brk1 and dcd1 act synergistically in subsidiary cell .../67531/metadc799473/m2/1/high_re… ·...

brk1 and dcd1 act synergistically in subsidiary cell formation in Zea mays

Divya Malhotra

Thesis Prepared for the Degree of

MASTER OF SCIENCE

UNIVERSITY OF NORTH TEXAS

August 2014

Approved By

Committee Members:

Amanda J. Wright (Major Professor)

Jyoti Shah (Committee Member)

Stevens Brumbley (Committee Member)

Malhotra, Divya. brk1 and dcd1 act synergistically in subsidiary cell formation in

Zea mays. Master of Science (Biochemistry and Molecular Biology), August 2014, 76

pp., 9 tables, 23 figures.

Subsidiary mother cell (SMC) divisions during stomatal complex formation in Zea

mays are asymmetric generating a small subsidiary cell (SC) and a larger epidermal

cell. Mutants with a high number of abnormally shaped subsidiary cells include the

brick1 (brk1) and discordia1 (dcd1) mutants. BRK1 is homologous to HSPC300, an

ARP2/3 complex activator, and is involved in actin nucleation while DCD1 is a

regulatory subunit of the PP2A phosphatase needed for microtubule generation (Frank

and Smith, 2002; Wright et al. 2009). Possible causes of the abnormal SCs in brk1

mutants include a failure of the SMC nucleus to polarize in advance of mitosis, no actin

patch, and transverse and/or no PPBs (Gallagher and Smith, 2000; Panteris et al 2006).

The abnormal subsidiary mother cell division in dcd1 is due to correctly localized, but

disorganized preprophase bands (PPBs; Wright et al. 2009). The observation that brk1

has defects in PPB formation and that the dcd1 phenotype is enhanced by the

application of actin inhibitors led us to examine the dcd1; brk1 double mutant (Gallagher

and Smith, 1999). We found that dcd1; brk1 double mutants demonstrate a higher

percentage of aberrant SCs than the single mutants combined suggesting that these

two mutations have a synergistic and additive effect on SC formation. Our observations

and results are intriguing and the future step will be to quantitate the abnormal PPBs

and phragmoplasts in the double and single mutants using immunolocalization of tubulin

and actin as well as observations of live cells expressing tubulin-YFP.

ii

Copyright 2014

By

Divya Malhotra

iii

ACKNOWLEDGEMENTS

First, I would like express my deepest thanks and gratitude to my major professor Dr.

Amanda J. Wright for giving me a platform to work in her lab and learn molecular

biology techniques. She is a great advisor. Her expertise, guidance, kindness, patience,

advice and her encouragement helped me achieve my goal. She has been a pillar of

support all through my failures and has been besides me as a true friend. I would like to

thank all my committee members for all their support and guidance. Dr. Lon Turnbull, I

thank you for all your time and training sessions on how to use the confocal microscope

as well as for all the extended help with the computer software. I thank all the past and

current members of the Wright lab for their support and help. Special thanks to my

husband, Vineet, and my lovely daughter, Janvi, for sticking by me through this entire

process and extending their support. My biggest thanks go to my parents, who live in

India, for all their love and encouragement. I couldn’t have progressed towards the

completion of my degree without their blessings.

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TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS...........................................................................iii

LIST OF TABLES........................................................................................v

LIST OF FIGURES.....................................................................................vi

Chapter

1. INTRODUCTION..............................................................................1

2. RELEVANCE OF PROJECT…………………………………………18

3. MATERIAL AND METHODS………………………..........................20

4. RESULTS…………………………………..........................................27

5. DISCUSSION...................................................................................61

6. REFERENCES……………...............................................................67

v

LIST OF TABLES

Page

1. Quantitative analysis of the number of abnormal subsidiary cells ……………….29

2. Percentage of subsidiary mother cells with polarized nuclei .…………………....34

3. Quantitative analysis of PPB orientation and organization in preprophase SMCs

adjacent to GMCs with a width < 6μm………………………………………….…...38

4. Quantitative analysis of PPB orientation and organization in preprophase SMCs

adjacent to GMCs with a width > 6μm………………………………………….…...42

5. Quantitative analysis of PPB orientation in dividing prophase SMCs…………...47

6. Quantitative analysis of PPB organization in dividing prophase SMCs………...49

7. Quantitative analysis of spindle orientation in dividing SMCs…………………...53

8. Quantitative analysis of phragmoplast orientation in dividing SMCs……….…..57

9. Summary ……………………………………………………………………………...62

vi

LIST OF FIGURES

Page

1. Cell cycle phases and cytoskeletal development………………….……………..2

2. Sequence of division in maize leaf epidermis…………………………………….6

3. Genotyping using dcd1 markers .......................................................................28

4. Graphical representation of abnormal subsidiary cells.....................................30

5. Cell outlines and stomatal complexes of wild type, dcd1, brk1, and brk1; dcd1

…………………………………………………………………………………………......31

6. Graphical representation of the overall mean number of SMCs with polarized

nuclei in the wild type, dcd1, brk1 and and brk1; dcd1………………………….35

7. Polarized and unpolarized representatives in wild type and mutants…………36

8. Graphical representation of PPB orientation in preprophase SMCs adjacent to

GMCs with width < 6 μm……………………………………………………………39

9. Graphical representation of PPB organization in preprophase SMCs adjacent to

GMCs with width < 6 μm……………………………………………………………41

10. Graphical representation of PPB orientation in preprophase SMCs adjacent to

GMCs with width > 6 μm……………………………………………………………43

11. Graphical representation of PPB organization in preprophase SMCs adjacent to

GMCs with width > 6 μm……………………………………………………………44

12. Preprophase PPB organization and orientation representatives in wild type and

mutants……………………………………………………………………………..45,46

13. Graphical representation of PBB orientation in dividing prophase SMCs……..48

vii

14. Graphical representation of PBB organization in dividing prophase SMCs…..50

15. Prophase PPB organization and orientation representatives in wild type and

mutants……………………………………………………………………………...51,52

16. Graphical representation of spindle orientation in dividing SMCs……………54

17. Spindle orientation in representatives in wild type and mutants ……............55,56

18. Graphical representation of phragmoplast orientation in dividing SMCs…...58

19. Phragmoplast orientation representatives in wild type and mutants...............59,60

1

CHAPTER 1

INTRODUCTION

1.1 Cell division and plant-based cytoskeleton structures: preprophase band and

phragmoplast

Cell division is an important process by which multicellular organisms multiply in

order to grow and expand. Cellular divisions are categorized as either symmetric in

which the mother cell divides equally into two daughter cells and asymmetric in which

the mother cell divides unequally into small and large daughter cells. Asymmetric cell

division introduces cellular diversity in an organism. The daughter cells generated after

an asymmetric division often adopt different fates (Gallagher and Smith, 1996; Scheres

et al 1999), and asymmetric divisions are associated with developmental events,

formation of new cell lineages and specialized cell types capable of carrying out

different functions. Earlier studies conclude that in animals and yeast, polarization of the

mother cell and specific orientation of an asymmetric division plane determines the

daughter cell fates (Rhyu and Knoblich, 1995; Tazikawa et al., 1997). During plant

development, cell divisions can be physically as well as developmentally asymmetric in

order to form daughter cells of different shapes, sizes, and functions (Gallagher and

Smith, 1997). To achieve synchronous working of all the cells within a tissue, the cells

have to be located correctly relative to one another and attain a specific shape and size.

Cell location, size and shape are defined by the inelastic cell wall that is laid down

during cytokinesis and fixes cells in position.

In plants, as in animals, microtubule and actin based structures play a vital role

2

during cell division. In an animal cell division, microtubule composed spindle governs

the orientation of division plane and the separation of the replicated chromosomes. Cell

division terminates with an invagination of the plasma membrane during cytokinesis,

which always occurs in a perpendicular to the plane of spindle axis and is mediated by

the actin cytoskeleton (Rappaport, 1986; Salmon, 1989).

In plant cells, cytokinesis is mediated by an actin and microtubule-based

structure called a phragmoplast. It guides Golgi-derived vesicles enclosing new cell wall

materials to a region between the daughter nuclei where they fuse to form a new cell

wall. The location of the phragmoplast and new cell wall is predetermined during

prophase by the position of a temporary cortical array of microtubules (composed of

tubulin) and microfilaments (composed of actin) called the preprophase band (PPB)

(Pickett-Heaps and Northcote, 1966; Palevitz, 1987; Traas et al., 1987; Wick, 1991).

The PPB defines the cortical division site (CDS) and determines in division plane

orientation in many plant cell types (Mineyuki 1999).

Figure 1: Illustrates cell cycle phases showing the variation in nuclear material and the

emergence of plant-specific cytoskeletal structures: the actin patch, preprophase band, and

phragmoplast.

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The assembly of the microfilament portion of the PPB is dependent on the microtubule

component (Palevitz 1987, McCundy and Gunning 1990). Formation of the PPB

depends on de novo microtubule nucleation at the cell cortex in combination with

microtubule stabilization in the PPB zone and microtubule destabilization outside of the

PPB zone (Cleary et al. 1992, Dhonukshe and Gadella 2003, Vos et al. 2004). Membes

of different microtubule-binding protein families are necessary for PPB formation and

localization. These proteins include MOR1, a homologue of XMAP125 which is part of a

highly conserved class of microtubule binding proteins that act to stabilize microtubules,

Arabidopsis CLASP, a microtubule binding protein that regulates microtubule dynamics,

and MAP65 which bundles microtubules by cross bridging individual microtubules.

