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P!!!!`a!!Figure 2.6: Dynamics of FtsZ1 and FtsZ2 expressed singly or together
. S. pombe
 cells expressing (A) FtsZ1
-eYFP (green), (B) FtsZ2
-eCFP (red), or (C and D) FtsZ1
-eYFP and FtsZ2
-eCFP were analyzed by FRAP
. Images from left to right represent 
fluorescence signals in photobleached regions (circled) just prior to bleaching (PB), 
at the time of bleaching (0 s), at the time closest to t
!, and at twice t
!. Representative 
plots of fluorescence recovery vs time are shown at right
. Data in each plot were 
normalized to the PB fluorescence signal (1 on the y
-axis) and the signal intensity
 at the time of bleaching (0 on the y
-axis)
. Boxes in the plots show the average t
1/2
 (also 
indicated by vertical lines) and average % recovery ± SD for 10
 independent FRAP 
datasets obtained for each strain
. Bars, 2 
"m. ! !``!!Figure 2.7: 
FtsZ1 and FtsZ2 dyna
mics were not altered over the range of protein 
levels analyzed during FRAP analysis. 
Graphs show individual half
-time and % 
recovery values
 versus photomultiplier tube (PMT) voltage settings for each of the ten 
cells analyzed by FRAP in all four single
-expression stains. With all other microscope 
acquisition settings remaining constant for all experiments, PMT voltage inversely 
correlates to fluorescence intensity 
(http://www.olympusmicro.com/primer/techniques/confocal/pmtintro.html
) and therefore 
with protein content (high voltage = lower protein and vice versa), though the 
relationship is not necessarily linear. Left y
-axis represents recovery half
-time (blue 
 !`N!Figure 2.7 (contÕd): 
bars); right y
-axis represents percent recovery (red bars). (A) 
FtsZ1, (B) FtsZ2, (C) FtsZ1 D275A, and (D) FtsZ2 D322A expressed alone.
!!![<!76?E!7??<??<;!@ER!GE<WI=<??:E8!EJ!"D?01!78;!"D?03!7JJ<GD<;!<7G@!EJ!D@<:=!
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!B,D7D:?D:G7665!?:H8:J:G78D!;:JJ<=<8G<!J=EC!D@<!GE==<?IE8;:8H!976><!:8!"D?03!?:8H6<!
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!D1Y3
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!"D?01
.33.10 ± 7.83.71.32 ± 11.51."D?01!23eN)
.49.79 ± 8.35a.53.91 ± 11.03a."D?03
.86.96 ± 22.08a.31.07 ± 7.70a."D?03!2a33)
.239.21 ± 271.60.12.09 ± 10.25b."D?01
.y!"D?03
.38.68 ± 9.88.73.74 ± 13.64.70.93 ± 13.37.74.82 ± 11.26b."D?01!23eN)
.y."D?03!2a33)
.46.93 ± 9.37.56.66 ± 10.31.123.21 ± 69.66.19.27 ± 12.47."D?01
.y."D?03!2a33)
.38.00 ± 11.01.46.83 ± 10.44a.72.11 ± 106.96.7.57 ± 8.53."D?01!23eN)
.y."D?03
.41.22 ± 11.10.50.37 ± 12.85.112.06 ± 49.74.30.74 ± 20.44.!`e!!Figure 2.8: Dynamics of GTPase
-deficient FtsZ1 and FtsZ2 expressed in various 
combinations. S. pombe
 cells expressing (A) FtsZ1 D275A, (B) FtsZ2 D322A (panel 3 
 !`d!Figure 2.8 (contÕd): 
is 100 s recovery, panel 4 is the image acquired closest to t
!), (C 
and D) FtsZ1 D275A and FtsZ2 D322A, (E and F) FtsZ1 and FtsZ2 D322A, and (G and 
H) FtsZ1 D27
5A and FtsZ2
. WT and mutant FtsZ1 and FtsZ2 proteins were fused to 
eYFP (green) and eCFP (red), respectively
. Images from left to right (except in B) 
represent fluorescence signals in photobleached regions (circled) just prior to bleaching 
(PB), at the tim
e of bleaching (0 s), at the time closest to t
!, and at twice t
!. Representative plots of fluorescence recovery vs time are shown at right
. Data in each 
plot were normalized to the PB fluorescence signal (1 on the y
-axis) and the signal 
intensity at the ti
me of bleaching (0 on the y
-axis)
. Boxes in the plots show the average 
t1/2
 (also indicated by the vertical lines) and average % recovery ± SD for 10 
independent FRAP datasets obtained for each strain
. Bars, 2 
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!!!!e3!Chapter 3
 Conservation of FtsZ 
Functionality Across the Photosynthetic Lineages
                   The pREP41
-SeFtsZ
-msfGFP
 and pREP41
-msfGFP
 expression constructs were 
generated and initially imaged by co
-author Joshua S. MacCready, Microbiology and 
Molecular Genetics Graduate Program, Department of Plant Biology, and Plant 
Research Laboratory at Michigan State University.
  !ea!Abstract
  FtsZ, a c
ytoskeletal GTPase, self
-assembles into a ring that is responsible for 
providing force for membrane constriction during cell division in bacteria and chloroplast 
division in photosynthetic eukaryotes. While bacteria accomplish cell division with a 
single F
tsZ, chloroplast division requires the coordinated action of two functionally 
distinct FtsZ isoforms, FtsZ1 and FtsZ2 in plants and green algae and FtsZA and FtsZB 
in red algae with primary chloroplasts. Recent functional studies have begun to 
elucidate th
e properties of FtsZ1 and FtsZ2 from Arabidopsis, yet very little functional 
data for FtsZ pairs from other photosynthetic organisms exist. In this study, we 
expressed fluorescent fusions of FtsZs from a monocot and dicot plant, green alga, red 
alga, and cyanobacterium in the heterologous 
Schizosaccharomyces pombe
 system to 
study their inherent filament morphology and turnover characteristics. 
We found that the distinct 
filament
 morphologies for the FtsZ1
/FtsZB
 and FtsZ2
/FtsZA
 families are highly 
conserved across a broad evolutionary distance
. When coexpressed, FtsZs from plants 
coassembled into FtsZ2
-like rings, while FtsZs from a red alga coassembled in 
filaments with characteristics of both FtsZ families. Additionally, FRAP 
experiments 
demonstrated that FtsZ1s have greater filament turnover properties than FtsZ2s and 
that the GTPase activity of FtsZ2/FtsZA, but not FtsZ1/FtsZB, is critical for promoting 
filament turnover. Interestingly, a cyanobacterial FtsZ displayed filamen
t morphology 
characteristics of both FtsZ families, but turnover properties highly similar to those of 
FtsZ2 and FtsZA. Taken together, our data suggest that the distinct properties 
previously described for Arabidopsis FtsZ1 and FtsZ2 extend to FtsZs from 
both green 
!e`!and red photosynthetic lineages and support the theory that FtsZ2 and FtsZA are the 
direct homologs of cyanobacterial FtsZ.
  !eN!Introduction
  FtsZ is a cytoskeletal GTPase that self
-assembles into a ring
-like structure that is 
responsible for drivi
ng cytokinesis in bacteria and chloroplast division in photosynthetic 
eukaryotes 
(Adams and Errington, 2009; Erickson et al., 2010; Mingorance et al., 2010; 
Osteryoung and Pyke, 2014)
. In bacteria, a single FtsZ protein assembles at the midcell 
to form the
 FtsZ ring (Z ring), which is attached to the cytoplasmic membrane via 
interaction with peripheral and integral membrane proteins 
(Addinall and Lutkenhaus, 
1996; Bi and Lutkenhaus, 1991; Liu et al., 1999)
. Once assembled, the Z ring acts as a 
scaffold for 
the recruitment of additional downstream division
-related proteins to the 
midcell before constriction occurs 
(Adams and Errington, 2009)
.  When assembled 
in vitro
, purified 
E. coli
 FtsZ normally assembles single
-stranded protofilaments, but also assembles 
into helices, sheets, and large bundles 
depending on assembly conditions 
(Adams and Errington, 2009; Erickson et al., 2010)
. FtsZ assembly stimulates its GTPase activity, which destabilizes the subunit interface 
and induces filament fragmentation and event
ually polymer turnover 
(Huecas et al., 
2007). Once assembled, FtsZ filaments alone are able to exert contractile force on 
membranes, which may be a result of a hinge
-opening mechanism induced by GTP 
hydrolysis and a fixed direction of curvature 
(Li et al.,
 2013; Osawa et al., 2008; Osawa 
et al., 2009). GTPase activity is correlated with both rates of polymer turnover and 
protofilament length and extent of bundling 
(Chen et al., 2007; Chen and Erickson, 
2009; Mukherjee and Lutkenhaus, 1998; Srinivasan et al.
, 2008). Additionally, it has 
been shown that GTPase activity is critical for sustaining Z
-ring constriction, as 
E. coli
 !ec!FtsZ assembled with a non
-hydrolysable GTP analog formed Z rings on tubular 
liposomes that began to constrict but the process is quickl
y halted 
(Osawa and 
Erickson, 2011)
.  In contrast to bacteria, the division of chloroplasts in plants and green algae 
requires the cooperation of two FtsZ families, FtsZ1 and FtsZ2 
(Osteryoung et al., 1998; 
Osteryoung and Vierling, 1995)
. FtsZ1 and FtsZ2 a
re highly conserved across all plants 
and green algae with sequenced genomes. Additionally, two FtsZ families, FtsZA and 
FtsZB, are highly conserved among red algae 
(Miyagishima and Kabeya, 2010; 
TerBush et al., 2013)
. FtsZ2 and FtsZA are believed to be mo
re ancestral, as they have 
retained a C
-terminal peptide, also present in bacterial FtsZs, that has been shown to 
anchor plant Z rings to the inner chloroplast membrane via interaction with the trans
-membrane protein ARC6 
(Maple et al., 2005; TerBush et al
., 2013; Vaughan et al., 
2004; Vitha et al., 2003)
. Therefore, both FtsZ2 and FtsZA likely evolved from a single 
FtsZ in the cyanobacterial ancestor of modern day chloroplasts, while FtsZ1 and FtsZB 
are thought to have evolved by duplication of the more an
cestral FtsZ2 in the green 
lineage and FtsZA in the red lineage, respectively 
(Miyagishima et al., 2004)
. FtsZ1 and 
FtsZ2 are nuclear
-encoded and translocated into the chloroplast stroma via an N
-terminal transit peptide, which is subsequently cleaved, whe
re they coassemble into the 
mid
-plastid Z ring 
(El-Kafafi et al., 2005; McAndrew et al., 2001; Vitha et al., 2001)
. Both 
FtsZ1 and FtsZ2 are critical for chloroplast division, as mutant plants lacking either FtsZ 
show severe defects in chloroplast morpholo
gy, indicated by larger and fewer 
chloroplasts per cell 
(Osteryoung et al., 1998)
. Other genetic evidence has shown that 
!ee!FtsZ1 and FtsZ2 from Arabidopsis are incapable of rescuing the otherÕs 
null mutant 
phenotype, indicating that they are not functionally
 redundant and possess distinct 
functions 
(Schmitz et al., 2009)
.  In vitro
, FtsZ1 and FtsZ2 assemble homopolymers and coassemble into thick 
bundles 
(El-Kafafi et al., 2005; Lohse et al., 2006; Olson et al., 2010; Smith et al., 2010)
. FtsZ1 and FtsZ2 both 
possess GTPase activity, with FtsZ1 activity being slightly greater 
than that of FtsZ2 
(Olson et al., 2010; Smith et al., 2010)
. Recently, we expressed FtsZ1 
and FtsZ2 fluorescent fusion proteins in the fission yeast 
Schizosaccharomyces pombe
 to analyze th
eir inherent assembly properties in a model cellular environment 
(TerBush 
and Osteryoung, 2012)
. We found that FtsZ1 and FtsZ2 have distinct filament 
morphology and dynamic properties. FtsZ1 filaments are more dynamic, with greater 
rates and extents of sub
unit exchange (turnover), and are able to increase the turnover 
of FtsZ2 in presumed heteropolymers, suggesting that FtsZ1 facilitates Z
-ring 
remodeling. FtsZ2 determines the morphology of coassembled filaments, suggesting 
that FtsZ2 has structural dominan
ce over FtsZ1 in mixed heteropolymers and therefore 
is the structural determinant of Z rings. Recent 
in vivo
 analysis of FtsZ1 and FtsZ2 
turnover in live chloroplast Z rings found that FtsZ1 has greater dynamic properties than 
FtsZ2 
(Johnson et al., 2015b)
, indicating that results in 
S. pombe 
accurately reflect 
behavioral trends 
in vivo
.  Until now, the vast majority of functional work on eukaryotic FtsZs has been 
performed on those from Arabidopsis. Due to the high conservation of two FtsZ families 
across 
both the green and red photosynthetic lineages, it is of great interest to study the 
!ed!properties of FtsZs from different species to better understand the functional 
significance of the FtsZ duplication events and whether both families of FtsZ possess 
conser
ved features across a broad evolutionary distance. Therefore, in this study, we 
continue to utilize 
S. pombe
 to investigate the assembly and dynamic properties of FtsZ 
proteins from species representing dicot and monocot plants, green algae, red algae, 
and cyanobacteria. We show that distinct assembly and dynamic properties for the 
FtsZ1 and FtsZ2 families are maintained across all species analyzed, including 
structural dominance of FtsZ2 and the higher turnover characteristics of FtsZ1. 
Additionally, we sh
ow that cyanobacterial FtsZ behavior is consistent with that of the 
eukaryotic FtsZ2 proteins, giving support to the theory that FtsZ2 is more ancestral and 
FtsZ1 arose from gene duplication and then diverged in functional properties.
   !e_!Results
 Conservation of distinct filament morphologies across the evolutionary lineages
  In our previous publication 
(TerBush and Osteryoung, 2012)
, we expressed 
FtsZ1 and FtsZ2 from Arabidopsis as fluorescent fusions to eYFP and eCFP, 
respectively. However, these
 fluorescent fusion proteins have a tendency to dimerize at 
high concentrations 
(Zacharias et al., 2002)
, potentially skewing the results away from 
the true inherent characteristics of FtsZ proteins. Therefore, in this study we express 
FtsZ1 and FtsZ2 repr
esentatives as C
-terminal fusions to a monomeric variant of the 
yellow fluorescent protein Venus (mVenus) and to the monomeric cyan fluorescent 
protein mCerulean, respectively. Both mVenus and mCerulean harbor the A206K 
mutation, which has been shown to el
iminate the dimerization potential of GFP
-derived 
fluorescent proteins 
(Zacharias et al., 2002)
, which should eliminate the possibility that 
fluorescent protein
-induced dimerization might affect the analyzed properties of FtsZs in 
S. pombe
.  To begin our a
nalysis, we expressed each FtsZ1 and FtsZ2 protein separately in 
S. pombe
 to analyze their intrinsic filament morphologies (Fig. 3.1). 
Arabidopsis thaliana
 FtsZ1 (AtFtsZ1), representing dicot plants, assembled into long filaments that followed 
the interior
 contours of the cell and annealed end
-to-end to form large rings that split 
into thinner filaments with a less intense fluorescence signal (Fig. 3.1 A), as seen 
previously when AtFtsZ1
-eYFP was expressed in 
S. pombe
 (TerBush and Osteryoung, 
2012). FtsZ1 f
rom the monocot plant 
Oryza sativa
 (OsFtsZ1), or rice, assembled into 
long cables that possessed flared ends (Fig. 3.1 B) and did not anneal into large rings to 
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SC%<=>6<78Q!X?"D?0)
SC%<=>6<78Q!78;!C
%<=>6<78!J6>E=<?G<8G<!
?:H876?!7=<!J76?<65!GE6E=<;!C7H<8D7P!27?@<;!6:8<?!=<I=<?<8D!D@<!G<66!E>D6:8<?P!47=?Q!N!
iCP
!!the same degree as AtFtsZ1. FtsZ1 from 
Ostreococcus tauri
 (OtFtsZ1), a green alga, 
assembled into long filaments that annealed to rings and la
terally associated into 
bundles that commonly split and frayed into loosely associated filaments (Fig. 3.1 C). 
FtsZB from 
Galdieria sulphuraria
 (GsFtsZB), a red alga, also assembled into long 
filaments and loosely associated bundled structures (Fig. 3.1 D)
 similar to those of 
OtFtsZ1. However, due to lower sequence similarity and poor transit peptide prediction 
tools,
 GsFtsZB and GsFtsZA, described below, expressed here were constructed 
without the N-terminal region.
  AtFtsZ2 assembled into large ring
-shape
d structures with smaller filaments
 breaking from the main ring (Fig. 3.1 E, arrow left panel). This is in contrast to the 
elaborate networks assembled by AtFtsZ2
-eCFP 
(TerBush and Osteryoung, 2012)
, which is likely due to the fusion to monomeric mCerulean
 used here. OsFtsZ2 
assembled into large rings with cross
-linking filaments protruding from the main ring 
structures, and into rings 
roughly 2
 "m in diameter
 (Fig. 3.1 F). OtFtsZ2 expressed in
 S. 
pombe
 failed to assemble any filamentous structures. Instead it adopted either a diffuse 
localization pattern or seemed to aggregate into spot
-like structures
 whether expressed 
alone or co
expressed with 
OtFtsZ1
 (Fig. 3.2 A &
 B). Western blot analysis
 with a !d3!mon
oclonal antibody recognizing GFP derivatives
 showed single bands for OtFtsZ1
-mVenus and OtFtsZ2
-mCerulean
 at their predicted molecular masses (Fig. 3.2 C), 
  9+KG@)'VISF'6R?@)BB+/0'/Q'
5)+"4'6'667).+#7"%
'9.BCS
'+0'12.('3$4
I!U)S4V!AI:J6>E=<?G<8G<!C:G=EH=7I@?!EJ!
'L#1%/2"#
G<66?!<WI=<??:8H!U)V!
(D"D?0
3SC%<=>6<78!E=!
U4V!(D
"D?01
SCb<8>?
!GE<WI=<??<;!R:D@!(D"D?03
SC%<=>6<78P!(D"D?01
SCb<8>?!78;!
(D"D?03
SC%<=>6<78!R<=<!J76?<65!GE6E=<;!H=<<8!78;!C7H<8D7Q!=<?I<GD:9<65P!&@<!R@:D<!
GE6E=!:8!D@<!C<=H<
;!:C7H<!:8;:G7D<?!=<H:E8?!R@<=<!D@<!J6>E=<?G<8G<!?:H876?!GE6EG76:T<P!
27?@<;!6:8<?!=<I=<?<8D!D@<!G<66!E>D6:8<P!
47=?Q!N!iCP
!U%V!
'CC>8EB6ED!78765?:?
!EJ!?E6>B6<!
B>6O!G>6D>=<!<WD=7GD
?!DE!;<D<=C:8<!D@<!CE6<G>67=!C7??<?
!EJ!)D"D?03
SC%<=>6<78!?:8H65!
<WI=<??<;!U6<JDV!78;
!(D"D?01
SCb<8>?!78;!(D"D?03
SC%<=>6<78
!GE<WI=<??<;
!UC:;;6<!
78;!=:H@DV
!:8!'L#1%/2"
P!A7G@!678<!=<I=<?<8D?!78!:8;<I<8;<8D65!H=ER8!G>6D>=<P
!&@<!B6ED!!!da!9+KG@)'VIS'W,/0.X*YF'
R7?!I=EB<;!R:D@!7!CE8EG6E876!uX"/P!
&@<!B78;?!=<I=<?<8D:8H!
(D"D?01
SCb<8>?!UmcN!O2Q!BEDDEC!7==ERV!78;!(D"D?03
SC%<=>6<78!Umc_!O2Q!DEI!7==ERV!
