasmid DNA from individual bacterial colonies. Protein Isolation The constructs, pET41 EK/LIC-ITB3, pET15b -ITB3, and pET41a-ITB3, were transferred into the host cell BL21 (DE3) (Novagen, Madiso n, W I) through chemical transformation for ITB3 expression. The single col ony with the target construct was inoculated into 500 ml Overnight Express Instant TB Medium (Novagen, Madison, WI) for overnight culture. The harvested host cells were lysed by 1x FasBreak Cell Lysis Reagent (Promega, Madison, WI). For pET15b-ITB3 with the 6xHis-tag, HisLink Protein Purification Resin (Promega) was used for His-tagged ITB3 binding. For pGEX-profilin, pGEX -ADF1 (provided by C. Staiger at Purdue University) and pET41-ITB3, which the proteins of interest were tagged with GST, the GST Binding Resin was used for the pr otein purification following the ma nufacturers instructions for

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53 the BugBuster GSTBind Purification Kit (Nova gen). The 6xHis-tag was removed from the fusion ITB3 with a Thrombin Cleavage Captur e Kit (Novagen) and the GST tag was removed using an Enterokinase Cleavage Capture Kit (Novagen) following the manufacturers instructions. The concentration of the purified proteins was meas ured using a DC Protein Assay Kit (Bio-Rad) following the manufacturers instructions. Pull-down Assay The purified proteins in 1x PBS wer e diluted to a 1g/l concentration with 1x PBS. The binding reaction was done in 100 l of binding buffer, which contained 5 mM Tris-HCl, 100 mM KCl, 1 mM MgCl, 1 mM CaCl2, 50 ng/l 6xHis-ITB3, and 100 ng/l ADF1 or profilin or BSA, pH 7.5. The reaction was allowed to pro ceed at room temperature for 80 minutes. After the binding was completed, 12.0 l of 10x nickel resin binding buffer containing 1 M HEPES and 100 mM imidazole (pH 7.5) and 8.0 l of 50% HisLink Protei n Purification Resin (Promega) were added to the binding buffer. The reaction tubes were rota ted at 12 rpm at room temperature for 60 min. The resin was spun down and washed three times for 10 minutes, with washing buffer containing 100 mM HEPES, 10 mM imidazole, and 0.1% NP-40. 3.0 l NuPAGE LDS Sample Buffer ( 4x) (Invitrogen, Carlsbad, CA) wa s added to the pellet resin suspension (~10.0 l). After a 5 minute heating at 95C, the samples were loaded into NuPAGE 12% Bis-Tris Gel (Invitrogen, Carl sbad, CA) for protein separation. Morphological Analysis SALK lines (SALK_073071, 015997, 019320, 008148, 001114, 001117, and 019328), which were ordered from Arabidopsis Biological Resource Center (The Ohio State University, Colum bus, OH) were examined under a diss ecting microscope, and lines SALK_073071 and 015997 showed segregation of plants with the itb3 trichome phenotype.

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54 Immunostaining of the Actin and Microtubule Cytoskeletons The immunostaining protocol for actin f ilam ents and microtubules, using specific antibodies against tubulin or actin, were de scribed in our previous studies (Zhang and Oppenheimer, 2004; Zhang et al., 2005b) Microscopy Fluorescen t images were collected with a Zeiss Axiocam HRm camera mounted on a Zeiss Axioplan 2 Imaging microscope (Jena, Germany). The following filter sets were used to collect fluorescent images: red fluorescence was obtained with Zeiss filter set 20 (excitation, 546/12; dichroic, 560 LP; emission, 575 to 640), green fluorescence was obtained with Zeiss filter set 10 (excitation, 450 to 490; dichroic, 510 LP; emission, 515 to 565). Optical sections were collected using the Zeiss Apotome and Axiovisi on 4.1 software. Light micrographs were collected with a Zeiss Axiocam MRc5 camera mounted on a Zeiss Stemi SV11 dissecting microscope. For scanning electron microscopy (SEM), previous ly described methods were used (Luo and Oppenheimer, 1999). Double Mutant Construction The double mutants were selected from individual F2 plants. The itb3-27 mutant was crossed with the T-DNA insertion lines, itb3l-4 and adf3 The selected putative double mutants were self pollinated. These F3 individuals were cr ossed with their original parents to check for mutations through exhibition of th eir specific mutant phenotypes. Results Cloning of the ITB3 Gene The m utation in the itb3-27 mutant was mapped between th e markers s2 (to BAC clone T5A14) and s3 (to BAC clone T30E16) on chromo some I (Zhang et al., 2005a). To clone the ITB3 gene, we further mapped the itb3 mutation relative to simple sequence length

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55 polymorphism (SSLP) markers (Bell and Ecker, 1994) to BAC clone F25P12 (Figure 2-1). Because the itb3-27 allele was isolated from a fast neutron mutagenized population, we screened BAC clone F25P12 for deletions by amplifying short regions spaced approximately every 1000 bp along F25P12. An approximately 42-kb deletion was found between the 92.4 and 134.4 kb positions in F25P12, but the deletion did not occur in wild t ype. In the deleted region, there were 16 putative genes (Figure 2-1). The SALK T-DNA insertion database was searched for insertions in these genes, and the resultant insertion lines (Alonso and Stepa nova, 2003) were screened for plants showing the itb3 trichome phenotype. Salk_073071 segreg ated plants that had strong itb3 phenotypes; Salk_15997 line also segregated plants with a weak itb3 trichome phenotype. These three lines all had a T-DNA insertion in At1g56580. The itb3-27 mutant was crossed to these three lines for complementation tests. The results indicated that the insertion mutations in the three lines were allelic to itb3 (data not shown), which demonstrated that ITB3 is At1g56580. The trichomes on the leaves of all three itb3 mutant alleles fit the criterion set for irregular trichome branching, where at least on e branch that is shorter than th e others in length, or at least two branch points positioned sepa rately on the stalk. Additionall y, the trichome size and branch number of itb3 mutants decreased when compared with wild type (Zhang et al., 2005a). To determine why these defects occur in itb3 mutants, we examined the cytoskeletons in itb3 and wild type trichomes by immunostaining with anti bodies against actin and tubulin. Compared to wild type, the microtubules in itb3 trichomes showed no apparent difference (data not shown), but the actin filaments in itb3 trichomes were disorganized (Figure 2-2). In trichomes of developmental stage 2-3, the actin cables in itb3 mutants accumulated ne ar the bottom of the stalk (Figure 2-2C), whereas in wild type more actin cables were distributed close to the top of

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56 the trichome (Figure 2-2B). At stage 3, the disorganization of actin cables in itb3 mutants was more pronounced. Most of the actin cables were pa rallel to the long axis of the stalk in the itb3 trichomes (Figure 2-2F), whereas in wild type actin cables were more abundant in the region between branches (Figure 2-2E). At stage 4, more disorganized actin cables were arranged under the trichome branch points in the itb3 mutant (Figures 2-2I, 2-3A ) compared with wild type (Figures 2-2H, 2-3B). The greatest differe nce was the formation of actin rings in itb3 mutants after stage 3, but these actin rings were rare ly found in wild type (Figures 2-3I, 2-4). ITB3 is a Plant-specific Gene ITB3 encodes an unknown protein with a m ass of about 18 KD and a pI of 6.64. No signal peptide or other known motif was found in the ITB3 protein sequence. Using ITB3 to search the complete Arabidopsis genome, we found a family consisting of 22 members. This family can be grouped into two distinct clades: the ITB3-Like ( AtITB3L-02-05 ) clade in the green square and ITB3 Related ( AtITB3R-01-13 ) clade in the yellow square, with 100% bootstrap support for their occurrence (Figure 2-5). The search of all availa ble genomes of other organisms indicated that ITB3 is a plant-specific gene that is present in all land plants, including moss (Figure 2-6). ITB3 Over-Expression Did Not Generate Novel Phenotypes Over-expression of ITB3 using the 35S prom oter completely complemented the defective trichome phenotype and no additional phe notypes were found. Over-expression of ITB3 with the same construct in wild type did not display any visible changes in the trichome shape or other phenotypes (data not shown). ITB3 Has No Specific Subcellular Location When the ITB3-GFP fus ion construct was transferred into onion epidermal cells through particle bombardment, the GFP signal was located in the cytoplasm and nucleus of transformed cells and was both indistinguishable from that of the control of GFP alone and remarkably

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57 different from the control of the nucleus-localiz ed GFP (Figure 2-7). The same gene fusion was used to stably transform itb3 mutants. The signal distribution in the trichomes was the same as that in the transgenic onion epidermal cells. Additionally, the itb3 mutant trichomes were rescued into wild type trichomes (data not show n), which indicated that the ITB3-GFP carried out normal functions in living cells. ITB3 Interacts With ADF3 in Yeast To search for ITB3 interactors, a yeast two-hybrid screen was perform ed using a rice cDNA library as the prey and ITB3 as the bait. On the plate without Trp, the yeast containing the bait plasmid, pDBLeu-ITB3, grew, but those co ntaining the prey plasmids, pPC86-ADF3 and DCD, did not. On the plate without Leu, the ye ast containing the prey plasmids pPC86-ADF3 and DCD grew, but those containing the bait plasmid, pDBLeu-ITB3, did not. On the plate lacking both Trp and Leu, the y east with both bait and prey plas mids grew. Additionally, this yeast also grew on the plate wit hout Trp, Leu, and His (Figure 2-8). This result indicated that His was produced by transcription activation thr ough ADF3 or DCD bound to ITB3. Therefore, rice ADF3 and DCD proteins are able to interact with Arabidopsis ITB3 in yeast cells. ITB3 Directly Binds with ADF3 in Vitro To f urther confirm the interaction between ITB3 and ADF3, we carried out a pull down experiment. The purified His-tagged ITB3 pu lled down GST-tagged AtADF1 in vitro, but did not pull down GST-tagged profilin (Figure 2-9). This data further supported the results from the yeast two-hybrid assay. The Trichomes are Defective in the Mutants of adf itb3 l-4 and Their Double Mutants To search for additional geneti c evidence that ADF is involved with ITB3 in controlling trichome development, the knockout lines of ADF and other ITB3 family members were examined for trichome phenotypes. A T-DNA insertion in ITB3L-4 gave rise to trichomes with

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58 fewer numbers of branches compared to wildty pe. Plants homozygous for a T-DNA insertion in ADF3 showed caused an increase in tric home branch number (Figure 2-10). Double mutants of adf3 and itb3 displayed trichomes with more branches than itb3 mutants and fewer branches than adf3 mutants. The mutation of ITB3 is additive to the mutation of ADF. However, double mutants of adf3 and itb3l-4 displayed trichomes with fewer branches than their parents. Double mutants of itb3 and itb3l-4 had much more severe decreases in branch number compared to either parent. Therefore, these two mu tations appear to be sy nergistic in controlling trichome branching. The transgenic plants of the itb3-27 mutants with 35S: ITB3GFP or 35S: ITB3CFP displayed the wild-type trichome phenotype. Howeve r, the transgenic plants of either the itb3l-4 mutants with 35S: ITB3L4CFP or the adf3 mutants with 35S: ADF3YFP exhibited mutant trichome phenotypes. These results led us to de vise the constructs that contain their own endogenous promoters for driving the fusion gene expression in the future (data not shown). Discussion Disruption of Actin Cytoskeleton Organ iz ation Leads to Misshapen Trichomes Pharmacological disruption of actin filaments with actin-specific drugs severely affects trichome morphogenesis and changes trichome shapes (Mathur et al., 1999; Szymanski et al., 1999). When the Arabidopsis trichomes were treated with the actin polymerization inhibitor Latrunculin B, no effect was f ound at the initial developmental stages, but after the branching events, the trichome cells rapidly expand. The inappropriately extended trichomes generate distortion, shortened branches, a nd separate branch points. Ge netic disruption of the actin filament dynamics through loss-of-function mutations of the distorted class of Arabidopsis genes gives rise to the aberrant tr ichome phenotypes, similar to those in the pharmacological experiments. The distorted class genes encode subunits of the Arp2/3 complex and its activation

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59 complex. Mutations in the distor ted class genes disrupt actin f ilament dynamics, which causes an altered distribution of cortical acti n cables (Mathur et al., 2003). The dis mutants display changes of overall trichome shape, a reductio n of branch length, and an increase in the distance between two branch points (Basu et al ., 2005; Zhang et al., 2005b). Thes e changes are similar to the trichome defects seen in itb3 mutants. Although the distortion defect was considered as the criterion for mutations in the DIS class genes (Hulskamp et al., 1994), it disappears in some genetic backgrounds. For example, the itb1-1 allele, ( dis3 ) does not display the distorted trichomes in the Wassilewskija (WS) genetic background. However, this mutant still exhibits the itb trichomes (Zhang et al., 2005b). In itb3 mutant trichomes, the actin cytoskeleton is severely disorganized. Therefore, our resu lts strongly support the idea that the actin cytoskeleton has a crucial role in the regulati on of trichome morphogenesis. The Precise Role of the Actin Cytosk eleton in T richome Morphogenesis A wealth of cellular observations indicates that a fine actin filament meshwork or diffuse actin patches promote cell expansion, whereas dense actin bundles or actin cables serve as structural scaffolds or tracks for myosin-based motors to transport organelles and vesicles. The precise role of actin filament s in cell expansion is dependent on actin types and subcellular locations. Our results show that th e fine actin filament meshwork is abundant close to the rapidly growing sites of normal trichomes. However, at rapid growth stages, the itb3 mutant trichomes did not show this meshwork. We hypothesize that these fine actin meshworks promote cell expansion. Similar actin patches have been obser ved in other cell types, such as root hairs (Baluska et al., 2000) and elonga ting lobes of epidermal pavement cells (Fu et al., 2005). Although dense actin cables were ob served in both wild type and itb3 mutants, many of the actin cables formed rings in itb3 mutants. These rings are unlikely to be normal tracks for vesicles. It is conceivable that the rings caused reduced and misdirected delivery of vesicles to the cell

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60 cortex, ultimately resulting redu ced growth. This might be a reason why the trichome size of itb3 mutants is reduced when compared to normal trichomes. Actin Filament Reorganization Is Required for Cell Expansion In response to internal and ex ternal signals for cell expans ion Actin filaments are rapidly remodeled by multiple regulators. Some of the regulators modulate the size and activity of the monomeric actin pool through intera ction with actin monomers. Others change the disassembly property of filamentous actin through binding to the filament sides (Staiger and Blanchoin, 2006). Actin and its regulators establis h a complex and adjustable syst em for plant cells in various environments. When the delicate balance is impaired by lo ss of function of some regulators, unusual consequences occur in actin assembly and cellular architecture. Additionally, recent findings suggest that a class-specific interacti on of actin with its regulators exist for proper remolding of the actin filaments (Kandasamy et al., 2007). ADF is a key regulator of actin. It binds with both actin monomers and filaments. ADF severs filaments, thereby generating new barbed ends for subunit addition (Staiger a nd Blanchoin, 2006). ADF severing activity is regulated by profilin (Didry et al., 1998), ACP1 (Bertling et al., 2004; Chaudhry et al., 2007), AIP1 (Okada et al., 2002; Mohri et al., 2006), phosphorylation (Huang et al., 2006), and pH (Gungabissoon et al., 2001; Chen et al., 2002). Our re sults show that plant ADF binds with ITB3. Additionally, ADF activity is inhibited by IT B3 binding (Oppenheimer and Grey, unpublished data). Our results provide new insight into diffe rences in actin filament dynamics between plant cells and animal cells, which in plant cells is the high ratio of monomeri c to filamentous actin (Snowman et al., 2002; Wang et al., 2005a). ITB3 is a Plant-Specific Regu lator of Actin Organi zation A search of all known organisms shows ITB3 is present only in the plant kingdom. Although actin and ADF both are co nserved in all eukaryotic cel ls, only 31 of the 67 animal

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61 actin-binding proteins appear to be conserved in plants (Hussey et al., 2002). In addition, plants have a higher percentage of m onomeric actin in the total ac tin pool (Gibbon et al., 1999; Snowman et al., 2002; Wang et al., 2005a). Therefore, plant-specifi c actin regulato rs like ITB3 may provide the functions of the animal proteins that are missing from plants. Future Perspectives Plant cell shapes are controlled by the cytoskel eton. Our data show that Irregular Trichom e Branch 3 (ITB3) is a novel regulator of actin cytoskelet on organization. Mutations in ITB3 caused disorganization of the actin cytoskeleton in trichomes resulting in an altered trichome shape. We showed that ITB3 interacts with and negatively regulates the function of actin depolymerizing factor (ADF) in vitro However, two important questions remain: First, how do ITB3 and ADF work together to regulate actin organization, and second, how does this interaction lead to site-sp ecific cell expansion to genera te normal trichome shape. To answer the first question, we need additional information on the molecular mechanism of ITB3 interaction with ADF. For example, doe s ITB3 regulate ADF binding to actin filaments? Does ITB3 regulate ADF phosphorylation or dephos phorylation? To address these questions, we can use actin depolymerization assays and actin filament-severing assays with purified ADF, in the presence or absence of IT B3. In vitro polymerized actin filaments can be labeled by including pyrene-labeled actin monomers in th e assay, and fluorescence microscopy can be used to visualize the actin filaments. The kinetics of F-actin depol ymerization can be monitored by continuous pyrene fluorescence measurements by using a Cary Eclipse fluorescence spectrophotometer (Yokata et al ., 2005). Although we have s hown convincingly that ITB3 interacts with ADF in vitro, it is important to confirm this interaction in planta To this end, Frster resonance energy transf er" (FRET) can be used. An ITB3-CYP fusion can be used as the donor; an ADF-YPF fusion can be used as the acceptor. The gene fusions are transferred into

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62 Arabidopsis leaf epidermal cells through biolistic bombardment, and the FRET efficiency can be determined. The higher the FRET efficiency, the higher the number of ITB3 and ADF interactions. Also, we know that both ADF and ITB3 are members of moderately sized families in Arabidopsis. It is possible that there are family member specific interactions between ITB3 and ADF proteins. Plant ADF family members are grouped into the vegeta tive and reproductive ADFs based on expression pattern. There are seque nce and activity differences between the two groups of ADFs. The ITB3 family has been di vided into two groups ba sed on protein sequence differences. Can ITB3 family members also be organized into vege tative and reproductive groups? This can be done by examining the ex pression of the ITB3 family members in microarray data sets. To address the binding of specific ITB3 members with specific ADF members, two approaches can be applied. The firs t approach is to determine expression patterns of ITB3 and ADF family members in trichomes using in situ hybridization. If two members are co-expressed in specific tissues or organs, they may be binding partners. The second approach is to use the yeast two-hybrid assay. Putative positives from this assay can be confirmed with pull down assays in vitro and in planta using FRET analysis.