(Whittington et al. 2001, Smertenko et al. 2004, Hussey et al. 2002, Mimori-Kiyosue et

al. 2005, Ambrose et al. 2007, Kirik et al. 2007). Other proteins needed for PPB

formation have homology to proteins that localize to animal centrosomes which function

as the main microtubule-nucleating center in animal cells. These include

FASS/TONNEAU2 (FASS/TON2) and TONNEAU1A and TONNEAU1B (TON1). All are

necessary for PPB formation. This interesting connection suggests that microtubule

nucleation may function via similar mechanisms in plants and animals despite the lack

of centrosomes in plant cells.

In addition to defining the CDS and determining the division plane, PPB

additionally contributes to a well-timed establishment of a normal bipolar spindle. It has

been observed that cells grown in culture that have no PPB or possess double PPB,

4

initially develop multipolar spindles which eventually ends up forming a bipolar spindle.

These cells also take longer to progress through metaphase (Chan et al., 2005; Marcus

et al., 2005). Timely progression of cell cycle and proper spindle assembly is

dependent on the microtubules connecting the nucleus and PPB (Ambrose and Cyr,

2008). During prophase, after nuclear envelope breakdown and after the initial

organization of the mitotic spindle, PPB disassembles (Dixit and Cyr, 2002). The role

of the mitotic spindle is to physically separate the duplicated chromosomes.

As previously discussed, the phragmoplast directs vesicles released by Golgi

bodies towards the generation and expansion of the new cell wall between the daughter

cells produced after cell division. The phragmoplast organizes the deposition of the new

cell membrane and immature cell wall. It has been suggested that the phragmoplast

arises from the remnants of the spindle (Jurgens 2005). It is in the shape of a donut and

expands centrifugally towards the area of the mother cell cortex formerly occupied by

the PPB at CDS (Galatis et al. 1984, Granger and Cyr 2001). The new cell wall joins the

mother cell cortex at the CDS previous established by the PPB (Wright and Smith 2007,

Van Damme 2009, Muller et al. 2009.) Phragmoplast expansion deposits new cell wall

material partitioning the daughter cells at the PPB defined cortical division site. The

dividing cell wall joins the mother cell at the site of the CDS, which is coincident with the

prior site of the PPB. Though these basics of division plane determination in plants are

well established, relatively few proteins have been identified as playing a specific role in

division plane orientation preventing a complete understanding of this process.

Because the cell walls that surround plant cells prevent cell migration and spatial

5

rearrangements, correctly oriented division planes are necessary for the normal

development of plant tissues, organs, and structures. Thus for a complete

understanding of many aspects of plant development, a complete understanding of

division plane orientation is necessary.

6

1.2 Understanding division plane orientation by studying subsidiary mother cell

divisions during stomatal cell complex formation in Zea mays

A schematic of the events that occur during the formation of stomatal complex in

the maize leaf is shown in Figure 2.

Figure 2: Sequence of division events in maize leaf epidermis. A) SMC polarization towards GMC

and formation of actin patch (yellow). B) Actin patch assists nuclear migration and spindle tethering

in the SMC. Microtubule composed PPB (green) localization occurs after nuclear migration, and

then the mitotic spindle forms as the PPB breaks down. C) The microtubule-based phragmoplast

mediates cytokinesis and assists in cell plate formation. It mediates the fusion of the new cell wall

with the site on the mother cell cortex previously occupied by the PPB.

D) SC are formed and longitudinal division of GMC occurs producing stomatal complex which consists of

two guard cells flanked by two SC

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The maize leaf is an excellent system for studying asymmetric cell divisions during

stomatal development. Linear rows of cells make up the maize leaf epidermis.

Transverse and longitudinal symmetric cell divisions contribute to maize leaf expansion

and growth (Sharman, 1942; Sylvester et al., 1990). Daughter cells produced as a

result of asymmetric division adopt specialized features such as gas exchange by

stomatal complexes and protection by silica and cork cell pairs.

The formation of a stomatal complex starts with a symmetric transverse division

of a cell referred as a guard cell progenitor (GCP), also called a stomatal precursor. It

further undergoes an asymmetric, transverse division to form an apical guard mother

cell (GMC) and basal interstomatal cell. Stomata lie between interstomatal cells forming

a repeating pattern. The final stomatal complexes are composed of four cells (a pair of

guard cells surrounded by a pair of subsidiary cells). The first asymmetric division

produces a guard mother cell (GMC), which further divides symmetrically and

longitudinally to form two guard cells. Before the GMC division takes place, cells

adjacent to the GMCs called subsidiary mother cells (SMC) undergo asymmetric

divisions. The SMC division starts with the polarization and migration of the nucleus

towards the GMC. Polarization of SMC nucleus is considered to be a response to an

extrinsic signal from the adjacent GMC (Stebbins and Shah, 1960). There are intrinsic

cues that also contribute to this event. A cytological marker of SMC polarization is the

formation of an actin patch. This patch is found along the SMC wall flanking the GMC

(Galatis and Apostolakos, 2004). Recent study in this direction suggests there is also an

accumulation of endoplasmic reticulum (ER) at SMC–GMC contact sites (Giannoutsou

8

et. al., 2011). The asymmetrically positioned PPB is subsequently formed near the GMC

to mark the future plane of division. During cytokinesis, a phragmoplast is formed from

the remnants of spindle to produce the portioning cell wall between the two daughter

nuclei. The phragmoplast expands centrifugally by guiding Golgi derived vesicles to

attach the new cell wall at the former location of the PPB. This division yields a lens-

shaped subsidiary cell and a much larger sister pavement cell.

9

1.3 Background information of mutants that have abnormal shaped subsidiary

cells

Studies have been conducted to investigate the effects of mutations that disrupt

SMC division on the differentiation of subsidiary cells. Although the basic role of the

cytoskeletal structures has been established, it is vital to find out answers regarding the

molecular entities that are involved in the regulation, spatial positioning and functions of

these structures. Previous reports describe the characterization and cloning of the

recessive mutations dcd mutants (discordia1/2); pan mutants (pangloss1/2) and brk

mutants (brick1/2/3).

Maize dcd1 mutants have defects in the orientation of asymmetric cell divisions.

DCD1 is a regulatory subunit of the PP2A phosphatase needed for microtubule

generation (Frank and Smith, 2002; Wright et al. 2009). Similar to wild-type stomatal

complex formation, SMCs in dcd1 mutants polarize and nuclear migration occurs

towards the GMC and cortical actin patches. The PPBs are formed at normal sites,

though the PPBs are often frayed and disorganized. As the mitosis progresses, the

spindle remains associated with the actin patch similar to wild type cells. Remnants of

spindle give rise to the phragmoplast at the normal location. During cytokinesis, unlike

in wild-type SMCs where the phragmoplasts move towards the actin patch and curve

tightly around the inner daughter nucleus to form a small lens-shaped daughter cell,

dcd1 mutant cells lack apical movement of the phragmoplast towards the actin patch.

10

Thus the new cell wall is not in lens shaped, but rather is oblique. It generally attaches

properly at one end but not to the other end. However, sometimes the new wall

attaches inappropriately at both ends. The resulting division is nearly longitudinal

(Gallanger and Smith, 1999).

In Arabidopsis, FASS encodes a putative regulatory B’’ subunit of the PP2A

phosphatase which is homologous to dcd1. fass mutants completely lack PPBs (Traas

et al., 1995; McClinton and Sung, 1997; Camilleri et al., 2002). The maize homologues

of FASS are DCD1 and a paralogue, ADD1. DCD1 and ADD1 share 96% similarity to

each other and are 85% identical to FASS. Loss of dcd1 disrupts asymmetric cell

divisions as described earlier whereas loss of add1 function does not cause a

phenotype in the maize leaf epidermis (Gallagher and Smith 1999, Wright et al. 2009).

Simultaneous loss of dcd1 and add1 function in maize results in embryonic lethality due

to the inability to from PPBs and the disorganized cell divisions that follow (Wright et al.

2009).

DCD1/ADD1/FASS encodes proteins homologous to regulatory subunits of

heterotrimeric PP2A phosphatase complex. The complex consists of three subunits, a

catalytic “C” subunit, a scaffolding “A” subunit, and a regulatory “B” subunit (Janssens

and Goris 2001). There are four B subunit gene families, B, B’, B’’, and B’’’. They all are

unrelated to each other. The C terminus of DCD1/ADD1/FASS is homologous to the C

terminus of regulatory subunit subgroup B’’ and the human protein PR72 (Hendrix et al.,

1993; Camilleri et al., 2002). The N-terminal domains of DCD1/ADD1/FASS are

11

interrelated to each other and other vertebrate proteins but are different to the N-region

of PR72. Yeast two-hybrid experiments suggested that these plant proteins likely

function as phosphatase subunits (Camilleri et al., 2002). A likely hypothesis is that

these proteins are required for PPB formation and they target the PP2A phosphatase

complex to a protein(s) for dephosphorylation so as to initiate PPB formation.

DCD1/ADD1 co-localizes with the PPB and remains at the CDS throughout

metaphase signifying a probable role in both CDS organization and in PPB formation

(Wright et al., 2009). The C. elegans homologue of DCD1/ADD1/FASS, RSA-1 co-

precipitates with PP2A catalytic and scaffolding subunits (Schlaitz et al., 2007). The

latter is found to be localized to centrosomes and is needed for microtubule outgrowth

establishing a parallel between proteins needed for PPB formation in plant cell and

centrosome organization in animal cell. Absence of DCD1/ADD1 during the cytokinesis

at the CDS, suggests that they do not play a role in phragmoplast regulation.

Overall proteins needed for PPB formation have homology to proteins that

localize to animal centrosomes which function as the main microtubule-nucleating

center in animal cells. This interesting connection suggests that microtubule nucleation

may function via similar mechanisms in plants and animals despite the lack of

centrosomes in plant cells.