=78!?6:H@D65!6ER<=!E8!D@<!B6ED!D@78!D@7D!EJ!)D"D?03
SC%<=>6<78!Ume3!O2Q!IE?:D:9<!GE8D=E6VP!
)II=EW:C7D<!CE6<G>67=!C7??<?!:8!O2!7=<!?@ER8!DE!D@<!6<JDP
! indicating 
the fluorescence 
signals
 represent
 full-length OtFtsZ2
-mCerulean
. The OtFtsZ2
 morphology is radically different from that of any FtsZ that has been previously 
visualized 
in vivo 
(Fu et al., 2010; Kiessling et al., 2004; Li et al., 2007; Mingorance et 
al., 2005; Miyagishima et al., 2012; Strauss et al., 2012; Vitha et al., 2001)
 or in 
S. 
pombe
 (Srinivasan et al., 2007; Srinivasan et al., 2008; TerBush and Osteryoung, 
2012). Therefore, we will not pursue OtFtsZ2 in further analysis, due to the unknown 
cause of 
its peculiar assembly characteristics. Finally, GsFtsZA assembled into large 
rings with filament splitting and cross
-linking (Fig. 3.1 G). In control experiments, 
unfused mVenus and mCerulean adopted completely diffuse localization patterns (Fig. 
3.1 H-I).
 Taken together, the single expression data indicate that, while there are slight 
differences in morphology from one species to the next, the distinct long cables for 
FtsZ1/FtsZB and rings of FtsZ2/FtsZA are well conserved from plants to green, and red 
algae. Colocalization of FtsZ pairs
  FtsZ pairs invariably colocalize to mid
-plastid Z rings in wild type plants and to 
abnormal filaments in various chloroplast division mutants 
(McAndrew et al., 2001; Vitha 
et al., 2001). Additionally, FtsZ1 and FtsZ2 from 
Arabidopsis coassembled into bundled 
!d`!heteropolymers 
in vitro
 and colocalized when previously coexpressed as FtsZ1
-eYFP 
and FtsZ2
-eCFP fusions in 
S. pombe
 (Olson et al., 2010; TerBush and Osteryoung, 
 2012). To assess if this behavior is consistent with Arabidopsis FtsZ1
-mVenus and 
FtsZ2
-mCerulean, and conserved among the additional FtsZ pairs being analyzed here, 
we coexpressed each FtsZ pair in 
S. pombe
 to analyze their coassembled filament 
morphology char
acteristics and degree of colocalization.
  When coexpressed, AtFtsZ1 and AtFtsZ2 coassembled into ring structures that 
had cross
-linking filaments and regions where the ring split into smaller filaments (
Fig. 3.3 A). We used the PearsonÕs correlation coeff
icient to quantify the degree of 
fluorescence signal overlap and how well 
the signal intensities were correlated 
(Bolte 
and Cordeli‘res, 2006)
. AtFtsZ1 and AtFtsZ2 displayed a high PCC of 0.92 
± 0.01 (mean 
± SEM, n = 23), which indicates that the fluoresce
nce signals were extensively 
colocalized and that the signal intensities were directly correlated to each other. 
Similarly, OsFtsZ1 and OsFtsZ2 also colocalized to large and small rings that were 
highly similar to those assembled when OsFtsZ2 was expressed
 alone (Fig. 3.3 B). 
OsFtsZ1 and OsFtsZ2 also displayed a very high PCC value of 0.92 
± 0.01 (n = 9). In 
contrast, GsFtsZA and GsFtsZB colocalized to large loosely bundled rings (
Fig. 3.3 C, arrow) and a network of interconnected filaments. The overall mor
phology of the 
colocalized filaments resembled a hybrid of between GsFtsZA and GsFtsZB expressed 
alone (Fig. 3.1 D & G). However, consistent with the other FtsZ pairs, GsFtsZA and 
GsFtsZB displayed a very high PCC value of 0.91 
± 0.01 (n = 17).
 ! !dN! Figure 3.3: 
FtsZ1 and FtsZ2 proteins from plants colocalize into FtsZ2
-like rings while FtsZA and FtsZB coassemble into a hybrid filament network.
 Epifluorescence 
micrographs of 
S. pombe
 cells coexpressing (A) AtFtsZ1
-mVenus and AtFtsZ2
-mCerulean, (B) OsFtsZ1
-mVenus and OsFtsZ2
-mCerulean, and (C) GsFtsZB
-mVenus 
and GsFtsZA
-mCerulean. AtFtsZ1
-mVenus, OsFtsZ1
-mVenus, and GsFtsZB
-mVenus 
were falsely colored green. AtFtsZ2
-mCerulean, OsFtsZ2
-mCerulean, and GsFtsZA
-mCerulean were falsely colored magenta. The white colo
r in merged images indicates 
 !dc!Figure 3.3 (contÕd): 
regions where the fluorescence signals colocalize. Dashed lines 
represent the cell outline. Bars, 5 
"m.   Our coexpression data indicate that, at least in plant FtsZ pairs, FtsZ2 maintains 
structural domin
ance over FtsZ1 when these proteins coassemble into presumed 
heteropolymers. The data also suggest that the FtsZ2 structural dominance may be lost 
in red algal FtsZA. 
However, 
due to the lack of the N
-terminal region in the GsFtsZ 
constructs analyzed here,
 the possibility remains that this region is required for GsFtsZA 
to exhibit structural dominance over GsFtsZB.
 Conservation of distinct FtsZ1 and FtsZ2 polymer turnover characteristics
  We also analyzed the subunit exchange properties of FtsZ filaments us
ing fluorescence recovery after photobleaching (FRAP) experiments. When expressed 
individually, all FtsZ proteins displayed fluorescence recovery back into the bleached 
region (
Fig. 3.4). AtFtsZ1 recovered with a half
-time, the time it takes for the 
fluore
scence signal to recover to one half of its total recovery (t
1/2
), of 51 s and to 53% 
of the pre
-bleach prebleach intensity (n = 16, 
Fig. 3.4 A). OsFtsZ1 fluorescence signal 
recovered with a t
1/2
 of 74 s and to 45% of the pre
-bleach intensity (n = 13, 
Fig. 3.4 B). 
OtFtsZ1 recovered with a t
1/2
 of 101 s and to a maximum recovery of 45% (n = 16, 
Fig. 3.4 C). GsFtsZB recovered with a t
1/2
 of 54 s and to a maximum recovery of 61% (n = 
14, Fig. 3.4 D). AtFtsZ2 filaments displayed much less turnover than those co
mposed of 
AtFtsZ1 with a t
1/2
 of 67 s and to 13% of the pre
-bleach intensity (n = 16, 
Fig. 3.4 E). 
OsFtsZ2 filaments recovered with a t
1/2
 of 95 s and with a total recovery of 35
 % (n = 12, 
  !de!Figure 3.4: 
Greater polymer turnover 
dynamics in FtsZ1 are conserved across the 
green and red photosynthetic lineages.
 Cells expressing (A) AtFtsZ1
-mVenus, (B) 
 !dd!Figure 3.4 (contÕd): 
OsFtsZ1
-mVenus, (C) OtFtsZ1
-mVenus, (D) GsFtsZB
-mVenus, 
(E) AtFtsZ2
-mCerulean, (F) OsFtsZ2
-mCerulean, (G) GsFtsZA
-mCeru
lean, and (H) 
GsFtsZB coexpressed with (I) GsFtsZA were analyzed by FRAP. From left to right, 
images represent fluorescence signal at the location of photobleaching (dashed circle) 
prior to bleaching (PB), at the time of bleaching (0 s), the time closest t
o the t1/2
, and the time closest to twice that of t
1/2
. The average fluorescence recovery vs time plots are 
show at right. Data in each graph were normalized to the PB fluorescence signal (1 on 
the y-axis) and the fluorescence signal at the time of photobl
eaching (0 on the y
-axis). 
Data shown in the boxes show the calculated t
1/2
 (also indicated by the vertical black bar 
in the graph), maximum extent of recovery, and the number of independent FRAP 
experiments performed for each expression strain. The red br
ackets represent SEM at 
each time point. AtFtsZ1
-mVenus, OsFtsZ1
-mVenus, OtFtsZ1
-mVenus, and GsFtsZB
-mVenus fluorescence signals are falsely colored green. AtFtsZ2
-mCerulean, OsFtsZ2
-mCerulean, and GsFtsZA
-mCerulean fluorescence signals are falsely colored
 magenta. 
Bars, 2 
"m.  Fig. 3.4 F). GsFtsZA recovered with a t
1/2
 of 70 s and a total extent of recovery of 16% 
(n = 12, 
Fig. 3.4 G).
 We also performed FRAP experiments on coassembled GsFtsZs to assess if 
GsFtsZB was able enhance the turnover of GsFtsZA, a
s previously shown for AtFtsZ1 
and AtFtsZ2 
(TerBush and Osteryoung, 2012)
. GsFtsZB recovered with a t
1/2
 of 52 s 
and to 75% of the pre
-bleach intensity (n = 10, 
Fig. 3.4 H). Similarly, GsFtsZA recovered 
!d_!with a t1/2
 of 74 s and to 73% of the pre
-bleach intensity (n = 10, 
Fig. 3.4 I). In this 
coexpression strain, the rates of GsFtsZA and GsFtsZB turnover were similar to those 
in the single
-expression strains. More importantly, the maximum extent of fluorescence 
recover
y for GsFtsZA was greatly increased in the presence of GsFtsZB, similar to the 
effect of AtFtsZ1 on AtFtsZ2 
(TerBush and Osteryoung, 2012)
. When coexpressed, 
OsFtsZ1 and OsFtsZ2 showed reduced expression in terms of the number of cells with 
visible coassem
bled filaments and fluorescence intensity, 
potentially 
due to codon 
usage in the rice FtsZ proteins not being optimal for 
expression
 in S. pombe
 (Fig. 3.3 B). Therefore, this coexpression strain was omitted from FRAP experiments due to 
these technical chal
lenges and lack of confidence in the filament turnover data
. Overall, the FtsZ dynamics data indicate that both faster rates and greater 
extents of total recovery observed for Arabidopsis FtsZ1 than FtsZ2 filaments are 
conserved traits across both the gree
n and red photosynthetic lineages (
Fig. 3.5). 
Additionally, the ability of GsFtsZB to enhance the turnover of GsFtsZA in coassembled 
filaments shows that this is a trait also conserved across both the green and red 
photosynthetic lineages. All FRAP data ar
e summarized in Table 3.1.
   !_M!!Figure 3.5: 
FtsZ1 
and FtsZB homo
polymers are more dynamic than FtsZ2
 and FtsZA homopolymers
. The average fluorescence recovery vs time plots comparing the 
normalized recovery of (A) AtFtsZ1
-mVenus and AtFtsZ2
-mCerulean, (B) O
sFtsZ1
-mVenus and OsFtsZ2
-mCerulean, and (C) GsFtsZA
-mCerulean and GsFtsZB
-mVenus. 
Data in each graph were normalized to the PB fluorescence signal (1 on the y
-axis) and 
 !_1!Figure 3.5 (contÕd): 
the fluorescence signal at the time of photobleaching (0 on the 
y-axis). AtFtsZ1
-mVenus, OsFtsZ1
-mVenus, and GsFtsZB
-mVenus data are represented 
by red circles. AtFtsZ2
-mCerulean, OsFtsZ2
-mCerulean, and GsFtsZA
-mCerulean data 
are represented by green diamonds. The brackets in the datasets represent the 
standard error o
f the mean (SEM) for the normalized recovery at every time
-point.  Protein
 t1/2
 (s)
 % Recovery
 n AtFtsZ1
-mVenus
 51 53% 16 OsFtsZ1
-mVenus
 74 45% 13 OsFtsZ1
D249A
-mVenus
 101 21% 13 OtFtsZ1
-mVenus
 101 45% 16 OtFtsZ1
D224A
-mVenus
 61 20% 14 GsFtsZB
-mVenus
 54 61% 14 GsFtsZB
D300A
-mVenus
 87 35% 13 AtFtsZ2
-mCerulean
 67 13% 16 OsFtsZ2
-mCerulean
 95 35% 12 OsFtsZ2
D300A
-mCerulean
 372 30% 11 GsFtsZA
-mCerulean
 70 16% 12 GsFtsZA
D322A
-mCerulean
 51 10% 11 SeFtsZ
-msfGFP
 113 29% 14 SeFtsZ
D239A
-msfGFP
 46 8% 15 GsFtsZB
-mVenus
 + GsFtsZA
-mCerulean
 52 75% 10 74 73 10  Table 3.1: FRAP data for all single and coexpression strains
. Time for the 
normalized and averaged half
-time of recovery (t
1/2
, s) and maximum percent recovery 
for each protein in each expression 
strain. n represents the number of independent 
FRAP experiments performed on each protein in each expression strain.
 !_3!Effect of GTPase activity on FtsZ filament morphology and turnover
 To further test 
the importance of
 GTPase activity 
for FtsZ 
filament 
morphology
 and turnover, we expressed mutant forms of each FtsZ from 
O. sativa
, O. tauri
, and G. sulphuraria
 in which the conserved T7 loop aspartate required for GTPase activity, but 
not GTP binding 
(Oliva et al., 2004; Scheffers et al., 2001)
 was changed
 to alanine. Mutation of this critical residue reduces GTPase activity of Arabidopsis and 
E. coli
 FtsZs 
to less than 10% and ~1% of their WT activities, respectively 
(Olson et al., 2010; Redick 
et al., 2005). Consistent with our previous findings for AtFts
Z1 and AtFtsZ2 
(TerBush 
and Osteryoung, 2012)
, all putative GTPase
-deficient FtsZs assembled into filamentous 
structures when expressed in 
S. pombe 
(Fig. 3.6). OsFtsZ1
D249A
 assembled into a single 
long filament that curved around the interior contours of t
he cell or multiple straight
 filaments that were loosely associated (
Fig. 3.6 A). Similarly, OtFtsZ1
D224A
 assembled
 multiple straight filaments that did not bend around the contours of the cell interior 
(Fig.
3.5 B). GsFtsZB
D300A
 assembled straight or curved filaments that did not loop 
around the cell interior and were commonly loosely associated filaments (
Fig. 3.6 C), 
 similar to those of OsFtsZ1
D224A
. OsFtsZ2
D300A
 assembled filaments and rings with 
regions of variable thickness
es and spot
-like structures (
Fig. 3.6 D), while GsFtsZA
D322A
 assembled large rings with regions of variable thickness, but were more frequently split 
and frayed into smaller filaments with less fluorescence intensity (
Fig. 3.6 E). [<!76?E!I<=JE=C<;!".)/!<W
I<=:C<8D?!DE!J>=D@<=!7??<??!@ER!X&/7?<!7GD:9:D5!
7JJ<GD?!D@<!J:67C<8D!D>=8E9<=!G@7=7GD<=:?D:G?!EJ!D@<!"D?0?!?D>;:<;!@<=<P!(?"D?01
23`_)
!=<GE9<=<;!R:D@!7!D
1Y3
!EJ!1M1!?!78;!7!DED76!<WD<8D!EJ!=<GE9<=5!EJ!31p!U
":HP!aPe!)Q!8!l!1aVP!
OtFtsZ1
D224A
 had a t1/2
 of 61 s and recovered to 20% of the pre
-bleach fluorescence 
  !_a! Figure 3.6: 
Putative GTPase
-deficient mutants display altered 
filament
 morphologies. Epifluorescence micrographs of 
S. pombe 
cells expressing (A) 
OsFtsZ1(D249A)
-mVenus, (B) OtFtsZ1
D224A
-mVenus, (C) G
sFtsZB
D300A
-mVenus, (D) 
OsFtsZ2
D300A
-mCerulean, and (E) GsFtsZA
D322A
-mCerulean. OsFtsZ1
D249A
-mVenus, 
OtFtsZ1
D224A
-mVenus, and GsFtsZB
D300A
-mVenus fluorescence signals are falsely 
colored green. OsFtsZ2
D300A
-mCerulean and GsFtsZA
D322A
-mCerulean fluorescence 
signals are falsely colored magenta. Dashed lines represent the cell outlines. Bars, 5 
"m. ! !_`! Figure 3.7: 
Putative FtsZ1 GTPase
-deficient mutants show reduced, yet not 
abolished, polymer turnover, while putative FtsZ2 GTPase
-defici
ent mutants lack 
polymer turnover.
 Cells expressing (A) OsFtsZ1
D249A
-mVenus, (B) OtFtsZ1
D224A
-mVenus, (C) GsFtsZB
D300A
-mVenus, (D) OsFtsZ2
D300A
-mCerulean, and (E) 
GsFtsZA
D322A
-mCerulean were analyzed by FRAP. From left to right, images represent 
fluorescen
ce signal at the location of photobleaching (dashed circle) prior to bleaching 
 !_N!Figure 3.7 (contÕd): 
(PB), at the time of bleaching (0 s), the time closest to the t
1/2
, and the time closest to twice that of t
1/2
. The average fluorescence recovery vs time p
lots are 
show at right. Data in each graph were normalized to the PB fluorescence signal (1 on 
the y-axis) and the fluorescence signal at the time of photobleaching (0 on the y
-axis). 
Data shown in the boxes show the calculated t
1/2
 (also indicated by the 
vertical black bar 
in the graph), maximum extent of recovery, and the number of independent FRAP 
experiments performed for each expression strain. The red brackets represent SEM at 
each time point. OsFtsZ1
D249A
-mVenus, OtFtsZ1
D224A
-mVenus, and GsFtsZB
D300A
-mVenus fluorescence signals are falsely colored green. OsFtsZ2
D300A
-mCerulean and 
GsFtsZA
D322A
-mCerulean fluorescence signals are falsely colored magenta. Bars, 2 
"m.  intensity (
Fig. 3.7 B, n = 14). GsFtsZB
D300A
 recovered with a t
1/2
 of 87 s and to a 
maximum extent of 35% (
Fig. 3.7 C, n = 13). OsFtsZ2
D300A
 filaments had severely 
reduced turnover with a t
1/2
 of 372 s and a maximum recovery of 30% (
Fig. 3.7 D, n = 
11). Likewise, GsFtsZA
D300A
 recovered with a t
1/2
 of 51 s and to 10% (
Fig. 3.7 E, n = 11).
  Taken together, these filament morphology and turnover experiments on putative 
GTPase
-deficient FtsZs show that GTPase activity is important for normal FtsZ 
assembly and turnover across various photosynthetic eukaryote lineages. More 
imp
ortantly, these data show that, consistent with our previous results for Arabidopsis 
FtsZ1 and FtsZ2 
(TerBush and Osteryoung, 2012)
, GTPase activity is critical for 
promoting filament turnover for FtsZ2s from diverse photosynthetic eukaryotes, but not 
sole
ly responsible for tur
nover of FtsZ1/FtsZB filaments.
