Download Expression of the Nucleus-Encoded Chloroplast Division Genes and

Expression of the Nucleus-Encoded Chloroplast Division
Genes and Proteins Regulated by the Algal Cell Cycle
Shin-ya Miyagishima,*,1 Kenji Suzuki,2 Kumiko Okazaki,à,2 and Yukihiro Kabeya1
1
Center for Frontier Research, National Institute of Genetics, Shizuoka, Japan
Advanced Science Institute, RIKEN, Saitama, Japan
àPresent address: Department of Life Sciences, Graduate School of Arts and Sciences, University of Tokyo, Komaba 3-8-1,
Meguro-ku, Tokyo 153-8902, Japan
*Corresponding author: E-mail: [email protected].
Associate editor: Charles Delwiche
2
Abstract
Key words: cell cycle, chloroplast division, endosymbiosis, endosymbiotic gene transfer.
Introduction
Mitochondria and chloroplasts function as energy-converting
organelles in eukaryotic cells. Both organelles arose from
endosymbiotic events in which bacterial cells became integrated into primitive eukaryotic cells (Archibald 2009;
Keeling 2011). In addition to these particular organelles,
there are many other endosymbiotic events, which have
integrated new functions into eukaryotic host cells. Specifically, all chloroplasts (and nonphotosynthetic plastids)
trace their origin to a primary endosymbiotic event that
took place more than a billion years ago in which an ancestral cyanobacterium was integrated into a previously
nonphotosynthetic eukaryote possessing mitochondria.
The ancient alga, which resulted from this primary endosymbiotic event, evolved into the Glaucophyta (glaucophyte algae), Rhodophyta (red algae), and Viridiplantae
(green algae, streptophyte algae, and land plants), which
together are grouped as the Plantae or Archaeplastida. After these primitive green and red algae had been established, chloroplasts then spread into other lineages of
eukaryotes through secondary endosymbiotic events in
which a red or a green alga became integrated into a previously nonphotosynthetic eukaryote (Archibald 2009;
Keeling 2011).
In a manner reminiscent of their free-living ancestor,
chloroplasts have retained the bulk of their bacterial biochemistry and replicate by the division of the preexisting
organelle. However, chloroplasts have also been substantially remodeled by the host cell so as to function as organelles within the eukaryotic host cell (Rodriguez-Ezpeleta
and Philippe 2006; Archibald 2009; Keeling 2011). In particular, most of the genes once present in the cyanobacterial
endosymbiont have either been lost or relocated to the nucleus (a process called endosymbiotic gene transfer). A protein import system was developed that translocates
proteins encoded by the nucleus from the cytosol to
the chloroplast. Several transporters that exchange metabolites between the cytoplasm and the chloroplast also
evolved. Chloroplast division ultimately came to be a process tightly regulated by the host cell, which ensured
© The Author 2012. Published by Oxford University Press on behalf of the Society for Molecular Biology and Evolution. All rights reserved. For permissions, please
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Mol. Biol. Evol. 29(10):2957–2970. 2012 doi:10.1093/molbev/mss102
Advance Access publication April 3, 2012
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Research article
Chloroplasts have evolved from a cyanobacterial endosymbiont and their continuity has been maintained by chloroplast
division, which is performed by the constriction of a ring-like division complex at the division site. It is believed that the
synchronization of the endosymbiotic and host cell division events was a critical step in establishing a permanent
endosymbiotic relationship, such as is commonly seen in existing algae. In the majority of algal species, chloroplasts divide
once per specific period of the host cell division cycle. In order to understand both the regulation of the timing of
chloroplast division in algal cells and how the system evolved, we examined the expression of chloroplast division genes
and proteins in the cell cycle of algae containing chloroplasts of cyanobacterial primary endosymbiotic origin (glaucophyte,
red, green, and streptophyte algae). The results show that the nucleus-encoded chloroplast division genes and proteins of
both cyanobacterial and eukaryotic host origin are expressed specifically during the S phase, except for FtsZ in one
graucophyte alga. In this glaucophyte alga, FtsZ is persistently expressed throughout the cell cycle, whereas the expression
of the nucleus-encoded MinD and MinE as well as FtsZ ring formation are regulated by the phases of the cell cycle. In
contrast to the nucleus-encoded division genes, it has been shown that the expression of chloroplast-encoded division
genes is not regulated by the host cell cycle. The endosymbiotic gene transfer of minE and minD from the chloroplast to
the nuclear genome occurred independently on multiple occasions in distinct lineages, whereas the expression of nucleusencoded MIND and MINE is regulated by the cell cycle in all lineages examined in this study. These results suggest that the
timing of chloroplast division in algal cell cycle is restricted by the cell cycle-regulated expression of some but not all of the
chloroplast division genes. In addition, it is suggested that the regulation of each division-related gene was established
shortly after the endosymbiotic gene transfer, and this event occurred multiple times independently in distinct genes and
in distinct lineages.