SMC division initiates with the polarization and migration of the SMC nucleus

towards the GMC. The mechanism of this migration is very poorly understood. The

mutant phenotype of pangloss1 (PAN1), a Leu- rich repeat receptor- like protein (LRR-

12

RLK) suggests that pan1 is needed for the polarization of subsidiary mother cell during

stomatal development in maize (Zea mays). PAN1 has an inactive kinase domain but is

required for the accumulation of an unidentified membrane-associated phosphoprotein,

signifying a function for PAN1 in signal transduction. It is proposed to act as a receptor

for GMC-derived polarizing cues (Cartwright et al., 2009). PAN1 localizes

asymmetrically in SMCs after the GMC formation. It is found at the site of GMC contact

to the neighboring SMC prior to nuclear polarization.

Another player in polarized cell growth is Type 1 ROPs (rho of plants) GTPases.

The significance of ROP activity in polarized cell growth and associations with RLKs

were previously established. The ROPs are involved in regulation of F-actin dynamics

and stimulate localized accumulation and fusion of vesicles needed for growth (Yang

and Fu, 2007; Yang, 2008; Fu, 2010). A ROP protein was previously isolated as a part

of a complex containing the Arabidopsis LRR-RLK CLAVATA1 involved in shoot apical

meristem maintenance (Trotochaud et al., 1999). Previous studies on ROP in various

model organisms such as Medicago imply that these interact with diverse members of

LRR-RLK family and are involved in signaling pathways (Molendijk et al.,

2008;Dorjgotov et al., 2009) In maize SMCs, ROPs interact with PAN1 to promote the

localized accumulation of F-actin at the GMC interface. This may be accomplished by

activating ROP maize homologs of proteins that promote F-actin assembly, such as

RIC4 (a protein mediating ROP stimulation of F-actin assembly in Arabidopsis), the

SCAR/WAVE complex (a regulator of the actin- nucleating ARP2/3 complex), or formins

(actin nucleators) (Yang, 2008; Campellone and Welch, 2010). The purpose of the

13

localized F-actin accumulation in maize SMCs is not well understood. Recent work

reports that ROP and PAN1 mutants do not show SMC polarizing defects towards GMC

suggesting a significant role of SMC F-actin patches that may be involved in polarized

vesicle trafficking (Humphries et. al., 2011). F-actin is found to be an essential

component in cellular polarity of many eukaryotic cell types (St Johnston and Ahringer,

2010). As nuclear polarization in SMCs is an actin- dependent process (Kennard and

Cleary, 1997; Panteris et al., 2006), ROPs may be involved in nuclear polarization via

regulation of F-actin dynamics or mechanisms that exclude actin involvement. In C.

elegans embryos and Drosophila melanogaster neuroblasts, the Rho family GTPase

Cdc42 stimulates the premitotic polarization of asymmetrically dividing cells via direct

interface with the polarity protein PAR6 (Siller and Doe, 2009; Nance and Zallen, 2011).

Although in plants no such homolog of PAR6 has been identified, this stresses the

possibility that ROPs may act via actin- independent as well as actin-dependent

mechanisms to promote premitotic SMC polarity in stomatal complex formation. Genetic

studies, protein localization assays, and biochemical observations of maize ROPs

suggests that ROPs play a vital role downstream of PAN1 in order to stimulate the

premitotic polarization of SMCs (Humphries et al., 2011). Co-immunoprecipitation

results indicate physical interaction between PAN1 and ROPs. Mild SMC polarization

defects have been observed due to partial loss of Type I ROP. Studies suggest that

Type I ROPs localize at the SMC and GMC contact site. The PAN1 patch is formed

during early prophase whereas ROP patches are formed at same site but later in the

cell division suggesting that PAN1 is involved in the recruitment of ROPs (Humphries

et.al., 2011)

14

Recent work in elucidating factors involved in polarization yielded a second LRR-

RLK promoting SMC polarization, PANGLOSS2 (PAN2). The two kinases PAN1 and

PAN2 are thought to function as receptors for the GMC-derived polarizing signal

needed for orienting the SMC asymmetric divisions (Stebbins and Shah, 1960). PAN1

and PAN2 have a homologous catalytically inactive kinase domain. Results suggest that

PAN2 is not a PAN1 co-receptor as it lacks few characteristics.PAN2 occurs

simultaneously rather than the one after the other in a sequence and they both interact

physically. Because, PAN2 localizes in the SMC prior to PAN1, it was concluded that

PAN2 lies upstream of PAN1 in the signaling cascade involved in perceiving GMC

signals to polarize. Thus, PAN2 component is first to act in the SMC-polarizing

mechanism identified to date (Zhang et al., 2012) suggesting that PAN2 is involved in

perceiving and amplifying the GMC-derived polarizing signals.

In addition to polarization defects in the development of the stomatal complex in

maize, the brick mutants also have defects in cell morphogenesis. As a plant cell's

shape is defined by the cell wall around them, cell morphogenesis is achieved by

guiding wall materials to expanding cells. Microtubule composed bands and F-actin

patches are involved in the guiding the new material to new cell walls. Drug analysis

has shown that F-actin plays a vital role in tip growth and in transporting secretory

vesicles containing cell wall material at the growth site (Geitmann and Emons, 2000;

Hepler et al., 2001). F-actin promotes the expansion of diffusely growing cells

15

established by genetic and pharmacological studies (Smith, 2003). Maize epidermal leaf

cells have a lobular structure and the establishment of this characteristic cell outline is

reliant on both microtubule and F-actin dependent processes. Lobes are formed as a

result of polarized outgrowths at numerous sites along the margins of cells which is

achieved by diffuse growth to increase the size overall. Isolation of a recessive

mutation, brk1 in maize suggested that it is involved in epidermal cell lobes in the maize

leaf as mutants lacked lobes completely. brk1 mutants expand similar to wild type but

fail to define the polar growth site which gives rise to lobes. While there was little

difference in microtubule organization, and absence of cortical F-actin at the tips of

emerging lobes was observed in these mutants when compared to the wild type.

Three brk mutants have been identified and are found to promote polarized cell

growth to produce lobes on the margins of leaf epidermal pavement cell thus

contributing towards the morphogenesis in maize leaf epidermis (Frank et al., 2002).

BRK1 is homologous to mammalian HSPC300, an ARP2/3 activator, and is involved in

actin nucleation (Djakovic et al., 2006). The brk1 gene encodes a small protein (8kD)

that is highly conserved in plants and animals. In addition to morphological defects in

epidermal pavement cell morphogenesis, brk1 mutants were reported to have 20-40%

abnormal stomatal subsidiary cell divisions (Gallagher and Smith, 2000). During wild-

type stomatal complex developmental, the SMCs polarization towards the GMC is

evident by the formation of the actin patch and nuclear migration. Loss of BRK1 causes

defects in the polarization of SMCs likely because BRK1 stimulates ARP2/3 complex-

dependent actin polymerization in the SMC (Gallagher and Smith, 2000). Concluding

events after polarization are affected and result into abnormally shaped subsidiary cells

16

due to division plane orientation defects along with the characteristic brick-shaped leaf

epidermal cells.

To summarize, dcd1 mutants exhibit no SMCs polarization defects, but the

division plane is often misoriented producing abnormally shaped SCs. A likely

explanation for the dcd1 SCs abnormalities is that the disorganized PPBs fail to

establish a robust cortical division site leading to misguided phragmoplasts during

cytokinesis. A percentage of SMCs in brk1 and pan1 mutants display either partial or no

polarization of the nucleus, the subsequent cell division is not asymmetric, and therefore

the SCs are abnormally shaped and fail to differentiate into subsidiary cells. Overall,

polarization defects in SMCs during preprophase of brk1 and pan1 mutants affect the

phragmoplast orientation during cytokinesis resulting in the development of transverse

or oblique cell walls.

In order to analyze and study the effects of dcd1 and brk1 mutations, microscopy

was used to visualize the abnormalities in the cytoskeletal structures involved in

stomata complex formation. The research is significant as it is focused on learning more

about the regulation of cell division orientation by evaluating asymmetric cells divisions

critical for stomata formation on maize leaves.

17

The observation that brk1 has defects in PPB formation and that the dcd1

phenotype is enhanced by the application of actin inhibitors led me to examine the brk1;

dcd1 double mutant (Gallagher and Smith, 1999). Possible causes of the abnormal SCs

in brk1 mutants include a failure of the SMC nucleus to polarize in advance of mitosis,

no actin patch, and transverse or no PPBs (Gallagher and Smith, 2000; Panteris et al

2006). The abnormal subsidiary mother cell division in dcd1 is due to correctly localized,

but disorganized PPBs (Wright et al. 2009). The objective of this research is to observe

the microtubule-based structures necessary for cell division in the dcd1 and brk1 single

mutants as well as in wild-type plants and brk1; dcd1 double mutants.

18

CHAPTER 2

RELEVANCE OF THE PROJECT

A cell wall surrounds a plant cell to give it shape and the ability to withstand internal

turgor pressure. The inability of plant cells to alter their structure after the completion of

cytokinesis places an important emphasis on the correct orientation of the division plane

during mitosis for overall plant growth and the development of plant structures, tissues,

and organs. Research in this direction is needed to elucidate the proteins and other

factors governing the two plant specific cytoskeletal structures (PPB and phragmoplast)

that guide division plane orientation. The following research is focused on determining

mechanisms that orient asymmetric cell divisions. The research uses stomata formation

in Zea mays as model system to investigate division plane orientation in asymmetric

divisions. Stomatal complexes in plants are necessary for gas exchange between the

plant and the atmosphere via the stomatal pore, which is surrounded by guard cells that

govern its opening and closing. In monocots, flanking each guard cell is a subsidiary

cell that acts to assist, reinforce, or protect the guard cells. Though plant cells are rigid

due to their cellulose cell walls, guard cells must expand and contract for gaseous

exchange so subsidiary cells afford a cushioning effect to protect the adjoining (more

rigid) cells from the guard cell expansions and contractions.