!!_c!Gnalysis of cyanobacterial FtsZ filament morphology and turnover
  To assess the inherent assembly and dynamic properties of cyanobacterial FtsZ, 
we expressed the single FtsZ from 
Synechococcus elongatus
 PCC 7942
 (SeFtsZ) in 
S. 
pombe
 as a fluorescent fusion to monomeric superfolder GFP (msfGFP) 
(Costantini et 
al., 2012) and analyzed
 its filament morphology and turnover characteristics. SeFtsZ 
readily assembled into filaments in the 
S. pombe
 cytosol (
Fig. 3.8 A). It commonly 
!!Figure 3.8: 
Cyanobacterial FtsZ 
filament
 morphology shows characteristics of 
FtsZ1 and FtsZ2 families, but polymer turnover is 
similar to
 that of FtsZ2
 and FtsZA
. (A-C) Epifluorescence micrographs of 
S. pombe 
cells expressing (A) 
SeFtsZ
- !_e!Figure 3.8 (contÕd): 
msfGFP, (B) SeFtsZ(D239A)
-msfGFP, or (C) msfGFP. Dashed 
lines represent the cell outlines. (D
-E) Cells expressing (D) SeFtsZ
-msfGFP or (E) 
SeFtsZ(D239A)
-msfGFP were analyzed by FRAP. From left to right, images represent 
fluores
cence signal at the location of photobleaching (dashed circle) prior to bleaching 
(PB), at the time of bleaching (0 s), the time closest to the t
1/2
, and the time closest to 
twice that of t
1/2
. The average fluorescence recovery vs time plots are show at ri
ght. Data in each graph were normalized to the PB fluorescence signal (1 on the y
-axis) and 
the fluorescence signal at the time of photobleaching (0 on the y
-axis). Data shown in 
the boxes show the calculated t
1/2
 (also indicated by the vertical black bar 
in the graph), 
maximum extent of recovery, and the number of independent FRAP experiments 
performed for each expression strain. The red brackets represent SEM at each time 
point. SeFtsZ
-msfGFP, SeFtsZ(D239A)
-msfGFP, and msfGFP fluorescence signals are 
falsely colored magenta. White bars, 5 
"m. Grey bars, 2 
"m. !formed long cables that looped around the cell (
Fig. 3.8 A, left panel) and also large 
rings that split into loosely associated filaments (
Fig. 3.8 A, right panel). We also 
expressed SeFtsZ as a puta
tive GTPase
-deficient mutant bearing the aspartate
-to-alanine mutation in the conserved T7 loop (SeFtsZ
D239A
) to assess how GTPase activity 
affects SeFtsZ assembly characteristics. SeFtsZ
D239A
 assembled straight filaments that 
commonly were the length of t
he cell, but did not curve around the ends (
Fig. 3.8 B). In 
control experiments, msfGFP adopted a completely diffuse localization pattern (
Fig. 3.8 C). FRAP experiments showed that SeFtsZ fluorescence recovered back into the 
bleached region with a t
1/2
 of 113 s and to 29% of the pre
-bleach fluorescence intensity 
!_d!(Fig. 3.8 D, n = 14). In contrast, SeFtsZ
D239A
 had nearly abolished filament dynamics 
with a t1/2
 of 46 s but only recovering to 8% of the pre
-bleach fluorescence intensity (
Fig. 3.8 E, n = 15). The
se filament morphology and turnover data show that SeFtsZ 
possesses filament morphology properties of both FtsZ1/FtsZB and FtsZ2/FtsZA, but its 
filament turnover properties are highly similar to those of FtsZ2/FtsZA. The overall 
greater consistency of SeFt
sZ filament morphology and turnover properties with those 
of eukaryotic FtsZ2/FtsZA supports the theory that the FtsZ2 and FtsZA families 
evolved from the probable single FtsZ in the ancestor of chloroplasts and that the FtsZ1 
and FtsZB families evolved la
ter by gene duplication events, in their respective 
photosynthetic lineages, and followed by functional divergence.
 Previous 
data have shown that reversible photobleaching can 
greatly 
influence
 fluorescence recovery kinetics of proteins fused to GFP variants
 For example
 ion chann
el proteins
 previously shown to be immobile
 displayed 65.9% fluorescence 
recovery with a time constant of 0.36 s
 when fused to eCFP
 U,:88<GO<=!<D!76PQ!3MMNV
. Therefore
, in control experiments, we photobleached the entire contents of cells 
expressing a representative for each fluorescence protein used (mVenus, mCerulean, 
and msfGFP) and recorded fluorescence intensity over time to assess for the 
spontaneous recovery of ble
ached fluorescence proteins. In all cases, there was no 
substantial fluorescence recovery in the cells (
Fig. 3.9). Combined with the low 
fluorescence recovery and long t
1/2
 values
 of FtsZ proteins fused to mCerulean
, these 
data indicate that reversible pho
tobleaching do
es not influence the FtsZ dynamics data 
in this study.
  !__! Figure 3.9: Photobleached FtsZ fusions do not spontaneously recover 
fluorescence. 
Cells expressing (A) AtFtsZ1
-mVenus, (B) AtFtsZ2
-mCerulean, and (C) 
SeFtsZ
-msfGFP were analyzed by FRA
P. At the photobleaching event, the entire 
content of the cells were bleached to assess if the fluorescence proteins could 
spontaneously recover their fluorescence intensity. From left to right, images represent 
fluorescence signal in the cell prior to ble
aching (PB), at the time of bleaching (0 s), 100 
s post
-bleaching, and 200 s post
-bleaching. The average fluorescence recovery vs time 
plots are show at right. Data in each graph were normalized to the PB fluorescence 
signal (1 on the y
-axis) and the fluor
escence signal at the time of photobleaching (0 on 
the y-axis). The recovery data were the average of 3 independent FRAP experiments 
for each expression strain (n = 3). The red brackets represent SEM at each time point. 
 !1MM!Figure 3.9 (contÕd): 
AtFtsZ1
-mVenus
 fluorescence signal is falsely colored green. 
AtFtsZ2
-mCerulean and SeFtsZ
-msfGFP fluorescence signals are falsely colored 
magenta. Bars, 2 
"m. ''!1M1!Discussion  Studies analyzing the functional behavior of FtsZs from photosynthetic 
eukaryotes have begun to emerge 
(El-Kafafi et al., 2005; Johnson et al., 2015a; 
Johnson et al., 2015b; Olson et al., 2010; Smith et al., 2010; Smith et al., 2011; TerBush 
and Osteryoung
, 2012). However, the vast majority of these data are for FtsZ1 and 
FtsZ2 from Arabidopsis, and there have been no studies to compare the behavior of 
FtsZ pairs from different photosynthetic organisms, especially between FtsZs from the 
green and red photos
ynthetic lineages. Performing such a comparative analysis 
in vivo
 would be highly challenging and time
-consuming, as chloroplast division is highly 
sensitive to FtsZ expression level, FtsZ fluorescent fusions expressed from their native 
promoters produce l
ow fluorescence signal amid high background chlorophyll 
autofluorescence in photosynthetic tissues, and transformation procedures for algal 
species are not well established. To date, 
S. pombe
 has proven to be a useful and 
robust expression system for gaini
ng insight into the chloroplast division machinery, 
especially into the assembly and dynamics of FtsZs and the roles of the regulatory 
proteins that govern their assembly and positioning 
in planta (TerBush and Osteryoung, 
2012; Zhang et al., 2013). We have
 found that FtsZ1 and FtsZ2 from Arabidopsis 
assemble filaments and coassemble into presumed heteropolymers that undergo 
dynamic turnover. These results are highly consistent with previous findings from 
in vivo
 and in vitro
 experiments on plant FtsZs 
(El-Kafafi et al., 2005; Johnson et al., 2015b; 
McAndrew et al., 2001; Olson et al., 2010; Smith et al., 2010; TerBush and Osteryoung, 
2012; Vitha et al., 2001)
. A recent study analyzing the turnover of AtFtsZ1 and AtFtsZ2 
!1M3!in Z rings and filaments from live Ara
bidopsis epidermal cells showed that turnover rates 
were significantly longer than those we measure in 
S. pombe
, likely the result of 
elaborate mechanisms for positioning and regulating Z
-ring assembly and constriction 
(Johnson et al., 2015b; TerBush and O
steryoung, 2012)
. Nevertheless, in plants FtsZ1 
still displayed greater dynamics than FtsZ2, which suggests that the results in 
S. pombe
 reflect the overall 
in vivo 
behavioral trends of FtsZ1 and FtsZ2. Therefore, the continued 
use of 
S. pombe
 to analyze t
he inherent filament assembly and turnover properties of 
FtsZs from various photosynthetic organisms, in order to assess functional conservation 
and variation, is highly advantageous, as fission yeast are easily transformed, grow and 
express proteins at a 
rapid rate, and possess none of the proteins that interact with and 
modulate FtsZ behavior 
in planta. FtsZ filament morphology
 When comparing the Arabidopsis FtsZ1
-mVenus and FtsZ2
-mCerulean filaments in 
this study with the Arabidopsis FtsZ1
-eYFP and FtsZ2
-eCFP filaments in our previous 
study 
(TerBush and Osteryoung, 2012)
, there is a distinct change in morphology, 
especially for FtsZ2. Previously, FtsZ1
-eYFP assembled into large rings and single 
cables that looped around the cell compartment, while FtsZ2
-eCFP assembled into 
small rings and intricate networks of filaments with regions of variable thickness. In this 
study, FtsZ1
-mVenus assembled multiple cables that looped around the cell and 
occasionally into large rings, which likely formed by the annealing
 of cable ends (Fig. 
3.1 A). Additionally, FtsZ2
-mCerulean had a drastic shift in morphology, where it 
assembled into very regular rings, with a diameter the length of the cell interior 
!1Ma!compartment, that had smaller filaments protruding off of the main str
ucture (Fig. 3.1 E, 
left panel arrow). This shift in filament morphology was likely caused by changing from 
eYFP and eCFP, which are prone to dimerize at high concentrations in cellular 
environments, to mVenus and mCerulean bearing the A206K mutation, whic
h eliminates the dimer interface and produces truly monomeric fluorescent proteins 
(Rizzo 
and Piston, 2005; Zacharias et al., 2002)
. This is consistent with the more loosely 
bundled appearance of FtsZ1
-mVenus and may represent a more accurate filament 
morp
hology than observed in our previous study. However, the dynamic and 
colocalization properties of the AtFtsZ1
-mVenus and AtFtsZ2
-mCerulean are consistent 
with those of AtFtsZ1
-eYFP and AtFtsZ2
-eCFP, suggesting that the inherent behaviors 
of FtsZs fused to 
mVenus and mCerulean are consistent with those fused to eYFP and 
eCFP.
  As we discussed in TerBush and Osteryoung (2012), we assume that the 
filaments observed in 
S. pombe
 represent bundles of individual polymers. With this in 
mind, we observe differences 
in bundling in FtsZ1 proteins from the various species 
analyzed. The representatives from 
A. thaliana and O. sativa
 (rice) form relatively tightly 
bundled cables, albeit OsFtsZ1 cables possess flared ends. However, 
Ostreococcus 
tauri
 (green alga) FtsZ1 sho
ws more loosely associated bundles (Fig. 3.1 C), while 
Galdieria sulphuraria 
(red alga) FtsZB shows even less ability for individual polymers to
 laterally associate into bundles (Fig. 3.1 D). This reduced bundling activity is also 
evident in GsFtsZA, but t
o a lesser degree (Fig. 3.1 H). A previous study of 
E. coli
 and B. subtilis
 FtsZ showed that the C
-terminal variable region (CTV), which includes the 
!1M`!amino acids downstream of the C
-terminal peptide required for FtsZ interaction with 
membrane
-associating p
roteins, promoted lateral assembly of individual polymers into 
bundles based on its degree of positive charge 
(Buske and Levin, 2012)
. Upon analyzing a multiple sequence alignment (
Fig. 3.10), there seems to be a correlation in 
the appearance of FtsZ filam
ent bundling in S. pombe
 and the number of positively 
charged amino acids in their CTVs (Fig. 1.3, 
Fig. 3.10 magenta box). Both OtFtsZ1 and 
GsFtsZA have a reduced CTV compared to those in AtFtsZ1/OsFtsZ1 and 
AtFtsZ2/OsFtsZ2, respectively, and display a moderate degree of loose bundling. 
GsFtsZB lacks the C
-terminal region downstream of the conserved GTP
-binding and hydrolysis domain all together (
Fig. 3.10) and shows less filament bundling. Further 
studies on assembly and bundling activity 
in vitro
 will be necessary to test this point 
further, but our data do support a role for the CTV in promoting lateral intera
ctions and 
bundling of polymers based on the degree of positive charge in this region.
  We commonly observed that OsFtsZ2 assembled rings of two sizes, those that 
seemed more constrained by the interior dimensions of the yeast cell and those that 
were roug
hly 2 
"m in diameter. However, rings of both diameters were evident in the 
coexpression strains for Arabidopsis and rice FtsZ. Similar small rings have been 
observed in transgenic plants 
(Johnson et al., 2015a; Zhang et al., 2013)
, suggesting 
that the smal
l-diameter rings may represent an inherent curvature of plant FtsZ2 
proteins and that plant FtsZ2s have two distinct states of polymer geometry, a straight 
and more curved conformation. Similar curvature patterns have been observed for 
E. 
coli
 FtsZ when 
polymerized with GDP 
(Erickson et al., 2010; Lu and Erickson, 1999; 
  !1MN!!Figure 3.10: Multiple Sequence alignment of FtsZ proteins included in this survey. 
Multiple
 protein
 alignment
 was performed
 for FtsZ
 homologs
 from
 Arabidopsis
 thaliana  !1Mc!Figure 3.10 (cont
Õd):
 (FtsZ1
 - NP_200339.1
 and FtsZ2
 - NP_565839.1),
 Oryza
 sativa
 japonica
 (FtsZ1
 - EEE61856.1
 and FtsZ2
 - EEE59586.1),
 Ostreococcus
 tauri
 (FtsZ1
 - XP_003080788.1
 and FtsZ2
 - XP_003080256.1),
 Galdieria
 sulphuraria
 (FtsZB
 - BAA82091.1
 and FtsZA
 - BAA82090.1)
, and Synechococcus
 elongatus PCC
 7942 (FtsZ
 - AAC26227.1)
 using
 MAFFT
 E-INS-i online server
 (Katoh
 and Standley,
 2013) and graphic
 generated
 using
 ESPript
 3.0 with a %Equivalent
 and global score
 of 0.7 sequence
 similarity
 depiction
 parameter
 (Robert
 and Gouet, 2014). Alignment
 identified a highly conserved
 GTP binding domain
 (blue
 bar),
 moderately
 conserved
 C-terminal
 domain
 (green
 bar),
 and conserved
 C-terminal
 peptide among
 FtsZ2/FtsZA
 and SeFtsZ
 (grey
 box).
 The later is consistent
 with previous
 reports
 highlighting the absence
 of the C-terminal
 peptides in FtsZ1Õs
 of land plants and green
 algae (TerBush
 et al., 2013). Illustrative
 of FtsZ
 divergence,
 Galdieria
 sulphuraria
 FtsZB
 lacks
 a large
 portion
 of the C-terminal
 region
 while FtsZA
 possesses
 the highly conserved
 C-terminal
 peptide. The C-terminal
 variable
 region
 (CTV)
 (Buske
 and Levin,
 2012) is indicated
 by a magenta
 box.
  Romberg et al., 2001)
, suggesting that GTP hydrolysis may be a step in the process of 
inducing polymer bending. This is supported
 by a recent study suggesting that 
Mycobacterium tuberculosis
 FtsZ undergoes a hinge
-opening mechanism in response 
to GTP hydrolysis that induces protofilament bending 
(Li et al., 2013)
. However, another 
study on 
E. coli
 FtsZ showed that Z rings devoid of 
GTP hydrolysis assemble on tubular 
liposomes and start the constriction process, but that constriction is halted shortly 
thereafter, indicating that GTPase activity is not required for inducing constriction but for 
!1Me!sustaining it 
(Osawa and Erickson, 2011)
. Further structural analysis into inherent 
curvatures, mechanisms of protofilament bending, and force generation will be needed 
to clarify these issues for FtsZs from photosynthetic eukaryotes as well as bacteria.
  As we observed previously, AtFtsZ1 and At
FtsZ2 coassembled into filaments that 
were similar to filaments formed by AtFtsZ2 alone. This was also observed for the 
coassembly of OsFtsZ1 and OsFtsZ2, suggesting that FtsZ2 structural dominance is a 
trait conserved across plant FtsZs. However, GsFtsZA 
and GsFtsZB coassembled into 
filament networks that had characteristics of both FtsZs but were more similar to those 
of GsFtsZB, including the ability to assemble large rings and loosely bundled filaments. 
However, due to lack of primary sequence conservat
ion and poor transit peptide 
prediction tools, GsFtsZA and GsFtsZB were expressed with the N
-terminal regions 
upstream of their conserved GTP
-binding and hydrolysis domains removed. The 
assembly data for these proteins suggest that FtsZ2 structural dominan
ce is a feature 
conserved among FtsZ pairs from both the green and red photosynthetic lineages. 
Additionally, they also suggest that the N
-terminal region of FtsZs may influence FtsZ2Õs 
structural dominance over FtsZ1.
 FtsZ 
filament
 turnover
  All FtsZs in 
this study assembled filaments that underwent dynamic turnover. 
Invariably, the FtsZ1 representative from each pair displayed greater turnover 
dynamics, i.e., more rapid rates and greater total extents of recovery, indicating that this 
is a conserved trait
 for FtsZ pairs across both the green and red photosynthetic lineages 
(Fig. 3.5). GsFtsZB possessed the greatest filament turnover dynamics of any FtsZ 
!1Md!studied here. The high dynamic character coincided with the weakest apparent lateral 
interactions, sugge
sting the possibility that lateral packing of polymers may have a 
significant impact on the diffusion of assembly subunits into and out of bundles. 
However, the extent to which lateral interactions between polymers affect subunit 
diffusion may not be signi
ficant, as AtFtsZ1 displayed the highest degree of bundling of 
any FtsZ1/FtsZB studied here, yet filament turnover was near that of GsFtsZB. These 
data suggest that FtsZ filament turnover may be insensitive to the extent of lateral 
association and more inf
luenced by another factor, such as FtsZ subunit interface 
stability. This hypothesis is also supported by GsFtsZA assembling filaments with the 
weakest apparent lateral interactions of any FtsZ2/FtsZA in this study, yet showed very 
low turnover properties.
 An alternative hypothesis is that the GsFtsZB studied here 
lacks its N
-terminal region upstream of the conserved GTP
-binding and hydrolysis 
domains and natively does not possess a C
-terminal tail, which is found in all other 
FtsZs studied. This idea sugge
sts the possibility that the N
- and C-terminal regions may 
be important for promoting polymer stability and lateral interactions leading to bundling, 
as discussed above. In either case, additional studies will be required to verify the 
validity of these hy
potheses by testing for subunit interface stability and the effect of the 
N- and C-terminal regions on FtsZ filament
 turnover and lateral association of individual 
polymers.
 All putative GTPase
-deficient FtsZs displayed reduced filament
 turnover 
characteri
stics compared to their WT counterparts (
Fig. 3.11). While differences in the 
specific t
1/2
 and total extent of recoveries were present in putative GTPase
-deficient 
  !1M_! Figure 3.11: Putative GTPase
-deficient FtsZ mutants have reduced polymer 
turnover dynamics than that of 
their WT FtsZ counterpart
. The average 
fluorescence recovery vs time plots comparing the normalized recovery of (A) OsFtsZ1
-mVenus and OsFtsZ1
D249A
-mVen
us, (B) OtFtsZ1
-mVenus and OtFtsZ1
D224A
-mVenus, 
and (C) GsFtsZB
-mVenus and GsFtsZB
D300A
-mVenus, (D) OsFtsZ2
-mCerulean and 
 !11M!Figure 3.11 (contÕd): 
OsFtsZ2
D300A
-mCerulean, (E) GsFtsZA
-mCerulean and 
GsFtsZA
D322A
-mCerulean, and (F) SeFtsZ
-msfGFP and SeFtsZ
D239A
-msfGFP. Data in 
each graph were normalized to the PB fluorescence signal (1 on the y
-axis) and the 
fluorescence signal at the time of photobleaching (0 on the y
-axis). OsFtsZ1
-mVenus, 
OtFtsZ1
-mVenus, GsFtsZB
-mVenus, OsFtsZ2
-mCerulean, GsFtsZA
-mCerulean, a
nd SeFtsZ
-msfGFP data are represented by red circles. OsFtsZ1
D249A
-mVenus, 
OtFtsZ1
D224A
-mVenus, GsFtsZB
D300A
-mVenus, OsFtsZ2
D300A
-mCerulean, GsFtsZA
D322A
-mCerulean, and SeFtsZ
D239A
-msfGFP data are represented by green diamonds. The 
brackets represent SEM a
t each time point.