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63 Table 2-1. Primers used in this study Primer name Sequence (5 3) Used for* F6D8-25F CTACATTTGTTCATAT ACAGGGAGTTC RLD 134/Ler 109 F6D8-25R GCCGAGATATACTTGGATCATACTG F12P19-26F CTGGAAATATCT GCGAAGTGGAA RLD 119/Ler 93 F12P19-26R CATGAACTGTTTGTGCATCTCTG F5I14-55F CGGATGCGGTTATA TAAATAGAGA RLD 231/Ler 176 F5I14-55R CCCTCCCTTTTCTTGCTACAAA T30E16-57F ACACTCTTTACT GGAAGATGCAA RLD 138/Ler 81 T30E16-57R AACACACCCATGCAAGTGAA F7F22-17F GCTCACACTTTCCAATGGTGT RLD 82/Ler 95 F7F22-17R CCTTGGAAGCGTAGACCCA F25P12-14F GCACGATCCTATGAGTTAGCA RLD 101/Ler 87 F25P12-14R TTACACGCGAGGAATGAAGA F23H11-39F TTTGATGGAGATTT TGCTGATT RLD 119/Ler 80 F23H22-39R ACCATTGACAGTGGAGCTACATT T14G11-12F TTTGGATGGATTTGTGCGTG RLD 81/Ler 69 T14G11-12R CGATGAGGTCAATCCTAAAGATCAG T5A14-14F GACCAATACAGAGATACAAAGCAA RLD 89/Ler 103 T5A14-14R TCCGCTAACTTATCCGACAA T6H22-11F GACAATTTTCTTCTATA TAAGGATGTGG RLD 127/Ler 116 T6H22-11R GGTCATCCTTGCAAGATATCAA ITB3enF GGGTGATTTCATACCACACCACC ITB3 over expression ITB3enR TGGCTATGAAGTAACCGCTGAGAT ITB3GSTpF ATGGGTTTGG TTACAGATGAAGTG GST-ITB3 protein ITB3GSTpR AGCATCTGTGACTGCAACAGCTTC ITB3HISpF ATGGGTTTGGTTACAGATGAAGTGAGAGC His-tagged ITB3 ITB3HISpR CTAAGCATCTGTGACTGCAACAGC *Note: A majority of primers are used for ITB3 positional cloning, for example, RLD 134/Ler 109 is a SSLP marker that has PCR products of 134 bp in RLD wild type and 109 bp in Ler wild type.

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64 nga248 nga240Chromosome 1 Recombinants and BACs about 42kb between 21187150 and 21228933 240 nga111 F7F22 F6D8 T5A14 F25P12 T30E16 F5I14 F12P19 H23H11 710 nga248 1770 256 52 20 7 0 20 34 128 157 Total T6H22 Genes in the deleted region Figure 2-1. Positional cloning of ITB3 The itb3 mutation was mapped near SSLP marker ATPase on chromosome 1. Additional molecular markers were used to map the itb3 mutation to BAC clone F25P12. The numbe rs of recombinants (out of 1770 chromatids screened) are given above BAC cl ones. The locations of all putative genes on BAC clone F25P12 are listed. The numbers inside the flags above specific genes are numbers of recombinants for that marker. ITB3 is indicated by the vertical arrow.

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65 Figure 2-2. Actin cytoskelet on is disorganized in the itb3 mutant. (A), (D), and (G) are wildtype trichomes showing diffe rent developmental stages through scanning electronic microscope (SEM). (B), (E), and (H) ar e wild-type trichomes showing the normal organization of actin cytoskeleton with the identical developmental stages with (A), (D) and (G). (C), (F), and (I) are the itb3 mutant trichomes at the same developmental stages as wild type showing disorganization of the actin cytoskeleton through immunostaining and fluorescent microscopy. Bar = 100 m in all images.

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66 Figure 2-3. Actin cable or ganization in the stalk of trichomes. (A) The itb3 mutant trichome stalks showing clear differenc es in actin organization compared to the wild type in (B). Bar = 10 m.

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67 Figure 2-4. Actin rings in the itb3 mutant. Bar = 5 m

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68 Figure 2-5. Phylogenic tree of the Arabidopsis ITB3 family members

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69 Figure 2-6. Alignment of ITB3 protein seque nce with its homologs in other plants

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70 Figure 2-7. ITB3-GFP is not specif ically localized to any subcellular structure in transformed onion epidermal cells. (A): GFP alone; (B ): GFP-NIa in nucleus; (C): ITB3-GFP. Bar = 5 m

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71 Figure 2-8. Yeast two-hybrid screen for ITB3 interactors. (1): Bait ITB3; (2): bait ITB3 and prey ADF; (3): prey ADF; (4): pre DCD; (5 ): bait ITB3 and prey DCD; (6): positive control.

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72 Figure 2-9. ITB3 directly interacts with ADF in vitro. (1): GST-ADF; (2): the GST-ADF is pulled down by His-ITB3; (3): the GST-profilin fails to be pulled down by His-ITB3; (4) His-ITB3 alone; (5): GST-profilin alone.

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73 Figure 2-10. Trichome shapes are defective in adf3 and itb3l-4 mutants. (A) and (D): Col wild type; (B) and (E): adf3 ; (C) and (F): itb3l-4 Bar = 100 m

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74 CHAPTER 3 IRREGULAR TRICHOME BRANCH 2 (ITB2) IS A PUTATIVE AMINOPHOSPHOLIPID TRANSL OCASE THAT REGULATES TRICHOME BRANCH ELONGATION IN ARABIDOPSIS Introduction Lipids are the m ajor components of all euka ryotic membranes. They mainly include phospholipids, sphingolipids, and sterols, whic h are distributed asymmetrically within bilayer membranes. Phospholipids form the main homogenous planar architecture, but sphingolipids and sterols are rich in microdomains called lipid shells/rafts, which theoretically float freely in the more flui d surrounding membranes analogous to the socalled liquid disordered phase. Distinct lipids also specifica lly localize the two leaflets of membranes. In general, the ami nophospholipid, phosphatidylserine (PS), and phosphatidylethanolamine (PE) are concentrat ed in the cytosolic leaflet, whereas phosphatidylcholine (PC) and sphingolipids are enriched in th e exoplasmic leaflet (Holthuis and Levine, 2005; Pomorski and Menon, 2006). The asymmetric distribution of lipids is generated by energy-dependent flippases that hydrolyze ATPs for energy to translocate speci fic lipids across the lipid bilayer (Pomorski and Menon, 2006). The P-type ATPase is such a flippase and its translocase activity was found not only in the plasma membrane (PM) (Pomorski et al., 2004), but also in the membranes of distinct vesicles (Zachowski and Gaudry-Talarmain, 1990; Alder-Baerens et al., 2006) and the trans Golgi network (Natarajan et al., 2004). In yeast, the DRS2 gene codes for Drsp, a member of the P4-ATPase family in the P-type ATPase superfamily (Ripmaster et al., 1993). Mutations in DRS2 caused an absence of low temperature uptake of a labeled PS analog at the PM (Tang et al., 1996; Gomes et al., 2000). Additionally, loss of the Golgi-associated P4-ATPases Dr s2p and Dnf3p abolished the asymmetric

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75 arrangement of endogenous PE in post-Golgi se cretory vesicles (Ald er-Baerens et al., 2006). These findings indicated an essential role for P4-ATPases in generating and maintaining lipid asymmetry during membrane flow through the Golgi. Additional P4ATPase family members also have been identif ied, and they are all a ssociated with lipid translocation in other species (Ujhazy et al., 2001; Perez-Vict oria et al., 2003; Wang et al., 2004), including plants (Gomes et al., 2000). P4-ATPase is involved in vesicle formation. During the biogenesis of intracellular transport vesicles, lipids need to translocat e from the inner leaflet to the outer one by flippases in membranes (Pomorski and Menon, 2006) In yeast, the absence of the two PM associated P4-ATPases, Dnf1p and Dnf2p, re sulted in a cold-sensi tive defect in the biogenesis of endocytic vesicles (Pomorski et al., 2003) and inactivation of Drs2p caused a decrease in clathrin-coate d vesicle budding from the tr ans-Golgi (Gall et al., 2002; Natarajan et al., 2004). Stimulation of PS a nd PE inward translocation induced the formation of endocytic vesicl es in red blood cells (Birchme ier et al., 1979; Muller et al., 1994). Conversely, enhancement of outward directed lipid transl ocation led to a defect in endocytosis (Kean et al., 1997; Decottignies et al., 1998). The role of flippases in vesicle biogenesis was considered a direct and mechanical action on vesicle budding. P4-ATPases interacted with such cytosolic proteins as guanine nucleotide excha nge factors (GEFs) and small GTPases, which are crucial for the recruitm ent of such coat proteins as clathrin at sites of the lipid translocation (Pomor ski and Menon, 2006; Liu et al., 2007). The membrane curvature, which is generated by lipid translocation that creates an area difference between the two leaflets, prom otes vesicle budding (Pomorski and Menon, 2006).

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76 Vesicle-mediated membrane trafficking play s a crucial role in cell expansion. Polar expansion such as elongation of pollen t ubes and root hairs requires vesicles for transporting materials to build cell walls (S amaj et al., 2006). Abundant vesicles are transported and deposited cell wall molecules at growing sites. Normal vesicle trafficking is also necessary for other cells to initiate or execute anisotropic expansion. Aberrant vesicle trafficking caused lobe reduction of Arabidopsis epidermal pavement cells and shape changes of trichomes (Z heng et al., 2005). In this st udy, we cloned the Arabidopsis ITB2 gene through a map-based strategy. ITB2 is identical to ALA3, a member of the putative aminophospholipid transloc ase (ALA) subfamily in the P4 ATPase family in the P-type ATPase superfamily. Mutations in itb2 mutants reduced the tr ichome branch length. We provide the evidence that plant P4 ATPases regulate cell expa nsion, likely through contribution to the vesicle formation. Materials and Methods Plant Materials and Growth Conditions The itb2-28, itb2-29, and itb2-12 ( 9412-12) m utants were isolated in the Rschew ( RLD) genetic background (Zhang et al., 2005). The itb2-4 (Salk_015929) mutant is a TDNA insertion line in the Columbia (Col) ecotype, based on the SALK T-DNA Expression Database (http://signal.salk.edu/ cgibin/tdnaexpress). The wild t ype used for construction of the mapping population is the Landsberg erecta (Ler) ecotype. Seeds of the SALK lines, SALK_015929, 006470, 067322, 139762, 129494, 133319, 082561, 066531, and 109350, were ordered from the Arabidopsis Biological Resource Center (T he Ohio State University, Columbus, OH). Seeds were sown on a soil-less potting medi um, Fafard 2 Mix (Conrad Fafard, Inc., Agawam, MA). Seedlings were grown at 24 C under constant light, provided by 40W cool

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77 white fluorescent tubes. Plants were watere d with PGP nutrient solution (Pollock and Oppenheimer, 1999) every two weeks. Positional Cloning of ITB2 The m apping population for cloning the ITB2 gene was generated as described by Zhang et al., (2005). The phenotypically itb2 mutant plants were selected from the F2 generation of plants. From each selected plant, one of the cotyledons was removed for DNA extraction using the RED Extract-N-Amp Plant PCR Kit (Sigma-Aldrich, St. Louis, MO). The isolated DNA was used to map the itb2 mutation relative to simple sequence length po lymorphisms (SSLPs) (Bell and Ecke r, 1994; Lukowitz et al., 2000). Because the itb2 mutant was isolated from a fast ne utron mutatgenized population, after the itb2 mutation was mapped to a relatively small region, the presence of a deletion was tested by using itb2-28 DNA as the template to amplify about 500 bp target fragments. After the deletion was found, all the genes in the dele ted region were sequenced to check for mutations using PCR products amplified from itb2-19 DNA. Plasmid Construction To com plement the itb2 mutant, the 2146 bp element at the 5 end of ITB2 replaced the 35S promoter on pAM-PAT-GW through Xho I and Asc I, and the 1380 bp-element at the 3 end of ITB2 was directionally cloned into the pAM-PAT-GW backbone using the Pst I restriction site. Because of the large size of the ITB2 genomic sequence, its cDNA was used for construction. The fu ll length coding sequence of ITB2 cDNA was amplified by RT-PCR from total RNA of from Col wild type. The PCR fragment was cloned into the pBluescript SK II (+) vector. The amplified PCR products were sequenced to identify any mutations that were intro duced during PCR. The mutati ons in the cloned cDNA of ITB2 were corrected in the Col wild-type version by multiple substitutions with the correct PCR

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78 fragments. The resultant wild type ITB2 cDNA was cloned into pENR1A (Invitrogen, Carlsbad, CA) using the Bam HI and Eco RI sites. Finally the ITB2 cDNA was transferred into the modified pAM-PAT-GW vector through an LR reaction. For overexpression of ITB2 the 35S:ITB2 and GL2:ITB2 constructs were made. The ITB2 cDNA of the Col wild-type version wa s transferred into pAM-PAT-GW (Bekir Ulker, Max Planck Institute for Plant Breeding, Cologne, Germany) for expression from the 35S promoter, or pCK86 (Arp Schnittger, Max Planck Institute for Plant Breeding) for expression from the GL2 promoter through an LR recombination. To localize ITB2, a 35S:ITB2-CFP gene fusion was made. Cyan fluorescence protein (CFP) was amplified by PCR using the pAVA574 (von Arnim et al., 1998) plasmid as a DNA template. The stop codon was removed from pENR1A-ITB2 through sequence substitution at the 3 end containing Nde I with the PCR product. The two PCR products were ligated together through blunt ends. The fused fragment was cloned into pENR1AITB2 using the Nde I and Eco RI sites. Finally the ITB2-CFP gene fusion was transferred into pAM-PAT-GW for expression from th e 35S promoter through an LR reaction. RNA Extraction and RT-PCR Total RNA was extra cted from six-week-old Col wild-type plants using the RNeasy Plant Mini Kit (Qiagen, Valencia, CA) accord ing to the manufacturers instructions. The full length ITB2 cDNA was amplified, following the instructions for the cMaster RT plus PCR System (Eppendorf AG, Hamburg, German y). The first strand DNA synthesis was primed using oligo (dT)20. The cDNA was amplified using specific primers of ITB2 The PCR products were sequenced by the Interdisciplinary Center for Biotechnology Research (ICBR) at the University of Florida.