19

Research in this direction will provide us with more details about the

organization and orientation of the preprophase bands that are responsible for

determining the location of the cortical division site and the final division plane. It will

also provide additional insights into phragmoplast orientation during cytokinesis. The

results will enable us to answer how this process is deployed during stomata formation

and aid in the refinement of experimental techniques in maize, a crop plant

20

CHAPTER 3

Material and Methods

3.1 Identification of Mutants for Analysis:

Around 100 kernels from ears segregating the brk1-O and dcd1-O mutations

were planted in a flat with 2 inches of Metro-mix 900 soil in the EESAT greenhouse

facility at University of North Texas. 1 tbsp of Osmocote fertilizer was sprinkled evenly

across the soil and worked into the top layer. Kernels were spaced over on soil in 10

rows and 10 columns evenly. Each seed (kernel) was pressed into the soil

approximately 2 cms deep. Seeds were watered thoroughly until the soil was completely

wet and the flat was heavy to pick up. Flats were monitored for water content regularly

and watered as needed. Seeds germinated in three to four days. When seedlings had

4-5 leafs, the plants were numbered and part of the leaf blade from leaf 3 was

detached. The detached leaf blades were used to perform initial screening of altered

epidermal cell shape by making impressions of the leaf surface in Loctite cyanoacrylate

glue. A minimum of five stripes of glue were made on a microscopic slide and a thin

portion of the leaf blade was pressed against it. The slide was turned upside down on a

waxed sheet of paper and light pressure was applied. After the glue dried, the leaf was

gently removed from the slide, leaving an impression of the leaf surface. The slides

were then observed under a light microscope to detect the subsidiary cell shapes. For

each set of experiments, a hundred plants were preliminarily screened and selected

based on the glue impression phenotype as wild type, brk1, or dcd1. Eight to ten plants

from each category were transplanted into pots with metro-mix 900 and osmocote

fertilizer for further growth.

21

3.2 DNA Extraction

Genomic DNA was extracted from leaf tissue taken from transplanted plants belonging

to each category (wild type, brk1, and dcd1). The initial preparation step for DNA

extraction involved warming up 20% SDS by placing it in a water bath 650C to ensure it

was in solution and placing Isopropanol and 5M potassium Acetate in the freezer to chill

prior to use. Extraction buffer was made and autoclaved for future use. To make 200 ml

of extraction buffer following ingredients were mixed: 1M Tris (20 ml); 0.5M EDTA (20

ml); 4M NaCl (5 ml); distilled water (155 ml).

Leaf tissue from the wild type, dcd1, and brk1 were taken and placed in a labeled, clean

Eppendorf tube. Just before use, 0.7 µl of β-mercaptoethanol (BME) was added per 1

ml of extraction buffer. 500 µl extraction buffer including BME was added to each

Eppendorf with leaf tissue in it. The tissue was ground using pestles until well shredded

and the extraction buffer became bright green in color. 35 µl 20% SDS was added to

each tube and the tubes were inverted to mix. The DNA extractions were incubated at

650C in the water bath for 10 minutes. 130 µl of ice-cold 5M potassium acetate was

added and mixed well by tapping and inverting the tubes. To attain a viscous gel like

fluid the tubes were placed on ice for 5 minutes. Next the tubes with samples were

centrifuged for 10 min at 13,000 rpm. After centrifugation two layers were formed. The

top, clear layer contains DNA and was transferred to the clean Eppendorf tube. An

equal volume of ice-cold isopropanol and 1/10 volume of 3M sodium acetate were

added to the supernatant containing tube. The tubes were incubated for 60 minutes or

overnight in –20oC. To pellet the precipitated DNA, tubes were centrifuged at 13,000

rpm for 10 min. The supernatant was then discarded and the pellet was cleaned with 1

22

mL 70% ethanol by centrifuging for 5 min at 13,000 rpm. All ethanol was completely

removed and the pellet was left to dry at room temperature for 30 min. The DNA pellet

was re-suspended in 50-100µl of RNase solution (10µg/ml) and kept overnight at 4oC to

ensure that it went into solution. Finally, the DNA solution was incubated at 37oC for 30

min then stored at -20oC for future use.

23

3.3 Setting up PCR: genotyping using dcd1 primers

DNA extracted from wild type, brk1, and dcd1 plants were used to genotype for dcd1.

PCR was setup using the dcdcapfor (GTGGTGACCTGGAGAATATCG) and

dcdcaprev2 (ATTAACAATAATTCCAGCTGGGATA) primers with B73 DNA, dcd1-O

DNA and water positive and negative controls. A single reaction included: 2 µl of 10X

Thermo Pol buffer (NE Biolabs), 2 µl 2.5mM dNTPs, 1 µl DMSO, 0.1 µl Taq DNA

polymerase (NE Biolabs), 1.25 µl of 100 ng/ul for primer, 1.25 µl of 100 ng/ul rev primer,

11.4 µl ddH2O, and 1 µl of genomic DNA. PCR conditions were set as follows: 94oC for

2 min, followed by three steps 94oC for 1 min, 56oC for 1 min, elongation at 68oC for

1min 30 sec, which was repeated 35 times, then 68oC for 10 min and a hold at 10oC.

The PCR products were digested and run on 4% agarose gel or 12% polyacrylamide

gels. For each digest, 0.4 µl EcoRV HF (NE Biolabs), 2 µl of NEB buffer #4, and 7.6 µl

of dd H2O were combined with 10 µl of PCR product. The tubes were placed on a heat

stable rack and incubated for 2-3 hours at 37oC in an enclosed incubator. Meanwhile, a

4% agarose gel was prepared by using Gene pure Hires Agarose gel dissolved into 1X

TAE. 4 µl of loading dye was added to each restriction digest product and a total of 24

µl of sample was loaded into each well and analyzed using 1Kb plus ladder (Life

Technologies). The products were electrophoresed on the gel for 2-3 hours at 95-105

Volts in 1XTAE. The DNA fragments were visualized by staining the gel in 1:10,000

dilution of Syber gold (Life Technologies) for 30-45 mins. The gel was observed and

photographed under blue light using a Cannon Photoshop digital camera. The resulting

gel image file was modified using iPhoto and ImageJ and printed out.

24

3.4 Immunolocalization of Tubulin and DNA Staining

The microtubule immunolocalization protocol used is similar to that described in Wright

et al. 2009. Necessary solutions include autoclaved 2XPHEM (120 mM PIPES, 50 mM

HEPES, 20 mM EGTA, 8 mM MgCl2, pH 6.9) and PBS (NaCl 80 gm; KCl 2 gm;

Na2HPO4 14.4 gm; KH2PO4 2.4 gm dissolved in 800 ml of dH2O, pH was adjusted to 7.4

with HCl and volume was brought up to 1L).

Tissue strips (2 mm wide) from an adult leaf (leaf #8-12) were cut from the basal 0 to

1.5 cm and 1.5 cm - 3.5cm of the developing leaf. Strips were fixed in a microtubule

fixing solution composed of 2.5ml (16%) formaldehyde, 10 µl (0.1%) Triton X-100, 5 ml

2XPHEM and 2.5ml H2O for 2 hours on a shaker. The strips washed three times in

1XPHEM with 0.05% Triton X-100 (PHEM/T) while shaking for 30 min. Cell walls were

permeabilized by digestion in 1% Driselase (Sigma-Aldrich) and 0.5% Pectolyase Y-23

(MP Biomedicals) in water for 15 min and then washed with PHEM/T. The strips were

extracted using 2XPHEM, 1% Triton X-100, 1% DMSO and H2O for 1h, rinsed in 1XPBS

three times for 30 min. The tissue strips were blocked in 5% normal goat serum (NGS)

diluted in 1XPBS for 30min. To visualize microtubules, the tissue strips were incubated

for 30 min under vacuum infiltration in an anti-α-tubulin antibody (clone B-5-1-2; Sigma-

Aldrich) diluted to 0.5 mg/ml in blocking solution (PBS/NGS) at room temperature and

later left to shake overnight. The next day, the tissue strips were washed three times

with 1XPBS with 0.05% Triton X-100 (PBS/T) for a total of 60 min. The tissue strips

were incubated in the secondary antibody, Alexa Fluor-488–conjugated anti-mouse IgG

(Life Technologies) diluted to 0.5 mg/mL in blocking solution (PBS/NGS) for 4 hrs in the

25

dark while shaking. While still in the dark, tissue strips were washed three times with

PBS/T. The last wash went overnight and samples in scintillation vial were stored in

4oC. To label nuclei, tissue strips were incubated in 10 mg/mL propidium iodide (Sigma-

Aldrich) in PBS or in water for 10 min and washed in PBS/T. Leaf strips are stored at

4oC for future use. They were mounted in Vectashield (Vector Laboratories) for

observation and imaging. Fluorescence was visualized on a Zeiss laser scanning

confocal microscope with using parameters recommended for visualizing FITC or

propidium iodide (PI).

3.5 Confocal Microscopy, Image Processing, and Analysis

Alexafluor 488, Alexafluor 568 and propidium iodide were excited at the appropriate

wavelengths with an argon (488-nm line), argon/krypton laser (568-nm line), or violet

blue laser (440-nm line). The confocal system was controlled using the Zeiss Zen

software. Z-projections of selected slices from stacks were assembled using Zeiss Zen

software. Image processing was performed using Image J. Measurements of the GMC

length and width was calculated using Image J. The measurements were converted

from pixels to microns.