  FtsZs from one species to the next, the overall findings were that FtsZ1/FtsZB filaments
 retained a significant degree of turnover, while FtsZ2/FtsZA filament
 turnover was 
nearly abolished across FtsZs from all species analyzed. These d
ata are highly 
consistent with our previous results for AtFtsZ1
-eYFP and AtFtsZ2
-eCFP 
(TerBush and 
Osteryoung, 2012)
 and suggest that the hypothesis that the FtsZ1
-FtsZ1 subunit 
interface is weak and prone to breakage, even in the absence of GTPase activit
y, may 
be extended to FtsZ1/FtsZB proteins across the green and red photosynthetic lineages. 
Similarly, these data suggest that the hypothesized FtsZ2
-FtsZ2 strong subunit interface 
that is highly dependent on GTP hydrolysis for destabilization may also be
 extended.
  In our previous study analyzing Arabidopsis FtsZ1 and FtsZ2, one of the critical 
findings was that AtFtsZ1 was able to enhance the turnover properties of AtFtsZ2 in 
presumed heteropolymers 
(TerBush and Osteryoung, 2012)
. Therefore, it was of gr
eat !111!evolutionary interest to assess if this is a conserved trait across the green and red 
photosynthetic lineages. In this study, we found that the extent of fluorescence recovery 
of the FtsZ2
-like protein in the red alga 
Galdieria sulphuraria
, GsFtsZA, wa
s greatly 
enhanced in the presence of GsFtsZB relative to when GsFtsZA was expressed alone. 
However, the rate of filament
 turnover was unaffected. This is highly consistent with our 
previous findings for the effect of AtFtsZ1 on AtFtsZ2 turnover. These dat
a indicate that 
the ability of FtsZ1 to augment the turnover of FtsZ2 in heteropolymers, and therefore 
potentially facilitate Z
-ring constriction by enhancing remodeling, is a conserved trait 
across photosynthetic eukaryotes, even with the presumed indepen
dent gene duplication events that produced FtsZ1 and FtsZB in the green and red photosynthetic 
lineages, respectively, after their divergence 
(Miyagishima et al., 2004)
. FtsZ evolution
 The prevailing evolutionary theory states that chloroplasts arose when 
a free
-living cyanobacterium was engulfed by a heterotrophic eukaryote and was subsequently 
enslaved, evolving into the organelles we see today 
(Gould et al., 2008)
. Further, it is 
believed that green
-lineage FtsZ2 and red
-lineage FtsZA are more ancient an
d are 
homologs of the single FtsZ in the common ancestor of chloroplasts. FtsZ1 and FtsZB 
arose later from independent gene duplication events in the green and red 
photosynthetic lineages, respectively 
(Miyagishima et al., 2004)
. These hypotheses are 
large
ly based on sequence homology and there has been no functional analysis for 
cyanobacterial FtsZ proteins to date. Therefore, this comparative analysis is a major 
!113!first step in elucidating the behaviors of cyanobacterial FtsZ and how they relate to 
FtsZs fr
om photosynthetic eukaryotes.
  SeFtsZ assembled well in 
S. pombe
, possessing filament morphology that 
resembled the characteristics of both FtsZ1/FtsZB and FtsZ2/FtsZA families. The long 
looping cables formed by the WT protein and straight filaments formed
 by the putative 
GTPase
-deficient mutant were highly consistent with filament morphologies for 
FtsZ1/FtsZB. SeFtsZ also assembled large rings with loosely associated bundles similar 
to those of FtsZ2/FtsZA. However, FRAP experiments show that SeFtsZ proper
ties are 
completely consistent with those of the FtsZ2/FtsZA, as shown by a slow rate coupled 
with a low extent of turnover of the WT protein and the loss of filament
 turnover by the 
putative GTPase
-deficient mutant. Overall, the higher functional similari
ty between 
SeFtsZ and FtsZ2/FtsZA is consistent with the theory that FtsZ2 and FtsZA are the 
direct homologs of the FtsZ present in the cyanobacterial endosymbiont and that 
FtsZ1/FtsZB arose later from gene duplication events.
 Conclusions
  Taken together, the comparative analyses presented in this study expand our 
understanding of how FtsZ function has been conserved across the photosynthetic 
lineages. Although minor differences in filament morphology and turnover exist from one 
species to t
he next, trends in the major behaviors of the FtsZ1/FtsZB and FtsZ2/FtsZA 
families persist, strongly suggesting that the distinct functions of FtsZ1/FtsZB promoting 
Z-ring constriction by enhancing remodeling and of FtsZ2/FtsZA as the structural 
determinan
ts of the Z ring are well conserved and important for of Z
-ring activity across 
!11a!the green and red photosynthetic lineages. Additionally, this work provides the first 
functional characterization of a cyanobacterial FtsZ and supports the current theory that
 the direct homologs of the FtsZ from the endosymbiotic ancestor of chloroplasts are 
FtsZ2 and FtsZA, for the green and red photosynthetic lineages, respectively. These 
data also support that FtsZ1 and FtsZB arose later in their respective lineages and 
underwent convergent evolution to develop their unique function in Z
-ring constriction.
  !11`!Materials and Methods
 Cloning of S. pombe Expression Constructs
 FtsZ genes were cloned into the 
pREP41X
 and pREP42X
 expression vectors 
under the control of the 
nmt1*
 promo
ter for analysis in 
S. pombe
 (Basi et al., 1993; 
Forsburg, 1993; Maundrell, 1993)
. Sequences encoding 
Arabidopsis thaliana
 FtsZ1
 (NP_200339.1) and 
FtsZ2
 (NP_565839.1) lacking the predicted transit peptide (the first 
57 and 48 amino acids, respectively) wer
e PCR
-amplified from corresponding 
S. pombe
 expression plasmids 
(TerBush and Osteryoung, 2012)
 using primers 
AT262/AT298 and 
AT264/AT299, respectively
. A monomeric variant of the yellow fluorescent protein 
Venus
 (mVenus) bearing the A206K mutation shown to
 eliminate the dimerization 
potential of GFP-derived fluorescent proteins 
(Nagai et al., 2002; Zacharias et al., 
2002), generated in a bacterial expression vector provided by Desmond A. Moore and 
Harold P. Erickson (Duke University, Durham, NC), was PCR am
plified with primers 
AT7/AT297
. mCerulean
 (Addgene, Cambridge, MA) was PCR
-amplified with primers 
AT7/AT297
. AtFtsZ1
 and mVenus
 were cloned into 
pREP41X
 digested with BamH1 by 
the Gibson Assembly method 
(Gibson et al., 2009)
 to create 
pREP41X
-AtFtsZ1
-mVenu
s. AtFtsZ2
 and mCerulean
 were cloned into 
pREP42X
 digested with BamH1 by 
the Gibson Assembly method to create pREP42X
-AtFtsZ2
-mCerulean
. We had the additional eukaryotic FtsZ 
CDS from 
Oryza sativa
, Ostreococcus tauri
, and Galdieria 
sulphuraria
 synthesized with codon optimization for expression in 
Arabidopsis thaliana
 from Integrated DNA Technologies (Coralville, IA) or Invitrogen (Carlsbad, CA)
. The sequence encoding 
OsFtsZ1 (EEE61856.1) lacking the predicted transit peptide (the 
!11N!first 35 amino
 acids) was PCR
-amplified with primers AT325/AT326 and cloned
 with mVenus
 into the same 
pREP41X
 digested with BamH1 with the Gibson assembly 
method to generate 
pREP41X
-OsFtsZ1
-mVenus
. The sequence encoding OsFtsZ2 
(EEE59586.1) lacking the 
predicted transit
 peptide (the first 20 amino acids) was PCR
-amplified with primers AT327/AT328 and cloned with 
mCerulean
 into the same 
pREP42X
 digested with BamH1 with the Gibson assembly method to generate 
pREP42X
-OsFtsZ2
-mCerulean
. The sequence encoding 
OtFtsZ1 (XP_0030
80788.1) 
lacking the predicted transit peptide (the first 18 amino acids) was PCR
-amplified with 
primers AT329/AT330 and cloned with 
mVenus
 into the same 
pREP41X
 digested with 
BamH1 with the Gibson assembly method to generate 
pREP41X
-OtFtsZ1
-mVenus
. The sequence encoding 
OtFtsZ2 (XP_003080256.1) lacking the predicted transit peptide 
(the first 35 amino acids) was PCR
-amplified with primers AT331/AT332 and
 cloned 
with mCerulean
 into the same 
pREP42X
 digested with BamH1 with the Gibson 
assembly method to gene
rate 
pREP42X
-OtFtsZ2
-mCerulean
. Due to the lack of 
conservation in the N
-terminal extension of red algal FtsZs and poor transit peptide 
prediction tools, we PCR
-amplified 
GsFtsZA
 and GsFtsZB
 beginning at the start of the 
conserved GTP
-binding domain. The s
equence encoding GsFtsZA (BAA82090.1) 
lacking the first 120 amino acids was PCR
-amplified with 
primers AT78/AT80 and joined 
to mCerulean
, amplified with AT7/AT81, by primers AT78/AT81 by splicing by overlap 
extension and asymmetric PCR 
(Warrens et al., 199
7). This fused gene fragment was 
cloned into 
pREP42X
 digested with Xho1 and BamH1 by conventional clon
ing techniques to create 
pREP42X
-GsFtsZA
-mCerulean
. The sequence encoding 
GsFt
sZB 
!11c!(BAA82091.1) lacking the first 90 amino acids was PCR
-amplified with pri
mers 
AT76/AT77 and joined to 
mVenus
, amplified with primers AT7/AT81, by primers 
AT76/AT81 by splicing
 by overlap extension and asymmetric PCR. This fused gene 
fragment was cloned into 
pREP41X
 digested with Xho1 and BamH1 by conventional 
cloning techniques
 to create 
pREP41X
-GsFtsZB
-mVenus
. Generation of 
pREP41
-SeFtsZ
-msfGFP
 was performed by PCR
-amplification of 
FtsZ
 (synpcc7942_2378) from 
Synechococcus elongatus
 PCC 7942 in two fragments. Fragment one, which consisted 
of nucleotides 1
-240, was PCR
-amplified
 with primers 
PSZ1F/PSZ1R
. Piece two, which 
consisted of nucleotides 241
-1182, was PCR
-amplified with primers 
PSZ2F/PSZ2R
. Amplification of 
msfGFP
, which also bears the A206K mutation to eliminate dimerization 
(Costantini et al., 2012)
, was performed with primers 
PGFPF/PGFPR
. All three PCR 
products were cloned into 
pREP41
 digested with Nde1 and BamH1 by the Gibson 
Assembly method.
 To generate the control constructs that expressed only mVenus or 
mCerulean, sequences 
encoding mVenus and m
Cerulean were PCR
-amplified from 
S. 
pombe
 expression constructs with primers AT310 and AT297 and cloned into
 BamH1
-digested
 pREP41X
 or 
pREP42X
, respectively, by Gibson Assembly.
 To generate the 
control construct 
pREP41
-msfGFP
, msfGFP
 was PCR
-amplified with
 primers PGF/PGR 
and cloned into 
pREP41
 digested with Nde1 and BamH1 by the Gibson Assembly 
method.
  Putative GTPase
-deficient mutants were created by introducing a single 
nucleotide substitution via overlapping primers and PCR amplification of the constru
cts 
described above. 
pREP41X
-OsFtsZ1
D249A
-mVenus
 was created by PCR 
amplification of
 !11e!OsFtsZ1
D249A
-mVenus
 in two pieces from 
pREP41X
-OsFtsZ1
-mVenus
 with primers 
AT325/AT340 and AT339/AT297 and cloned into 
pREP41X
 digested with BamH1 by 
Gibson Assembly
. pREP
42X-OsFtsZ2
D300A
-mCerulean
 was created by PCR 
amplification of
 OsFtsZ2
D300A
-mCerulea
n in two pieces from
 pREP42X
-OsFtsZ2
-mCerulean
 with primers AT327/AT342 and AT341/AT297 and cloned into 
pREP42X
 digested with BamH1 by Gibson Assembly
. pREP41X
-OtFtsZ1
D224A
-mVenus
 was 
created by PCR amplification of 
OtFtsZ1
D224A
-mVenus
 in two pieces from 
pREP41X
-OtFtsZ1
-mVenus
 with primers
 AT329/AT344 and AT343/AT297 and cloned into 
pREP41X
 digested with BamH1 by Gibson Assembly
. pREP42X
-GsFtsZA
D322A
-mCerulean
 was created by PCR amplification of
 GsFtsZA
D322A
-mCerulean
 in two pieces 
from
 pREP42X
-GsFtsZA
-mCerulean
 with primers AT296/AT294 and AT293/AT297 and 
cloned into 
pREP42X
 digested with BamH1 by Gibson Assembly
. pREP41X
-GsFtsZB
D300A
-mVenus
 was created by PCR
 amplification of
 GsFtsZB
D300A
-mVenus
 in two pieces from
 pREP41X
-GsFtsZ1
-mVenus
 with primers AT295/AT292 and 
AT291/AT297 and cloned into 
pREP41X
 digested with BamH1 by Gibson Assembly
. pREP41X
-SeFtsZ
D239A
-msfGFP
 was created by PCR amplification of
 SeFtsZ
D239A-msfGFP
 in two pieces from
 pREP41
-SeFtsZ
-msfGFP
 with primers AT351/AT354 and 
AT353/AT352 and cloned
 into pREP41X
 digested with BamH1 by Gibson Assembly.
  An expression construct was made to coexpress both GsFtsZA
-mCerulean and 
GsFtsZB
-mVenus from a sing
le vector. The expression cassette from 
pREP41X
-GsFtsZB
-mVenus
, including the 
nmt
1* promoter, 
GsFtsZB
-mVenus
, and nmt
1 terminator was PCR
-amplified with 
primers AT109/AT110 and
 cloned into 
pREP42X
-!11d!GsFtsZA
-mCerulean
 digested with AatII by Gibson Assembly. A
ll primers used for 
cloning are listed in Table 3.2.
  Table 3.2: List of primers used 
for
 cloning. Column 1: Primer designation. Column 2: 
Nucleotide sequence of each individual primer. Column 3: Indication of the presence or 
absence of a restriction site 
sequence engineered into the primer. See materials and 
methods section for additional details.
 Primer
 Sequence
 Restriction 
Site
 AT7
 )&X!X&X!)X%!))X!XX%!X)X!X)X!%&X
!N/A
 AT76
 &&&!&&&!
%&%!X)X!
)%%!)&X!%)X!&X%!))X!)&&!))X!X&&!X&X!XX)!X&&!X
!Xho1
 AT77
 X%%!%&&!
X%&!%)%!%)&!%&X!%)&!)X)!X))!&%%!XX&!)X%!%)%!%)%!)X&!
))%!N/A
 AT78
 &&&!&&&!
%&%!X)X!
)%%!)&X!%%&!&X%!)&&!)&%!))X!X&&!X&X!XX)!X&&!X
!Xho1
 AT80
 X%%!%&&!X%&!%)%!%)&!%&X!%)&!%&&!X&&!%&%!&&X!%&X!X))!%%&!&%&!
))X!X
!N/A
 AT81
 &&&!&&&!
XX)!&%%
!&&)!%&&!X&)!%)X!%&%!X&%!%
)&!X%%!X)X
!BamH1
 AT109
 X)&!))&!))&!XX&!&&%!&&)!X)%!X&
X!&%X!)&%!X)%!&%&!)X)!XX)!&%)!
X))!))&!&)&!%
!N/A
 AT110
 X))!))X!&X%!%)%!%&X!)%X!&
X%!)&&!)%&!))&!)X)!))X!X)&!&)&!&&%!
)%&!&%&!))&!&)%!)%)!))&!&%%!X
!N/A
 AT262
 GTT AAA TCA TAC CTC GAG 
GGA TCC 
ACC ATG AGG 
TCT AAG TCG ATG 
CGA TTG AGG
 BamH1
 AT264
 X&&!)))!&%)!&)%!%&%!X)X!
XX)!&%%!
)%%!)&X!X%%!X%&!%)X!)))!&%&!
X))!&%&!&%&
!BamH1
 AT291
 X&X!X)&!&&%!X%)!
X%&
!X&&!)X)!)%&!X&X!)&X
!N/A
 AT292
 %&!))%!
)X%
!&X%!X))!)&%!%)%!)&&!%)%!X)X
!N/A
 AT293
 X&X!X)&!&&%!X%&!
X%&
!X&X!)X)!&%&!)&%!)&X
!N/A
 AT294
 %&!%)%!
)X%
!)X%!X))!)&%!%)%!)&&!X)&!X)X
!N/A
 AT297
 X)%!)&&!%%&!&&&!)%%!%XX!
XX)!&%%!
&&)!%&&!X&)!%)X!%&%!X&%!%)&!
X%%!X)X
!BamH1
 !11_!$-LD)'VIS'W,/0.
!*Y'AT298
 %&%!X%%!%&&!X%&!%)%!%)&!
%&X!%)&
!X))!X))!))X!&%&!)%X!XXX!)X)!
)X)
!N/A
 AT299
 %&%!X%%!%&&!X%&!%)%!%)&!
%&X!%)&
!X)%!&%X!XXX!)&)!)%X!)X)!X%&
!N/A
 AT325
 X&&!)))!&%)!&)%!%&%!X)X!
XX)!&%%!
)%%!)&X!X%)!&%X!X&&!%X&!&X%!
)X&!&&&!X%&!%%
!BamH1
 AT326
 %&%!X%%!%&&!X%&!%)%!%)&!
%&X!%)&
!)%&!)))!%))!&%&!%%&!%X)!%%)!
XX&!)X%!&X
!N/A
 AT327
 X&&!)))!&%)!&)
%!%&%!X)X!
XX)!&%%!
)%%!)&X!X%)!X%%!X%)!X))!X%&!
)X)!)%)!%&X
!BamH1
 AT328
 %&%!X%%!%&&!X%&!%)%!%)&!
%&X!%)&
!))%!&%X!&XX!X))!%%&!XX)!XXX!
&%!N/A
 AT329
 X&&!)))!&%)!&)%!%&%!X)X!
XX)!&%%!
)%%!)&X!X%&!%X&!X%%!))%!X%&!
)))!)&)!))X!X&)!%
!BamH1
 AT330
 %&%!X%%!%&&!X%&!%)%!%)&!
%&X!%)&
!&%&!&%&!X))!)X)!%%%!))X!)))!
%%%!%%
!N/A
 AT331
 X&&!)))!&%)!&)%!%&%!X)X!
XX)!&%%!
)%%!)&X!X%X!)X)!X)&!X)&!X%&!
%X)!X&)!X)&!X%
!BamH1
 AT332
 %&%!X%%!%&&!X%&!%)%!%)&!
%&X!%)&
!%%&!X%%!%)&!)%X!)%X!&&&!&))!
X))!XX%
!N/A
 AT339
 X&X!X)%!&&%!X%&!
X%&
!X&&!)))!X%&!X&X!)&X
!N/A
 AT340
 &&&!))%!
)X%
!)X%!X))!X&%!%)%!X&&!))%!&))!)%%
!N/A
 AT341
 %!X&&!X)%!&&%!X%&!
X%%!
X&&!%X&!X%
!N/A
 AT342
 )%X!))%!
XX%!)X%!X))!X&%!))%!X&&!X)%
!N/A
 AT343
 X&X!X)%!&&&!X%)!
X%%
!X&X!)XX!)%&!X&)
!N/A
 AT344
 %%&!%)%!
XX%!&X%!)))!X&%!%)%!X&&!%)%
!N/A
 AT351
 X&&!)))!&%)!&)%!%&%!X)X!