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79 Plant Transformation The constructs, 5 untranslated region ( UTR ):IT B2 : 3UTR, 35S: ITB2 GL2 : ITB2 and 35S: ITB2-CFP, were transferred into itb2-28 mutants by the floral dip method (Clough and Bent, 1998). The transgenic plants were select ed using a 1000X dilution of Finale (Farnam Companies, Phoenix, AZ) with 5.78% glufosinate-ammonium 5.78%. Results Characterization of the itb2 Mutants ITB2 controls the trichom e shape mainly through regulati on of branch expansion. All the itb2 mutant alleles displayed defective tricho mes in which one branch is longer than others (Zhang et al., 2005a). The itb2-12, itb2-19, and itb2-28 mutant alleles are in the RLD genetic background; and itb2-4 is in the Col genetic background. All mutants display identical trichome shape that is a weak distortion and at leas one branch longer than others (Figure 3-1G-L). In the RLD genetics backgr ound, cotyledon shape al so was altered in itb2 mutants. A kidneyshape d cotyledons are sh own in Figure 3-1A-F. In addition to this effect, the genetic background also influe nces the frequency of the phenotypically itb2 mutant plants in F2. The segregation of the itb2 allele does not fit the monogenic model for Mendelian segregation (Table 3-1). In the RLD and Col ecotypes, the ratio of wild-type and mutant plants in F2 was approximately 15: 1, which is closer to the segregation ratio expected for two loci segregating independent ly. However, in the Ler genetic background, the ratio was much lower, which also seems to indicate that the itb2 phenotype results from multiple independent lesions (Table 3-1). Cloning of the ITB2 Gene ITB2 was mapped between ITB3 and ITB4 on chrom osome I (Zhang et al., 2005a). To clone the ITB2 gene, we executed its fine mapping re lative to SSLP markers. Because

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80 the itb2-28 mutant is in the RLD genetic b ackground, additional SSLPs markers were developed between the RLD and Ler ecotypes. In addition, we screened for DNA deletions in the itb2-28 mutant. One set of PCR primers did not produce a product when itb2-28 DNA was used as the template, but produced products when wild-type DNA from RLD or Ler was used. Additionally, the primers in this reaction are complementary to the sequences that are located to the middle of the mapped region, the BAC clone F23H11 (Figure 3-2A). To determine the exact position of the detected deletion, we screened F23H11 through amplification of short regions spaced approximately every 500 bp along this BAC clone. Finally, an approximately 19-kb deletion was found between positions 62757 bp and 81830 bp on F23H11. The sequencing data showed that the breakpoint was TTTAAGCCATGACGCTGAGCGAT//AAAAAGCTTTGATCGTCTTTGAT. The right deletion breakpoint includes 779-bp cDNA of the At1g59780 gene. The letf side of the deletion covered a 1336 bp deletion of th e At1g59820 gene. Additionally, the deleted region covered only five genes (Figure 3-2B), all of which were sequenced using the genomic DNA from the itb2-19 allele as the template for PCR amplification. We identified an approximately 800 bp deletion in the ge ne At1g59820 in this mutant and further sequenced the itb2-12 allele. The same deletion as the itb2-19 allele was found. These results indicated that ITB2 was identical with At1g59820, and that the itb2-19 and itb2-12 alleles were likely to derive from the same allele. For an allelism tests, the itb2-28 mutant was crossed with itb2-19 and itb2-12 mutants. Their F1 generation all displayed the itb2 mutant trichome phenotype. To search for more alleles of the itb2 mutant, the Salk T-DNA insertion lines were examined. The Salk_015929 line segregated into two kinds of phenotypic plants, one of which displayed

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81 the itb2 mutant trichomes. The PCR analysis indicated the two plants with the itb2 mutant trichomes were homozygous for th e T-DNA insertion in At1g59820. The itb2-28 mutant was crossed with the homozygous T-DNA lines of Salk_015929 as a complementation test. The F1 plants all displayed the itb2 mutant trichome phenotype indicating that the plants from SALK_015929 with the itb2 phenotype were itb2 alleles. The ITB2 gene is relatively large, with an 8425-bp genomic coding sequence and a 3642-bp cDNA sequence. The coding sequence consists of 27 exons (Figure 3-2C). Compared to the gene in the Col ecotype the RLD ecotype has six single nucleotidealterations were identified. Among the SNPs de tected, five alterations were located in introns and one alteration was located in the last exon, but this transition causes no change in the protein sequence. Through RT-PCR, the same size the ITB2 cDNA was amplified using the total RNA from Col wild type. The results from sequencing the ITB2 cDNA indicated that the ITB2 structure is identical with th e annotation of The Arabidopsis Information Resource (TAIR) in Figure 3-2C. Complementation of the itb2 Mutant and Over-expression of the IT B2 Gene To complement itb2 mutants and over-express ITB2 we made constructs using the ITB2 cDNA. One reason for using the ITB2 cDNA is that it is much smaller size than the genomic DNA. The ITB2 cDNA that was cloned into the pBluescript SK II (+) vector (SKITB2) had 9 base changes (Figure 3-3) co mpared to the Col wild type. Among these mutations, five cause amino acid changes: at 878 (the position in the cDNA beginning with 1 at A of ATG), A to G (the amino acid transi tion, K to R); at 2554, A to G (M to V); at 2690, T to C (L to P); at 3265, A to G (M to V); and at 3289, A to G (T to A). The transition of L to P and T to A is likely to affect protein function and others are not, because similarity of mutated amino acids with the wild type in structure and property.

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82 The re-cloning of the full length ITB2 cDNA was unsuccessful and no full length ITB2 cDNA clones were avai lable from the stock center. To correct the mutations in the resultant clone of the ITB2 cDNA, two EST clones from the ITB2 gene, G7B4 and M65O03, were ordered from the ABRC (The Ohio State University, Columbus, OH). These two clones both contai n about a 1 kb sequence at the 3 end region of the ITB2 cDNA, which can be used for substitution of the mutated part of SK-ITB2. The host cells that contain G7B4 grew well and the plasmids could be isolated through mini-preparation. However, the host cells that contain M65O023 gr ew slowly and the plasmids could not be isolated through standard metheds. The isol ated plasmids were sequenced and a TCdeletion was found in the G7B4 clone at the 2946-bp position of the ITB2 cDNA. The target fragments were amplified through PCR using the M65O03 DNA as the template and cloned into pBluescript SK. The cells that hosted the resultant construct also grew slowly and compared to the G7B4 clone, much le ss plasmid DNAs could be isolated using standard plasmid isolation procedures. Therefor e, in the later experiments, we added 0.5% glucose to the medium to prevent expression of the genes of intere st in the host cells. Through multiple substitutions, all the changed bases in SK-ITB2 were corrected into the Col wild-type version (Figure 3-3). To complement the itb2 mutant, both the 2146-bp element of the ITB2 gene at the 5 end of ITB2 and the 1380-bp element at the 3 end drove the ITB2 cDNA coding sequence of the Col wild type version. The transgenic plants in the itb2-28 background displayed the wild type trichome phenotype (Table 3-2 and Figure 3-4). This result provided further evidence that the ITB2 gene is At1g59820.

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83 To over-express ITB2 the ITB2 cDNAs were cloned into either pAM-PAT-GW for expression by the 35S promoter or pCK86 for expression by the GL2 promoter through a homologous recombination. The transgen ic plants containing the mutated ITB2 cDNA in the itb2-28 mutant background displayed the itb2 mutant trichome phenotype (data not shown). However, the transgenic plants of the same genetic background as the Col wildtype ITB2 cDNA version exhibited w ild-type trichomes (Table 3-2 and Figure 3-4). This result indicated that the C-terminus is crucial for functional ity. Additionally, no novel visible phenotypes were observed for these transgenic plants. To localize ITB2, we constructed the ITB2 gene fusion with CFP, using the Col wild type ITB2 cDNA. The transgenic plants in the itb2 mutant background with the 35S: ITB2CFP construct displayed the itb2 trichome phenotype. This result further supports the hypothesis that the C-terminus of IT B2 is crucial for functionality. Discussion Mem brane trafficking is essential for es tablishment and maintenance of plant cell polarity, especially for such tip growth as the elongation of pollen tubes and root hairs. At the cellular level, tip growth is achieved through polar-speci fic and cell domain-specific trafficking of vesicl es (Hepler et al., 2001) ; at the molecular level, the tip growth machinery is assembled mainly by small GTPases in the Rab, Arf, and Rop/Rac families and their regulatory proteins such as Rho gua nine nucleotide exchange factors (GEFs) and GTPase activating proteins (GAPs) (Samaj et al., 2005; Yang and Fu, 2007). A trans-Golgi network (TGN) was further proposed as a tiplocalized vesicular compartment integrating targeted secretion and endoc ytosis within the growing tip (Samaj et al., 2006). When pollen tubes are growing, endomemb rane trafficking activity transports secretory vesicles along the flank of the tube to the tip; meanwhile, endocytic recycling

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84 vesicles move back distally along the center of the tube, which forms a reverse-fountain cytoplasmic streaming pattern (Hepler et al ., 2001). The secretory vesicles accumulate within the tip clear zone where they form cl usters and fuse with the apical membrane, depositing new membrane, proteins and cell wall materials to support growth (Samaj et al., 2006). The behavior of vesicles in the growing root hairs is re miniscent of that in growing pollen tubes (Hepler et al ., 2001; Voigt et al., 2005). Although trichome branching is not consider ed to belong to the tip growth, its process is a typical anisotr opic cell expansion. Endomembrane trafficking also plays an important role in plant epid ermal cell expansion, such as Arabidopsis leaf pavement cells and trichomes (Smith and Oppenheimer, 2005). The defective endocytic membrane traffic can cause changes in the shapes of these ce lls. Using FM4-64 to track endocytic membrane traffic, the larger vesicle clusters aggregated and were surrounded with an ring of Golgi stacks in the leaf pavement cells of the cer10 mutant (Zheng et al., 2005). These defects resemble the compartments found in the bref eldin A (BFA)-treated cells. BFA is an inhibitor that disrupts ex ocytosis (Baluska et al., 2002; Samaj et al., 2004). The cer10 mutant leaf pavement cells were considerab ly smaller with less pr onounced lobes, and the cer10 trichomes were smaller with short, crooke d, and aberrant swollen stalks and branches (Zheng et al., 2005). The alterations of these cell shapes indicated th at their anisotropic expansion was compromised, because of def ects in endocytic membrane trafficking. A further molecular model was proposed after analysis of Kinesin-13A, ZWI, and AN functions (Lu et al., 2005; Smith and Oppe nheimer, 2005). The plant Golgi apparatus and secretory vesicles are trans ported by myosin(s) from the perinuclear region to the cell cortex, dispersed to cell expanding sites by Kinesin-13A along the cortical microtubules.

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85 Mutations of AtKinesin-13A increased the trichome branch number and a compromised anisotropic trichome cell expans ion. In the trichomes of the kinesin-13a-1 mutant, the Golgi stacks aggregated (Lu et al., 2005). Further support for this modelcomes from the aberrant trichome shapes found in m yosin mutants (Ojangu et al., 2007). ITB2 is putative aminophospholipid translocas e 3 (ALA3) in the P4 ATPase family. Although 12 members (ALA1-12) in th is family were identified in Arabidopsis their biological functions are less known (Axelsen and Palmgren, 2001). Evidence suggests that ALA1 is involved in generating membrane lipid asymmetry. Down-regulation of ALA1 results in a significant reducti on in plant size and an altera tion of plant morphology at low temperatures (Gomes et al., 2000). It is likel y that the aberrant plants resulted from defective cell expansion. ITB2 regulates an isotropic expansion in trichome morphogenesis most likely through its role in generating memb rane lipid asymmetry for vesicle formation during endomembrane trafficking. To confir m our explanation, we will track membrane trafficking in the itb2 mutants. Future Perspectives ITB2 is a putative flippase that transl ocates am inophospholipids from one leaflet to another. It belongs to the P4 ATPase family. Our results show that mutations in ITB2 cause a defective trichome shape phenotype and a sl ight change in cotyledon shape. These mutants provide good experimental materials fo r further studies on function of ITB2 in controlling plant cell shape. Recently publis hed data showed that aminophospholipid translocase 3 (ALA3) is identi cal to ITB2 (Poulsen et al., 20 08). It was shown that ALA3 localizes to the Golgi. We predict that defec tive Golgi localization can also be observed in itb2 mutant trichomes.

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86 To observe Golgi localization in deve loping trichomes, we can cross the ST-YFP construct to itb2 mutants. The ST-YFP construct is a fusion of the Golgi-localized sialyltransferase enzyme to yellow fluorescent pr otein. This construct labels Golgi stacks in transformed plant cells. Once this construct is introgressed into the itb2 mutant background, localization of Golgi can be obs erved in living, developing trichomes using confocal microscopy. By comparing the locali zation pattern and dynamics of Golgi stacks in itb2 mutants with that of wild type trichom es, we can determine if Golgi dynamics and localization is affected in itb2 mutants. If this is found, th en it suggests that proper Golgi localization is a key to dir ectional cell expansion. Also th is result would suggest that flippase activity is impor tant for Golgi function. The key question that remains to be answ ered is whether or not ALA3/ITB2 is actually a flippase. For this to be shown, flippa se activity has to be reconstituted in vitro. Many groups have attempted this, but have not been successful.

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87 Table 3-1. Segregation of the mutant plan ts in F2 with different genetic background Cross name Number of wt plants Number of mutant plants X2 value* P value* itb2-28 /RLD wt 424 28 0.0022 >0.900 itb2-28 /Col wt 867 52 0.5495 >0.500 itb2-28 /Ler wt 1441 46 25.2859 <0.001 is for 15:1 ratio. Table 3-2. Trichome shapes of the transgenic plants Construct name Promoter% irregular% normalTotal XG61 ITB2 10.7 89.3 799 XG62 35S 8.2 91.8 972 XG63 GL2 5.4 94.6 815 RLD 3.5 96.5 713 itb2-28 89.3 10.7 651

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88 Figure 3-1. Defects in leaf tr ichome and cotyledon shape of itb2 mutants. (A), (G), and (J): RLD wild type; (D), Col w ild type; (B), (H), and (K), itb2-28; (C), itb2-19; (E), (I), and (L), itb2-4 ; (F), itb2-12. Bars = 1000 m in (A)-(F); 200 m in (G)(I); 100 m in (J)-(L)

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89 25 9 5 3 1 1 0 4 8 12 Chromosome 1 Recombinants and BACs F7F22 F6D8 T5A14 F25P12 T30E16 F5I14 F12P19 F23H11 nga248 nga111Genes in the deleted region nga248 nga111 76 Total At1g 59790 At1g 59820 At1g 59810 At1g 59800 At1g 59780 Target geneITB2 : At1g59820 779 bp 1336 bp 19074 bp-deletion A B C Figure 3-2. Positional cloni ng and gene structure of ITB2 The itb2 mutation was mapped near SSLP markers between nga248 and nga111 on chromosome 1. Additional molecular markers were used to map the itb2 mutation to BAC clone F23H11. The numbers of recombinants (out of 76 chromatids screened) are given above BAC clones. All putative genes inside the identified deletion on BAC clone F23H11 are listed. ITB2 structure is shown in (C ). The thick bars in (C) represent exons.

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90 2554 A G 3265 A G 3618 A G 3073 A G 2690 T C 2662 T C 2205 T C 878 A G 3289 A G K R M V M V L P T A ATG TAAPCR product A T M65O03 A PCR product T substitution 1 substitution 2 substitution 3Point mutation Amino acid change Figure 3-3. Mutations and corrections of ITB2 cDNA. All mutations of the cloned ITB2 cDNA are listed in the vectanglar boxe s. The numbers on the above bases are the positions of mutated bases. The respective amino acids encoded by the change genetic codes are also list ed below the thick line representing ITB2 cDNA. Approximate positions of the P CR primers used for substitution are shown by the black arrows with the corrected base above them.

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91 Figure 3-4. Transgenic plants with ITB2 cDNA. (A) and (C), RLD wild type; (B) and (D) the transgenic plant. Bars = 500 m in (A) and (B); 100 m in (C) and (D)

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92 CHAPTER 4 DISPROPORTIONATE (DPP) E NCODES A KETOACYL REDUCTASE INVOLVED IN TRICHOME CELL EXPANSION Introduction The control of plant cel l shape has been suggested to o ccur in three se quential steps: First, cell polarity is established by intrac ellu lar mechanisms and/or extracellular cues. Second, using the established polarity, cytoskel etal rearrangements take place. Third, the cytoskeletal changes enable polarized cell expansion, which includes the inco rporation of membrane and cell wall material at defined ar eas of the cell periphery (Hulskamp et al., 1998; Smith and Oppenheimer, 2005). Arabidopsis trichomes are and excellent mode l for studies on the control of plant cell shape. Through a forward genetic approach, mo re than 20 genes that regulate trichome development have been cloned (Marks, 1997; Schellmann and Hulskamp, 2005). The products of these trichome genes are diverse; their functions include transcription initiation, cytoskeletal organization and ve sicle trafficking. For example, GL1 GL2 GL3 EGL3 TTG1 TRY, ETC MYB23 and CPC all function as transcripti onal regulators (Schellmann and Hulskamp, 2005). DIS1, DIS2, DIS3/ ITB1 WURM CRK GRL, ATRK1 and PIR / KLK belong to the distorted group of trichome ge nes that encode regulators of the actin cytoskeleton (Schellmann and Hulskamp, 2005; Szymanski, 2005). AN ZWI MYA2 and KINESIN-13A are known or predicted to be involved in vesicle trafficking (Smith and Oppenheimer, 2005). Recently, a group of genes id entified by wax and cuticle phenotypes whose products are involved in wax synthe sis and transport (Kunst and Samuels, 2003) have been shown to be involved in trichome morphogenesis (see below). Wax is derived mainly from very long chain fatty acids (VLCFAs), which are required for sphingolipid synt hesis. VLCFA moieties in sp hingolipids are essential for

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93 determining the physical propert ies and characteristics of me mbranes. Sphingolipids are an important class of lipids in the plasma membrane and the endomembrane system (Simons and Toomre, 2000). In yeast and mammalian cells sphingolipids are concentrated in lipid rafts, which are involved not only in cellular trafficking of certain plasma membrane proteins, but also play important roles in signal transduction and ge neration or maintenance of cell polarity (Rajendran and Simons, 2005). Plant lipid rafts are also enriched in sphingolipids, but their role in generation or maintenance of cell polarity has rarely been reported (Bhat and Pans truga, 2005; Grennan, 2007). VLCFA synthesis is a complex process including two stages in different cellular compartments (Kunst and Samuels, 2003). The de novo fatty acid synthesis of C16 and C18 acyl chains occurs in the stroma of pl astids by the soluble enzyme complex called the fatty acid synthase (FAS). The synthesized fatty acyl precursors are further extended to C34 VLCFA chains through the same reactions as the de novo fatty acid synthesis, but these reactions are catalyzed by membrane-bound enzyme complexes called fatty acid elongases (FAE) located in the endoplasm retic ulum. FAE is composed of four enzymes that catalyze four sequentia l reactions: These are 1) c ondensation of malonyl-CoA to acetyl-CoA by 3-ketoacyl-CoA synthase (KCS ), 2) reduction of 3-ketoacyl-CoA by 3ketoacyl-CoA reductase (KCR), 3) dehydr ation of 3-hydroxyacyl-CoA by 3-hydroxyacylCoA dehydrase (DCH), and 4) reduction of trans-2-enoyl-CoA by enoyl-CoA reductase (ECR). The resultant VLCFAs are finally modified into diffe rent kinds of waxes (Kunst and Samuels, 2003). Wax monomers are exported to the cell surface by the ABC transporters such as ABCG12/CER5 (Pighi n et al., 2004) and ABCG11/WBC11 (Bird et al., 2007).