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Miyagishima et al. · doi:10.1093/molbev/mss102
permanent inheritance of the chloroplasts during the
course of cell division and from generation to generation.
The majority of algae (both unicellular and multicellular)
have just one or at most only a few chloroplasts per cell, and it
is obvious that a direct and precise relationship must exist
between the host cell and chloroplast division. That is, chloroplasts must divide once per cell cycle before the host cell
completes cytokinesis (Possingham and Lawrence 1983). In
contrast, land plants and certain algal species contain dozens
of chloroplasts per cell that divide nonsynchronously, even
within the same cell. Because land plants evolved from algae
(streptophyte algae/charophytes) (Becker and Marin 2009;
Delwiche and Timme 2011), there should have been a linkage
between the cell cycle and chloroplast division in their algal
ancestor that was subsequently lost during land plant evolution. Thus, it is probable that the continuity of chloroplasts in
host cells was originally established by the synchronization of
endosymbiotic cell division with host cell division in an ancient alga, as is seen in existing algae. However, the mechanism underlying the regulation of chloroplast division is
poorly understood. Nevertheless, recent progress on the understanding of the chloroplast division machinery has allowed
an examination of how the host cell regulates the timing of
chloroplast division (Yang et al. 2008; Maple and Moller 2010;
Pyke 2010; Yoshida et al. 2010; Miyagishima et al. 2011).
Recent studies have shown that chloroplast division is
performed by the constriction of a protein complex that
forms at the division site, spanning both the inside and
the outside of the two chloroplast envelope membranes
(Yang et al. 2008; Maple and Moller 2010; Pyke 2010;
Yoshida et al. 2010; Miyagishima et al. 2011). Consistent
with the endosymbiotic origin of chloroplasts, the complex
has retained certain components of the cyanobacterial division machinery, such as FtsZ and ARC6, inside the chloroplast (i.e., on the inner envelope membrane and the
stromal side). In addition, proteins and structures of eukaryotic host origin, such as DRP5B and the PD rings,
are involved in the division complex both inside and outside of the chloroplasts. The formation of the FtsZ ring is
the first known event that occurs at the chloroplast division site. As in bacteria, positioning of the FtsZ ring formation is, at least in part, regulated by Min proteins of
cyanobacterial origin. Normally, all the chloroplast division
genes of cyanobacterial origin are encoded in the nuclear
genome, indicating that these genes were translocated
from the endosymbiont to the host genome. However,
there are exceptions in which a few of the division genes,
such as FtsI, FtsW, SepF, and MinD, are still retained by the
chloroplast genome in certain algal lineages.
The identification of the chloroplast division proteins
provides a strategy for elucidating how the timing of chloroplast division is coupled with the host cell cycle. This requires a determination of when the chloroplast division
genes and proteins are expressed and when the chloroplast
division complex is formed during the algal cell cycle. Previous studies examined the expression patterns of chloroplast division genes using the red alga Cyanidioschyzon
merolae (Takahara et al. 2000; Fujiwara et al. 2009; Yoshida
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et al. 2010), the green alga Chlamydomonas reinhardtii
(Adams et al. 2008), and the diatom Seminavis robusta
(which possesses chloroplasts of a red algal secondary endosymbiotic origin) (Gillard et al. 2008), the cell cycles of
which are synchronized by a 24-h light–dark cycle. It was
shown that transcripts of certain nucleus-encoded chloroplast division genes accumulate during a specific period of
the synchronous cultures when chloroplasts divide. These
studies raised the possibility that the cell cycle–based expression of the chloroplast division genes allows the chloroplast to divide once in a specific phase of the host cell
cycle. Because cells were synchronized by a 24-h light and
dark cycle and chloroplast division occurred early in the
dark period, it is still unclear whether this expression of
chloroplast division genes is directly controlled by the cell
cycle or instead is regulated by light and/or circadian
rhythms.