XX)!&%%!
)%%
!)&X!X%%!)%%!X)%!%%&!)&X!
%%X!)&%!))%!))&!&%
!BamH1
 AT352
 X)%!)&&!%%&!&&&!)%%!%XX!
XX)!&%%!
%&)!XXX!&%X!X&&!&&X!))&!&&&!
%%X!&&X!&)X!X
!BamH1
 AT353
 X)%!&&&!X%%!
X%%
!X&&!%X%!X%%
!N/A
 AT354
 X%X!))%!
XX%!XX%!)))!X&%!X)%!X
!N/A
 PSZ1F
 GACTTATAGTCGCTTTGTTAAATCAATGACGGATCCGATGCCAATTA
 N/A
 PSZ1R
 ACTACCGCTGCCACTTCCCGCGGAATGCAGCAACGCCTGC
 N/A
 PSZ2F
 GCACGGCATGGATGAACTCTACAAAGCCCCCAAGCGGATGCAGTTGGG
 N/A
 !!13M!$-LD)'VIS'W,/0.
!*Y'PSZ2R
 AAGGGAGACATTCCTTTTACCCGGGCTAGGGTCGGTTTTGAATTTTCC
 N/A
 PGFPF
 TTCCGCGGGAAGTGGCAGCGGTAGTATGAGTAAAGGTGAAGAACTG
 N/A
 PGFPR
 TTTGTAGAGTTCATCCATGCCG
 N/A
 PGF
 GACTTATAGTCGCTTTGTTAAATCAATGAGTAAAGGTGAAGAACTG
 N/A
 PGR
 AAGGGAGACATTCCTTTTACCCGGGTCATTTGTAGAGTTCATCCATG
 N/A
 !D.-$,:%./-5*%$#*$5%#'L#1%/2"
# FtsZ constructs were transformed into 
S. pombe
 using a lithium acetate 
procedure 
(http://www.sanfordburnham.org/labs/wolf/ 
Protocols/Protocols/Fission%20Yeast/Nurse%20Lab%20Manual.htm)
. S.pombe
 (strain 
MBY192 [h
- leu1-32 ura4
-D18]) cultures were grown in 50 mL of yeast extract plus 
supplements media (YES, 
http://www-bcf.usc.edu/~forsburg/media.html
) for ~40 hours 
at 32¡C. 5x10
8 cells (50 mL at OD
600 = 0.5
) were harvested by centrifugation at 4,000
g for 10 minutes at room temp. The resulting cell pellet was washed in 25 mL of TE 
(10 
mM Tris
-HCl, 1 mM EDTA, pH 7.5). The cells were pelleted again at 4,000
g for 10 
minutes at room temp. The cells were resuspend
ed in 1 mL of TE and LiAc (100 mM 
lithium acetate, pH 7.5) and allowed to incubate at 30
¡C for 30 min. 200 
"L of cells were 
aliquoted to a 2.0 mL microfuge tube that contained 
20 "L of 10 "g/"L carri
er sperm 
DNA (Agilent Technolo
gies) and 1 
"
g of plasmid D
NA (
~2-3 "L in 2 mM Tris
-HCl, pH 8.5) that had been incubating on ice. Cells were 
mixed by vortexing
. For strains 
coexpressing 2 FtsZs from separate plasmids, 1 
"g of each plasmid DNA was added to 
the transformation reaction to co
-transform both plasmids i
nto S. pombe
 at the same 
time. 1.2 mL of PEG solution (40% PEG, TE pH .5, and LiAc) was added to each tube 
and vortexed for 10 s. The tubes were incubated at 30
¡C for 30 min with shaking at 250 
!131!rpm. DMSO was added to 5% total volume (71
 "L), the cells were
 vortexed again for 10 
s, and then the tubes were incubated at 42
¡C for 15 min. The cells were pelleted at 
7,000g for 30 s, and the supernatant was pipetted off. The cells were resuspended in 
300 "L of TE and plated onto solid PMG with selection for each p
lasmid (
-leucine for 
pREP41
 and pREP41X
, -uracil for 
pREP42X
, or 
-leucine/
-uracil for strains with both 
pREP41X
 and pREP42X
) and 15 
"M thiamine to repress protein expression 
(Maundrell, 
1990). The plates were allowed to grow at 32
¡C until colonies formed.
 Growth and Expression of Transformed Cell Lines
  Yeast strains containing the FtsZ fluorescent fusions described above were 
streaked for isolation on solid PMG with 15 
"M thiamine, to repress expression. The 
plates also lacked leucine, uracil, or both to 
select for constructs pREP41/pREP41X, 
pREP42X, or both pREP41X and pREP42X, respectively. Plates were incubated at 
32¡C until colonies formed. Individual colonies were picked and used to inoculate 3 mL 
of liquid culture without thiamine to induce expressio
n of the fusion proteins. These 
cultures were allowed to grow at 32
¡C with 250 rpm shaking for ~40 h to reach steady 
state conditions for polymer assembly and turnover.
 Fluorescence Microscopy and FRAP Analysis
  S. pombe
 cells grown in liquid PMG media und
er selection, as described above, 
were imaged by differential interference contrast (DIC) and epifluorescence microscopy, 
using a microscope (model DMRA2; Leica, Wetzlar, Germany) with an HCX PL 
FLUOTAR 100X (1.30 NA) oil
-immersion objective (Leica) and a 
CCD camera (retiga 
Exi; QImaging, Burnaby, BC, Canada). 2 
"L of liquid culture was pipetted onto a glass 
!133!poly-lysine coated slide and covered with a No. 1.5 coverslip. All imaging was 
performed at room temperature. Z stacks were taken with 0.5 
"m increment
s. Images 
were acquired and subsequently de
-blurred using nearest neighbor deconvolution at 
70% haze removal using Image
-Pro 7.0 software (Media Cybernetics). Additional image 
processing and analysis for epifluorescence micrographs was performed using Fiji
 software (ImageJ; http://fiji.sc/Fiji). Single
-plane projections were generated from Z 
stacks using the maximum intensity algorithm and were falsely colored green for all 
FtsZ1 and FtsZB fluorescent signals and magenta for all FtsZ2, FtsZA, and 
cyanobacte
rial FtsZ fluorescent signals. Coexpression overlays were created with the 
merge channel tool, where colocalized fluorescence signals appear in white. 
Colocalization of fluorescence signals in coexpressing strains was quantified by creating 
a composite ima
ge of the 2 fluorescence signals from a deconvoluted Z stack, cropping 
the image to contain only the single cell being analyzed, unmerging the 2 channels, 
using the Coloc2 plugin within Fiji to calculate a PearsonÕs Correlation Coefficient (PCC) 
(Bolte and
 Cordeli‘res, 2006)
, and averaging all PCC values for each coexpression 
strain, 
± SEM.  Fluorescence Recovery After Photobleaching (FRAP) experiments were 
performed at room temperature on a laser
-scanning confocal microscope (Fluoview 
1,000 Spectral, Olymp
us) with a Plan FLN 60X (1.42 NA) oil
-immersion objective with a 
3.2X zoom. Just prior to performing each FRAP experiment, the high voltage setting for 
the photomultiplier tube was set so the greatest fluorescence signal in the image was 
just below saturat
ion. FRAP data were collected with FV1000 ASW software (Olympus). 
!13a!2 "L of liquid culture was pipetted onto a glass poly
-lysine coated slide and covered 
with a No. 1.5 coverslip. All polymers were bleached
 for 20 ms at a % laser intensity so 
that one-half to two-thirds of the fluorescence signal was bleached, and continuous 
imaging was performed at the minimal laser intensity to minimize photobleaching but 
still provide adequate fluorescence signal. mVenus, mCerulean, and msfGFP fusion 
constructs were imaged
 and bleached with a 515, 458, and 488 nm laser, respectively. 
All time
-course imaging for FRAP experiments were performed with 10 s intervals 
between time points. Three images were acquired before bleaching to calculate the 
average pre
-bleach fluorescence
 intensity, and subsequent imaging occurred for an 
additional 260 s. Fluorescence intensity measurements were acquired for the region of 
photobleaching, a background sample, and an area of fluorescence signal that was 
away from the bleach region. Raw FRAP 
data was processed to produce 
photobleaching corrected and normalized recovery datasets 
(Rabut and Ellenberg, 
2005). All datasets for each fluorescent fusion were averaged together to produce the 
average recovery dataset. Additionally, standard error of th
e mean (SEM) was 
calculated for normalized recovery at each time point. This average FRAP recovery 
dataset was used for curve
-fitting using pro Fit software (QuantumSoft), where the data 
were fit to the single exponential equation f(t) = A(1
-e-kt). The tim
e for one
-half recovery 
of the fluorescence signal (t
1/2
) was calculated as t
1/2
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'!!1a1!Results
 FtsZs lacking their N
- and/or C
-terminal regions show altered filament morphologies
  To begin this comparative analysis, I expressed various forms of FtsZ1 and 
FtsZ2 C
-terminally fused to monomeric fluorescence proteins in 
S. pom
be and analyzed 
their filament morphology characteristics. As described in Chapter 3, full
-length FtsZ1 
lacking the predicted transit peptide (FtsZ1
FL) (Olson et al., 2010)
 assembled cables, 
presumably representing bundles of individual polymers 
(Erickson 
et al., 2010; Mukherjee and Lutkenhaus, 1999; Popp et al., 2009)
, that looped around the cell and 
annealed to form large rings
 conforming to the interior compartment of the yeast cell 
(Fig. 3.1 A, Fig. 4.2 A). These cables displayed minimal fraying of smal
ler filaments from 
the main cable structure. FtsZ1 lacking the N
-terminal region (FtsZ1
#NT, Fig. 4.1) also 
assembled cables (Fig. 4.2 B). However, there were more numerous cables, which also 
possessed a higher frequency of smaller filaments splitting off o
f the main structure. 
FtsZ1 lacking the C
-terminal region (FtsZ1
#CT) assembled straight or bent filaments and 
a large number of thinner filaments, with weaker fluorescence signal, fraying off of the 
main filament (Fig. 4.2 C). FtsZ1 lacking both the N
- and C-terminal regions (FtsZ1
#NCT) showed the most drastic change in filament morphology from FtsZ1
FL. FtsZ1
#NCT assembled multiple long filaments that looped around the cell, but these filaments were 
frequently frayed into several loosely associated filaments, suggesting a reduced level 
of lateral interactions and bundling between individual polymers (Fig. 4.2 D).
  Full length FtsZ2 lacking the predicted transit peptide as described in Chapter 3 
(FtsZ2
FL) (Olson et al., 2010)
 assembled large rings and an overall even fluorescence 
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iCP! distribution along the ring structure (Fig. 4.2 E). Similar to FtsZ1
FL, the dimensions of 
these rings appeared to 
conform to
 the interior dimensions of the yeast cell
 (Fig. 4.2 E). 
In contrast, FtsZ2 lacking the N
-terminal region (FtsZ2
#NT) assembled into straight 
filaments that had even fluorescence distribution and were up to the length of the cell 
(Fig. 4.2 F). A small numbe
r of the assembled filaments displayed forked and branching 
ends, suggesting that lateral interactions in this truncated form of FtsZ2 were partially 
inhibited. FtsZ2 lacking the C
-terminal region (FtsZ2
#CT) assembled more highly 
branched rings accompanied
 by several cross
-linking filaments (Fig. 4.2 G). 
Additionally, FtsZ2
#CT assembled smaller rings of 
roughly 2 
im in diameter. FtsZ2 
lacking both the N
- and C-terminal regions (FtsZ2
NCT) assembled large rings, but these 
possessed cross
-linking filaments, a
reas of loose bundling with smaller filaments 
splitting from the main structure, and small rings
 roughly 2 
im in diameter (Fig. 4.2 H). 
In control experiments, unfused mVenus and mCerulean displayed completely diffuse 
localization patterns
 (Fig. 4.2 I
-J).  !1a`!Coassembly of FtsZ1 and FtsZ2 is influenced by their N
- and C-terminal flanking 
regions
  To further analyze the effect of the N
- and C-terminal regions on FtsZ filament 
morphology, I coexpressed FtsZ1
FL with full-length and all truncated forms of FtsZ2 
(Fig. 
4.3). As shown in Chapter 3 (
Fig. 3.3 A), FtsZ1
FL and FtsZ2
FL coassembled into ring 
structures that had cross
-linking filaments and regions where the ring split into smaller 
filaments (Fig. 4.3 A). These proteins also assembled smaller rings similar 
to those 
assembled by FtsZ2
CT. I also assessed the degree of colocalization by calculating the 
Pearson
gs correlation coefficient (PCC), which gives a measure of the degree of overlap 
between the two fluorescence signals and how the intensities correlate 
(Bolte and 
Cordeli
vres, 2006)
. FtsZ1
FL and FtsZ2
FL colocalized to a high degree with a PCC of 
0.92 ± 0.01 (mean 
± SEM, n = 23), as described in Chapter 3.
 In contrast, FtsZ1
FL and FtsZ2
NT coassembled into long, looping cables that resembled those formed by FtsZ1
FL when expressed alone (Fig. 4.3 B). However, the FtsZ1
FL fluorescence signal was 
slightly diffuse in this strain, resulting in a reduced PCC of 0.88 
± 0.01 (n = 12), a 
statistical
ly significant decrease compared to when 
FtsZ1
FL and FtsZ2
FL were 
coassembled 
(P < 0.01). Similar to FtsZ1
FL and FtsZ2
FL, FtsZ1
FL and FtsZ2
CT coassembled large rings with cross
-linking filaments and small rings, with a high PCC of 
0.92 ± 0.01 (n = 11, 
Fig. 4.3 C). FtsZ1
FL and FtsZ2
NCT coassembled into FtsZ1
FL-like 
cables. In this strain, FtsZ1
FL showed a more apparent diffuse localization pattern than 
in the FtsZ1
FL and FtsZ2
NT coexpression strain (Fig. 4.3 B and D). Because of this, 
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D@<!DRE!J6>E=<?G<8G<!?:H876?!E9<=67I!78;!GE6EG76:T<P!27?@<;!6:8<?!=<I=<?<8D!G<66!
E>D6:8<?P!47=?Q!N!
iCP! these proteins had a statistically significa
nt reduction in their PCC (0.74 
± 0.02, n = 17, 
P < 0.01) compared to coassembled structures of 
FtsZ1
FL and FtsZ2
FL. I also coexpressed FtsZ2
FL with the three truncated forms of FtsZ1 (Fig. 4.4). 
FtsZ1
NT and FtsZ2
FL coassembled into large rings with cross
-linking filaments and 
small rings with a PCC of 0.91 
± 0.01 (n = 7, Fig. 4.4 A). 
FtsZ1
CT and FtsZ2
FL coassembled into similar structures with a high PCC
 of 0.90 ± 0.01 (n = 13, Fig. 4.4 B). 
FtsZ1
#NCT and FtsZ2
FL also coassembled large and small rings, but the large rings were 
more frequently split with smaller filaments fraying off of the main structure. However, 
FtsZ1
#NCT and FtsZ2
FL were also highly colocalized with a 
PCC of 0.89 
± 0.01 (n = 15, 
Fig. 4.4 C). Al
though there was slightly more filament splitting and fraying for 
coassembled filament structures of FtsZ1
#NCT and FtsZ2
FL, all combinations of FtsZ2
FL  with the truncated forms of FtsZ1 coassembled into structures resembling those 
coassembled by 
FtsZ1
FL and FtsZ2
FL.  !1ae! 9+KG@)'ZIZF'&/-BB)PLD)*'Q+D-P)0.'P/@?>/D/K+)B'Q/@'9.BCS
95',/)R?@)BB)*'N+.>'.>)'
.@G0,-.)*'Q/@PB'/Q'9.BCAI'
AI:J6>E=<?G<8G<!C:G=EH=7I@?!EJ!
'L#1%/2"
!G<66?!
GE<WI=<??:8H!U)V!"D?01
$&SCb<8>?!78;!"D?03
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%&SCb<8>?!78;!
"D?03
"*SC%<=>6<78Q!78;!U%V!"D?01
$%&SCb<8>?!78;!"D?03
"*SC%<=>6<78P!"D?01
$&SCb<8>?Q!"D?01
%&SCb<8>?Q!78;!"D?01
$%&SCb<8>?!J6>E=<?G<8G<!?:H876?!7=<!J76?<65!
GE6E=<;!H=<<8Q!R@:6<!"D?03
"*SC%<=>6<78!J6>E=<?G<8G<!?:H876!:?!J76?<65!GE6E=<;!C7H<8D7P!
!!1ad!9+KG@)'ZIZ'W,/0.X*YF'
&@<!R@:D<!GE6E=!:8!D@<!C<=H<;!:C7H<?!=<I=<?<8D!=<H:E8?!R@<=<!D@<!
DRE!J6>E=<?G<8G<!?:H876?!E9<=67I!78;!GE6EG76:T<P!27?@<;!6:8<?!=<I=<?<8D!G<66!E>D6:8<?P!
47=?Q!N!
iCP! Taken together, these filament morphology data show that the N
- and C-terminal 
regions of F
tsZ1 and FtsZ2 play critical roles in normal FtsZ assembly, including 
promoting lateral association of individual FtsZ polymers into bundles and inducing 
filament curvature. The coexpression data are consistent with our previous finding that 
FtsZ2 has a la
rge influence on the morphology of coassembled filaments 
(TerBush and 
Osteryoung, 2012)
. However, these data expand our understanding of how FtsZ1 and 
FtsZ2 interact by showing that the N
-terminal region of FtsZ2 contributes strongly to 
FtsZ2
gs structural 
dominance over FtsZ1, that the N
- and C-terminal regions of FtsZ2 
influence the extent of colocalization between FtsZ1 and FtsZ2, and that the N
- and C-terminal regions of FtsZ1 may enhance bundling of coassembled structures.
 65,7,#4-(B*$A#5)"*.#M
>#-$!#3>5"./*$-4#."A*%$,#-,,"/24"#285#@*5)#*$)*2*5"!#4-5".-4#
*$5".-(5*%$,
# The results of the filament morphology experiments in 
S. pombe
 suggest that the 
N- and C-terminal regions influence FtsZ1 and FtsZ2 assembly properties. To further 
address this, purified ful
l-length and N- and C-terminally truncated forms of FtsZ1 and 
FtsZ2 were purified and assembled in the presence of GTP, and the resulting structures 
were imaged by
 transmission electron microscopy (TEM, Fig. 4.5). FtsZ1
FL readily 
assembled into polymers th
at laterally interacted to form bundled structures (Fig. 4.5 A). 
  !1a_! Figure 4.5: 
In vitro
 assembly of full
-length and truncated forms of FtsZ1 and 
FtsZ2.
 Transmission electron micrographs of assembly reactions containing (A) FtsZ1
FL  !1`M!Figure 4.5 (contÕd): 
with GTP, (B) FtsZ2
FL with GTP, (C) FtsZ1
#NCT with GTP, (D) 
FtsZ2
#NCT with GTP, (E) FtsZ1
FL with GDP, (F) FtsZ2
FL with GDP, (G) FtsZ1
#NCT with GDP, and (H) FtsZ2
#NCT with GDP. All reactions contained 5 
"M FtsZ and 500 
"M nucleotide. Assembly was allowed to
 proceed for 5 minutes before the protein was 
loaded onto carbon
-coated grids and stained with 2% uranyl acetate. (C
-D) Arrows 
indicate polymers ~5
-7 nm in width, which are likely single
-stranded protofilaments. 
Bars, 200 nm.
  Although the 3
-dimensional ge
ometry of these structures is not known, they appear to 
be small sheets of FtsZ1
FL polymers that ranged from ~55
-600 nm in length and ~30
-90 nm in width. FtsZ2
FL extensively assembled into large bundles that frequently split into 
smaller ones (Fig. 4.5 B).