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94 LACERATA (LCR ) encodes a monooxygenase, which catalyzes -hydroxylation of fatty acids ranging from C12 to C18. The trichomes on lcr mutant leaves exhibited underdevelopment with a variety of aberra nt shapes (Wellesen et al., 2001). The FIDDLEHEAD (FDH) gene codes for a KCS. Mutations in FDH have a deleterious effect on trichome differentiation because leaf trichom e number was reduced 2-fold in fdh mutants (Yephremov et al., 1999). ECERIFERUM10 ( CER10) encodes an ECR; mutations in this gene caused defective leaf trichomes, which had short, crooked, and abnormally swollen stalks and branches (Zheng et al., 2005). The BODYGUARD ( BDG ) gene encodes a putative extracellular synthase responsible for the formation of the cuticle. The bdg mutants displayed many misshapen leaf trichomes incl uding ones with flat, bent, and collapsed shapes (Kurdyukov et al., 2006). The maize ( Zea mays) GLOSSY1 (GL1) gene codes for a component in the pathway leading to cuticular wax biosynthesis in seedling leaves. The gl1 mutation results in leaf trichomes that ar e smaller than normal (Sturaro et al., 2005). Mutations in YORE-YORE ( YRE), a putative GL1 homolog in Arabidopsis, also led to small trichomes. In addition, the trichome shape of yre cer1 double mutants was greatly deformed (Kurata et al., 2003). Genetic lesions in ( DESPERADO ) DSO / AtWBC11 an ATP binding cassette (ABC) transporter for wax export, led to a dramatic alteration in wax load and trichome development. The dso/atwbc11 mutants had waxless stems and collapsed and underdeveloped trichomes (Bird et al., 2007; Lu o et al., 2007; Panikashvili et al., 2007). Here, we report the cloning of the DPP gene through a novel strategy amenable to the mapbased cloning of other dominant, homoz ygous lethal mutations. We show that DPP codes for a KCR, and dpp mutations lead to a waxless phe notype in addition to dramatically altered trichome shape.

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95 Materials and Methods Plant Materials and Growth Conditions The dpp m utant was isolated in a genetic screen of fast-neutron mutagenized seeds, in the Rschew (RLD) genetic background, whic h were purchased from Lehle Seeds (Round Rock, TX). The Landsberg erecta (Ler) ecotype was used as wild type for construction of the mapping population. Wild-type plants of th e Columbia (Col) ecotype were used for transformation. Seeds were sown on a soil-l ess potting medium, Fafard 2 Mix (Conrad Fafard, Inc., Agawam, MA). Seedlings were grow n at different temperat ures (18 C, 20 C, 22 C and 24 C) in growth chambers under 16 hr light and 8 hr darkness. Light was provided by 40W, cool white fl uorescent tubes. Plants were watered with PGP nutrient solution (Pollock and Oppenheimer, 1999) every two weeks. Positional Cloning For cloning DPP, a m apping population was genera ted from a cross between the dpp mutant and Ler. Because the dpp mutant was dominant and homozygous lethal, 646 phenotypically wild type plants were selected from the F2 population for mapping. The first leaf pair from each individual was used for DNA extraction, following a standard protocol (Edwards et al., 1991). The isolated DNA was used as a template for PCR to map the dpp mutation relative to simple sequence length polymorphism s (SSLPs) (Bell and Ecker 1994, Lukowitz et al. 2000) and cleaved amplifie d polymorphic sequences (CAPS) (Konieczny and Ausubel, 1993). After the dpp mutation was mapped into a narrow region, all the candidate genes in the mapped region were amplified by PCR of genomic DNA from the dpp mutant to check for deletions, as expected in a mutant isolated from fast neutron mutagenesis. Primers sequences are listed in Table 4-1.

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96 To determine the lesion in the dpp mutant, we crossed dpp plants to SALK lines that were homozygous for T-DNA insertions in the candidate genes located within the mapped region. The SALK lines were obtained from th e Arabidopsis Biologica l Resource Center (The Ohio State University, Columbus, OH). Plants homozygous for the T-DNA insertions were identified through PCR with T-DNA and gene-specific primers designed using the SALK T-DNA insertion primer design website (http://signal.salk.edu/tdnaprimers.2.html). The homozygous T-DNA insertion lines were used as the female and the dpp mutant was used as the male for the crosses. The F1 pr ogeny were grown at temperatures below 20C to ensure that the dpp tricho me phenotype was visible. The segregation of the trichome phenotype in the F1 plants was recorded. DNA wa s extracted from the F1 plants using the DNeasy Plant Mini Kit (Qiagen, Valencia, CA), following the manufact urers instructions. The candidate genes were amplified by PCR, an d the products were sequenced to check for mutations. The sequences were compar ed to that of RLD wild type. Plasmid Construction Geno mic DNA from dpp mutants was used as the template for amplification of At1g67730 using primers DEd7 and DEd8. PCR reac tion conditions were as follows: 94 C, 3 minutes; 25 cycles of 94 C, 30 seconds ; 64 C, 30 s; 68 C, 3 minutes. KOD XL DNA polymerase (Novagen, Madison, WI) was used to amplify the entire At1g67730 coding sequence, which comprised 4967 bp. The PCR pr oducts were cloned into pBluescript SK using the Pst I and Eco RV sites. Positive clones were sequenced to check for mutations in the cloned gene. The GC-deleted dpp allele was also cloned into pAM-PAT-GW (Bekir Ulker, Max Planck Institute for Plan t Breeding, Cologne, Germany) using the Asc I and Pst I sites. This clone was used to transform Col and RLD wild type plants to regenerate the dominant dpp trichome phenotype.

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97 Plant Transformation Plants were transform ed by the flor al dip method (Clough and Bent, 1998). Transgenic plants were grow n at 16 C, and sprayed with a 1000X dilution of Finale (5.78% glufosinate-ammonium) (Farna m Companies, Inc., Phoenix, AZ). Results Characterization of dpp Mutants To gain additional insight in to trichom e morphogenesis, we screened fast neutronmutagenized Arabidopsis populations for plants that disp layed defects in trichome shape. One of the mutants isolated in this screen showed shorter branches and longer stalks (Figures 4-1B, 4-1C) than wild type (Fi gures 4-1E, 4-1F). This mutant was named disproportionate ( dpp). During routine plant growth, we noticed that the dpp phenotype was temperature sensitive; at temperatures below 22C, plants displayed the dpp phenotype, while at temperatur es above 24C, the trichomes developed normally. At temperatures between 22C and 24C, most of the trichomes on the first leaf pair had a dpp phenotype, but later leaves had a higher pr oportion of normal trichomes. In addition, dpp leaf blades adhered to each other at an early developmenta l stage (Figure 4-1A), whereas the wild type leaves are open at the same stage (Figure 4-1D). When dpp plants were crossed to other wild type plants, the resulting F1 plants segregated for the dpp phenotype. This result showed that the dpp mutation was dominant. The resulting F2 progeny also displayed segregation of both dpp and wild type plants in a ratio of 2:1, which fits a monogenic model for dpp (Table 4-2), assuming that the dpp homozygotes are lethal. We also noticed that all dpp plants segregated both dpp and wild type plants in the next generation. This phenomenon was examined in more detail by scoring the phenotypes of at leas t 50 progeny from 108 individual dpp plants. The plants in

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98 all the observed populations segregated for dpp and normal trichomes. This result indicated that the populations were derived fr om the homozygous plants. Therefore, dpp plants are homozygous embryo lethal because no homozygous plants were found. To test transmission of the dpp allele, the dpp mutant was reciprocally crossed with RLD wild type or Salk_143503. When the dpp mutant was used as the female parent, a higher frequency of F1 plants with the dpp phenotype occurred than when the dpp mutant was used as the male parent (Table 4-3). Because dpp mutants are he terozygous, when dpp is used as the male parent, both dpp and wild type pollen are tran sferred to the stigma. This result indicated that the dpp pollen was less competitive than wild type pollen during pollination and/or fertilization. Positional Cloning of DPP To clone the DPP gene, we generated an F2 m apping population consisting of 646 wild type plants. Wild type plants were used because the dpp homozygotes do not survive, and known homozygous plants need to be ge notyped to identify recombinants. The DPP locus was mapped relative to SSLP markers to two BAC clones, T1F15 and F12A21. It was further mapped relative to CAPS markers to an 82 Kb region on F12A21. According to the TAIR database, this region covers 34 putative genes (Figure 4-2). Because the dpp mutant was isolated from fast neutron-mutagenized seeds, we expected this mutation was caused by a deletion. We designed primers to amplify 0.5 Kb to 3 Kb fragments that cover the region of F12A21 that contained dpp. PCR products were amplified from both wild type and dpp mutants, and the sizes of the PCR products were compared with each other to determine the deletion position. Results showed that PCR products that differed in size from those predicted for wild type were generated by nonspecific binding of the primers, and not due to a deletion in the dpp mutant. The PCR

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99 products were further sequenced through amp lification of the codi ng sequences of the putative genes in this region us ing the genomic DNA from the dpp mutant plants. This strategy identified single nucle otide polymorphisms (SNPs) between the of RLD and Ler ecotypes (Table 4-4 and Figure 43), and these SNPs were used as the markers for further mapping the dpp mutation. Many sequencing results we re equivocal because the genomic DNA used for sequencing was extracted from the heterozygous dpp plants. For example, see sequencing results of At1g67760 (Figur e 4-4) and At1g67730 (Figure 4-5, 4-6). At1g67760 encodes a chaperonin and At1g67730 encode s a ketoacyl reductase. There were no clear connections between these two gene products and the regulation of trichome shapes. Therefore, they were not immediately pursued further. Identification of DPP The hom ozygous lethality of the dpp mutation presented a challenge to identify the molecular lesion in the dpp allele because all dpp plants were heteroz ygous. Therefore, to identify the lesion in the dpp allele, we developed a strategy whereby we could specifically amplify the dpp allele from the heterozygous dpp plants, which was accomplished by crossing the dpp mutant to a homozygous SA LK lines (Figure 4-7). The dpp mutation is dominan, the heterozygous plants grow normally except for the dpp trichome phenotype, and the homozygous dpp plants are lethal. If the T-DNA insertion in the Salk lines causes a decrease in DPP gene expression, the F1 plants may display a novel phenotype. The result of phenotypic segreg ation in the F1 plants from crosses of dpp to homozygous SALK lines confirmed this prediction. Among the 10 homozygous SALK lines (with T-DNA insertions in 7 distinct genes) two F1 families showed lower transmission of the dpp mutation (Table 4-5). These two SALK lines had a T-DNA inserted in the 5 UTR of At1g67730. The F1 plants from one of these combinations segregated for

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100 a novel phenotype: crinkled leaves, collapsed tr ichomes, waxless stems, and fused floral organs (Figure 4-9). Some of these phe notypes are similar to those displayed by deadhead ( ded ) mutants (Lolle et al., 1998). The crinkled leaves and severe defective trichomes led us to focus our attention on At1g67730. We re-sequenced the PCR products using the DNA from the dpp plants as the template. The result remained equivocal (F igure 4-10). However, the forward reaction worked well with the same DNA sample (F igure 4-11), suggesting that a deletion or insertion likely occured in this gene. When the F1 plants that had the novel phenotypes grew larger, DNA was isolated from their leav es for further analysis. By designing primers that flank the T-DNA insertion, such as primers 720S1 and 720S2 or 730S1 and 730S2 (see Table 4-1 and Figure 4-12), we could specifically amplify the dpp allele, using the isolated DNA from the F1 plants. Sequencing of the PC R product revealed a 2 bp deletion (GC) at position 1595 and 1596 of the coding seque nce of gene At1g67730 (Figure 4-13). This deletion was not observed in the sequence of PCR products from DNA extracted from F1 plants that had a wild type trichome phenotype (Figure 4-14). To confirm that the GCdeletion also exsits in the dpp mutant and that DPP is At1g67730, the entire coding sequence of At1g67730 including upstream a nd downstream intergenic regions was amplified from DNA isolated from dpp plants using primers DEd7 and DEd8 (Figure 412). The PCR products were cloned into the pBluescriptSK vector. The plasmid DNAs from two individual colonies were sequenced. One of them showed the GC-deletion (Figure 4-15) (and other was wild ty pe (Figure 4-16). The GC-deleted DPP was used to transform wild type plants. Plants transforme d with this GC-deleted transgene showed the same trichome defect as the original dpp mutant (Figure 4-17). These results further

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101 confirm that DPP is At1g67730, which encodes a ketoacyl reductase, basing on the gene annotation (Arabidopsis Information Resource). Discussion DPP Encodes a -ketoa cyl Reductase We isolated the dpp mutant during a screen for mutants with altered trichome shape. We found that the dpp mutation was dominant and homozygous lethal. In addition, the dpp mutants had a temperature sensitive phenotype ; at 24C and above, the trichomes appeared perfectly normal, whereas at 22C and below, the trichomes displayed an elongated stalk and short branches. To clone DPP, we combined microarray information from lasercapture microdissected trichome cells with traditional positional cloning methods to identify the most likely candidate gene, At1g67730, for further analysis. At1g67730 encodes a -ketoacyl reductase (KCR) that is invol ved in cuticular wax biosynthesis. To overcome the problems associated with identifying the dpp mutation from DNA isolated from heterozygous plants, we devised a novel strategy using primers flanking a T-DNA insertion in At1g67730. Using these primers a nd DNA from F1 plants from a cross of dpp with At1g67730 T-DNA insertion, only the dpp allele was amplified by PCR and the dpp mutation unequivocally identi fied. Recapitulation of the dpp phenotype in wild type plants containing the mutant dpp transgene confirmed that At1g67730 is DPP. The straightforward cloning of dpp demonstrates the efficacy of our approach for the positional cloning of other dominant, hom ozygous lethal alleles. DPP Has Pleiotrop ic Functions in Cell Expansion and Wax Synthesis The mutation in the dpp mutant caused a 31amino acid-de letion at the C-terminus of KCR. The mutated KCR resulted in plant le thality, but heterozygous plants in the RLD ecotype background, at the restrictive temperature, exhibited a trichome phenotype.