26

3.6 Toulidine Blue O staining on epidermal peels

The epidermal TBO (stains starch) staining protocol was taken from Gallagher and

Smith, 1999. Prior to beginning, the following solutions were prepared. The fix solution

included: 20 ml of 0.5M Na phosphate buffer (pH7.2), 4 ml of 0.5 M EDTA (pH 8), 0.8

gm of 2% saponin and H2O for a total volume of 40 ml. The acetate buffer recipe

included: 847 ml 0.1M acetic acid and 153 ml 0.1M sodium acetate (trihydrate). TBO

staining solution was made by mixing 0.02 g TBO and 40 ml acetate buffer and can be

stored at room temperature for future use.

TBO staining was performed on adult leaves (defined here as leaves numbering 8-12,

counting the first leaf to be initiated as leaf number 1) from the plants sacrificed for

tubulin staining. Leaf blades were cut into 1 cm squares and fixed in 1 ml 40%

formaldehyde, 1 ml 10X fix stock solution and 8 ml H2O for at least 2 hours at room

temperature (RT). Tissue pieces were then washed 2-5 times in dH2O, digested in 0.1%

pectolyase (Sigma, St. Louis) in dH2O for at least 2 hours at RT then rinsed in dH2O.

The epidermis was then peeled from the rest of the leaf using dissecting microscope.

Peels were incubated in 0.05% TBO pH 4.0 until evenly stained and rinsed with water

multiple times to rinse off the extra stain. Peels were mounted in water and

photographed under bright-field conditions on a light microscope with a 20× objective.

.

27

CHAPTER 4

RESULTS

4.1 Identification of brk1; dcd1 (double) mutant plants

Due to the previously reported enhancement of the dcd1 phenotype with actin

cytoskeleton inhibitors (Gallagher and Smith, 1999), maize ears segregating the dcd1

and brk1 mutant alleles were obtained. brk1 encodes a protein needed for full ARP2/3

function and thus is required for normal organization of the actin cytoskeleton. dcd1

encodes a protein needed PPB organization and is thus required to modulated the

microtubule cytoskeleton. Individual and double mutants were identified via a

combination of phenotypic analysis and genotyping. Glue impressions of the epidermal

surface of each plant were taken. Using the glue impressions, the epidermal cell

outlines of each plant were examined and plants were placed in "wild-type" (normal

epidermal cell lobing; no abnormally shaped subsidiary cells), "brk1" (no epidermal cell

lobing; abnormal subsidiary cells), or "dcd1" (normal epidermal cell lobing; abnormal

subsidiary cells) categories. The plants were then genotyped for the presence/absence

of the dcd1 allele. These genotyping results were used to confirm the identity of the

dcd1 mutants and to unambiguously distinguish brk1 single mutants from the dcd1; brk1

double mutants (Figure 3).

28

Figure 3: Genotyping using dcd1 markers. The red boxes marked in the figure give the

interpreted genotype of the wild-type (WT) and dcd1 controls. DNA from wild type, brk1,

dcd1, and the brk1; dcd1 plants were amplified using dcd1 specific primers, digested

with Eco RV HF, and electrophoresed. Amplification products from the wild-type allele

are not cut by the restriction enzyme while the dcd1mutation in combination with the

dcdcapfor primer introduces an EcoRV cut site. The 230 bp band corresponds to the

wild-type allele and the 200 bp band corresponds to the dcd1 allele.

dcd

1

230bp 200bp

230bp 200bp

29

4.2 Quantitative analysis of abnormal subsidiary cells in mature leaf tissue.

To ascertain if the brk1; dcd1 double mutant has an enhanced phenotype relative

to the brk1 and dcd1 single mutants, epidermal peels were stained with Toluidine Blue

O to highlight the cell outlines and visualized under light microscope. The percentages

of abnormal subsidiary cells were calculated for each genotype (Table 1). Analysis of

subsidiary cell shape in the wild type, brk1, dcd1, and the brk1; dcd1 mutants showed

that the double mutants demonstrate a higher percentage of aberrant SCs than either of

the single mutants (Figure 4 and 5). The percentage of abnormal subsidiary cells in the

brk1; dcd1 mutant was much higher than expected on the basis of the single mutant

phenotypes alone suggesting that these two mutations have a synergistic effect SMC

division rather than additive one.

Genotype Total no. of abnormal

subsidiary cells

Total no. of subsidiary

cells

% of abnormal

subsidiary cells

wild type 3 675 0.44

dcd1 310 1188 26.09

brk1 172 978 17.59

brk1; dcd1 1139 1162 98.02

Table 1: Quantitative analysis of the number of abnormal subsidiary cells in wild type,

dcd1, brk1 and brk1; dcd1 plants observed by staining epidermal peels with Toluidine

Blue O.

30

0.00

20.00

40.00

60.00

80.00

100.00

wt dcd1 brk1 brk1;dcd1

0.44

26.09 17.59

98.02

% o

f A

bn

orm

al S

ub

sid

iary

cel

ls

Genotype

Percentage of abnormal SCs

%abnormal

Figure 4: Graphical representation of the number of abnormal subsidiary cells in

wild type, dcd1, brk1 and brk1; dcd1 plants observed by staining epidermal peels

with Toluidine Blue O.

31

Figure 5: Cell outlines and stomatal complexes of wild type (A), dcd1 (B), brk1 (C) and

brk1; dcd1 (D) observed by staining epidermal peels with Toluidine Blue O. Arrows

indicate normal subsidiary cells in the wild-type panel and abnormally shaped subsidiary

cells in the dcd1, brk1, and brk1; dcd1 panels.

C: brk1 D: brk1; dcd1

B: dcd1 A: Wild Type

32

4.3 Analysis of the organization of the microtubule cytoskeleton (PPB, spindle

and phragmoplast) and nuclear position in dividing subsidiary mother cells in

wild type, brk1, dcd1, and brk1; dcd1 mutants

To understand the cause of the abnormally shaped subsidiary cells in the brk1;

dcd1 mutant, the organization of the microtubule cytoskeleton was examined in

subsidiary mother cells (SMCs) undergoing cell division. Tubulin immunolocalization

was used to visualize the microtubule cytoskeleton and propidium iodide staining was

used to visualize nuclear position and chromatin status in immature, adult leaf tissues

during stomatal complex formation. The correct zone of cells was identified by looking

for rows of GMCs that contained the characteristic interphase microtubule rings.

Subsidiary cells adjacent to GMCs were examined in detail and the position and stage

of the nucleus/chromatin, organization of the cytoskeleton, and width of the associated

GMC were recorded. Data was collected for a range of SMCs in the division zone, not

just those obviously in mitosis. For each genotype, dividing cells from 3 individual plants

were evaluated. Some of the observed characteristics are very common and sufficient

numbers were accumulated to consider each individual plant as a replicate and

calculate standard error of the mean (SEM) for my results. Other characteristics are

very rare and to get an idea of the pattern I grouped the data from the 3 plants from

each genotype together and no SEMs could be calculated since n=1. Finally GMC

width has been correlated with the competence of the GMC to send the polarizing signal

(Humphries et al 2009). Guard mother cells < 6 µm in width are less likely to have sent

a signal and are considered immature versus mature GMCs which are <6 μm in width.

33

When appropriate, the data was binned to evaluate SMCs adjacent to the immature

GMCs (<6 µm in width) and mature GMCs (>6 µm) in width.

A) Nuclear migration

Polarization of the subsidiary mother cell nucleus towards the adjacent GMC is

the first step in the asymmetric cell division of a SMC. The migration of the SMC

nucleus is in response to an extrinsic signal from the adjacent GMC (Stebbins and

Shah, 1960). Table 2 and Figure 6 shows the percentage of polarized subsidiary cell

nuclei in relation to the GMC width. In total the nuclear position of SMCs in 281 wild

type, 380 dcd1, 873 brk1, and 617 brk1; dcd1 in interphase/preprophase or prophase

cells were evaluated. A cell was judged to be in prophase if the chromosomes were

condensed while all other SMCs adjacent to GMCs were considered to be in interphase

or preprophase. A SMC nucleus was considered polarized only if it was found directly

adjacent to the GMC. All other nuclear positions were considered not polarized. The

data reported in Table 2 shows the average number of polarized cells per plant as well

as the average over all plants of the same genotype. Figure 6 represents the data in

graph with SEM plotted as error bars.

34

GMC Width

(μm)

% of wild-type

SMC nuclei

polarized

% of dcd1 SMC

nuclei polarized

% of brk1 SMC

nuclei polarized

% of brk1; dcd1

SMC nuclei

polarized

GMC <6

plant 1 30 (n=101) 18 (n=22) 70 (n=54) 19 (n=63)

GMC <6

plant 2 44 (n=43) 37 (n=135) 12 (n=405) 12 (n=67)

GMC <6

plant 3 78 (n=85) 23 (n=180) 43 (n=353)

GMC <6

mean 37 (SEM = 5) 44 (SEM = 14) 35 (SEM = 15) 25 (SEM = 8)

GMC >6

plant 1 54 (n=50) 59 (n=54) 59 (n=41) 36 (n=76)

GMC >6

plant 2 59 (n=56) 71 (n=62) 16 (n=49) 38 (n=43)

GMC >6

plant 3 90 (n=31) 86 (n=22) 42 (n=144) 73 (n=15)

GMC >6

mean 68 (SEM = 9) 72 (SEM = 6) 39 (SEM=10) 49 (SEM= 10)

Table 2: Percentage of subsidiary mother cells with polarized nuclei. SEM = standard error

of the mean while n refers to the total number of cell evaluated for each condition.