 These bundles ranged from ~570
-6,300 nm in length and 
~40
-180 nm in width. These bundled structures are in contrast to the single or double
-stranded protofilaments previously assembled by full
-length Arabidopsis FtsZ1 and 
FtsZ2 
(Olson et al., 2010; Smith 
et al., 2010). However, those proteins were either 
purified out of inclusion bodies, which required denaturation and refolding, and only 
assembled in the presence of calcium as a stabilizing agent or had misplaced transit 
peptide cleavage sites. In contras
t to full-length FtsZ1, FtsZ1
NCT assembled thin 
protofilament bundles, indicating reduced lateral assembly (Fig. 4.5 C). These 
structures ranged from ~220
-550 nm in length and ~6
-40 nm in width. The thin filaments 
of ~6 nm in width likely represent single
 or double
-stranded protofilaments, based on 
dimensions of bacterial FtsZ proteins 
(Romberg et al., 2001)
 (Fig. 4.5 C, arrows). 
!1`1!FtsZ2
NCT also assembled into much smaller structures than its full
-length counterpart 
(Fig. 4.5 D). FtsZ2
NCT structures ranged
 from ~80
-430 nm in length and ~5
-35 nm in 
width. Protofilaments of ~5 nm in thickness were observed (Fig. 4.5 D, arrows), 
suggesting that FtsZ2
NCT also assembled single or double
-stranded protofilaments, but 
were not as prevalent as in FtsZ1
NCT assembly
 reactions. In control reactions, FtsZs 
were assembled in the presence of GDP. Consistent with previous results 
(Olson et al., 
2010), neither FtsZ1 nor FtsZ2 assembled polymerized structures when GDP was the 
only nucleotide present (Fig. 4.5 E
-H). These TE
M data show that full
-length FtsZ1 and 
FtsZ2 assemble highly bundled structures and that their N
- and C-termini greatly 
promote lateral association of polymers into bundled structures.
 The N- and C-terminal regions influence FtsZ filament 
turnover
  To cont
inue my analysis, I performed fluorescence recovery after photobleaching 
(FRAP) experiments on the various truncated forms of FtsZ1 and FtsZ2 to assess their 
steady
-state filament turnover characteristics. All constructs displayed recovery of 
fluorescence 
into the photobleached regions (Fig. 4.6). As shown in Chapter 3, the 
FtsZ1
FL fluorescence signal recovered with a half
-time (t
1/2
) of 51 s and to 53% of the 
pre
-bleach intensity (
Fig. 3.4 A, n = 16). All three of the truncated FtsZ1 forms showed 
altered f
ilament
 turnover characteristics. The FtsZ1
NT fluorescence signal showed 
reduced recovery, with a t
1/2
 of 95 s and to 53% of the pre
-bleach intensity (Fig. 4.6 A, n 
= 13). The FtsZ1
CT fluorescence signal displayed a t
1/2
 of 90 s and a maximum 
recovery of 21% (Fig. 4.6 B, n = 14), while the FtsZ1
NCT fluorescence signal recovered 
with a t1/2
 of 93 s and to a maximum extent of 37% (Fig. 4.6 C, n = 14).
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iCP!  As shown in Chapter 3, the FtsZ2
FL fluorescence signal recovered with a t
1/2
 of 67 s and to 13% of the pre
-bleach intensity (
Fig. 3.4 E, n = 16). Similar to the truncated 
FtsZ1 forms, all truncated forms of FtsZ2 showed altered filament turnover properties. 
The FtsZ2
NT fluorescence signal
 recovered with the slowest t
1/2
, 163 s, and to 33% of 
the pre
-bleach intensity (Fig. 4.6 D, n = 14). The FtsZ2
CT fluorescence signal recovered 
with a t1/2
 of 118 s and to 43% of the pre
-bleach intensity (Fig. 4.6 E, n = 12), the 
greatest percent recovery
 among the FtsZ2 truncated forms. Finally, the FtsZ2
NCT fluorescence signal recovered with a t
1/2
 of 92 s and to 21% of the pre
-bleach intensity 
(Fig. 4.6 F, n = 11). Taken together, the FRAP data show that all truncated forms FtsZ1 
!1``!and FtsZ2 display redu
ced filament
 turnover dynamics compared to their full
-length counterparts. Additionally, FtsZ1 and FtsZ2 proteins lacking their N
- and C-terminal 
regions are more similar to each other in their turnover properties than are their full
-length counterparts. T
hese data indicate that the N
- and C-terminal flanking regions 
promote FtsZ1 and FtsZ2 filament
 turnover and contribute to their distinct dynamic 
properties. However, these regions do not fully account for these differences.
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!!!!1eN!Appendix
 Functional Analysis of the Chloroplast Division Complex Using 
Schizosaccharomyces pombe
 as a Heterologous Expression System
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!!!1ec!Abstract
 Chloroplast division is driven by a macromolecular complex that assembles at the mid
-plastid
. The central structure in this complex is the cytoskeletal FtsZ ring (Z ring), which 
is composed of the functionally distinct proteins FtsZ1 and FtsZ2. Recent studies in the 
heterologous 
Schizosaccharomyces pombe 
system showed that Arabidopsis FtsZ1 and 
FtsZ2 polymers have distinct assembly and turnover characteristics. To further analyze 
these FtsZs, we employed this system to compare the assembly and dynamic 
properties of FtsZ1 and FtsZ2 lacking their divergent N
- and C-termini with those of their 
full-length counterparts. Our data provide evidence that the N
- and C-termini promote 
polymer bundling and turnover in both FtsZs, but that they also contribute to the distinct 
behaviors of FtsZ1 and FtsZ2 filaments. We also assessed the effect of the FtsZ2 
binding protein ARC6 on FtsZ2 polymer dynamics, and found that ARC6 interacts with 
and stabilizes FtsZ2 filaments in 
S. pombe
 independent of its presumed Z
-ring tethering 
function 
in planta. Finally, we generated FtsZ1
-FtsZ2 coexpression constructs to 
facilita
te reconstitution of more complex interaction networks. Our experiments yield 
new insight into factors influencing FtsZ assembly and dynamics and highlight the utility 
of S. pombe
 for analysis of chloroplast FtsZs and their assembly regulators.
   !1ee!Introduct
ion  Chloroplasts arose from the endosymbiosis of an ancient cyanobacterium and 
are present in high numbers in several green tissues of plants 
(Gould et al., 2008)
. To maintain these high population numbers as plant cells grow and divide, chloroplasts 
undergo growth and binary fission, where a constriction forms at the chloroplast 
midpoint and narrows until the two daughter chloroplasts are formed 
(Osteryoung and 
Pyke, 2014)
. Chloroplast division is driven by a large macromolecular complex that 
includes the
 formation of ring structures on both the stromal and cytosolic sides of the 
chloroplast membranes 
(Osteryoung and Pyke, 2014)
. Consistent with chloroplasts
g origin from an engulfed cyanobacterium by a heterotrophic eukaryote, many of the 
division proteins
 in the chloroplast interior (stroma) are homologs of bacterial cell 
division components. However, over evolutionary time, chloroplasts have acquired 
additional division proteins of eukaryotic origin.
  The centerpiece of the chloroplast division machinery 
is the cytoskeletal protein 
FtsZ, which assembles into the mid
-plastid FtsZ ring (Z ring). The Z ring establishes the 
chloroplast division site and likely provides at least some of the constriction force in the 
early stages of chloroplast division, as show
n for bacterial Z rings 
(Osawa et al., 2008)
. The chloroplast Z ring is composed of two functionally distinct FtsZ isoforms, FtsZ1 and 
FtsZ2 
(Miyagishima et al., 2001; Osteryoung et al., 1998; Osteryoung and Vierling, 
1995; Schmitz et al., 2009; Vitha et a
l., 2001). FtsZ1 and FtsZ2 are tubulin
-like 
GTPases that are capable of self
-assembly as homopolymers 
in vitro
 but are always 
colocalized 
in vivo
 and preferentially coassemble into heteropolymers 
in vitro
 (El-Kafafi 
!1ed!et al., 2005; Lohse et al., 2006; 
McAndrew et al., 2001; Olson et al., 2010; Smith et al., 
2010; Vitha et al., 2001)
. Several stromal regulatory proteins of both bacterial and 
eukaryotic origin function together to antagonize FtsZ assembly at the chloroplast poles, 
thereby only allowing Z
-ring formation at the chloroplast midpoint 
(Colletti et al., 2000; 
Glynn et al., 2009; Itoh et al., 2001; Maple et al., 2002; Nakanishi et al., 2009; Shimada 
et al., 2004; Zhang et al., 2013). Although FtsZ1 and FtsZ2 are soluble proteins in the 
chloroplas
t stroma, Z rings are tethered to the chloroplast membrane via interaction of 
FtsZ2 with the integral membrane protein ARC6 
(Glynn et al., 2008; Maple et al., 2005; 
Schmitz et al., 2009; Vitha et al., 2003)
. ARC6 plays a major role in coordinating the 
stro
mal Z ring and cytosolic dynamin
-like DRP5B ring across the envelope membranes
 (Gao et al., 2003; Glynn et al., 2008)
.  Much work has gone into understanding the interactions between the chloroplast 
division components as well as their mechanistic function
s. Most of our understanding 
has come from various genetic experiments observing chloroplast morphology by light 
microscopy and FtsZ assembly patterns by immunofluorescence microscopy in various 
mutant backgrounds, and from
 yeast two
-hybrid and other inter
action assays to 
elucidate the interaction networks
 (Glynn et al., 2008; Glynn et al., 2009; Maple et al., 
2005; Maple et al., 2007; Marrison et al., 1999; McAndrew et al., 2001; Miyagishima et 
al., 2001; Mori et al., 2001; Osteryoung et al., 1998; Schmitz
 et al., 2009; Vitha et al., 
2003; Vitha et al., 2001; Yoshida et al., 2010; Zhang et al., 2013)
. One of the necessary 
next steps is analysis of the division proteins in live cells to assess the mechanistic 
function of each and understand how these protein
s work together to drive division. 
!1e_!Conducting such experiments 
in planta requires fluorescence microscopy of tagged 
chloroplast division components, which can be challenging for several reasons, 
including the intolerance of some division components to fluo
rescent protein tagging, 
sensitivity of chloroplast division to altered levels of division proteins, transgene 
instability, long generation times, weak fluorescence signals from constructs driven by 
native promoters amid high background noise due to chloro
phyll autofluorescence, and 
chloroplast movement in response to illumination on a microscope. An expression 
system in a cellular environment in which analysis of the inherent behaviors of FtsZ1 
and FtsZ2 could be easily observed, as well as how FtsZ1 and F
tsZ2 properties are 
modulated by various regulatory proteins, can enhance our understanding of chloroplast 
division mechanisms.
  Srinivasan et al. (2008) expressed various bacterial FtsZs as fluorescent fusions 
in the fission yeast 
Schizosaccharomyces pomb
e and studied their polymer assembly 
and dynamic properties. They found that FtsZ from 
Escherichia coli
 and Mycobacterium 
tuberculosis
 assembled filaments in the yeast cytosol that displayed turnover 
characteristics consistent with previous 
in vivo
 and in vitro
 findings 
(Anderson et al., 
2004; Chen et al., 2007; Chen and Erickson, 2005)
, suggesting that FtsZ behavior in 
S. 
pombe
 accurately reflects its 
in vivo
 behavior. This study paved the way for TerBush 
and Osteryoung (2012) to study 
Arabidopsis thaliana
 FtsZ1 and FtsZ2 in fission yeast, 
where they found that FtsZ1 and FtsZ2 have distinct filament morphology and turnover 
characteristics. Their data, viewed in the context of prior genetic studies 
(Osteryoung et 
al., 1998; Schmitz et al., 2009; Yoder et al.
, 2007), suggested that FtsZ2 is the structural 
!1dM!determinant for chloroplast Z rings while FtsZ1 promotes Z
-ring constriction by 
enhancing turnover. Additional experiments in fission yeast were instrumental in 
establishing that the FtsZ
-positioning factor A
RC3 acts to directly antagonize FtsZ 
assembly 
(TerBush and Osteryoung, 2012; Zhang et al., 2013)
. These initial studies 
have shown fission yeast to be a robust tool for analysis of FtsZ organization and 
dynamics, and provided evidence on how chloroplast di
vision regulatory proteins 
modulate FtsZ behavior. 
S. pombe
 cells can be grown quickly and are easily 
transformed, and, as 
S. pombe
 and chloroplasts share no division components, 
individual chloroplast division proteins can be studied in isolation or in co
mbination with 
specific regulatory proteins in a cellular environment. Thus 
S. pombe
 can be a valuable 
complement to other experimental tools for elucidating the functions and interactions of 
the chloroplast division components.
  Here, we utilize 
S. pombe
 to analyze the role of specific regions of FtsZ1 and 
FtsZ2 on their filament morphology and turnover characteristics as well as assessing 
the effect ARC6 has on FtsZ2 filament stability. These studies illustrate that the N
- and C-terminal regions of FtsZ1 
and FtsZ2 promote lateral interactions and polymer 
turnover, while ARC6 directly stabilizes FtsZ2 polymers in the absence of its membrane 
tethering function. We also discuss technical limitations and advantages of performing 
experiments in fission yeast.
   !1d1!Results
 Effect of N
- and C-terminal truncations on FtsZ filament morphology
 FtsZs possess a highly conserved GTP
-binding and hydrolysis domain and N
- and C-terminal regions with far less sequence conservation 
(TerBush et al., 2013; 
Vaughan et al., 2004)
. We hypothesized that the poorly conserved N
- and C-terminal 
flanking regions play a role in determining the distinct behavior of FtsZ1 and FtsZ2. 
Therefore, we compared the polymer assembly characteristics of chloroplast FtsZs 
lacking the N
- and C-terminal
 regions with those of full length FtsZs expressed in 
S. 
pombe
. Full-length FtsZ1 lacking its predicted transit peptide (amino acids 58
-433) 
(Olson et al., 2010)
 and the truncated form of FtsZ1 (amino acids 74
-377) were fused at 
their C
-termini to mVenus, 
a monomeric version of the yellow fluorescence protein, to 
create FtsZ1
58-433-mVenus and FtsZ1
74-377-mVenus. The equivalent FtsZ2 proteins were 
fused at their C
-termini to mCerulean, creating FtsZ2
49-478-mCerulean and 
FtsZ2
119-424-mCerulean. Consistent wit
h previous experiments in 
S. pombe
 (TerBush and 
Osteryoung, 2012)
, FtsZ1
58-433-mVenus assembled multiple cables, presumably 
representing bundles of individual polymers 
(Erickson et al., 2010; Mukherjee and 
Lutkenhaus, 1999; Popp et al., 2009)
, that looped around the cell and annealed to form 
large rings conforming to the interior compartment of the yeast cell (Fig. A.1 A). These 
cables displayed minimal fraying of smaller filaments from the main cable structure. 
FtsZ1
74-377-mVenus also assembled multiple lo
ng filaments that looped around the cell, 
but these filaments were frequently frayed into several loosely associated filaments, 
  !1d3!!Figure A.1: Polymer morphologies of full
-length 
FtsZ1 and FtsZ2 
and constructs 
lacking their N
- and C-terminal regions
 in S. pombe. Epifluorescence micrographs of 
(A) FtsZ1
58-433-mVenus, (B) FtsZ1
74-377-mVenus, (C) FtsZ2
49-478-mCerulean, (D) 
FtsZ2
119-424-mCerulean, (E) mVenus, and (F) mCerulean fluorescence signals in 
expressing yeast cells. FtsZ1
58-433-mVenus, FtsZ1
74-377-mVenus, and mVenus 
fluorescence signals are falsely colored green. FtsZ2
49-478-mCerulean, FtsZ2
119-424-mCerulean, and mCerulean fluorescence signals are falsely colored magenta. Dotted 
line represents the cell outline. Bars, 5 
"m. ! !1da!suggesting a reduced l
evel of lateral interactions and bundling between individual 
polymers (Fig. A.1 B).
 FtsZ2
49-478-mCerulean assembled large rings and an overall even fluorescence 
distribution along the ring structure (Fig. A.1 C). Similar to FtsZ1
58-433-mVenus, the 
dimensio
ns of these rings appeared to be controlled by the interior dimensions of the 
yeast cell. 
FtsZ2
119-424-mCerulean
 also assembled large rings, but these possessed 
cross
-linking filaments, areas of loose bundling with smaller filaments splitting from the 
main
 structure, and small rings
 roughly 2 
im in diameter (Fig. A.1 D). In control 
experiments, unfused mVenus and mCerulean displayed completely diffuse localization 
patterns (Fig. A.1 E
-F).
 Effect of N
- and C-terminal truncations on s
teady-state turnover of F
tsZ polymers
  Another important aspect of FtsZ behavior is dynamic, steady
-state turnover of 
polymers 
(Anderson et al., 2004; Chen and Erickson, 2005; Osawa and Erickson, 2011; 
Srinivasan et al., 2008; Stricker et al., 2002)
. Therefore, we performed fluore
scence 
recovery after photobleaching (FRAP) experiments on the full
-length and truncated 
forms of FtsZ examined above to determine the effect the N
- and C-terminal regions 
have on FtsZ polymer turnover.
 The fluorescence signal in FtsZ1
58-433-mVenus
 filaments 
exhibited an average recovery half
-time (t
1/2
) of 51 s and a maximum percent recovery 
of 53% of the pre
-bleach intensity (n = 16 cells, Fig. A.2 A). The truncated FtsZ1
74-377-mVenus
 was less dynamic, with a t
1/2
 of 93 s and a maximum recovery of
 37% (n = 14, 
Fig. A.2 B). For FtsZ2
49-478-mCerulean
, the fluorescence signal recovered with a t
1/2
 of 67 s and to 13% of the pre
-bleach intensity (n = 16, Fig. A.2 C). 
FtsZ2
119-424-mCerulean
   !1d`!!9+KG@)'!ISF'!0-DJB+B'/Q'?/DJP)@'*J0-P+,B'Q/@'9.BCA'-0*'9.BCS
'?@/.)+0B'D-,O+0K'
.>)+@'2
T'-0*'&T.)@P+0-D'@)K+/0BI
!%<66?!<WI=<??:8H!U)V!"D?01
e`SaeeSCb<8>?!78;!U4V!
"D?03
11_S`3`SC%<=><678!R<=<!78765T<;!B5!".)/P!"=EC!6<JD!DE!=:H@DQ!D@<!I78<6?!=<I=<?<8D!
D@<!J6>E=<?G<8G<!:8D<8?:D5!:8!D@<!B6<7G@<;!=<H:E8!U;7?@<;!G:=G6<V!I=:E
=!DE!B6<7G@:8H!U/4VQ!
7D!D@<!D:C<!EJ!B6<7G@:8H!UM!?VQ!7D!D@<!D:C<!G6E?<?D!DE!D
1Y3
Q!78;!7D!D@<!D:C<!G6E?<?D!DE!DR:G<!
D@7D!EJ!D1Y3
P!&@<!H=7I@?!DE!D@<!=:H@D!?@ER!D@<!8E=C76:T<;!79<=7H<!J6>E=<?G<8G<!=<GE9<=5!
9?!D:C<!U?VP!&@<!;7D7!7=<!8E=C76:T<;!DE!D@<!I=<
SB6<7G@!J6>E=<?G<8G<!:8D<8?:D5!U1!E8!D@<!
5S7W:?V!78;!D@<!J6>E=<?G<8G<!:8D<8?:D5!7D!D@<!D:C<!EJ!B6<7G@:8H!UM!E8!D@<!5
S7W:?VP!&@<!
;7D7!:8!D@<!BEW!7=<!D@<!G76G>67D<;!C7W:C>C!J6>E=<?G<8G<!=<GE9<=5!78;!D
1Y3
!U76?E!