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102 Otherwise the plants grew normally. Expressi on of the truncated KCR with the endogenous promoter in Col wild type plants exhib ited novel phenotypes such as waxless stems, crinkled leaves, fused organs and severe aberrant trichome shapes. These phenotypes are similar to those seen in the F1 plants from the cross of the dpp mutant with the T-DNA insertion line, except that the phe notype of the transgenic plants is less severe. This may be due to the fact that only one mutant copy of dpp is present along with twocopies of wild type DPP in the transgenic plants. This suggests that the severity of the phenotype is dependent on the dosage of the DPP gene relative to the mutant allele. The truncated DPP protein encoded by the dpp allele is likely to have less function compared to wild type. The more dpp protei n that gets assembled into FAE, the less VLCFAs are synthesized. The dpphomozygous plants are leth al because of the VLCFA deleption. Therefore, these long-chain fatty acids are essential for plant viability. The possibility that the dpp protein is a gain -of-function allele cannot be excluded. For example, the dpp protein might interfere with wax synthesis or transport. Plants with the dpp allele in the Col genetic back ground lacked cuticular wax and had fused leaves and floral organs. These phenotype s are typical of the defects resulting from mutations in genes that are i nvolved in wax synthesis. The GL8 gene in maize (Zea mays L.) codes for a KCR. gl8 mutants reduced the amount and altered the composition of seedli ng cuticular waxes (Xu et al., 1997). The disruption of CER6/CUT1 coding for a KCS reduced wax accumulation on stems (Millar et al., 1999; Fiebig et al., 2000). The FIDDLEHEAD (FDH) gene encodes another KCS (Fiebig et al., 2000; Pruitt et al., 2000). This mutant disp layed defects in organ fusion (Lolle et al., 1992). Mutation of the CER10 gene, which encodes an ECR, caused a

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103 reduction of wax abundance (Zheng et al., 2005). However, the dpp mutation in the RLD background did not cause a reductio n in wax accumulation or organ fusion. The reason for this difference is not known, but this difference was also observed in othe r studies. For example, the appearance of crinkled and fused leaves in the abnormal leaf shape1 ( ale1 ) mutant depended on the genetic background, a nd the mutant phenotypes could be observed in the Landsberg erecta background but not in the Columb ia and Wassilewskija genetic backgrounds (Watanabe et al., 2004). Mutations in CER10/ECR caused a change in the VLCFA content of sphingolipids (Zheng et al., 2005). Sphingolipids play a ro le in generation and maintenance of cell polarity. Genetic or pharmacological inactiv ation of sphingolipid synthases not only prevents polarized hyphal growth, but it also abolishes cell polarity establishment (Cheng et al., 2001). This result was further confirmed by distinct sphingolipid synthases with a chemical genetic approach (Li et al., 2006). The epidermal leaf pavement cells in the cer10 mutants displayed a three-fold reduction in size and less pronounced lobing, when compared to these cells in wild-type leaves. Aberrant cell shapes were caused by a disruption of trafficking since the Golgi stacks aggregated, forming ring-like structures in the cer10 mutants (Zheng et al., 2005). The dpp mutant in the RLD background also exhibited a change in the trichome shape at the restrictive temperature, but the wax and other phenotypes were normal. DPP is Vital for Plant Viability DPP codes for a KCR, a subunit of FAE for VLCFA synthesis. In distinct FAEs, specificity of each elongation reaction on different chain length substrates is determined by the selectivity of a KCS (Millar and K unst, 1997). A large family of 21 KCS-like sequences in the Arabidopsis genome contributes to wax biosynthesis, which takes place

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104 in several different tissues at different stag es of plant development (Kunst and Samuels, 2003; Costaglioli et al., 2005; S uh et al., 2005). In contrast, tw o KCRs apparently have no particular acyl chain length specificity and are shared by distinct FAEs (Kunst and Samuels, 2003). In the yeast genome, because only one gene was found to code for each KCR, TSC13 coding for ECR is essential for yeast viability (Kohlwein et al., 2001), but YBR159w coding for a KCR is not essential (B eaudoin et al., 2002). In the Arabidopsis genome, five genes code for ECRs (Costag lioli et al., 2005). Muta tions in CER10/ECR disrupted normal shoot development and cell ex pansion, but plants were viable (Zheng et al., 2005). The genes coding for KCR are At1g67730/DPP and At1g24470. These two proteins are 44% identical and 68% similar to each other (Kunst and Samuels, 2003). Based on microarray analysis, At1g67730 was expressed to significant levels, but At1g24470 expression is low (Costaglioli et al., 2005). Therefore, it is likely that DPP is the major component of FAE. The Salk-096487 line having a T-DNA in serted in an exon of At1g24470 displayed no vi sible phenotype (Xiaoguo Zhang, Unpublished data). DPP is Likely to be DEADHEAD The deadhead mutation was mapped to chromosome I. It maps to the same location as the dpp mutation. More importantly, among deadhead bulkhead and hothead only the deadhead mutant displayed no wax on stems (Lolle et al., 1998). The F1 from the combination of the dpp mutant and Salk line143505 also displayed this waxless phenotype on stems. Therefore, it is possible that the DPP gene is the DEADHEAD gene. Unfortunately, a complementation test between dpp and ded is not possible, because dpp is dominant.

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105 Future Perspectives DPP encodes a ketoacyl reductase that is one m ajor component of the fatty acid synthesis complex. The dpp mutation causes disproportionate trichomes, which have longer stalks and shorter branches as compared to wild type trichom es. Interestingly, the dpp trichome phenotype is temperature sensitiv e: below 22C, the trichomes showed the dpp phenotype, whereas at temperatures above 24C, the trichomes appeared normal. Additionally, the dpp mutation was monogenic, dominant, and homozygous lethal. A twobase (GC) deletion in the dpp gene causes a 30 amino acid-truncation from the C terminus of encoded ketoacyl reductase. Based on these re sults, several interesti ng questions come to mind. For example, why does this truncated ketoacyl reductase generate a dominant trichome phenotype? Is it a ga in-of-function mutation or a loss-of-function mutation? Why and how is this truncated protein so sensitiv e to temperature? To answer these questions, biochemical assays in vitro need to be performed. For example, activity of the wild type and truncated DPP can be compared in vitro at different temperatures. Plants heterozygous for dpp have only defective trchom es. However, our results shows the F1 plants from the cross of th e dpp mutant with the T-DNA insertion line containing a T-DNA in 5 UTR of DPP display pleitropic phe notypes including collapsed trichomes, crinkled leaves, fused floral or gans, and waxless stems. Some of these phenotypes are similar to the phenotypes seen in deadhead ( ded ) mutants. In addition, the ded mutation has been roughly mapped to the south end of chromosome 1, which is the same location of DPP. Currently it is not known if DPP and DED are same gene. A simple complementation test between dpp and ded cannot be done, because dpp is a dominant mutation. To address this question, the DPP genes from multiple alleles of ded mutants

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106 need to be sequenced. If all the ded alleles have mutations, then it is likely that ded and dpp are the same gene. Published data show that mutations in the gene encoding an enoyl reductase, which is another major subunit of the fatty acid synt hesis complex, caused defects in the Golgi apparatus. Leaf epidermal cells of the mutant display many clustered Golgi stacks. It would be interesting to examine the organization of the Golgi stacks in the dpp mutant. If the dpp mutant shows altered Golgi organization, then this result would s upport the idea that the Golgi apparatus plays an important role in cel l expansion, and that fatty acid synthesis is crucial for proper membrane trafficking. To visualize the Golgi a pparatus in live plant cells, the Golgi-labeled markers such as ST-YFP can be used for this study.

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107 Table 4-1. Primers sequence used in this study Primer name Sequence(5 3) Use for* dCAPS3F GTAGTCGCCTTGAGAAAATCTTC A dCAPS3R CCATTGCCTTTGTTAAAGTTTCA Nde I at At1g68060 dCAPS6F GGTCGCTTCGAGAACAACA T TA dCAPS6R GGTGTGGTCAGGAGTCCTTTA MseI at At1g67865 dCAPS7F GAGAGAATCACACGAATTCAAAAGAAA C C dCAPS7R GGTGATAGCAGAAAGGCCAAAA Mnl I At1g68140 dCAPS9F GGGGTTCTGTCTACTGTGGTAACTCCA T dCAPS9R GGTATTGGATCTTATTTAGAAGCCTC aTagI at AT1G67850 dCAPS10F CCACTCTTTAAATGGAAAATCTGGTCATCATC T A dCAPS10R TGCTTGCAATTGTGATCATCTTG XbaI at BAC F12A21 T30E16-57F ACACTCTTTACTGGAAGATGCAA T30E16-57R AACACACCCATGCAAGTGAA RLD 138/Ler 81 F12P19-26F CTGGAAATATCTGCGAAGTGGAA F12P19-26 R CATGAACTGTTTGTGCATCTCTG RLD 119/Ler 93 T1F15-42F GCTGATAAGCGTATCATCACACA T1F15-42R GGTGCGCCATCAAATAATGT RLD 153/Ler 111 F5A8-15F TGGAGTT AACATATTTTTAATTTATCC F5A8-15R GTGGTCAACATCACATTAAAAACA RLD 143/Ler 128 T22E19-6F CCCAATCTAACGGATTTGAAT T22E19-6R GGGCTTTGTTTCTTGTGAAAT RLD 92/Ler 86 T26J14-42F GTCTTTC AACTGGTTTCAAATTTGT T26J14-42R GTTCCATTTTGGTACTTAGTAATGGAC RLD 102/Ler 102 T6C23-20F CGCTACTAAATTTGGTGGGGGTT T6C23-20R TGAGCCTAAAACTTTAACTTCTGC RLD 118/Ler 98 720S1GCGACCTATAGAGGAGGCATTATTGCG720S2CCTTTTGTTCTGTCTCAAGTTACAGGIdentification of DPP 730S1GGCAACAGCAACCAAGTGCATGTCTC730S2CCTTGCTTACTAGCTTCCTCGAGCIdentification of DPP 730cod1CCTTGAAGAGACGCAAACCAT730cod2TTCTATCCACCTTCGTCCCTTIdentification of DPP DEd7CGTCTTCTCTTCCCTCAGCTADEd8 CACTAGACTGGCTAACTCGGCIdentification of DPP The primers here are used for the DPP mapping and identification. For mapping, dCAPS and SNP markers are listed, for example, tw o primers, dCAPS3F and dCAPS3R, are a primer pair for a dCAPS marker which is located at the gene At1g68060. When the PCR products from this primer pair are digested by Nde I a difference in the size of the digested DNAs between the ecotypes of RLD and Ler w ill be observed. The degenerated bases are bold and underlined. Similarly, T30E16-57F a nd T30E16-57R is a primer pair for a SNP marker. The PCR products from them display a difference in size. For the RLD ecotype, a 138-bp DNA is seen, whereas for the Ler ecotype the size is 81 bp.

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108 Table 4-2. Segregation of tr ichome phenotypes in F2 of the dpp mutant crossed to wildtype plants Genotype dpp Wt Total X2 value* P value* dpp/ + RLD wild type 403 148 551 10.3662 >0.001 dpp/ + Col wild type 178 65 243 4.7407 >0.01 dpp/ + Ler wild type 322 135 457 2.8981 >0.05 is the ratio of 2:1. Table 4-3. Segregation of tr ichome phenotypes in F1 of the dpp mutant reciprocally crossed to wild type plants Genotype dpp Wt Total X2 value (1:1) P value (1:1) LD wild type/ dpp 72 79 151 0.3245 >0.50 dpp/RLD wild type 75 106 181 5.3094 >0.01 Salk_143503/ dpp 23 35 58 2.4828 >0.20 dpp/Salk_143503 44 114 158 31.0127 < 0.001 Table 4-4. Single nucleotide polymorphism identified between the Ler and RLD ecotypes Gene name Position* Ler RLD At1g67790 ATTCCGTATGACGATACAT A / T GACCGATTCTTT A T At1g67760 ATTACTCAGCTCCATTAAT T / G TTCAACTTCATC T G At1g67760 ATCAACCTTTTGTCTTTTA G / A CTGTCTTCACCT G A At1g67670 ATAGGATTTGTCGAGACTT G / T TTTTTGTTTAT G T Red bold letters represent single nucleotide polymorphisms between the Ler and RLD ecotypes

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109 Table 4-5. Segregation of phenotypes in the F1 of the dpp mutant crossed to the Salk lines Gene name Salk line Position dpp Wt Total % dpp C At1g67730 Salk_143503 300-UTR5 11 41 52 0.21 20 At1g67730 Salk_143503 300-UTR5 9 22 31 0.29 24 At1g67730 Salk_039982 300-UTR5 14 41 45 0.31 20 At1g67730 Salk_039982 300-UTR5 10 40 50 0.20 24 At1g67750 Salk_095735c 300-UTR5 6 13 19 0.32 20 At1g67750 Salk_017335c 300-UTR5 34 28 62 0.55 24 At1g67750 Salk_017336 300-UTR5 36 50 86 0.42 24 At1g67680 Salk_025786 Exon 11 11 22 0.50 24 At1g67770 Salk_129146 Exon 34 47 81 0.42 24 At1g67620 Salk_100543 Exon 26 32 58 0.49 24 At1g67630 Salk_017965 300-UTR3 20 40 60 0.33* 24 *The hybrid seeds were collected 9 days afte r pollination, whereas others were 15 days after pollination.

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110 Figure 4-1. The dpp mutant trichomes in the RLD genetic background. (A), (B), and (C): dpp mutants; (D), (E), and (F): RLD wild type. Bar = 1000 m in (A) and (D), 100 in others.

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111 ATPaseChromosome 1 Recombinants and BACs The mutated gene T30E16 F12P19 F5A8 T1F15 F12A21 T26J14 T6C23 T22E19 1292 107 31 5 4/0 0/1 8 10 38 123 ATPase 0 1 1 Total Figure 4-2. Positional cloning of DPP The dpp mutation was mapped near SSLP marker ATPase on chromosome 1. Additional molecular markers were used to map the dpp mutation to BAC clone F12A21. The num bers of recombinants (out of 1292 chromatids screened) are given above BAC clones. The locations of all putative genes on BAC clone F12A21 are listed. The numbers inside the flags above specific genes are recombinant event numb ers at that the sp ecific location. DPP is indicated by the vertical arrow.

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112 Tin RLD; Ain Ler Figure 4-3. Single nucleotide polymorphism between RLD and Ler wild types. N is indicating a single nucleo tide polymorphism in the gene At1g67790 between RLD and Ler wild types. N = T in the RLD wild type; N = A in the Ler wild type A T CC T T A Figure 4-4. Equivocal sequenci ng result using the DNA template from plants heretozygous for the dpp mutation. The overlapping peaks at an exon in At1g67760 are showing in the recta ngle, using DNA from dpp mutant plants.

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113 T ? C ? Figure 4-5. Sequencing re sult of At1g67730 using the dpp mutant DNA as a template. The indistinguishable base s are showed inside the rectangles, but they are distinguishable in the repeated experiment in Figure 4-11. ? A G ? Figure 4-6. Sequencing re sult of At1g67730 using the dpp mutant DNA as a template. The bases of interest are showing indi stinguishable inside the rectangles.

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114 Salk lines Homozygous for T-DNA insertion Pollen The dpp mutant F1 F F R R T-DNA Amplified genes of interest The dpp mutant plant cells Figure 4-7. Schematic explanation of DPP identification. The cross of dpp mutants with T-DNA insertion lines for DPP identification. The F1 is generated by a cross with dpp mutants as the male parent a nd Salk lines homozygous for T-DNA insertion as the female parent. DNA is amplified by PCR with primers flanking the inserted T-DNA. No PCR product is am plified because of a large size of TDNA between the regions, parts of which end sequences are identical with the designated primer pairs.

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115 F1 plantsdpp wt dpp wt Larger deletion Smaller deletion 50 bp 400 bp 100 bp 800 bp 2000 bp 3000 bp 5000 bp 50 bp 400 bp 100 bp 800 bp 2000 bp 3000 bp 5000 bp Figure 4-8. Schematic explanati on of deletion identification in dpp mutants. F1 seedlings are segregated into the dpp and wild-type (wt) plants. DNA is extracted from these two kinds of plants, using for PCR as the template with primers indicated in Figure 4-7. The size of PCR products from dpp mutants and wt DNA are identified by running gels. The deletion of dpp mutations is roughly determined by band positions.

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116 Figure 4-9. Novel phenotypes in the F1 of dpp mutants and T-DNA insertion lines. (A), (B), (C), and (D): RLD wild type; (E), (F), and (G): dpp mutants heterozygous for the dpp mutation and wild type. I, J, and K: F1 heterozygous for the dpp mutation and T-DNA insertion. (H) Showing the wax stem of the Col wild type (left) and the waxless stem of the F1 pl ant (right). (L) Showi ng fused organs of floral meristems. Bar = 1000 m in (A ), (E), and (I); 100 m in others.

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117 G/ A C/ A G A G A G Figure 4-10. Equivocal sequencing result of At1g67730, using the DNA from dpp mutants as the template by the reward primer in the repeated experiment. The bases of interest are showing indisti nguishable in the rectangle. NotT YesC Figure 4-11. Unequivocal sequencing resu lt of At1g67730, using the same DNA as in Figure 4-10 in the repeated experiment by the forward primer. The bases of interest, indicating two i ndistinguishable bases in Figure 4-5, are showing unequivocal.

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118 730S2 730S1 4202 bp 720S2 720S1 7095 bp 730cd1 720cd2 2055 bp T-DNA insertion 4967 bp DEd8 DEd7 Figure 4-12. Identification of DPP. Gene structure of DPP and its neighboring genes are showing exons in green segmental lin es. The T-DNA insertion is showing between the coding sequences of DPP and its neighbor genes. The specific primers used in the DPP identification are showing their positions.

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119 G C Figure 4-13. Unequivocal sequence re sult of At1g67730, using the DNA from ded plants of F1 in the combination of dpp mutants and Salk_143503 as the template. The deleted bases of interest are showing inside the rectangle. G C Figure 4-14. Unequivocal sequencing resu lt of At1g67730, using the DNA from the wildtype plants of F1 in the combination of dpp mutants and Salk_143503 as the template. The bases of interest are showing inside the rectangle.

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120 G C deletion Figure 4-15. GC deletion in dpp cloned into pBluescript SK Wild type Figure 4-16 Wild-type DPP cloned into pBluescript SK

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121 Figure 4-17. Transgenic plants with the mutated DPP in distinct genetic backgrounds. (A), (E), and (I): RLD wild type; (B), (F) (J), and (M): transgenic plants with the mutated DPP in the RLD wild-type ecotype; (C) (G) (K), and (O): transgen ic plants with the mutated DPP in the Col wild-type ecotype; (D), (H) (L), and (P ): Col wild type. (N): F1 between dpp mutants and Salk_143503. Bar = 500 m in (A), (B), (C) and (D); 200 m (E), (F) (G), and (H); 100 m in (I), (J), (K), and (L); 1000 m in (M), (N), (O), and (P).