35

Our data shows that when the adjacent GMCs are immature with width less than 6 μm,

less than 50% of the nuclei had migrated in all the samples and the degree of nuclear

migration was similar across genotypes. Whereas the nuclei in SMCs flanking GMCs

with a width more than 6 μm showed a greater degree of polarization, especially in the

wild type and dcd1 plants. SMCs in brk1 and brk1; dcd1 plants showed a similar

reduction in nuclear migration. Figure 7 shows the representative images of polarized

and unpolarized nuclei in wild type, dcd1, brk1, and brk1;dcd1 plants.

0%

10%

20%

30%

40%

50%

60%

70%

80%

90%

GMC <6 GMC >6

% o

f SM

Cs

GMC width (µm)

Nuclear Migration

wt polarize

dcd1 polarize

brk polarized

double polarize

Figure 6: Graphical representation of the overall mean number of SMCs with

polarized nuclei in the wild type, dcd1, brk1 and brk1; dcd1 (double) plants. Error

bars report the standard error of the mean.

36

Figure 7: Microtubules (green) and DNA (red) shown in (A) wild type, (B) dcd1, (C) brk1

and (D) brk1; dcd1 SMCs. Asterisks indicate the adjacent GMC and arrows point

towards polarized nucleus in wild type and unpolarized nucleus in dcd1, brk1 and brk1;

dcd1 mutant SMCs.

37

B) The organization and orientation of preprophase PPBs

After nuclear migration, the SMC enters into S phase (replication of DNA material

within the nucleus) and then G2 of cell cycle. During G2 phase of cell cycle, the PPB

band forms and cells are considered to be in "preprophase". SMCs in preprophase

were examined to record the variation in the orientation and organization of PPB in wild

type, dcd1, brk1 and brk1; dcd1 cells.

Table 3 illustrates the orientation and organization of PPBs, which are vital for

determining the orientation of the upcoming cytokinesis.

38

Genotype

PPB

orientation: %

normal

PPB

orientation:

% transverse

PPB

organization:

% normal

PPB organization:

% disorganized

PPB

organization:

% no PPB

present

wild type

plant 1 81 (n=26) 19 (n=26) 79 (n=28) 14 (n=28) 7 (n=28)

wild type

plant 2 88 (n=8) 13 (n=8) 38 (n=16) 13 (n=16) 50 (n=16)

wild type

plant 3

wild type

mean 84 (SEM = 2) 16 (SEM = 2) 58 (SEM=15) 13 (SEM=1) 29 (SEM=15)

dcd1

plant 1

dcd1

plant 2 97 (n=36) 3 (n=36) 11 (n=45) 69 (n=45) 20 (n=45)

dcd1

plant 3 83 (n=36) 17 (n=36) 12 (n=60) 48 (n=60) 40 (n=60)

dcd1

mean 90 (SEM=5) 10 (SEM=5) 11 (SEM=0) 59 (SEM=7) 30 (SEM=3)

brk1

plant 1 100 (n=12) 0 (n=12) 22 (n=36) 11 (n=36) 67 (n=36)

brk1

plant 2 59 (n=29) 41 (n=29) 7 (n=57) 44 (n=57) 49 (n=57)

brk1

plant 3 90 (n=20) 10 (n=20) 20 (n=40) 30 (n=40) 50 (n=40)

brk1


brk1; dcd1

plant 1 100 (n=7) 0 (n=7) 0 (n=12) 58 (n=12) 42 (n=12)

brk1; dcd1

plant 2 33 (n=3) 67 (n=3) 0 (n=13) 23 (n=13) 77 (n=13)

brk1; dcd1

plant 3 74 (n=69) 26 (n=69) 5 (n=157) 39 (n=157) 56 (n=157)

brk1; dcd1


Table 3: Quantitative analysis of PPB orientation and organization in SMC's adjacent to GMCs

with a width < 6μm. The SMCs are in preprophase since the nucleus has migrated or a PPB is

visible, but the chromosomes are not condensed. SEM = standard error of the mean while n=

refers to the total number of cells evaluated for each condition.

39

Normal (asymmetrically positioned) and transverse PPBs in the SMCs adjacent to

GMCs less than 6 μm width in all the representative genotypes were counted and the

results are shown in Figure 8. No differences were detected amoung the different

genotypes.

0%

10%

20%

30%

40%

50%

60%

70%

80%

90%

100%

normal transverse

% o

f S

MC

s

PPB orientation in SMCs

Preprophase PPB orientation

wild type

dcd1

brk1

brk1; dcd1

Figure 8: Graphical representation of PPB orientation in preprophase SMCs adjacent to

GMCs with width <6 μm. Error bars report the standard error of the mean.

40

Likely the SMCs cells with transverse PPBs are due to the proximity of the SMCs

to an immature GMC that failed to polarize the SMC. One of the daughters of these

transverse divisions will then be polarized by the mature GMC allowing the creation of

the needed SC.

Figure 9 illustrates the organization of PPB, which delineates the CDS and the place of

future phragmoplast fusion as well as contributes to the initial orientation the spindle, in

SMCs adjacent to GMCs with a width < 6 μm. PPBs observed in wild type cells were

well defined; that is the microtubules formed a tight ring that circumvented the

subsidiary mother cell. Disorganized PPBs were frayed and disorganized in structure

such that the PPB microtubules spread across the cell surface instead of encircling the

cell in a tight ring. Cells that had polarized nuclei, but no obvious PPB were placed into

the "no PPB" category. The percentage of preprophase SMCs (adjacent to immature

GMCs) with normal or disorganized PPBs or with no PPB are graphically shown in

Figure 9.

41

Disorganized PPBs were commonly seen in the dcd1 SMCs adjacent to immature guard

mother cells and while SMCs in preprophase with no obvious PPBs were observed in

across all genotypes, the brk1 and brk1; dcd1 plants had a greater percentage of SMCs

adjacent to immature GMCs lacking PPBs entirely.

Observations were also made regarding the PPB in SMCs adjacent to the GMC width

>6 μm. SMCs adjacent to GMCs with a width > 6 μm are more likely to have responded

to the GMC polarizing signal. Table 5 reports the PPB orientation and organization in

SMCs adjacent to GMCs with a width > 6 μm. These are considered mature GMCs.

0%

10%

20%

30%

40%

50%

60%

70%

80%

normal disorganized PPB no PPB

% o

f SM

Cs

PPB organization in SMCs

Preprophase PPB structural organization

wild type

dcd1

brk1

brk1; dcd1

Figure 9: Graphical representation of PPB organization in preprophase SMCs

adjacent to GMCs with width <6 μm. Error bars report the standard error of the

mean.

42

Genotype

PPB

orientation:

% normal

PPB

orientation:

% transverse

PPB

organization:

% normal

PPB

organization:

% disorganized

PPB

organization:

% no PPB

present

wild type

plant 1 100 (n=23) 0 (n=23) 85 (n=26) 4 (n=26) 12 (n=26)

wild type

plant 2 100 (n=12) 0 (n=12) 39 (n=31) 0 (n=31) 61 (n=31)

wild type

plant 3 95 (n=20) 5 (n=20) 63 (n=24) 21 (n=24) 17 (n=24)

wild type


dcd1

plant 1 64 (n=25) 36 (n=25) 9(n=32) 69 (n=32) 22 (n=32)

dcd1

plant 2 100 (n=36) 0 (n=36) 7 (n=42) 79 (n=42) 14(n=42)

dcd1

plant 3 92 (n=13) 8 (n=13) 28 (n=28) 44 (n=28) 28 (n=28)

dcd1

mean 74 (SEM=9) 15 (SEM=9) 15 (SEM=5) 64 (SEM=8) 21 (SEM 3)

brk1

plant 1 87 (n=15) 13 (n=15) 46 (n=24) 17 (n=24) 38 (n=24)

brk1

plant 2 67 (n=3) 33 (n=3) 13 (n=8) 25 (n=8) 63(n=8)

brk1

plant 3 88 (n=40) 13 (n=40) 29 (n=58) 40 (n=58) 31 (n=58)

brk1


brk1; dcd1

plant 1 100 (n=13) 0 (n=13) 0 (n=29) 45 (n=29) 55 (n=29)

brk1; dcd1

plant 2 60 (n=10) 40 (n=10) 6 (n=18) 50 (n=18) 44 (n=18)

brk1; dcd1

plant 3 100 (n=6) 0 (n=6) 10 (n=10) 50 (n=10) 40 (n=10)

brk1; dcd1

mean

87 (SEM=

11) 13 (SEM=11) 5 (SEM= 2) 48 (SEM=1) 47 (SEM=4)

Table 5: Quantitative analysis of PPB orientation and organization in SMCs adjacent to GMCs with a

width > 6μm. The SMCs are in preprophase since the nucleus has migrated or a PPB is visible, but the

chromosomes are not condensed. SEM = standard error of the mean while n= refers to the total

number of cells evaluated for each condition.

43

Table 5 and Figure 10 shows that with the increase in the adjacent GMC width, the

percentage of normally oriented PPBs increased in the wild-type plants. However, in

both single and the double mutants there remained a higher percentage of transverse

PPBs compared to wild-type.

0%

20%

40%

60%

80%

100%

120%

normal transverse

% o

f S

MC

s


Preprophase PPB orientation

wild type

dcd1

brk1

brk1; dcd1

Figure 10: Graphical representation of PPB orientation in preprophase SMCs

adjacent to GMCs with width >6 μm. Error bars report the standard error of the

mean.

44

The organization of PPBs observed in SMCs adjacent to GMCs with a width

more than 6 μm (Figure 11) shows that dcd1 has the highest percentage of

disorganized PPBs and low percentage of normal PPBs, which is similar to what was

previously reported (Wright et al 2009). The brk1; dcd1 SMCs have the next greatest

amount of disorganized PPB while the brk1 SMCs have even less. The brk1 and brk1;

dcd1 preprophase SMCs with polarized nuclei failed to form PPBs the most often, but

due to variability in the wild-type data, the amount of missing PPBs are not significantly

different from wild type.