In this study, we examined the relationship between the
cell cycle and expression of the chloroplast division genes
and proteins in various algal lineages that possess primary
chloroplasts (glaucophytes and red, green, and streptophyte algae). In addition, the expression patterns of the
division-related genes and proteins that are encoded in some
of these algal chloroplast genomes were analyzed. The results show that most but not all the nucleus-encoded chloroplast division genes and proteins, regardless of origin, are
expressed during the S phase and that the expression of the
chloroplast-encoded division genes is not regulated by
the host cell cycle. It is also shown that endosymbiotic
gene transfer of minD and minE occurred on multiple independent occasions in distinct lineages, even though the
expression of nucleus-encoded MIND and MINE and
the corresponding proteins are restricted to S phase in
these distinct lineages (Note that the genes in the chloroplast genome are indicated in lowercase, whereas those in
the nuclear genome are in uppercase.). Our results suggest
that the division synchronization of the host cell and chloroplasts relies on the cell cycle–based expression of a certain
set of chloroplast division genes. The results also suggest
that the cell cycle–based regulation of a division gene expression was established shortly after the gene was transferred into the host nuclear genome and that this event has
occurred on multiple occasions.
Materials and Methods
Algal Culture
Cyanophora paradoxa UTEX555 (NIES-547), Chlorella vulgaris C-27 (NIES-2170), Nephroselmis olivacea NIES-484, and
Mesostigma viride NIES-296 were obtained from the Microbial Culture Collection at the National Institute for Environmental Studies in Japan (NIES). C. paradoxa, C. vulgaris, and
M. viride cells were cultured in C medium. C. reinhardtii 137c
cells were cultured in Sueoka’s high salt medium (HSM).
N. olivacea cells were cultured in AF-6 medium. C. merolae
10D cells were cultured in Allen’s medium. All media were
prepared according to the description provided by the NIES
(http://mcc.nies.go.jp/02medium-e.html).
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For the synchronization of C. merolae, the cells in stationary culture were subcultured to ,1 107 cells/ml
and subjected to a 12-h light/12-h dark cycle (100 lmol
photons m2s1) at 42°C under aeration with ordinary
air (Suzuki et al. 1994). The cells in the second cycle were
used for further analyses. To arrest the cells in the S phase,
a 1/50,000 volume of 10 mg/ml camptothecin solution in
dimethyl sulfoxide (DMSO) was added 8 h after the onset
of the second light period. To arrest the cells in the M
phase, 1/500 volume of 50 mM MG132 solution in DMSO
was added at the onset of the second dark period.
For the synchronization of C. reinhardtii, the cells in log
phase were subcultured to ,2 105 cells/ml and subjected to a 12-h light/12-h dark cycle (100 lmol photons
m2s1) at 24°C under aeration with 1% CO2 in air
(Surzycki 1971). The cells in the second cycle were used
for further analyses. To arrest the cells in the S phase, 2deoxyadenosine was added to the culture at a concentration of 5 mM 8 h after the onset of the second light period.
For the synchronization of C. vulgaris, the cells in log
phase were subcultured to ,5 105 cells/ml and subjected to a 16-h light/8-h dark cycle (100 lmol photons
m2s1) at 24°C under aeration with 2% CO2 in air
(Tamiya et al. 1953). The cells in the third cycle were used
for further analyses.
For the synchronization of C. paradoxa, the cells were
cultured to a cell density of 1 106 cells/ml at 24°C under
continuous light (40 lmol photons m2s1) under aeration with ordinary air. To arrest the cells in the S phase,
a1/1,000 volume of 5 mg/ml aphidicolin solution in DMSO
was added to culture and cells were cultured for 24 h. To
remove aphidicolin, cells were washed twice with fresh medium by centrifugation at 200 g for 10 min and then
cultured under the same conditions as above.