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122 CHAPTER 5 IRREGULAR TRICHOME BRANCH 4 IN ARABIDOPSIS ENCODE S THE PLANT HOMOLOG OF THE 64 KDA SUBUNIT OF CLEAVAGE STIMULATION FACTOR AND REGULATES TRICHOME MORPHOGENESIS AND FLORAL DEVELOPMENT Introduction Polyadenylation includes tw o sequential reactionsendonuc leolytic cleavage and adenylate-residue addition at sp ecific poly(A) sites in 3 non -coding sequences of pre-mRNAs. During eukaryotic developm ent, polyadenylation pl ays an important role in the regulation of gene expression through contributions to tran scription (Zorio and Bentley, 2004), mRNA export to the cytoplasm (Vinciguerra and Stutz, 2004), mRNA stability, and tr anslation efficiency (Wilusz et al., 2001). The specific cleavage at th e 3-end of pre-mRNAs is performed mainly by two complexes: cleavage and polyadenylation speci ficity factor (CPSF) a nd cleavage stimulation factor (CstF) (Zhao et al., 1999). In mammalian cells the former is a heterotetramer consisting of the CPSF-160, CPSF-100, CPSF-73, and CPSF-30 subunits, which are necessary for both cleavage and polyadenylation (Murt hy and Manley, 1992), while the latt er is a heterotrimer that is composed of CstF-77, CstF-64, and CstF-50 su bunits (Takagaki et al., 1990). A a recent model hypothesized that each component in the CstF complex is a homodimer (Bai et al., 2007). The CstF complex is required for cleavage, but is di spensable for the synthesi s of the poly(A) tail. CPSF-73 is an endoribonuclease that cleaves pr e-mRNAs at poly(A) site s (Mandel et al., 2006). CstF-77 functions as a bridge between CstF64 and CstF-50 (Takagaki and Manley, 1994), and also interacts with the CPSF complex th rough CPSF-160 (Murthy and Manley, 1995). CPSF-160 specifically binds to the canonical AAUAAA el ement (Murthy and Manley, 1992). CstF-64 recognizes and binds to a conserved Uor G/U-rich downstream sequence element (DSE) in premRNAs (Takagaki and Manley, 1997). The CstF complex cooperates with the CPSF complex to

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123 facilitate stable binding of both complexes to a pre-mRNA, and enhances polyadenylation efficiency (Murthy and Manley, 1992). CstF-64 is the limiting component for assembly of the active CstF complex (Takagaki et al., 1996). There is increasing evidence that shows th at altered levels of CstF-64 expression has a significant impact on 3 mRNA pr ocessing, and thus regulates sp ecific gene expression, which modulates eukaryotic development. For example, decreases in CstF-64 expression had a variety of effects on B-lymphoma cells (Takagaki a nd Manley, 1998). Decreased CstF-64 expression in chicken B cells caused lower IgM heavy chai n mRNA accumulation as well as an isoform change due to the utilization of an alternative poly(A) site. Also the cell cycle prolonged, cells arrested in G0/G1, and eventually entered apopt osis. In contrast, incr eased CstF levels in differentiating mouse and human B-cells had the opposite effect; the cells transitioned from G0 to S phase and were induced to proliferate (Tak agaki et al., 1996; Mar tincic et al., 1998). In mouse macrophages following lipopolysacchride s timulation, a 10-fold increase in CstF-64 expression significantly altered th e expression of 51 genes (Shell et al., 2005). The change in gene expression was due to alternative polyadeny lation through the choice of either a strong (or weak) poly(A) site, which removed (or retained) instability elements in the mature transcripts (Shell et al., 2005). Five poly(A) sites, L1 to L5, in the adenovirus major late transcri ption unit (MLTU) are used for generation of distinct mRNAs through a lternative polyadenylati on (Larsson et al., 1992; Mann et al., 1993). During the course of adenovirus in fection, the activity of CstF in HeLa cells varies (Mann et al., 1993). In th e later phases of infec tion, the activity of Cs tF was substantially decreased. Additionally, the interaction of CstF with the L3 poly(A) site of the MLTU was found to be more stable than the interaction of CstF w ith the L1 poly(A) site and the L3 site was used

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124 three times more frequently than the L1 site. Thes e events are essentially a reverse of the events observed during the early phases of infection (Mann et al., 1993). CstF-64 expression was also reported to vary in other cell type s. Mouse testicular cells contai n at least 250-fold more CstF-64 mRNA than liver cells (Dass et al., 2001). Male ge rm line cells in meiosis, have no detectable CstF-64 mRNA (Wallace et al., 1999). The Arabidopsis homologs of mammalian CstF have been cloned, and were named AtCstF-77, AtCstF-64 and AtCstF-50. Additionally, biochemical assays in vitro show that AtCstF-64 binds to mRNA 3 non-coding regions, and interacts with AtCstF -77 similarly to the mammalian CstF-64 (Yao et al., 2002). However, th e role of AtCstF in the control of plant development has not been demonstrated. Here, we report the identification of a developmental role for AtCstF-64 in Arabidopsis. Our re sults show that AtCstF-64, encoded by the ITB4 gene, is highly expressed in growing and proliferating cells, and it is required for normal trichome morphogenesis and floral development. Although th e basic mechanism is widely conserved in all eukaryotic cells, results of our f unctional analysis of CstF-64 suggest that key differences exist in the specific mechanisms of mRNA 3-end pro cessing among yeast, plants and mammals, Materials and Methods Plant Materials and Growth Conditions The itb4-1 and zwi-3 single m utants and the itb4-1 zwi-3 double mutant are in the Columbia (Col) genetic backgr ound (Zhang et al., 2005b). A novel itb4 mutant allele ( itb4-2 ) is also in the Col genetic background. Seeds were sown on a soil-less pott ing medium, Fafard 2 Mix (Conrad Fafard Inc. Agawam, MA 01001, USA). Seedlings were grown at 24C under constant light, which was provided by 40W cool white fluorescent tubes. Plants were watered with PGP nutrient solution (Pollock and Oppenheimer, 1999) every two weeks.

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125 The itb4-2 Mutant Isolation and ITB4 Cloning The F2 m apping population for cloning ITB4 was generated as desc ribed by Zhang et al., (2005). A total of 2310 phenotypically itb4 zwi-3 double mutant plants were selected from the F2 population. One of the cotyledons was removed from each selected plant for DNA extraction with the RED Extract-N-Amp plant PCR kit (Sigma-Aldrich, St. Louis, MO). The isolated DNA was used to map the itb4-1 mutation relative to simple sequence length polymorphisms (SSLPs) (Bell and Ecker, 1994; Lukow itz et al., 2000). After the itb4-1 mutation was mapped to a narrow region where the SSLP markers were not available, cleaved amplified polymorphic sequence (CAPS) markers were used (Koni eczny and Ausubel, 1993). Landsberg er genomic information provided by Monsantos Cereon SNP database ( http://www.arabidopsis.org/Cereon/index.jsp ), was used to design the SSLP and CAPS prim ers for fine scale mapping. Finally the itb4 mutation was narrowed down to an approximately 41-kb region, where no additional markers were available. All of the putative genes in this region were examined for mutations by sequencing PCR products amplified from itb4-1 mutant DNA. Putative mutations were re-checked through sequencing or enzymatic digestion of the PCR pr oducts from different template DNAs isolated from 20 individual double mutant plan ts from the F2 mapping population, the zwi-3 single mutant and Col wild type. The primers used for these gene amplifications are listed Table 1. To confirm that we had identified ITB4 the SALK T-DNA insertion line, SALK_131655, was ordered from Arabidopsis Biological Reso urce Center (The Oh io State University, Columbus, OH). The seeds were sown on plates c ontaining MS medium with 1% sucrose and 50 g/ml kanamycin. Kanamycin resistant seedlings were transferred into pots, and seeds of individual plants were collected separately and sown into diffe rent pots. The phenotypes of the resulting plants were characterized. The T-DNA insertion in SALK_131655 was confirmed by

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126 PCR and sequencing using the ITB4 -specific primers. Loss of detectable full-length ITB4 transcript in this allele was verified by RT-PCR. Transgene Construction For ITB4 overexpression, the ITB4 genom ic region was expressed from either the constitutive 35S promoter, or the trichome and root-hair specific GL2 promoter (Szymanski et al., 1998). The resulting constructs were named 35S:ITB4 and GL2:ITB4, respectively. The genomic sequence was amplified by PCR from Col wild type genomic D NA. The PCR fragment was cloned into pENTR1A (Invitrogen, Carlsbad CA) and transferred into either pAM-PATGW (a gift from B. Ulker, Max Planck Institute for Plant Breeding, Cologne, Germany) for expression from the 35S promoter or pCK86 (a gift from A. Schnittger, Max Planck Institute for Plant Breeding) for expression from the GL2 promoter through an LR recombination. To localize ITB4, 35S:ITB4-GFP and GL2:ITB4-GFP were constructed. The P CR fragment was amplified using primer pairs that introduced an NcoI site at both ends of th e PCR products. The digested PCR products were cloned into the NcoI site on the GFP fusion v ector, pAVA319 (von Arnim et al., 1998). The resulting gene fusions were liberated by digestion with Sma I and XhoI, and transferred to either pAM-PAT-GW or pCK86, which were digested by Pst I, filled in using Klenow, and then digested with XhoI. To produce Gal1:ITB4 the full-length cDNA of ITB4 was amplified by RT-PCR using total RNA from si x-week-old Col wild-type plants. The PCR product was cloned into pYES-DEST52 (Invitrogen, Carlsbad, CA) through an LR recombination. To produce 35S:hCstF-64, 35S: hCstF-64 ,GL2:hCstF-64, and GL2 : hCstF-64 the same 5-leader sequence (TL) and 3-t erminator (Ter) as in the construct of 35S:ITB4-GFP were used. The coding sequences of hCstF-64 and hCstF-64 were amplified by PCR from the plasmids containing hCsF-64 or hCstF-64 (gifts from C. C. Macdonald, Texas Tech University

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127 Health Sciences Center, Lubbock, TX). The PC R products were cloned into the modified pNENR1A with TL and Ter elements using Kpn I and Bgl II. The resulting TLhCstF-64Ter and TL-hCstF-64Ter fusions were transferred into either pAM-PAT-GW or pCK86 through an LR recombination. All primers used for the construc ts are listed in Table 1, and all PCR products were sequenced to confirm that no mutations were introduced. Sequencing was performed by the Interdisciplinary Center for Biotechnology Re search (ICBR) at the University of Florida. RNA Extraction and RT-PCR Total RNA was extra cted from six-week-old Co l wild-type plants using the RNeasy Plant Mini Kit (Qiagen Inc. Valencia, CA) according to the manufactures instructions. The full length ITB4 cDNA was amplified using the cMaster RT plus PCR System (Eppendorf AG, Hamburg, Germany). First strand cDNA synthe sis was primed using oligo (dT)20. The cDNA was amplified using the ITB4 -specific primers (see Table 5-1). The PCR products were sequenced by ICBR at the University of Florida. Plant and Yeast Transformations For ITB4 subcellula r localization, the 35S:ITB4-GFP construct was used to transiently transf orm onion epidermal cells by particle bombardment using the Biolistic PDS-1000/He Particle Delivery System (Bio-Rad, Richmond, CA ). The transformation was carried out using the manufacturers instructions. Briefly, 5 L DNA (1 g/L) was precipitated onto 3 mg gold microcarriers (Bio-Rad) of 0.6 m in diameter, by adding 50 L 2.5 M CaCl2 and 20 L 0.1 M spermidine. After the precipitated DNA was washed once each with 140 L 70% and 100% ethanol, it was resuspended in 50 L 100% ethanol. Ten microliters of this DNA suspension solution was spread on one 1,100 psi rupture di sk. The onion epidermal tissue was placed on solid MS medium for bombardment. Fluorescence was visualized after a 36-hour incubation at room temperature in darkness.

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128 The 35S: ITB4, GL2 : ITB4 35S: ITB4-GFP and GL2: ITB4-GFP constructs were used to transform itb4-1 zwi-3 double mutants and Col wild type plants. The 35S: hCstF-64, 35S: hCstF64, GL2 : hCstF-64 and GL2 : hCstF-64 constructs were used to transform itb4-1 zwi-3 double mutants. Transformation was accomplished using the floral dip method (Clough and Bent, 1998). The transgenic plants were selected using a 1000X dilution of Finale (5.78% glufosinateammonium) (Farnam Companies, Inc., Phoenix, AZ). The Gal1:ITB4 construct was used to transf orm yeast using a Yeast Transformation Kit (Sigma-Aldrich, St. Louis, MO) according to the manufactures instructions. The yeast strains transformed were rna14-1, rna15-1, and w303 wild type. These yeast strains were a generous gift from Franois Lacroute, Centre de Gntique Mo lculaire, Yvette, France. The rna14-1 strain contains a mutation in RNA14 gene, a homolog of CstF-77 in mammals and AtCstF-77 in Arabidopsis; the rna15-1 strain contains the mu tation in RNA15 gene, a homolog of CstF-64 in mammals and AtCstF-64 in Arabidopsis (http://www.uky.edu/~aghunt00/polyA2010.html). Morphological Analysis Arabidopsis trichom es were isolated from l eaves and stained with Toluidine Blue as previously described (Zhang and O ppenheimer, 2004). About 50 plants of itb4-2 and Col wild type were grown in constant light. After one week when the first flower opened on each plant, 50 flowers for each genotype were selected from different plants, and dissected under the microscope to determine the number of sepals, petals and stamens. Microscopy GFP i mages were obtained with a Zeiss Axiocam HRm camera mounted on a Zeiss Axioplan 2 Imaging microscope using Zeiss Filter set 10 (excitation: 450-490, dichroic: 510 LP, emission: 515-565). Zeiss Filter Set 02 (excitation: 365, dichroic: 395 LP, emission: 420 LP) was

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129 used to collect fluorescent signal from DAPI stained tissue. A Zeiss Axiocam MRc5 camera mounted on a Zeiss Stemi SV11 dissecting micros cope was used to obtain light micrographs. Environmental Scanning electron microscopy wa s carried out at the National High Magnetic Field Laboratory (Tallahassee, FL), in an Electroscan Model E-3 environmental scanning electron microscope. Tissue samples were mounted on moist paper towels and scanned at 20 kV under 1-2 torr pressure. in situ Hybr idization Fixation, dehydration, and embedding of Arab idopsis inflorescences and young siliques were performed as previously descri bed (Zhang et al., 2005a). RNA probes were made using the DIG RNA Labeling Kit (Roche Diagnostics, Indianapolis, IN) following the manufacturer's instructions. The gene-specific primer pairs and their antisense primer pairs were designed such that a T7 promoter was introduced (Table 5-1) and the transcription templates were prepared by PCR. The steps of in situ hybridization were e ssentially those described by the Meyerowitz's laboratory at http://www.its.caltech.edu/~plantlab/htm l/protocols.htm l, except for the following changes: the 50% Denhardts Solution (in the hybridization solution) was replaced with 10% Blocking Reagent (Roche Diagnostics, GmbH, Mannheim, Germany), the hybridization temperature was 45 C, and the washing temperature was 50 C. Results ITB4 Encodes AtCstF-64 To further understand the role of ITB4 in trichome development, we cloned the ITB4 gene using positional cloning methods We generated an F2 mapping population through a cross of the itb4-1 zwi-3 double mutant with Ler wild type. The itb4-1 zwi-3 double mutant was used as a parent for the following reasons: first, the itb4-1 mutant trichomes have three branches with a subtle change only in tricho me branch length; second, the zwi-3 mutant trichomes have two

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130 branches and its mutation was known to be located on chromosome V, unlinked to itb4-1 ; finally, the itb4-1 zwi-3 double mutant displayed an unequivocal trichome phenotype, i.e., unbranched trichomes. Therefore, we were able to unambiguously identify itb4 plants in the F2 mapping population. A total of 2310 plants with unbranch ed trichomes were selected from the F2 population. Through the use of SSL P markers, we mapped the itb4-1 mutation to a region between BAC clones F23N20 and F28P 5 BAC. Using CAPS markers, itb4-1 was further narrowed down to an approximately 41-kb region on the BAC clone F14O23 according to the TAIR database (Figure 5-1A). This region contains 10 putative gene s. All open reading frames in this region were amplified by PCR using genomic DNA from itb4-1 seedlings. The PCR products were sequenced, and a C to T transition at base 1896 (starting at the A in the ATG start codon) was found in gene Atg71800 The mutation, which was located in the last second exon of At1g71800, created a new TAA stop codon, causing an 88 amino acid truncation from the Cterminus of the putative ITB4 protein (Figure 5-1A). To confirm that this mutation also exists in the itb4-1 zwi-3 double mutants, we randomly selected 20 DNA samples from the F2 mapping population for amplification of the target sequence that contains the mutated ba se. Because the mutation creates an Mse I cut site in the amplified target sequence, the PCR products were digest with MseI. Our results showed all the itb4-1 zwi-3 double mutants indeed had the mutated base in the ITB4 gene (Figure 5-1B). To confirm that the phenotype observed in the itb4-1 mutant is caused by the mutation in At1g71800 the wild type At1g71800 gene was expressed in itb4-1 zwi-3 double mutant plants. The construct used for transformation was a 2800-bp genomic fragment, which contained the 5UTR (250 bp), the At1g71800 coding sequence (2245 bp) and the 3 UTR (300 bp), and was expressed from either the 35S pr omoter or the GL2 promoter. Ov er 30 independent transgenic