Figures 12 and 13 highlight the PPB organization and orientation respectively in SMCs

0%

10%

20%

30%

40%

50%

60%

70%

80%

normal disorganized PPB no PPB

% o

f S

MC

s


Preprophase PPB Organization

wild type

dcd1

brk1

brk1; dcd1

Figure 11: Graphical representation of PPB organization in preprophase SMCs

adjacent to GMCs with width >6 μm. Error bars report the standard error of the

mean.

45

adjacent to GMCs with a width < or > 6µm in wild type, dcd1, brk1 and brk;dcd1 plants.

Figure 12: PPB organization in preprophase SMCs in wild type, dcd1, brk1 and brk1;

dcd1Asterisks indicate GMC and arrows points towards the PPB organization. Mid view (A) and

top view (B) of normally organized PPB in a wild type SMC. Mid view (C) and top view (D) of no

PPB in in brk1. Mid view (E) and top view (F) of disorganized PPB in brk1; dcd1. Microtubule

structure (green) and DNA (red).

A B

E F

46

Figure 13: PPB orientation in preprophase SMCs in wild type, dcd1, brk1 and brk1;

dcd1 SMCs. Asterisks indicate GMC and arrows points towards the PPB orientation.

Mid view (A) and top view (B) of normally oriented PPB in wild type SMCs. Mid view

(C) and top view (D) of SMCs with transverse PPB in brk1; dcd1. Microtubule structure

(green) and DNA (red).

47

C) The organization and orientation of prophase PPBs

As the SMCs move into prophase, the chromosomes are condensed and

become distinct. PPB analysis at this phase of the cell cycle is very important as it

defines the cortical division site which determines where the phragmoplast will mediate

new cell wall fusion to the mother cell. PPB organization and orientation is also

important for the initial orientation of spindle. SMCs in prophase were identified by

condensed chromosomes and the PPB organization and orientation for each cell was

noted. Because of the rarity of these events, cells from all three plants were considered

together so standard error of the mean could not be calculated. Table 6 shows

percentage normal and transversely oriented PPB in all the categories.

Genotype

Number of

prophase SMCs

observed

PPB orientation:

% normal

PPB orientation:

% transverse

wild type 10 100 0

dcd1 14 75 25

brk1 16 64 36

brk1; dcd1 15 40 60

Table 6: Quantitative analysis of PPB orientation in dividing SMCs in prophase in

wild type, dcd1, brk1 and brk1; dcd1 plants.

.

48

As shown in table 6, the data indicated that the highest percentage of transversely

oriented PPBs are in the brk1; dcd1 SMCs. Summation of the percentage of transverse

PPBs in brk1 and dcd1 SMCs is equivalent to what is seen in the double mutants

suggesting that the effect of mutations in double is additive of the single mutants. The

PPB orientation defects during SMC prophase in the brk1; dcd1 double likely contribute

to aberrations observed in SC shape.

0%

20%

40%

60%

80%

100%

120%

normal transverse

% o

f SM

Cs


Prophase PPB orientation

wild type

dcd1

brk1

brk1; dcd1

Figure 14: Graphical representation of PBB orientation in dividing SMCs in

prophase in wild type, dcd1, brk1 and brk1; dcd1 plants.

.

49

The organization of PPBs in prophase SMCs was also observed across all genotypes

and the results are shown in Table 7 and Figure 15. The organization of the brk1 PPBs

was similar to wild-type while the dcd1 and brk1; dcd1 SMCs had the highest

percentage of disorganized PPBs.

Genotype

Number of

prophase

SMCs

observed

PPB

organization:

% Normal PPB

PPB

organization:

% disorganized

PPB

PPB

organization:

% no PPB

wild type 12 83 0 20

dcd1 14 29 65 6

brk1 17 79 21 0

brk1; dcd1 15 13 87 0

Table 7: Quantitative analysis of PPB organization in dividing SMCs in prophase in wild

type, dcd1, brk1 and brk1; dcd1 plants.

50

Figures 16 and 17 shows representative images of PPB organization and

orientation in prophase SMCs.

0%

10%

20%

30%

40%

50%

60%

70%

80%

90%

100%

normal disorganized absent

% o

f SM

Cs


Prophase PPB organization

wild type

dcd1

brk1

brk1; dcd1

Figure 15: Graphical representation of PBB organization in dividing

prophase SMCs in wild type, dcd1, brk1 and brk1; dcd1 plants.

51

Figure 16: PPB organization in SMCs in prophase in wild type, dcd1, brk1 and brk1;

dcd1. Asterisks indicate GMC and arrows points towards the PPB organization. Mid

view (A) and top view (B) of normally organized PPB in SMCs. Mid view (C) and top

view (D) of disorganized PPB in SMCs in brk1; dcd1 . Mid view (E) and top view (F)

of no PPB in SMCs in dcd1. Microtubule structure (green) and DNA (red)

C D

E F

52

Figure 17: PPB orientation in prophase SMCs in wild type, dcd1, brk1 and brk1; dcd1

SMCs. Asterisks indicate GMC and arrows points towards the normally or

transversely oriented PPB. Mid view (A) and top view (B) of normally oriented PPB in

wild type. Mid view (C) and top view (D) of transverse PPB in brk1; dcd1. Microtubule

structure (green) and DNA (red)

C D

53

D) Spindle orientation in SMCs

The organization and orientation of spindle determines the initial positioning of

the phragmoplast because the latter arises from the remnants of spindle. It was thus

important to observe the orientation of spindle to determine the probable position of

initial phragmoplast formation. Normal spindles are those whose long axis forms a 90

degree angle with the side of the SMC adjacent to the GMC. Tilted spindles are those

that deviate from this 90 degree angle while transverse spindle are parallel to the long

axis of the SMC. Table 8 illustrates the variations observed in the orientation of

spindles in wild-type, brk1, dcd1, and the brk1; dcd1 SMCs.

Genotype Total no. of

subsidiary cells

% Normal

Spindle

% Tilted

Spindle

% Transverse

spindle

wild type 10 100 0 0

dcd1 20 65 35 0

brk1 17 47 35 18

brk1;dcd1 17 0 71 29

Table 8: Quantitative analysis of spindle orientation in dividing SMCs in wild


9: Quantitative analysis of spindle structure and orientation in subsidiary cells

54

The data in Table 8 and Figure 18 shows that none of the SMCs in the brk1; dcd1

mutant had normally positioned spindles, though no defects in spindle organization

were seen. The high percentage of tilted and transverse spindles seen in the double

mutant likely contributes to a misorientation of the phragmoplast and the subsequent

cell division. The single dcd1 and brk1 mutants also had a much higher percentage of

tilted spindles while transverse spindles were seen in the brk1 and the double mutant

but not in wild type or dcd1 SMCs. Figures 19 and 20 shows examples of tilted and

transverse spindle in wild type and mutant genotypes.

0%

20%

40%

60%

80%

100%

120%

Normal Spindle Tilted Spindle Transverse spindle

% o

f SM

Cs

Spindle orientation in SMCs

Spindle Orientation

wild type

dcd1

brk1

brk1; dcd1

Figure 18: Graphical representation of spindle orientation in dividing SMCs in


55

A

*

B

C D

Figure 19: Spindles in wild type, dcd1, brk1 and brk1; dcd1 SMCs. Asterisks indicate GMC and arrows

points towards orientation of spindle. (A) normal spindle in wild type, (B) titled spindle in dcd1, (c) tilted

spindle in brk1 and (D) tilted spindle in brk1; dcd1 mutants. Microtubule structure (green) and DNA (red).

56

C

B

D

Figure 20: Spindles wild-type, dcd1, brk1 and brk1; dcd1 SMCs. Asterisks indicate GMC and

arrows points towards orientation of spindle. (A) normal spindle in wild type, (B) transverse

spindle in dcd1, (c) transverse spindle in brk1 and (D) transverse spindle in brk1; dcd1

mutants. Microtubule structure (green) and DNA (red).

57

E) Phragmoplast orientation observed in SMCs

Structural aspects and the orientation phragmoplasts in dividing SMCs were

evaluated in all genotypes. In wild type, during cytokinesis, it has been observed that

the phragmoplast develops from the remnants of the spindle and expands towards the

CDS (cortical division site) which was established by the PPB during preprophase and

prophase. Table 9 and Figure 21 illustrate the variation seen in phragmoplast

orientation in SMCs. Normal, on-track phragmoplasts are those in the process of

expanding towards the normal CDS position in SMC cells. Off-track phragmoplasts are

wandering elsewhere in the cell while transverse phragmoplasts are those that are

expanding in a straight line directly across the cell.

Genotype Total number

of SMCs

% of Normal on

track phragm.

% of Off track

phragm.

% of Transverse

phragm.

wt 12 100 0 0

dcd1 26 65 27 8

brk1 16 75 19 6

brk1;dcd1 11 18 64 18

Data collected indicates the highest percentage of off-track and transverse

phragmoplast in the double mutants followed by the single mutants. The wild-type

SMCs had no off track or transverse phragmoplasts.

Table 9: Quantitative analysis of phragmoplast orientation in dividing SMCs in wild


58

Phragmoplast positioning is important as it determing the final position of the cell

plate the forms between the daughter cells. Thus the abnormally oriented

phragmoplasts (transverse or off-track) observed in double reflects the higher

percentage of abnormal SCs associated with the GMC in their stomatal complex.

Figure 28 and 29 shows representative examples of phragmoplast orientation in wild-

type and mutant SMCs.

0%

20%

40%

60%

80%

100%

120%

Normal phragm. Off track phragm. Transverse phragm.