M. viride cells were cultured to a cell density of 2 106
cells/ml at 24°C under continuous light (40 lmol photons
m2s1) on a rotary shaker. One microliter of the culture
was subjected to a l Cell Sorter (Advance, Japan) to fractionate the smaller and larger cells.
Gene Cloning
In this study, we newly cloned C. paradoxa MIND and MINE,
C. vulgaris FTSZ1, MINE, and DRP5B, M. viride PCNA
(AB673435), FTSZ2, DRP5B, and N. olivacea FTSZ2 genes
(The GenBank accession numbers other than that of
M. viride PCNA are listed in supplementary table 1, Supplementary Material online). All the genes were cloned by reverse transcription polymerase chain reaction (RT–PCR)
using cDNA as the template. cDNA was synthesized from
total RNA using a random hexamer with ThermoScript
RT (Invitrogen). For 3#-rapid amplification of cDNA ends
(RACE), cDNA was synthesized using the primer 3race1
(poly-T and an adapter sequence). RNA was extracted as described below.
A partial cDNA sequence of C. paradoxa MIND
(EH035858) and a full-length cDNA sequences of MINE
(EC659992) were found in the expressed sequence tag
(EST) data (Reyes-Prieto et al. 2006). Expression of MINE
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was confirmed by RT–PCR using the primers Cp-MINEf1 and Cp-MINE-r1. The upstream flanking sequence of
the MIND fragment was amplified by degenerate RT–
PCR using Cp-MINE-f1 and Cp-MINE-r1 for the first PCR
and Cp-MINE-f1 and Cp-MINE-r2 for the second nested
PCR.
Partial cDNA sequences of C. vulgaris FTSZ1, MINE, and
DRP5B were obtained by degenerate PCR using the primers
Cv-FTSZ-f1 and Cv-FTSZ-r1 for FTSZ1, Cv-MINE-f1, and CvMINE-r1 for MINE and Cv-DRP5B-f1 and Cv-DRP5B-r1 for
DRP5B, respectively. The downstream flanking sequences
of ftsZ1 were obtained by 3#-RACE using the primers
Cv-FTSZ-race-f1 and 3race2 for the first PCR and CvFTSZ-race-f2 and 3race2 for the second nested PCR. The
downstream flanking sequences of MINE were obtained
by 3#-RACE using Cv-MINE-race-f1 and 3race2 for the first
PCR and Cv-MINE-race-f2 and 3race2 for the second nested
PCR.
Partial cDNA sequences of M. viride PCNA, FTSZ2, and
DRP5B were obtained by degenerate PCR using the primers
Mv-PCNA-f1 and Mv-PCNA-r1 for PCNA, Mv-FTSZ-f1 and
Mv-FTSZ-r1 for FTSZ2, and Mv-DRP5B-f1 and Mv-DRP5Br1 for DRP5B, respectively. The downstream flanking
sequences of DRP5B were obtained by 3#-RACE using
Mv-DRP5B-race-f1 and 3race2.
A partial cDNA sequence of N. olivacea FTSZ2 was obtained by degenerate PCR using the primers No-FTSZ-f1
and No-FTSZ-r1.
Semiquantitative RT–PCR
Cells were harvested from 1 to 10 ml culture by centrifugation and stored at 80°C until RNA extraction. Cells
were disrupted with TRIzol reagent (Invitrogen). For the
disruption of C. reinhardtii, C. vulgaris, and C. paradoxa,
cells were homogenized in TRIzol with acid-washed glass
beads (425–600 lm) by vigorous shaking. After the addition of chloroform and phase separation, total RNA was
extracted from the upper aqueous phase by using an
RNeasy Plant Mini Kit (QIAGEN). After DNaseI treatment,
cDNA was synthesized from the RNA using random hexamer with ThermoScript RT (Invitrogen).
