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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
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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
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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.
3
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
7
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
mean 83 (SEM=10) 17 (SEM=10) 16 (SEM=8) 28 (SEM=5) 55 (SEM=8)
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
mean 69 (SEM=16) 31 (SEM=16) 2 (SEM=1) 40 (SEM=8) 58 (SEM=8)
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
mean 98 (SEM=1) 2 (SEM=1) 62 (SEM=11) 8 (SEM=5) 30 (SEM=13)
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
mean 80 (SEM=6) 20 (SEM=6) 29 (SEM=8) 27 (SEM=5) 44 (SEM=8)
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
PPB orientation in SMCs
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
PPB organization in SMCs
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
PPB orientation in SMCs
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
PPB organization in SMCs
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
type, dcd1, brk1 and brk1; dcd1 plants.
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
wild type, dcd1, brk1 and brk1; dcd1 plants.
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
type, dcd1, brk1 and brk1; dcd1 plants.
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
wild type, dcd1, brk1 and brk1; dcd1 plants.
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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