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131 plants for each construct displayed trichomes w ith two branches identical to the trichomes on zwi-3 mutants (Figures 5-2C, 5-2D). Occasionall y, three-branched trichomes were observed on some transgenic plants (Figure 5-2E). The rescue of the itb4-1 mutant phenotype by the At1g71800 coding sequence demonstrates that ITB4 is At1g71800. Loss-of-function Mutations in ITB4 Cause Aberrant Development of Trichomes and Flow ers ITB4 contains three functional domains: the RNA recognition motif (RRM) that directly binds to mRNA; the Hinge domain that interacts with CstF-77 in mammals and Arabidopsis; and the PC4/sub1/res1 domain that interacts with polymerase processivity factors in mammals and yeast (Herr et al., 2006 ). The mutation in itb4-1 causes a truncation of 88 amino acids at the Cterminus of ITB4. The truncated part include s the KIWI/KELP domain (Cormack et al., 1998), which is homologous to PC4 in mammals and Sub1 in yeast (Herr et al., 2006). The deletion of the KIWI/KELP domain in ITB4 causes only a sli ght alteration in trichome branch length and branch position. To further analyze the function of ITB4 in trichome morphogenesis, we used a reverse genetic approach to seek a co mplete loss-of-function mutation in the ITB4 gene. By searching the SALK T-DNA Express Database (http ://signal.salk.edu/cgi-bin/tdnaexpress), we identified eight lines that contained insertions in the At1g71800 gene: Salk_131655, Salk_088885, Salk_088876, Salk_088877 Salk_150929, Salk_133589, Salk_038729, and Salk_150929. After selection for kanamycin resistance, the progeny from line Salk_131655 segregated plants showing a defective trichome phenotype. Additionally, the segregation ratio of wild type to mutant was nearly 3:1. ITB4 in Salk_131655 is interrupted by a T-DNA insertion in its last exon (Figure 5-1A). Plants homozygous for the T-DNA insertion in this line were identified by PCR-based screening (Figur e 5-1C). This mutant allele was named itb4-2 ITB4

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132 expression was not detectable in the itb4-2 mutant by RT-PCR (Figure 5-1D), indicating that itb4-2 is likely to be a complete lo ss-of-function mutant allele of ITB4 The mutant phenotype caused by the T-DNA insertion mutation in the itb4-2 allele is considerably more severe than the itb41mutant. Because the latter exhibits shorter trichome branches and separate branch pos itions, it probably represents a partial loss-of-function mutation; however, the former shows dramatic defects in trichome morphoge nesis and floral development. The itb4-2 plants displayed changes in both tric home shape and trichome cell fate. The branch length and branch positions of the itb4-2 trichomes were clearly distinguished from wild type. In Col wild type trichomes, 96% trichomes ha ve three branches (Table 5-2) and the lengths of the branches are almost equal. Additionally, th e positions of the primary and secondary branch are adjacent or close to each other (Figure 5-3A). However, the branch number of the itb4-2 trichomes covered a wide range from one branch (Figures 5-3D, 5-3F, Table 5-2) to five branches (Figure 5-3J, Table 5-2) and the percen tage of three-branched trichomes is notably decreased. In contrast, the percentage of two-bran ched trichomes is prominently increased (Table 5-2). The branch lengths and pos itions of the trichomes on the itb4-2 mutant show the same characteristics as the trichomes on the itb4-1 mutant, i.e., unequal bran ch lengths and separated branch positions (Figures 5-3B, 5-3G-J) (Zhang et al., 2005b). Interestingly, the itb4-2 mutation produced trichome cluste rs, which appeared as twins on itb4-2 plants. The twin frequency was 2.52% in the itb4-2 mutant, but never observed in Col wild type (Table 5-2). The branch numbers of th e twin trichomes varied (Figures 5-3C, 5-3F, 53K-N), and they all seem to be conjoined at the bottom of their st alks (Figures 5-3C, 5-3E, 5-3F, 5-3K-J). Separation of the twin trichomes was resistant to treatments with both EGTA and

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133 pectinase (Figure 5-3E) (Zhang and Oppenheim er, 2004), which suggests that the trichomes share part of a cell wall. The other major defect of the ibt4-2 mutant is abnormal development of flowers. Beginning with stage 3 during fl oral development on wild type plants, the abaxial sepal primordia arise first, followed by the adaxial primordia. Entering stage 4, they elongate, curve inward and cover the dome-shaped meristem befo re petals arise (Figure 5-4A). The developing primordia rapidly enlarge from 30 m at stage 3 to 70 m at stage 4 (Smyth et al., 1990). Compared with wild type, the floral primordia on the itb4-2 mutant displayed aberrant development. First, the sepals elongate slowly and do not cover the floral organ primordia after the petals arise (Figure 5-4B, large arrow). Second, the abaxial sepals preferentially grow (Figures 5-4B, 5-4F) and often fuse with adjacent sepals (Figure 4B, small arrow). Additionally, the numbers of sepals and petals is significantly increased (Figures 5-4B, 5-4C, 5-4F, 5-5); floral buds with seven sepals were often observed (F igure 5-4C). Throughout flower development, the defective sepals were unable to completely en close the stamens and carpels as in wildtype (Figures 5-4D, 54E). Third, the stigmatic papill ae were aberrant (Figures 5-4H, 5-4I). On wildtype stigmas, the papillae stand straight and are regularly arranged, but the papillae on the itb4-2 mutant displayed irregular shap es and clusters (Figures 5-4H 5-4I). The anthers of the itb4-2 mutant contained few pollen grains and the one s that did not form, were mostly unviable (Figures 5-4K, 5-4L). In spite of the extra floral organs, the ove rall floral bud size was smaller than wild type (Figures 5-4B, 5-4E). The itb4-2 mutant plants also showed a relatively minor alteration of rosette leaf color and shape. Col wild-type plants produce green rosette leaves with smooth edges (Figures 5-5A, 5-5C, 5-5E). However, itb4-2 plants produced yellow -and-green mosaic rosett e leaves with serrated

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134 edges (Figures 5-5B, 5-5D). These defects are most prominent in the first l eaf pair (Figure 5-6B), while subsequent leaves di splay a gradual transition toward normal leaf morphology during vegetative growth (Figures 5-6D, 5-6F). ITB4 is Highly Expressed in Growing and Proliferating Cells To gain insight into why itb4 mutations caused the defects in specific cell types and organs, we examined the ITB4 expression pattern by in situ hybridization. We found that ITB4 is highly transcribed in developing trichome cells, embryos, meristems and floral primordia during plant vegetative growth and reproductive develo pment (Figure 5-7). At the globular stage of embryo development in Col wild type, ITB4 was highly expressed in the developing embryo, but not in cells of the suspensor (Fig ure 5-7A). At the heart stage, ITB4 expression became stronger (Figure 5-7B), but in mature embryos, ITB4 expression decreased (d ata not shown). During germination, ITB4 was expressed in the apical merist em and provascular cells. The actively dividing cells in the meristem showed higher ITB4 expression than the gr owing provascular cells (Figure 5-7C). Developing tric home cells displayed a higher ITB4 expression level than other epidermal cells (Figure 5-7E). At stage 1 of trichome development, ITB4 is strongly expressed. The nuclei of trichomes at this stage undergo endoreduplication from 2C to an average of about 8C with a concomitant in crease in nuclear volume. This increase in nuclear size distinguishes cells committed to the trichome fate from othe r epidermal cells (Hulskamp et al., 1994). The strong expression of ITB4 remains until stage 3 or 4, at which time the trichome size increases rapidly from about 20 m to 400 m. During flower development, ITB4 is strongly expressed in floral meristems (Figure 5-7F). At later developmental stages, strong expression of ITB4 can be observed in developing stamens and carpels, but weak expression is observed in developed sepals (Figure 5-7G).

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135 Loss of ITB4 Function Alters the Expression P attern of Perianth Organ Identity Genes Complete loss-of-function of ITB4 caused an increase in the number of sepals and petals, but no difference for stamen and carpel numbers. To understand how ITB4 influences perianth development, we examined the expression of the floral organ identity genes, AP1, AP3 and PI in the itb4-2 mutant and Col wild type fl owers through in situ hybridization. At stage 3 of floral development, AP1 expression domain is restricted to th e outer two whorls because AG represses AP1 expression in the inner two whorls of wild type flowers (Figure 5-8A) (Gustafson-Brown et al., 1994). However, in the itb4-2 mutant, in addition to its strong expression in the outer two whorls at stage 3, AP1 is ectopically expressed in the inner two whorls at stage 3 and even in stamens and carpels at stage 7 (Figure 5-8B). In wildtype, AP3 is expressed from stage 3 in the presumptive second and third whorls After floral stages 5 and 6, AP3 is expressed throughout the developing petals and stamens, and at the adaxial base of sepals (Figure 5-8C) (Jack et al., 1992). In itb4-2 mutant flowers, AP3 expression appears precocious ly and ectopically in the inner two whorls at the stage 3 (Figur e 5-8D). The PI e xpression pattern in ibt4-2 mutants is indistinguishable from that in the wild type (Figures 5-8E, 5-8F). ITB4 Localizes to Nuclei, but Does Not Func tionally Complement Tts Homolog in Yeast ITB4 was annotated as a cleavage stimula tion factor by both the Arabidopsis Genom e Initiative ( http://arabidopsis .org/in fo/agi.htm l) and GSF-MIPS ( http://mips.gsf.de/proj/plant/jsf/athal/sear chjsp/index.jsp). The full length cDNA o f ITB4 was amplified by RT-PCR using Col wild type RNA as a template. The PCR product is identical with the computer-based prediction of both of the data bases mentioned above. In silico translation of the ITB4 coding sequence predicts a pr otein of 461 amino acids with an isoelectric point of 9.32. This protein sequence is identical to that reported by Yao et al (2002) for AtCstF-64. Therefore, we renamed the ITB4/At1g71800 protein to AtCstF-64.

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136 AtCstF-64 functions in the cl eavage reaction for mR NA 3 end formation. It contains the same three functional domains as its homologs in yeast and mammals. It is generally believed that the CstF complex in plants functions in a similar way to the animal counterparts for mRNA 3'-end processing in nuclei (Yao et al., 2002; He rr et al., 2006). To s upport this idea, a GFPtagged version of AtCstF-64, 35S:ITB4-GFP, was transiently expressed in onion epidermal cells through biolistic transformation. The ITB4-GFP signa l was found in nuclei (Fig. 5-9). This result is consistent with the nuclear location of RNA15 in y east (Bonneaud et al., 1994) and CstF-64 in mammals (Schul et al., 1996). To te st the functionality of ITB4-GFP in vivo 35S: ITB4-GFP and GL2 : ITB4-GFP were used to transform itb4-1 zwi-3 double mutant plants through Agrobacterium -mediated stable transformation. Both constructs rescued the unbranched trichome phenotype of the itb4-1 zwi-3 double mutant, and transformants showed the twobranched trichome phenotype of the zwi-3 mutant. However, the nuclear localized ITB4-GFP signal was weaker than that obser ved in onion cells (data not show n). The nuclear localization of the fusion protein supports a function for AtCstF -64 in pre-mRNA processing in Arabidopsis. To determine if AtCstF-64 function is conserve d, we examined the ability of AtCstF-64 to complement yeast CstF subunit mutants. GAL1 : ITB4 was used to transform three different yeast strains: rna151, rna14-1 and w303 (wild type). The rna15-1 strain contains a mutation in RNA15, the counterpart of CstF-64 in mamm als and AtCstF-64 in Arabidopsis; the rna14-1 strain contains a mutation in RNA14, the counterpart of CstF-77 in mammals and AtCstF-77 in Arabidopsis. The mutations in both strains caus ed a temperature-sensitive phenotype. At the permissive temperature, 28 C, the rna15-1 and rna14-1 strains grow. However, at the restrictive temperature, 33 C, their growth stops. The transformed rna15-1 or rna14-1 strains that contained the GAL1:ITB4 construct could not grow at 33 C, but the transformed wild type strain

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137 grew normally (data not shown). These results indicate that AtCstF-64 does not functionally complement its counterpart in yeast. To further examine the functional conservation of AtCstF-64, we attempted to rescue the itb4 phenotype using CstF-64 homologs from mammals. 35S:hCstF-64 35S: hCstF-64, GL2 : hCstF-64 and GL2 : hCstF-64 were used to stably transform itb4-1 zwi-3 double mutant plants. The human counterpart of AtCstF-64, hC stF-64, exhibits 32% identity and 44% similarity to the Arabidopsis protei n sequence. The human h CstF-64 protein is a paralog of hCstF-64 that exhibits 74% protein sequence iden tity with hCstF-64 and is expr essed in male germ cells to maintain normal spermatogenesis (Dass et al., 20 01; Dass et al., 2002). None of the transgenic plants showed rescue or any other change in the itb4-1 zwi-3 double mutant trichome phenotype (data not shown). These results indicate th at the mammalian CstF64 does not functionally complement its counterpart in Arabidopsis. Discussion Polyadenylation is a co mmon event that occurs in the nuclei of all eukaryotic cells, during which a majority of mRNAs receive a string of A residues. Additionally, 25% mRNAs in Arabidopsis use alternative polyadenylati on sites (Meyers et al., 2004). Alternative polyadenylation of a number of mRNAs has been shown to affect plant development (Cheng et al., 2003; Quesada et al., 2005). Although there is increasing data to sugg est that alternative polyadenylation contributes to the c ontrol of gene expression in anim als, its role in the regulation of development in plants is not well understood. The results presen ted in this paper are the first report that a plant homolog of a subunit of the Cs tF complex influences specific events during plant development.

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138 ITB4 Plays a Crucial Role in Trichome Morphogenesis and Floral Development The itb4-1 mutation leads to a truncated AtCstF -64 protein that lacks the KI WI/KELP domain, and itb4-1 plants display only a relatively mild trichome shape defect. The truncated AtCstF-64 protein retains the other important functional domains such as the RRM and Hinge. The function of the KIWI/KELP domain is not well understood in Arabidopsis. It has been proposed that the KIWI/KELP domain interacts w ith general transcriptio n factors for activation of gene transcription (Cormack et al., 1998). Because AtCstF-64 is single copy gene in Arabidopsis and lack of the KIWI/KELP doma in produces few phenotypic effects compared with the likely null itb4-2 mutation, it is likely that the KIWI/KELP domain is nonessential for AtCstF-64 function. The relatively mild phenotype produced by the itb4-1 mutation greatly contrasts with the phenotype of the itb4-2 mutation, which led to profound changes in trichome, leaf, and flower development. These phenotypic defects suggest an important developmental role for AtCstF-64. The occurrence of twin trichomes in the itb4-2 mutant suggests that proper trichome cell fate specification requires AtCs tF-64 function. Normally, once an epidermal cell acquires the trichome fate, division of that cell ceases. The twin trichomes seen in itb4-2 mutants are reminiscent of the trichome clusters seen in siamese ( sim ) mutants, where incipient trichomes still divide due to a failure to properly ente r the endoreduplication cy cle (Walker et al., 2000). This phenotype suggests that in itb4-2 mRNAs encoding proteins involved in the control of endoreduplication may have altered in expression le vels, and hence be direct or indirect targets of AtCstF-64. Plants homozygous for itb4-2 also displayed a significant incr ease in the number of sepals and petals, compared to wild type, although the or gans were relatively normal in appearance. These results suggest that early developmental events are more se nsitive to loss of At CstF-64

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139 function than later differentiation events. This idea is supported by our finding that ITB4 is most highly expressed in actively pro liferating tissue and organ primo rdia. A similar developmental role for CstF-64 is seen during mammalian ce ll differentiation. High expr ession of CstF-64 is required for normal development of B-lymphocytes. Reduced expression of CstF-64 gave rise to aberrant differentiation and apoptotic cell d eath (Takagaki and Manley, 1998) Our results support the finding in mammals that the slowly gr owing or inactively dividing cells may be able to tolerate lower levels of CstF than the rapidly growing or actively dividing cells (Takagaki et al., 1996). In contrast to the CstF-64 muta tions in other organisms, the itb4-2 homozygous plants are viable. Depletion of CstF-64 in chicken and mo use B lymphocytes caused apoptotic cell death (Takagaki and Manley, 1998). RNA15, the homolog of CstF-64 in yeast, is also essential for cell viability (Minvielle-Sebastia et al., 1991). These differences between plant and animal CstF-64 mutants suggest that plants are more tolerant of loss of CstF-64 function than other eukaryotes. The recent identification of ESP1 (At1g73840) in Arabidopsis s uggests that there are at least two complexes that contain CstF-64 homologs in Arabidopsis (Herr et al., 2006). The enhanced silencing phenotype ( esp ) mutants affect gene silencing and are involved in RNA metabolism. ESP1 encodes an AtCstF-64 like protein that lacks the RNA-binding RRM domain. It has been postulated that the standard CstF complex contains AtCstF -64 that uses the RRM domain to bind pre-mRNAs. The other putative complex contains ESP1 and uses a separate RNA-binding protein to recognize pre-mRNAs (Herr et al., 2006). The two complexes function redundantly in mRNA 3 end formation, wh ich is likely to be a reason why the itb4-2 mutation is not lethal.