% o

f SM

Cs

Phragmoplast orientation in SMCs

Phragmoplast orientation

wild type

dcd1

brk1

brk1; dcd1

Figure21: Graphical representation of phragmoplast orientation in dividing SMCs in


59

Figure 22: Phragmoplast in wild type, dcd1, brk1 and brk1; dcd1 SMCs. Asterisks indicate GMC

and arrows points towards phragmoplasts. (A) Normal phragmoplast in wild type , (B) off track

phragmoplast in dcd1, (c) off track phragmoplast in brk1 and (D) off track phragmoplast in

brk1; dcd1 mutants. Microtubule structure (green) and DNA (red).

C

B

D

60

D

*

A B

*

C

Figure 23: Phragmoplast in wild type, dcd1, brk1 and brk1; dcd1 SMCs. Asterisks indicate GMC

and arrows points towards phragmoplasts oreintation. (A) Normal phragmoplast in wild type , (B)

Transverse phragmoplast in dcd1, (c) transverse phragmoplast in brk1 and (D) transverse

phragmoplast in brk1; dcd1 mutants. Microtubule structure (green) and DNA (red).

61

. CHAPTER 5

DISCUSSION

Subsidiary mother cell (SMC) divisions during stomatal complex formation in Zea

mays are asymmetric generating a small subsidiary cell (SC) and a larger epidermal

cell. Mutants with a high number of abnormally shaped subsidiary cells include the

brick1 (brk1) and discordia1 (dcd1) mutants. brk1 mutants have defects in actin

polymerization with dcd1 mutants have defects in microtubule nucleation/organization.

Possible causes of the abnormal SCs in brk1 mutants include a failure of the SMC

nucleus to polarize in advance of mitosis, no actin patch, and transverse and/or missing

PPBs (Gallagher and Smith, 2000; Panteris et al 2006). The abnormal subsidiary

mother cell division in dcd1 is due to correctly localized, but disorganized PPBs (Wright

et al. 2009). The reported observations that 1) brk1 mutants have defects in PPB

formation and 2) the dcd1 phenotype is enhanced by the application of actin inhibitors

led us to examine the brk1; dcd1 double mutant to see if we could uncover a role for

actin in PPB organization (Gallagher and Smith, 1999).

Experiments were conducted to determine the number of abnormally shaped SC

the single mutants (dcd1 and brk1) and in the double mutant (brk1; dcd1) and it was

found that brk1; dcd1 double mutants have a higher percentage of aberrant SCs than

the single mutants combined suggesting that these two mutations have a synergistic

rather than additive effect on the orientation of SMC divisions.

62

wild type dcd1 brk1 brk1; dcd1

% Polarized

(immature GMCs) 37 44** 35* 25

% Polarized

(mature GMCs) 68 72 39** 49**

% Polarized

(prophase) 100 95 67* 47**

% Normal PPB

(immature GMCs) 62 12** 17** 3**

% Normal PPB

(mature GMCs) 55 12** 29** 5**

% Normal PPB in

prophase 91 37** 64 7**

% Normal spindle 100 65 47** 0**

% Normal

phragmoplast 100 65 75 18**

% Normal SC 99.6 73.9 82.4 2

Table 10: Summary table illustrating the percentage of SMCs with polarized nuclei,

normal PPBs, normal spindles, normal phragmoplast and normal subsidiary cells.

Immature GMCs refers to SMCs adjacent to GMCs < 6 μm while mature GMCs refers to

SMCs adjacent to GMCs > 6 μm. * indicates a P-value < 0.05 and ** indicates a P-value <

0.01 suggesting the mutant condition is significantly different from wild type. P values were

calculated using the Fisher exact test.

63

To evaluate whether the defects seen in double mutants are synergistic or

additive, tubulin immunolocalization was used to visualize the organization of the

microtubule cytoskeleton (PPB, spindle and phragmoplast) in dividing subsidiary mother

cells in wild type, brk1, dcd1, and brk1; dcd1 mutants. By observing these

intermediate cell division steps, I wanted to learn where the defects arose that

contributed to the strong SC phenotype in double mutants. Moreover, the DNA was

stained with propidium iodide so that the position of the nucleus and progression

through the cell cycle could also be evaluated in all genotypes.

The SMC division process starts with nuclear migration towards the adjacent

GMC. The results show that wild type, dcd1, brk1 and brk1; dcd1 all exhibit the same

degree of nuclear migration when the polarizing GMC was immature (width <6 µm). The

SEM was calculated by taking the averages of the percentages in all the 3 rounds of

experiments to find out the mean deviation. The SEM was plotted as error bars and the

high degree of error bar overlap reflects that all the four genotypes lie in region of

standard mean thus single mutants and double mutants do not differ dramatically from

wild type SMCs adjacent to immature GMCs with respect to nuclear polarization events.

However, Fisher's exact test of independence identified a significant difference between

the extent of nuclear polarization between the single mutants (dcd1 and brk1) and wild

type in SMCs adjacent to immature GMCs (Table 10).

Data collected on SMCs near to mature GMC (width > 6 µm) and SMCs in

64

prophase indicates that wild type and dcd1 plants were more likely to have undergone

nuclear migration towards the GMC with the brk1 and brk1; dcd1 double SMCs equally

deficient in nuclear migration (Table 10). Earlier studies reported that brk1 mutants

have defects in actin patch formation and since this actin patch is thought to be involved

in migration. Therefore the above results related to nuclear migration in brk1 matches

well with previous studies. The lack of dcd1 activity does not seem to enhance the brk1

migration defects suggesting that microtubules (MTs) in general and dcd1 in specific are

not needed for nuclear migration.

The orientation of PPBs in preprophase SMCs adjacent to immature and mature

GMCs as well as SMCs in prophase was evaluated. When adjacent to an immature

GMC, the percentage of SMCs in preprophase with normal and transverse PPBs were

similar in all genotypes. A possible explanation of high percentage of transverse PPBs

could be due to SMCs progressing though the cell cycle without the influence of the

polarizing GMC signal. When the dividing SMCs are adjacent to mature GMCs, wild-

type SMCs almost never had a transverse PPB while all the mutants had a low

percentage. The same factor that contributes to the failure of brk1 plants to form actin

patches and undergo nuclear migration could also result in incorrectly orientated PPBs.

It is unclear why the dcd1 mutants have a low level of transverse PPBs though perhaps

the dcd1 SMC population that was evaluated was more likely to be near GMCs that

were not actively signaling to the adjoining SMC even though the GMC width exceeded

6 µm. By the time the SMCs were in prophase however, many more brk1; dcd1 double

mutants had transverse PPBs. It is unclear what could cause this dramatic increase

65

though perhaps small sample numbers of prophase stage cells could be a contributing

factor.

Organizational analysis of PPBs in SMCs in preprophase phase adjacent to

GMCs with widths < 6 µm and > 6 µm as well as in prophase showed that dcd1 mutants

have the highest percentage of disorganized PPBs at all stages. brk1 and brk1; dcd1

also showed high levels of disorganized PPBs relative to wild-type. The calculated P

values suggest that the number of SMCs with abnormal PPBs is significant in the single

and double mutants until prophase when the number of abnormal PPBs ceased to be

significant in the brk1 mutant only (Table 10). This suggests that perhaps PPBs

organize slower in brk1 plants, but by the time the cell cycle has advanced to prophase,

the brk1 PPB organization has caught up to wild type.

After prophase, the dividing cells form the mitotic spindle to distribute the nuclear

material into two daughter cells. Additionally, spindle orientation plays an important role

in the initial orientation of the cytokinetic cytoskeletal structure, the phragmoplast.

Spindle orientation in wild type, dcd1, brk1, and brk1; dcd1 showed the single and

double mutants had a greater percentage of misorientated spindles (Table 10). The

spindle misorientation in brk1 single mutant was due likely due to the lack of actin patch,

which helps tether the spindle in the correct orientation. The spindle misorientation in

the dcd1 single mutant is likely due to the disorganized PPB since MT polymerization on

the surface of the nucleus during spindle formation is inhibited by the PPB thus allowing

for formation of a distinct bipolar spindle orientated perpendicular to the PPB. A

66

disorganized PPB could result in initially disoriented spindles. The double mutants had

no normally oriented spindles since all the spindles were either tilted or transverse. The

higher percentage of affected spindles in the double mutant could be due to the additive

effects of the defects seen in the single mutants. P values suggested that the difference

between wild type and brk1 as well as between wild type and brk1; dcd1 were

significant.

Finally, phragmoplast orientation was observed in wild type, dcd1, brk1, and

brk1; dcd1 dividing SMCs. All the single mutants as well as the double mutant had a

significantly larger number of off-track or transverse phragmoplasts relative to wild type

(Table 10). Less than 20% of the double mutant SMCs were observed with a normally

oriented phragmoplast.

While the initial observation of the SC cell shape defect in the double mutant

suggested that the defects were synergistic, analysis of all the intermediate stages of

cell division suggests that the defects in SMC division in the double mutant are caused

by additive effects of the brk1 and dcd1 single mutations. We did not detect a role for

BRK1 in microtubule organization. Instead the dcd1 single mutants are able to

overcome the large percentage of disorganized PPBs while brk1 single mutants are

able to overcome the lack of actin patch formation which leads to poor nuclear migration

and tilted spindles, but the combination of these defects in the double mutant cannot be

overcome. This observation suggests a degree of redundancy with the functioning of

the actin and microtubule cytoskeletons to promote a high fidelity, asymmetric SMC

division.

67

The above observations are intriguing and the future directions include

increasing the numbers of prophase PPBs, spindle and phragmoplasts observed for

each genotype and actin patch evaluation in all the four genotypes. Additionally,

observing live cells expressing tubulin-YFP to could clarify the timing defects suggested

by our data.

68

CHAPTER 6

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