=<I=<?<8D<;!B5!D@<!9<=D:G76!B7=V!7?!R<66!7?!D@<!8>CB<=
!EJ!:8;<I<8;<8D!".)/!
!!1dN!9+KG@)'!IS'W,/0.X*YF'
<WI<=:C<8D?!I<=JE=C<;!U$VP!&@<!=<;!B=7GO<D?!=<I=<?<8D!,AF!7D!
<7G@!D:C<!IE:8DP!&@<!"D?01
e`SaeeSCb<8>?!78;!"D?03
11_S`3`SC%<=><678!J6>E=<?G<8G<!
?:H876?!7=<!J76?<65!GE6E=<;!H=<<8!78;!C7H<8D7Q!=<?I<GD:9<65P!47=?Q!3!iCP
! recovered with a t
1/2
 of 92 s and to a maximum of 21% (n = 11, Fig. A.2 D). The slower 
recovery rates for the truncated proteins relative to those of their full
-length counterparts, as well as the decreased extent of recovery of FtsZ1
74-377-mVenus
 compar
ed to that of FtsZ1
58-433-mVenus, indicate that FtsZ1
74-377-mVenus and FtsZ2
119-424-mCerulean homopolymers are more stable than those composed of the full
-length proteins. These data suggest that the N
- and/or C
-terminal regions of FtsZ1 and FtsZ2 
directly
 promote polymer turnover.
 All FRAP data are summarized in Table A.1.
 Protein
 t1/2
 (s) % Recovery
 n FtsZ1
58-433-mVenus
 51 53% 16 FtsZ1
74-377-mVenus
 93 37% 14 FtsZ2
49-478-mCerulean
 67 13% 16 FtsZ2
119-424-mCerulean
 92 21% 11 ARC6
68-614-mVenus
 + FtsZ2
49-478-eCFP 64 44% 14 130 28% 11  Table A.1: Summary of FRAP data from all single and coexpression strains.
 Time 
for the normalized and averaged half
-time of recovery (t
1/2
, s) and maximum percent 
recovery for each protein in each expression strain. n 
represents the number of 
independent FRAP experiments performed on each protein in each expression strain.
 !1dc!Effect of ARC6 on FtsZ2 filament dynamics
  The chloroplast division complex includes over a dozen different protein 
components that form an intricate
 interaction network 
(Osteryoung and Pyke, 2014)
. Several of the stromal proteins directly interact with and regulate the assembly of FtsZ1 
and FtsZ2. One such critical interaction is between FtsZ2 and ARC6, an inner envelope 
transmembrane protein that int
eracts specifically with FtsZ2 and is thought to tether the 
Z ring to the membrane 
(Glynn et al., 2008; Maple et al., 2005; Vitha et al., 2003)
. Previous genetic experiments in Arabidopsis suggest that ARC6 promotes assembly of 
and stabilizes FtsZ polymers
 at the division site 
(Vitha et al., 2003)
. However, these 
conclusions came from immunolocalization of FtsZ2 in fixed leaves in 
arc6
 null and ARC6 overexpression lines. To further investigate the effect of ARC6 on FtsZ2 polymer 
stability, we coexpressed th
e stromal region of ARC6 
(Vitha et al., 2003)
 fused at its C 
terminus to mVenus (ARC6
68-614-mVenus)
 with full length FtsZ2 C
-terminally fused to 
eCFP (FtsZ2
49-478-eCFP) 
(TerBush and Osteryoung, 2012)
 and analyzed FtsZ2
49-478-eCFP polymer morphology and ste
ady-state turnover characteristics. When expressed 
independently, ARC6
68-614-mVenus adopted a completely diffuse localization pattern 
(Fig. A.3 A), while FtsZ2
49-478-eCFP assembled an intricate network of interconnected 
filaments (Fig. A.3 B), as described
 previously 
(TerBush and Osteryoung, 2012)
. When ARC6
68-614-mVenus and FtsZ2
49-478-eCFP were coexpressed, they colocalized into a 
network of interconnected filaments with a Pearson
gs Correlation Coefficient (PCC) of 
0.92 q 0.01 (mean 
q SEM, n = 18, Fig. A.
3 C), consistent with previous yeast two
-hybrid 
and biomolecular fluorescence complementation assays 
(Glynn et al., 2009; Maple et 
  !1de! 9+KG@)'!IVF'9.BCS'?/DJP)@'P/@?>/D/KJ'-0*'*J0-P+,B'+0'.>)'?@)B)0,)'/Q'.>)'
B.@/P-D'@)K+/0'/Q'!%&_I'
U)S4V!AI:J6>E=<?G<8G<!
C:G=EH=7I@?!EJ!G<66?!<WI=<??:8H!U)V!
).%c
cdSc1`SCb<8>?Q!U4V!"D?03
`_S`edSC%<=>6<78Q!78;!U%V!).%c
cdSc1`SCb<8>?!78;!
"D?03
`_S`edSC%<=>6<78P!U2
SAV!%<66?!<WI=<??:8H!U2V!).%c
cdSc1`SCb<8>?!78;!UAV!"D?03
11_S`3`S<%"/!R<=<!78765T<;!B5!".)/P!"=EC!6<JD!DE!=:H@DQ!D@<!I78
<6?!=<I=<?<8D!D@<!
J6>E=<?G<8G<!:8D<8?:D5!:8!D@<!B6<7G@<;!=<H:E8!U;7?@<;!G:=G6<V!I=:E=!DE!B6<7G@:8H!U/4VQ!7D!
D@<!D:C<!EJ!B6<7G@:8H!UM!?VQ!7D!D@<!D:C<!G6E?<?D!DE!D
1Y3
Q!78;!7D!D@<!D:C<!G6E?<?D!DE!DR:G<!D@7D!
EJ!D1Y3
P!&@<!H=7I@?!DE!D@<!=:H@D!?@ER!D@<!79<=7H<!J6
>E=<?G<8G<!=<GE9<=5!9?!D:C<!U?VP!&@<!
;7D7!7=<!8E=C76:T<;!DE!D@<!I=<
SB6<7G@!J6>E=<?G<8G<!:8D<8?:D5!U1!E8!D@<!5
S7W:?V!78;!D@<!
!!1dd!9+KG@)'!IV'W,/0.X*YF'
J6>E=<?G<8G<!:8D<8?:D5!7D!D@<!D:C<!EJ!B6<7G@:8H!UM!E8!D@<!5
S7W:?VP!
&@<!;7D7!:8!D@<!BEW!7=<!D@<!G76G>67D<;!C7W:
C>C!J6>E=<?G<8G<!=<GE9<=5!78;!D
1Y3
!U76?E!
=<I=<?<8D<;!B5!D@<!9<=D:G76!B7=V!7?!R<66!7?!D@<!8>CB<=!EJ!:8;<I<8;<8D!".)/!
<WI<=:C<8D?!I<=JE=C<;!U$VP!&@<!=<;!B=7GO<D?!=<I=<?<8D!,AF!7D!<7G@!D:C<!IE:8DP!
).%c
cdSc1`SCb<8>?!J6>E=<?G<8G<!?:H876!:?!J76?<65!GE6E=<;!H=<<8
Q!R@:6<!"D?03
11_S`3`S<%"/!J6>E=<?G<8G<!?:H876!:?!J76?<65!GE6E=<;!C7H<8D7P!U)
S%V!27?@<;!6:8<?!=<I=<?<8D!G<66!
E>D6:8<?P![@:D<!47=?Q!N!iCP!X=<5!47=?Q!3!iCP
! al., 2005). These networks highly resembled those assembled by FtsZ2
49-478-eCFP 
when expressed alone (
Fig. A.3 B).
  We used FRAP to investigate the steady
-state polymer dynamics of ARC6
68-614-mVenus
 and FtsZ2
49-478-eCFP in this coexpression strain. ARC6
68-614-mVenus displayed 
fluorescence recovery back into the bleached region with an average t
1/2
 of 64 s 
and total recovery of 44% (n = 14, Fig. A.3 D). FtsZ2
49-478-eCFP fluorescence recovered with 
an average t
1/2
 of 130 s and total recovery of 28% (n = 11
, Fig. A.3
 E). The t1/2
 for 
FtsZ2
49-478-eCFP in the presence of ARC6
68-614-mVenus
 was significantly greater than 
that reported previously for FtsZ2
49-478-eCFP expressed alone (P < 0.05, average t
1/2
 of 87 s and total recovery of 31%)
 (TerBush and Osteryoung, 2012)
. The total fluorescence recovery was slightly decreased compared to that
 when FtsZ2
49-478-eCFP 
was expressed alone (Fig. A.4). However, the difference in total recovery was not 
 ! !1d_! 9+KG@)'!IZF'7@-?>',/P?-@+0K'QDG/@)B,)0,)'@),/E)@J'/Q'9.BCS'-D/0)'-0*'N>)0'
,/)R?@)BB)*'N+.>'!%&_I
!&@<!8E=C76:T<;!79<=7H<!J6>E=<?G<8G<!=<GE9<=5!9?!D
:C<!
H=7I@!JE=!"D?03
`_S`edS<%"/!76E8<!U=<;!G:=G6<V!78;!:8!D@<!I=<?<8G<!EJ!).%c
cdSc1`SCb<8>?!
UH=<<8!;:7CE8;VP!27D7!:8!<7G@!H=7I@!R<=<!8E=C76:T<;!DE!D@<!I=<
SB6<7G@!J6>E=<?G<8G<!
:8D<8?:D5!U1!E8!D@<!5
S7W:?V!78;!D@<!J6>E=<?G<8G<!:8D<8?:D5!7D!D@<!D:C<!EJ!I@EDEB6<
7G@:8H!UM!
E8!D@<!5S7W:?VP!&@<!B=7GO<D?!:8;:G7D<!,AF!7D!<7G@!D:C<!IE:8DP
! statistically significant (P > 0.05). These data indicate that ARC6
68-614-mVenus
 significantly decreases the rate of FtsZ2 polymer turnover without altering the mobile 
and immobile fractions in 
S. pombe
, consistent with the stabilizing 
effect of ARC6 
overexpression on FtsZ filament formation 
in vivo
 (Vitha et al., 2003).
 Generation of
 coexpression plasmids for analysis of multiple proteins
  One drawback to expression in 
S. pombe
 is the presence of only two selectable 
markers in the MBY192 strain based on complementation of leucine and
 uracil 
!1_M!auxotrophy
 (Srinivasan et al., 2008)
. As the chloroplast division machinery is composed 
of multiple proteins in addition to FtsZ1 and FtsZ2, the ability to reconstitute more 
elaborate interaction networks in 
S. pombe
 would be highly advantageous for increasing 
our understanding of chloroplas
t division. Therefore, we generated a construct to 
coexpress both FtsZ1
58-433-eYFP and FtsZ2
49-478-eCFP from the same plasmid 
backbone (Fig. A.5 A). As seen previously 
(TerBush and Osteryoung, 2012)
, FtsZ1
58-433-eYFP and FtsZ2
49-478-eCFP expressed from sep
arate plasmids coassembled into an 
FtsZ2
-like network of interconnected filaments with a PCC of 0.91 
± 0.01 (n = 14) (
Fig. A.5 B). When expressed from the coexpression plasmid, FtsZ1
58-433-eYFP and FtsZ2
49-478-eCFP also coassembled into an FtsZ2
-like netwo
rk of filaments (Fig. A.5 C), which 
was morphologically indistinguishable from the filament network observed when the 
proteins were coexpressed from separate plasmids (Fig. A.5 B). In the strain with the 
coexpression plasmid, the FtsZ1
58-433-eYFP and FtsZ2
49-478-eCFP fluorescence signals 
colocalized with a PCC of 0.93 
± 0.01 (n = 16), which was not statistically different from 
when FtsZ1
58-433-eYFP and FtsZ2
49-478-eCFP were expressed from separate plasmids 
(P > 0.05). Thus, these coexpression plasmids will 
allow for the analysis of 3 or 4
-part 
interaction networks and greatly increase our understanding of the regulation of 
chloroplast division.
 !!!1_1!!Figure A.5: Construction of an FtsZ coexpression plasmid and comparison to 
coexpression from separate plasmids.
 (A) A diagram displaying how the single 
plasmid to express both FtsZ1
74-377-eYFP and FtsZ2
49-478-eCFP was generated. 
  !1_3!Figure A.5 (cont
Xd):
  (B-C) Epifluorescence micrographs of cells expressing FtsZ1
74-377-eYFP and FtsZ2
49-478-eCFP 
from (B) separate plasmids or (C
) from the coexpression plasmid. The FtsZ1
74-377-eYFP and FtsZ2
49-478-eCFP fluorescence signals are falsely colored green and 
magenta, respectively. Regions of white color in the merged panels represent overlap 
between the FtsZ1
74-377-eYFP and FtsZ2
49-478-eCFP fluorescence signals. Dashed lines 
represent cell outlines. Bars, 5 
im. ! !1_a!Discussion Technical aspects of the S. pombe system
  Analysis of the inherent behaviors of various chloroplast division components in 
S. pombe
 has advanced our understanding of how chloroplast division occurs. 
However, there are technical aspects to expression in fission yeast 
of which 
one must 
be aware. For example, the 
pREP
 series of plasmids does not integrate into the yeast 
genome 
(Maundrell
, 1993). Therefore, expression from cell to cell is variable. Upon 
expression of fluorescent proteins from 
pREP41X
 and pREP42X
, a range of protein 
levels will be observed, as indicated by no or little fluorescence signal to quite intense 
fluorescence signa
l. This trait is both advantageous and disadvantageous at the same 
time. In our experiments, the variation in expression has allowed for the effect of protein 
level on polymer morphology and turnover to be assessed in a single culture, as 
performed in TerB
ush and Osteryoung (2012), where fluorescence intensity was used 
as a measure of relative protein content. However, this trait also makes it more 
challenging to assess culture
-wide traits, such as average protein content per cell. More 
elaborate quantifica
tion techniques are required to perform this type of analysis, which 
may include the use of a construct that integrates into the genome and results in more 
even protein expression. Such an approach was used to quantify the number of 
molecules per cell for 
28 different cytoskeletal and signaling proteins in 
S. 
pombe
 cells 
using a combination of quantitative western blotting and fluorescence microscopy 
(Wu 
and Pollard, 2005)
. While there are 
pREP
 plasmids that yield varying degrees of 
expression due to stepwi
se deletions of the TATA box in the 
nmt1
 promoter 
(Basi et al., 
!1_`!1993; Forsburg, 1993)
, the use of these vectors to analyze the effect of protein level on 
FtsZ polymer morphology and dynamics requires comparison of separate cultures.
  Another consideration 
is the effect of FtsZ polymer morphology on quantification 
of colocalization using PearsonÕs correlation coefficients (PCC), which measure the 
extent of fluorescence signal overlap and how the two fluorescence intensities correlate 
(Bolte and Cordeli‘res, 
2006). We have used colocalization analysis in 
S. pombe
 to assess the interaction of FtsZ1 and FtsZ2 with various proteins that regulate their 
assembly 
(TerBush and Osteryoung, 2012; Zhang et al., 2013)
(Fig. A.3). 
However, one 
must be careful when interpre
ting colocalization data, as FtsZ filament morphology may 
influence quantified data output. For example, FtsZ1
58-433-eYFP assembles single 
cables and large rings
, while FtsZ2
49-478-eCFP assembles extensive networks of 
filaments 
(TerBush and Osteryoung, 201
2). If these proteins were coexpressed with a 
diffusely localized fluorescent protein
 that does not interact with FtsZ1 or FtsZ2
, then FtsZ2
49-478-eCFP would naturally have more fluorescence signal overlap with the 
coexpressed protein due to the FtsZ2
49-478-eCFP networks filling more of the cell volume 
than the FtsZ1
58-433-eYFP cables.
 Even though the proteins do not inter
act, t
his would 
artificially 
result in a greater
 PCC value 
between the fluorescent protein
 and FtsZ2
49-478-eCFP 
than FtsZ1
58-433-eYFP.
 Thus, the filament morphology of FtsZs must be taken into 
account when performing colocalization analysis with proteins that show weak 
interaction and diffuse localization patterns. Including appropriate controls, such as 
coexpressing each form of FtsZ wi
th a diffuse fluorescence protein, provides excellent 
controls for such experiments.
 !1_N!Effect of the N
- and C-termini on FtsZ polymer morphology and turnover
  The coassembly of FtsZ1 and FtsZ2 into the mid
-plastid Z ring is a critical aspect 
of chloroplast F
tsZ function 
in vivo
. Previous 
in vivo
 and in vitro
 data show that FtsZ1 
and FtsZ2 assemble homopolymers and coassemble heteropolymers that display active 
polymer turnover 
(El-Kafafi et al., 2005; Johnson et al., 2015; McAndrew et al., 2001; 
Olson et al., 
2010; Smith et al., 2010; Vitha et al., 2001)
. For 
S. pombe
 to be a valid 
system for the analysis of FtsZ behavior, chloroplast FtsZ fluorescent fusions must also 
possess these properties in the yeast cytosol. Our initial work with 
S. pombe
 demonstrated th
at FtsZ1
58-433-eYFP and FtsZ2
49-478-eCFP assemble as homopolymers 
and coassemble into heteropolymers that undergo active subunit exchange, and 
furthered our understanding on the distinct behaviors of FtsZ1 and FtsZ2 
(TerBush and 
Osteryoung, 2012)
. However,
 these new discoveries led to new questions, including 
how the divergent N
- and C-termini in these proteins affect their assembly and dynamic 
properties and whether or not FtsZ1
-FtsZ1 or FtsZ1
-FtsZ2 interfaces are inherently less 
stable than those of FtsZ2
-FtsZ2. Our finding that FtsZ1
74-377-mVenus
 and FtsZ2
119-424-mCerulean assemble more loosely bundled filaments than full
-length proteins in yeast 
(Fig. A.1) suggests that the N
- and/or C
-terminal regions enhance polymer bundling. 
FtsZ2
119-424-mCerulean fil
aments showed a higher degree of bundling than FtsZ1
74-377-mVenus
 filaments, suggesting that the conserved GTP
-binding and hydrolysis domain 
of FtsZ2 by itself has a greater capacity for lateral interactions than that of FtsZ1, which 
may contribute to the 
greater stability of FtsZ2 polymers 
(TerBush and Osteryoung, 
2012) (Fig. A.2). A recent study of 
E. coli
 and B. subtilis
 FtsZs showed that the C
-!1_c!terminal variable region (CTV), which comprises the amino acids following the 
conserved C
-terminal peptide requ
ired for FtsZ interaction with the membrane
-tethering 
protein, was important for promoting lateral assembly of FtsZ polymers into bundles 
based on the degree of positive charge 
(Buske and Levin, 2012)
. The loose bundling of 
FtsZ1
74-377-mVenus
 and FtsZ2
119-424-mCerulean are consistent with these data, as both 
FtsZ1 and FtsZ2 possess CTVs with a positive charge. Therefore, their CTVs could 
potentially promote bundling by charge shielding between the negatively charged GTP
-binding and hydrolysis domains of Fts
Z1 and FtsZ2 in filaments. An alternative 
explanation is that the FtsZ N
- and C-terminal regions make direct contacts to other 
polymers in the bundle to stabilize lateral interactions, as shown for the bacteriophage
-encoded FtsZ
-like protein PhuZ 
(Zehr et 
al., 2014). Several studies on bacterial FtsZs have shown that FtsZ subunit exchange is widely 
conserved and is important for sustaining Z
-ring constriction, although the dynamics 
vary from species to species and under different assembly conditions, 
(Chen 
et al., 2007; Chen and Erickson, 2009; Osawa and Erickson, 2011; Srinivasan et al., 2008)
. Consistent with our previous findings in 
S. pombe
 (TerBush and Osteryoung 2012), a 
recent study in live chloroplasts showed that FtsZ1 and FtsZ2 bearing fluorescent 
tags 
undergo active turnover in Z rings and polymers in transgenic Arabidopsis plants, and 
that FtsZ1 has greater turnover than FtsZ2 
(Johnson et al., 2015)
. However, performing 
comparative 
in planta analyses of truncated FtsZs would be challenging on seve
ral 
fronts (as discussed in the Introduction).