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140 Loss of AtCstF-64 Function Influences the Ex pression of Multiple Genes that C ontrol Floral Organ Development In mammalian cells, altered levels of CstF-64 expression has been shown to influence the expression of at least 51 genes and induce alternative poly(A) s ite selection (Shell et al., 2005). This is also likely to also occur in plant cells. Our results showed that loss-of-function mutations in ITB4 caused changes in trichome fate, shape, and fl oral structure. It is likely that these phenotypic defects are due to cha nges in the expression of the ge nes involved in control of trichome morphogenesis and floral development. There are at least 30 genes known to regulate trichome morphogenesis (Schellmann and Hulskamp, 2005). For example, STI, A N, ZWI, FRC14 and GL3 function as positive regulators that promote trichome branching (Hulskamp et al., 1994; Oppenheimer et al., 1997; Luo and Oppenhe imer, 1999); loss of function mutations in these genes cause a decrease in the trichome branch number. Conversely, TFCA, RFI KAK PYM and SUZ4 function as a negative regulators that suppress trichome branching; mutations in these genes result in an increase in trichom e branch number (Krishnakumar and Oppenheimer, 1999; Perazza et al., 1999; Kiri k et al., 2002). Mutations in ITB2 and ITB3 affect trichome branch length (Zhang et al., 2005b), and mutations in TRY cause trichome clusters (Hulskamp et al., 1994). It is likely that the pre-mRNAs from the above-mentioned genes may be substrates of ITB4. Without ITB4 function, may not be corr ectly polyadenylated, and therefore, their expression levels are likely to be altered. Similar events may oc cur in the genes that regulate floral development. Regulators of floral organ identity genes may have altered expression leading to observed changes in floral architecture. Differences Exist in the Mechanism of mRNA 3 End Formation amon g Plants, Yeast and Mammals Formation of mRNA 3 ends is carried out by multiple trans -factors including CstF, CPSF, cleavage factors (CF) and the poly(A) polymeras e (PAP). Homologs of subunits for each of

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141 these factors have been identified in yeast, pl ants and mammals (Zhao et al., 1999; Yao et al., 2002; Elliott et al., 2003; Herr et al., 2006). It has been wide ly accepted that mRNA 3 end formation is similar in all eukar yotic cells, based on the high level of protein sequence identity of these factors, and the cooperative RNA-protein an d protein-protein inte ractions within the processing machinery (Yao et al., 2002; Elliott et al., 2003; Delaney et al., 2006; Herr et al., 2006; Xu et al., 2006). The results of our in vivo functional assays indicated that the homologs of CstF-64 in yeast, plants and mammals are not functionally equi valent. It is possible th at this is due to the differences in the sequences of the poly(A) sign als in pre-mRNAs in yeasts, plants and mammals (Mogen et al., 1990; Mogen et al., 1992; Li and Hunt, 1997; Zhao et al., 1999; Loke et al., 2005; Herr et al., 2006; Ji et al., 2007) The common minimal poly(A) signa l is composed of an A-rich sequence, a U-rich element, and a PyA cleavag e site in all eukaryot es. Nonetheless, the requirement for specific sequence elements differs greatly between yeast, plants, and animals (Zhao et al., 1999). In mammals, a single copy of the AAUAAA elem ent is highly conserved and absolutely necessary for precise 3 end formation. The AAUAAA element is located about 15 nucleotides upstream of the poly(A) site; and the DSE is located w ithin 50 nucleotides (Zhao et al., 1999). CPSF-160 and CstF-64 specifically recognize and bind to the AAUAAA element and DSE in pre-mRNAs, respectively (Murthy and Manley, 1992; Takagaki and Manley, 1997), and cleavage occurs preferentially at CA (Zhao et al., 1999). However, the plant poly(A) signal lacks a consensus element as in mammals. It ha s a wide distribution of multiple AAUAAA-like sequences, and the GU-rich elements are located up stream of the poly(A) site, not downstream as in mammals, and cleavage occurs preferentially at PyA(Wu et al., 1995; Li and Hunt, 1997;

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142 Zhao et al., 1999). Previous work has shown that plant cells do not properly recognize animal polyadenylation signals (Hunt, 1987). It is therefore likely, that human CstF-64 does not recognize plant polyadenylation signals. This may explain why the human CstF-64 was not able to functionally complement the itb4-2 mutant in Arabidopsis. The yeast poly(A) signal seems to be more similar to that in pl ants. In yeast, usually redundant A-rich and U-rich elements are located upstream of the cleavage site, and no una mbiguous DSEs have been identified. As in plants, the preferential cleavage site is PyA. Ho wever, yeast and plants do not share a consensus element in their poly(A) signal sequences (Zhao et al., 1999). This may explain our result that ITB4 could not complement the yeast CstF-64 mutant. In summary, we have shown that ITB4 encodes AtCstF-64, which is highly expressed in growing and proliferating cells, and is require d for normal trichome morphogenesis and floral development. Loss of ITB4 results in aberrant fl oral architecture due, in part, to the altered expression of floral organ identity genes. The fi nding that ITB4 is not functionally equivalent to its yeast counterpart, and that the mammalian CstF-64 could not functionally complement the itb4-2 mutant, supports the idea that there exists key differences between polyadenylation in yeast, plants, and animals even though the basic mechanism is conserved in all eukaryotic cells (Zhao et al., 1999; Yao et al., 2002 ; Elliott et al., 2003; Delaney et al., 2006; Herr et al., 2006; Ji et al., 2007). Given the importance of 3 end pr ocessing for a host of cellular processes including gene regulation and cell prolifer ation (Danckwardt et al., 2008) understanding these differences is needed to unravel the role of CstF in development. Future Perspectives Polyadenylation is a co mmon event that occurs in all eukaryotic nuc lei, but alternative polyadenylation events that affect plant development have rarely been reported. Our results show that absence of the plant ho molog of CstF-64 encoded by ITB4 causes pleiotropic effects in plant

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143 development such as severe defect s in trichomes and floral organs. ITB4 is highly expressed in these rapidly expanding and proliferating cells. ITB4 protein was localized to nuclei. The itb4 mutants had altered gene expressi on of important floral developm ental regulators. These results provide a link between Cst-64 and plant development. However, the function of ITB4 is still far away from being fully understood. Multiple questi ons need to be answered. The most important is which genes are affected by either the aberrant alternative polyadenylat ion or the failure of poly(A) addition in the itb4 mutants. To answer these questio ns, the following approach can be used. Through microarray experi ments using mRNA isolated from itb4-2 mutants and wild type, genes whose expression is altered in the mutant can be identified. The poly(A) site of these genes can be examined by using 3 rapid amp lification of cDNA ends (3 RACE). Although polyadenylation has been considered as a conserved mechanism in all eukaryotic cells, and the 3 end formation machinery is found in mammalian, yeas t and plant cells, our results suggest that there may be important differences in 3 end processing between these groups. We were unable to rescue a yeast Cs tF-64 mutant using th e Arabidopsis coding sequence, and the mammalian CstF-64 could not functionally complement itb4 mutants. However, alternative explanations exist. First, the mammalian CstF-64 gene may not be properly expressed in plants. To check this, we could us e western blotting of pr oteins extracted from transformants using an antibody to the mammalian CstF-64 protein. Likewise, we have to rule out lack of expression before we conclude that ITB4 cannot rescue the yeast mutant.

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144 Table 5-1. Primers used in this study Primer name Sequence (5 3) Used for itb4m F TGGCAAAAGAATAAACGAGGG itb4m R ATTCAGGGCATTCTAAGCGA itb4-1 mutation identification ITB4g F CTCCTATCGACGACGAATACGAAAG ITB4g R AGGGGCCACAGGATTAAAACCA ITB4 expression construct ITB4cDNA F CTCCTATCGACGACGAATACGAAAG ITB4cDNA R CTATGAAGGCTGCATCATGTGGTCTTGC ITB4 cDNA amplification ITB4GFP F ATGGCTTCAT CATCATCCCA ACGACGC ITB4GFP R TGAAGGCTGCATCATGTGGTCCTTGCTTG ITB4-GFP expression construct CstF64 F ATGGCGGG TTTGACTGTGAGAGACCC CstF64 R TACAGGTGCTCCAGTGGATTTCTGTATTTGTTCC CstF64 expression construct CstF64 F ATGTCGAGTTTGGCGGTGAGAGACCC CstF64 R GGAGGAGGGAAACCCT AATCCAAGTGTGGG CstF64 expression construct ITB4i1 F ATGGCTTCATCATCATCCCAAC ITB4i1 R GTGCCTTTGTCATTCTCAGCAA ITB4 in situ hybridization ITB4ic F ATGGCTTCATCATCATCCCAAC ITB4ic R GTGCCTTTGTCATTCTCAGCAA ITB4 in situ hybridization ITB4i2 F CGCCAAATATTGTTCAGGCCC ITB4i2 R TTGGTAATGCTTGGTGGGG ITB4 in situ hybridization ITB4i3 F AAGCAGA TTGGAGGGCCAGTAGATT ITB4i3 R TTTGCGTAAACTGCGAACCGA ITB4 in situ hybridization AP1i F GGGAAGGGGTAGGGTTCAATTGAAGA AP1i R GACAACAAGAGCAACTTCAGCATCAC AP1 in situ hybridization AP1ic F GGGAAGGGGTAGGGTTCAATTGAAGA AP1ic R GACAACAAGAGCAACTTCAGCATCAC AP1 in situ hybridization AP3i F CGAGAGGGAAGATCCAGATCAAGA AP3i R GCTAGAGAACATGATAATCGAAACCC AP3 in situ hybridization PIi F GGAGGAATGGA TTGGTGAAGAAGGCT PIi R GCCAGATAACTTCTGGTATTGGTCCA PI in situ hybridization Primers used for in situ hybridizatio n included the T7 promoter sequence, 5taatacgactcactataggg3 at the 5 end, for example: ITB4i1 R, 5taatacgactcactatagggGTGCCTTTGTCATTCTCAGCAA3 for the antisense RNA probe, and ITB4ic F, 5taatacgactcactatagggATGGCTTCA TCATCATCCCAAC3 for the control sense RNA probe.

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145 Table 5-2. Alteration of trichome cell shape in the itb4-2 mutant Trichome branches % a Strain 0 2 3 4 5 Total number b %Twin cluster c Col wt 0 0.1 96.0 3.9 0 1120 0 itb4-2 3.4 18.0 70.3 7.8 0.5 1540 2.52 a Numbers represent percentages of the total number of trichomes with the indicated number of branches. b Total number of trichomes counted on at least ten leaves. c Numbers represent percentages of the total number of trichomes that were present as twins.

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146 Figure 5-1. Positional cloning of ITB4 (A) Positional cloning strategy to identify ITB4 gene. (B) Confirmation of the C to T transition in the itb4-1 allele by digestion of PCR products with MseI. The C to T transition creates an MseI site in the itb4-1 allele. PCR products amplified using ITB4 specific primers (see Table 1) were digested with MseI and subjected to electropho resis through an agarose gel. (C) Identification of the itb4-2 mutant homozygous for the T-DNA inse rtion. Genomic DNA from wildtype, heterozygous or homozygous itb4-2 mutants was amplified using ITB4 and T-DNA specific primers (see Table 1), and the products were subjected to electrophoresis through an agarose gel. (D) Results of RT-PCR using ITB4 specific primers showing similar levels of ITB4 transcript in itb4-1 and wild type plants, and no detectable transcript in itb4-2 mutants.

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147 Figure 5-2 Rescue of the itb4-1 zwi-3 double mutant phenotype by At1g71800. (A) Light micrograph showing unbranched trichomes on the itb4-1 zwi-3 double mutant. (B) Light micrograph showing the tw o-branched trichomes on the zwi-3 single mutant. (C) Light micrograph of an itb4-1 zwi-3 double mutant transformed with the GL2:ITB4 construct. The transgenic double mutant shows the same phenotype as the zwi-3 single mutant, demonstra ting rescue of the unbran ched trichome phenotype. (D) Light micrograph of an itb4-1 zwi-3 double mutant transformed with the 35S:ITB4 showing two-branched trichomes. (E) Ma gnified image from panel D showing a three-branched trichom e. Scale bar = 200 m.

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148 Figure 5-3. The itb4-2 mutants display the trichome shape defects. (A) (D ) Scanning electron micrographs. (E) (H) Light micrographs of isolated trichomes. (A) Symmetrical Col wild type trichomes with equal bran ch length. (B) Irregular trichomes on an itb4-2 mutant showing unequal branch length and multiple branch points. (C) Developing twin trichome on an itb4-2 mutant showing a trichome cluster. (D) Developing unbranched trichome on an itb4-2 .mutant. (E) Twin trichome still attached following treatment with EGTA an d pectinase. (F) Twin unbranched trichome on an itb4-2 mutant. (G) (J) Irregular trichomes on itb4-2 mutants showing di fferent numbers of branches and separated branch positions. (K) (N) Twin trichomes with different numbers of branches.Scale bar = 100 m in (A) (F), and 50 m in (G) (N).

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149 Figure 5-4. Floral defects of itb4-2 mutants. (A), (D), (G), a nd (J) SEMs of Col wild type flowers. (B), (C), (E), (F), (H), (I ), (K), and (L) SEMs of flowers from itb4-2 mutants. (A) Wildtype developing flower buds s howing four sepal primordia. (D) Older wildtype buds showing the se pal completely enclosing the floral organs. (G) Wildtype stigma with normal papillae. (J) Wildtype anther containing normal pollen. (B) Developing itb4-2 flower buds showing the developi ng petal primordia (large arrow). Small Arrow indicates fused sepal primordia. (C) itb4-2 flower bud with extra sepals. (E) Developing itb4-2 flower buds showing the exposed stigmas. (F) Preferentially growing sepal on an itb4-2 flower bud (H) and (I) Abnormal itb4-2 stigma showing malformed papillae. (K) Abnormal itb4-2 anther with abortive pollen. (L) Abnormal itb4-2 anther lacking polle n. Scale bar = 100 m.

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150 Figure 5-5. Increased number of floral organs itb4-2 mutants. (A) Floral organ numbers for Col wild type and itb4-2 mutant flowers. (B) Representati ve flowers from wild type, left, and an itb4-2 mutant, right. Scale bar = 1 mm.

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151 Figure 5-6. Leaf shape and color defects in itb4-2 mutants. (A), (C), and (E) Col wild type plants. (B), (D), and (F) itb4-2 mutant plants. (A) Wild type plant showing normal first leaf pair. (B) itb4-2 mutant showing yellow first leaf pair. (C) Wild type seedling showing smooth leaf edge. (D) itb4-2 mutant seedling showing serrated leaf edge and less yellow leaf color. (E) Wild type mature plant. (F) itb4-2 mutant plant showing normal color leaves. Scale bar in (A) and (B ) = 0.1 mm; Scale bar in (C) and (D) = 5 mm; Scale bar in (E) and (F) = 10 mm;

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152 Figure 5-7. ITB2 expression pattern in Col wild type (A) Globular stage embryo showing strong expression. (B) Heart stage em bryo showing strong expression. (C) Germinating seed showing strong expressi on in the root cap and shoot apical meristem. (D) Germinating seed hybridized with the negative control sense RNA probe. (E) Longitudinal section of a six-w eek-old seedling showing strong expression in the developing leaves and trichomes. (F) Longitudinal section of the floral meristem showing strong expression in th e floral meristem and developing flower buds. (G) Floral organ primordia showi ng strong expression in the developing stamens and carpals and weak expression in the developed sepals. (H) Floral organ primordia hybridized with the negative c ontrol sense RNA probe. Scale bar = 10 m in (A) and (B), 50 m (B) a nd (C), 100 m in (E) (H).

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153 Figure 5-8. Altered expression patterns of floral organ identity genes in itb4-2 mutants. (A), (C) and (E) Longitudinal sections of Col wild t ype flower primordia. (B), (D), and (F) Longitudinal sections of flower primordia of the itb4-2 mutant. (A) and (B) AP1 antisense probe. (C) and (D) AP3 antisense probe. (E) and (F) PI antisense probe. Scale bar = 50 m.

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154 Figure 5-9. ITB4 is localized to the nucleus. Onion epidermal ce lls were transiently transformed with either 35S:ITB4-GFP (A) (C), or 35S:GFP. (D) (F). (A) GFP localization confined to the nucleus. (B) Same cell as in (A), but stained with DAPI to show position of the nucleus. (C) DIC image of the same cell as in (A) showing the position of the nucleus (arrow). (D) GFP localizati on in both the cytoplasm and the nucleus. (E) The same cell as in (D) stained with DAPI to show the position of the nucleus. (F) DIC image of the same cell as in (D) s howing the position of the nucleus (arrow). Scale bar = 100 m

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178 BIOGRAPHICAL SKETCH Xiaoguo Zhang was born in Hubei province, China. After completing his college education at Huazhong Agricultural University, he attended China Agricultural University for his m asters degree in crop genetics and breed ing. After his graduation, he work ed at Wuhan University. He started his Ph.D. program in plant molecular and cellular biology in 2005 at University of Florida.