 Therefore, performing polymer turnover 
experiments in 
S. pombe
 facilitates comparative studies to assess intrinsic behaviors of 
!1_e!FtsZ proteins. 
In our previous study 
(TerBush and Osteryoung, 2012)
, we found that FtsZ1 homopolymers had a faster turnover rate and much greater total fluorescence 
recovery than FtsZ2 homopolymers. In the current study, FtsZ1
74-377-mVenus
 and FtsZ2
119-424-mCerulean exhibited nearly identical polymer turnover rates and their 
recovery p
ercentages, though still different, were more similar than those of their full
-length counterparts (Fig. A.2). Taken together, these data suggest that the N
- and C-terminal regions enhance bundling while also promoting polymer turnover. Importantly, 
the mo
re consistent, yet still different, behavior of FtsZ1
74-377-mVenus
 and FtsZ2
119-424-mCerulean homopolymers than of full
-length FtsZ1
58-433-mVenus
 and FtsZ2
49-478-mCerulean homopolymers also suggest that the N
- and/or C
-terminal regions contribute 
to the distinct polymer assembly and turnover characteristics of FtsZ1 and FtsZ2, but 
are not solely responsible for them.
 Effect of ARC6 on FtsZ2 polymer stability
  When coexpressed, 
ARC6
68-614-mVenus
 and FtsZ2
49-478-eCFP colocalized to a 
high degree, indicating t
hat these proteins directly interact in the yeast cytosol. This 
finding is consistent with previous Y2H experiments 
(Glynn et al., 2008; Maple et al., 
2005). Previous attempts to complement an Arabidopsis 
ftsZ2
 null mutant with a C
-terminal FtsZ2 fusion pr
otein failed (Johnson et al., 2015; TerBush and Osteryoung, 
2012), we presumed because the C
-terminal tag interfered with the interaction between 
the FtsZ2 C
-terminal peptide and ARC6 
(TerBush and Osteryoung, 2012)
. However, the 
fact that ARC6
68-614-mVenus
 directly interacts with FtsZ2
49-478-eCFP in 
S. pombe
 suggests that the lack of complementation 
in vivo
 may not be due to lack of interaction 
!1_d!with ARC6. One alternative explanation could be that the C
-terminal fusion to FtsZ2 
interferes with ionic interact
ions between individual FtsZ polymers in the Z ring. 
Additional experiments will be needed to clarify this issue.
 Our FRAP experiments on cells coexpressing ARC6
68-614-mVenus
 and FtsZ2
49-478-eCFP showed that ARC6
68-614-mVenus
 had a faster rate and greater 
extent of 
fluorescence recovery than FtsZ2
49-478-eCFP. The fact that ARC6
68-614-mVenus
 does 
not polymerize into large filamentous structures but adopts a diffuse localization pattern 
when expressed alone, even though BiFC data suggest that ARC6 may self
-interact 
and form homodimers 
(Maple et al., 2005)
, indicates that the only factor binding 
ARC6
68-614-mVenus
 to the network of interconnected filaments in the coexpression 
strain is FtsZ2
49-478-eCFP. Therefore, we presume that any ARC6
68-614-mVenus
 not bound to FtsZ2
49-478-eCFP is freely diffusible, resulting in a greater ability to diffuse in 
and out of the filament. These data also suggest that not all of ARC6
68-614-mVenus
 is 
bound to FtsZ2
49-478-eCFP and that the interaction between the two proteins may be 
somewhat transient. Further analysis into the kinetics of the ARC6
-FtsZ2 interaction will 
be needed to elucidate these details.
 More importantly, we found that FtsZ2
49-478-eCFP fluorescence recovery in the 
presence of ARC6
68-614-mVenus was significantly sl
ower than when FtsZ2
49-478-eCFP 
was expressed alone. However, there was no difference in the total extent of 
fluorescence recovery in the presence or absence of ARC6
68-614-mVenus (Fig. A.4). 
These data indicate that ARC6 directly stabilizes FtsZ2 polymers,
 as suggested by 
previous genetic work 
in planta (Vitha et al., 2003)
. However, in 
S. pombe
, stabilization 
!1__!by ARC6 occurred independently of its 
in vivo
 membrane
-tethering function. One 
potential explanation for this effect is that ARC6 stabilizes the FtsZ
2-FtsZ2 subunit 
interface, reducing the rate of polymer fragmentation. In this situation, the rate at which 
polymers break down into diffusible subunits would decrease and more of the FtsZ2 
pool would be locked into the immobile faction. As a result, one w
ould see a decrease in 
both the rate of polymer turnover and total fluorescence recovery, as seen previously for 
the GTPase
-deficient mutant FtsZ2
D322A
-eCFP that assembles static filaments
 (TerBush 
and Osteryoung, 2012)
. However, total fluorescence recover
y of FtsZ2
49-478-eCFP is 
unaffected by the presence of ARC6
68-614-mVenus. Therefore, a more likely possibility is 
that ARC6 stabilizes bundles by increasing the packing of or promoting more favorable 
lateral interactions between individual FtsZ2 polymers. 
In this situation, the frequency of 
FtsZ2 polymer breakage would be unaffected, but the increased interactions between 
FtsZ2 polymers may reduce diffusion of FtsZ2 subunits in and out of filaments. This 
situation is consistent with our data. However, more 
experiments are needed to 
understand how ARC6 specifically stabilizes FtsZ2 assembly and bundling.
 Conclusions
 The use of 
S. pombe 
has allowed us to elucidate inherent polymer assembly and 
dynamic properties of chloroplast FtsZ1 and FtsZ2, as well as ident
ify potential 
contributors to their distinct behaviors. We have also begun to understand the functional 
interplay between FtsZ1 and FtsZ2 and how several regulatory proteins modulate their 
assembly. While protein behavior in this system must be interpreted
 with the in vivo
 biological context in mind, as with any 
non-native
 system, the resulting data have
 been !3MM!valuable for enhancing our understanding of the chloroplast division machinery, 
particularly the assembly and dynamics of the FtsZs and roles of their
 assembly 
regulators. Thus, 
S. pombe
 represents an advantageous and robust tool for studying 
the workings of the chloroplast division complex. Previous studies of bacterial FtsZ and 
MreB in 
S. pombe
 (Srinivasan et al., 2007; Srinivasan et al., 2008)
 indica
tes this system 
may also be broadly useful for analysis of an array of cytoskeletal elements from 
prokaryotes as well as organelles.
 ! !3M1!Materials and Methods
 Cloning and transformation into S. pombe
  FtsZ genes were cloned into the 
pREP41X
 and pREP42X
 expression vectors 
under the control of the 
nmt1*
 promoter for analysis in 
S. pombe
 (Basi et al., 1993; 
Forsburg, 1993; Maundrell, 1993)
. All primers
 used for cloning are listed in 
Supplemental Table 1. Sequences encoding 
Arabidopsis thaliana
 FtsZ1
 (AtFtsZ1
-1, At5g55280, NP_200339.1) and 
FtsZ2
 (AtFtsZ2
-1, At2g36250, NP_565839.1) lacking 
their predicted transit peptides (the first 57 and 48 amino acids, respectively 
(Olson et 
al., 2010)) were PCR
-amplified from the corresponding 
S. pombe
 expression p
lasmids 
(TerBush and Osteryoung, 2012)
 using primers 
AT262/AT298 and AT264/AT299, 
respectively
. A monomeric variant of the yellow fluorescent protein 
Venus
 (mVenus) 
bearing the A206K mutation shown to eliminate the dimerization potential of GFP
-derived flu
orescent proteins 
(Nagai et al., 2002; Zacharias et al., 2002)
, generated in a 
bacterial expression vector provided by Desmond A. Moore and Harold P. Erickson 
(Duke University, Durham, NC
), was PCR
-amplified
 with primers AT7/AT297
. The sequence encoding 
mCerulean (
Addgene, Cambridge, MA) was PCR
-amplified with 
primers AT7/AT297
. FtsZ1
58-433 and mVenus
 were cloned into 
pREP41X
 digested with 
BamH1 
using
 the Gibson Assembly method 
(Gibson et al., 2009)
 to create 
pREP41X
-FtsZ1
58-433-mVenus
. FtsZ2
49-478 and mCer
ulean were cloned into 
pREP42X
 digested 
with BamH1 by Gibson Assembly to create 
pREP42X
-FtsZ2
49-478-mCerulean
. To generate 
pREP41X
-FtsZ1
74-377-mVenus
, FtsZ1
74-377 was PCR
-amplified from a previous 
S. pombe
 expression vector with primers 
AT266 and AT285
. mVenus was PCR
-!3M3!amplified from an 
S. pombe
 expression vector with 
primers AT7 and AT297. These 
gene fragments were cloned into 
pREP41X
 digested with BamH1 by Gibson Assembly. 
To generate 
pREP42X
-FtsZ2
119-424-mCerulean
, FtsZ2
119-424 was PCR
-amplified from 
another 
S. pombe
 expression construct with 
primers AT268 and AT288. 
mCerulean
 (Addegene, Cambridge, MA) 
(Rizzo and Piston, 2005)
 was PCR
-amplified from an 
S. 
pombe
 expression vector with primers AT7 and AT297
. These gene fragments w
ere 
cloned into 
pREP42X
 digested with BamH1 by Gibson Assembly. To generate the 
control constructs that expressed only mVenus or mCerulean, sequences 
encoding 
mVenus and mCerulean were PCR
-amplified from 
S. pombe
 expression constructs with 
primers AT310 an
d AT297 and cloned into
 BamH1
-digested
 pREP41X
 or 
pREP42X
, respectively, by Gibson Assembly. The sequence encoding the stromal region of 
Arabidopsis thaliana
 ARC6 (At5g42480) was 
amplified from a bacterial
 expression 
plasmid by PCR with primers AT168 and A
T139. mVenus
 was PCR
-amplified with 
primers AT95 and AT147. These gene
 fragments were cloned into 
pREP41X
 digested 
with Xho1 and BamH1 by Gibson Assembly to create 
pREP41X
-ARC6
68-614-mVenus
. pREP41X
-FtsZ1
58-433-eYFP
 and pREP42X
-FtsZ2
49-478-eCFP were the sa
me constructs 
as described in TerBush and Osteryoung (2012). 
A construct for coexpressing FtsZ1
58-433-eYFP and FtsZ2
49-478-eCFP from two expression cassettes in a single plasmid 
backbone was generated by PCR amplification of the entire expression 
cassette 
from 
pREP41
X-FtsZ1
58-433-eYFP
, including the 
nmt1*
 promoter, 
FtsZ1
58-433-eYFP
 gene, and the nmt
 terminator, with primers AT109 and AT110 and cloned into 
pREP42
X-FtsZ2
49-!3Ma!478-eCFP digested with AatII 
using
 Gibson Assembly.
 All primers used for cloning are 
summarized in Table A.2.
  All constructs were transformed into 
S. pombe
 (strain MBY192 [
h- leu1-32 ura4
-D18] (Srinivasan et al., 2008)
) by a modified lithium acetate protocol 
 (http://www.sanfordburnham.org/labs/wolf/ 
Prot
ocols/Protocols/Fission%20Yeast/Nurse%20Lab%20Manual.htm)
. S. pombe
 cells 
were grown in 50 mL of yeast extract with supplements media (YES, 
http://www-bcf.usc.edu/~forsburg/media.html
) for ~40 h a
t 32¡C. 5x10
8 cells (50 mL at OD
600 = 0.5) 
were collected by centrifugation at 4,000
g at room temp for 10 min. The supernatant 
was removed and the cell pellet was resuspended and washed in 25 mL of TE (
10 mM Tris
-HCl, 1 mM EDTA, pH 7.5). The cells were 
pelleted again at 4,000g at room temp 
for 10 min. The supernatant was removed and the cell pellet was resuspended in 1 mL 
of TE + LiAc (100 mM lithium acetate, pH 7.5). The cells were incubated in a water bath 
at 30¡C for 30 min. 200 
"L of cells were added
 to a 2.0 mL microfuge tube already 
containing 20 
"L of 10 "g/"L carri
er sperm DNA (Agilent Technolo
gies) and 1 
"
g of 
plasmid DNA (
~2-3 "L in 2 mM Tris
-HCl, pH 8.5) that was previously incubating on ice. 
Cells were mixed by vortexing. For 
S. pombe
 strains 
being transformed by 2 plasmids at 
the same time, 
1 "g of each plasmid was added to the transformation reaction. 1.2 mL 
of PEG solution (40% PEG, TE pH 7.5, and LiAc) was added to each transformation 
reaction and the solution was mixed by vortexing for 10 
sec. The cells were then 
incubated at 30
¡C with shaking at 250 rpm for 30 min. DMSO was added to 5% of the 
total volume (71 
"L). The transformation reactions were place
d in a 42¡C water bath for 
 !3M`! Table A.2: List of primers used for cloning.
 Column 1: Primer designation. Column 2: 
Nucleotide sequence of each individual primer. Column 3: Indication of the presence or 
absence of a restriction site sequence engineered into the primer. See materials and 
metho
ds section for additional details.
  $7C<
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&)&!%X&!&)%!)&X
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.%&%!X%%!%&&!X%&!%)%!%)&!
%&X!%)&
!&&%!&%&!)))!XX)!)X)!)X)!
XXX!&%&!&%&!&X&!)X
.$Y)
.)&3_e
.X)%!)&&!%%&!&&&!)%%!%XX!
XX)!&%%!
&&)!%&&!X&)!%)X!%&%!X&%!
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XX)!&%%
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XX%!X)X!X)X!%&X
.47C-1
.!3MN!15 min to heat
-shock the cells. The cells were pelleted at 7,000
g for 30 s
, and the supernatant was discarded. The cell pellets were resuspended in 300
 "L of TE and 
plated on solid PMG media with selection (
http://www-bcf.usc.edu/~forsburg/media.html
, -leucine for 
pREP41X
, -uracil for 
pREP42X
 and the coexpression construct, or 
-leucine/
-uracil for 
pREP41X
 and pREP42X
). Plates contained 15 
"M thiamine to repress protein 
expression from the 
nmt1*
 promoter 
(Maundrell, 1990)
. Cell growth conditions and expression from transformed S. pombe lines
  S. pombe
 expression strains containing the constructs described above were 
streaked for isolation on solid PMG with selection and containing 15 
"M thiamine to 
repress protein 
expression. Plates were incubated at 32
¡C until colonies formed, which 
was usually after 3
-5 days. Individual cells were picked and used to inoculate liquid 
PMG with selection but lacking thiamine. Liquid cultures were incubated at 32
¡C with shaking at 250
 rpm for ~40 h to reach full expression and presumed steady
-state 
conditions for FtsZ polymer assembly and turnover before fluorescence microscopy 
experiments were performed.
 Epifluorescence microscopy
  Cells grown in liquid PMG were imaged by differential
 interference contrast (DIC) 
and epifluorescence microscopy with a Leica microscope (model DMRA2; Leica, 
Wetzlar, Germany) equipped with an HCX PL FLUOTAR 100X (1.30 NA) oil
-immersion 
objective (Leica) and a CCD camera (retiga Exi, QImaging, Burnaby, BC, C
anada). 2 
"L of cells were pipetted onto a glass slide coated with poly
-lysine and covered with a No. 
1.5 coverslip. Epifluorescence imaging was performed at room temperature. Z
-stacks 
!3Mc!were collected with 0.5 
"m increments and deblurred by performing Nearest Neighbor 
deconvolution at 70% haze removal with Image
-Pro 7.0 software (Media Cybernetics). 
Additional image processing for epifluorescence micrographs was performed using Fiji 
(ImageJ; 
http://fiji.sc/Fiji
). Single
-plane projections were generated from Z
-stacks using 
the maximum intensity algorithm and were falsely colored green for FtsZ1
-mVenus, 
ARC6
-mVenus, and mVenus fluorescence signals. Images depicting FtsZ2
-mCerulean
 or mCerulean fluorescence signals were falsely colored magenta. Colocalization 
overlays were created with the merge tool in Fiji and the white color represents regions 
where the two fluorescence signals overlap. The degree of colocalization in 
coexpressio
n strains was calculated by creating a composite deconvoluted Z
-stack of 
both fluorescence signals, cropping the image to contain only the individual cell being 
analyzed, unmerging the Z
-stacks, and using the Coloc2 tool in Fiji to calculate a 
PearsonÕs co
rrelation coefficient (PCC) for a number of cells 
(Bolte and Cordeli‘res, 
2006). The individual PCC values for each coexpression strain were averaged and a 
standard error of the mean (SEM) was calculated for each coexpression strain.
 Fluorescence Recovery 
After Photobleaching
  Fluorescence Recovery After Photobleaching (FRAP) experiments were 
performed at room temperature on an Olympus FV1000 spectral laser
-scanning 
confocal microscope equipped with a Plan FLN 60X (1.42 NA
) oil
-immersion objective 
and a 3.2X zoom. Immediately before each image acquisition, the high voltage for the 
photomultiplier tube was set so the greatest fluorescence signal in the cell was just 
below saturation. FRAP data were collected with FV1000 ASW 
software (Olympus). 2 
!3Me!"L of liquid S. pombe
 culture expressing a fluorescent fusion protein was pipetted onto 
a glass slide coated with poly
-lysine and covered with a No. 1.5 coverslip. All 
fluorescence signals were photobleached for 20 ms at a laser inten
sity so that one
-half to two-thirds of the fluorescence signal was bleached. Time
-lapse imaging was 
performed with the minimal laser intensity that allowed for adequate fluorescence signal 
intensity but minimized photobleaching. mVenus and mCerulean fluore
scence signals 
were imaged and bleached with the 515 nm and 458 nm lasers, respectively. Three 
images were acquired before the photobleaching event with an interval of 10 s to 
calculate the average pre
-bleach fluorescence intensity. Imaging following the 
photobleaching event occurred for 260 s with an interval of 10 s between image 
acquisitions. Fluorescence intensity measurements were taken at the location of 
photobleaching, at a region of background signal to account for random noise, and at a 
region of f
luorescence signal away from the photobleached location to account for 
photobleaching due to the continual imaging. Each raw FRAP dataset was processed to 
generate a photobleach
-corrected and normalized
 fluorescence recovery dataset, with 1 
being the avera
ge pre
-FRAP fluorescence intensity and 0 being the fluorescence 
intensity at the time of photobleaching 
(Rabut and Ellenberg, 2005)
. Each processed 
FRAP dataset for each fluorescence protein in each expression strain was averaged 
together to generate an av
erage fluorescence recovery dataset. SEM was calculated for 
the normalized average fluorescence recovery at each time point. The normalized 
average FRAP recovery dataset was used for curve
-fitting in pro Fit software 
(QuantumSoft). The data were fit to the
 single exponential f(t) = A(1
-e-kt) equation. The 
!3Md!time it took for fluorescence signals to recover to one
-half of the maximum recovery (t
1/2
) was calculated as t
1/2
 = ln(1/2)/
-k. For statistical comparison of FtsZ2
49-478-eCFP 
turnover properties in the pr
esence and absence of ARC6
68-614-mVenus, the individual 
raw FRAP datasets of FtsZ2
49-478-eCFP in the presence of ARC6
68-614-mVenus were 
normalized and used for curve
-fitting to create t
1/2
 and % recovery data for each FRAP 
run. This dataset was compared to our previous FRAP t
1/2
 and % recovery data, 
prepared in the same manner, for when FtsZ2
49-478-eCFP was expressed alone
 (TerBush and Osteryoung, 2012)
 by StudentÕs t
-test.
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