The cDNA synthesized from 5 ng of total RNA was used
for the PCRs. The PCRs were performed using the primers
listed in supplementary table 2 (Supplementary Material
online).
Preparation of Antibodies
The antibodies against C. paradoxa FtsZ, MinD, MinE, and
SepF, C. reinhardtii FtsZ and DRP5B, and C. vulgaris MinD
and MinE were raised in rabbits using the respective recombinant proteins as described previously (Miyagishima
et al. 2003). The cDNA sequence encoding the full-length or
a partial fragment of the respective protein was amplified
by PCR using the primers listed in supplementary table 2
(Supplementary Material online). These PCR products were
cloned into a pET100 expression vector (Invitrogen) and
6xHis fusion polypeptides were expressed in Rosetta
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(DE3) Escherichia coli cells, purified, and used as antigens.
Antibodies were affinity-purified from the respective antisera by using respective recombinant proteins coupled to
a HiTrap NHS-activated HP column (GE Healthcare). To
prepare antibodies against C. vulgaris and C. reinhardtii
Drp5B, both recombinant proteins were mixed and injected into the same rabbits.
Immunoblot Analyses
Cells were harvested from 5 to 50 ml of culture by centrifugation and stored at 80°C until immunoblot analysis.
Cells were suspended in the protein extraction buffer (50
mM Tris, pH 7.5, 8 M Urea, and 0.1% Triton X-100). To
disrupt C. reinhardtii, C. vulgaris, and C. paradoxa, cells were
homogenized in the protein extraction buffer with acidwashed glass beads (425–600 lm) by vigorous shaking
for 10 min. Protein content was determined with Bradford
assay and equal amounts of the total protein were subjected to a immunoblot analysis. Immunoblotting assays
were performed as previously described (Nakanishi et al.
2009). Antibodies against C. merolae Drp5B (Miyagishima
et al. 2003) (1:1000 to detect C. merolae Drp5B), C. reinhardtii FtsZ1 (1:2000 to detect C. reinhardtii and C. vulgaris
FtsZ1) and DRP5B (1:2000 to detect C. reinhardtii and
C. vulgaris Drp5B), C. vulgaris MinD (1:5000 to detect
C. reinhardtii and C. vulgaris MinD) and MinE (1:5000 to
detect C. reinhardtii and C. vulgaris MinE), C. paradoxa FtsZ
(1:2000 to detect C. merolae FtsZ2-1 and C. paradoxa FtsZ),
MinD (1:1000 to detect C. paradoxa MinD), MinE (1:1000
to detect C. paradoxa MinE), and SepF (1:2000 to detect
C. paradoxa SepF) were diluted as indicated. The primary
antibodies were detected by horseradish peroxidase–
conjugated goat anti-mouse or anti-rabbit antibody diluted
to 1:20,000. The signal was detected by SuperSignal West
Pico Chemiluminescent Substrate (Thermo Scientific) and
the VersaDoc 5000 imaging system (Bio-Rad).
Immunofluorescence Microscopy
For the immunofluorescent detection of FtsZ1 in C. reinhardtii, cells were fixed in methanol at 20°C for 10 min and
washed twice with phosphate-buffered saline, or PBS-T
(0.1% Tween 20 in PBS). The cells were permeabilized for
30 min at room temperature with 10% DMSO in PBS-T. After
blocking with 2% bovine serum albumin in PBS-T (blocking
buffer) for 30 min, cells were labeled at room temperature for
2 h with anti-C. paradoxa FtsZ antibodies diluted to 1:2000 in
blocking buffer (anti-C. reinhardtii FtsZ1 antibodies were not
used because of the high background level on immunofluorescence microscopy). Cells were then washed twice with
blocking buffer and incubated with Alexa Fluor 488 goat
anti-rabbit IgG (H þ L) (Invitrogen) diluted in the blocking
buffer at a concentration of 1:1000 at room temperature for
1 h. After washing twice with PBS-T, cells were observed under fluorescence microscopy (BX-51; Olympus).
Immunofluorescent detection of FtsZ in C. paradoxa
and M. viride was performed using a method modified from
that used for C. reinhardtii. Cells were fixed in the fixation
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buffer (3% paraformaldehyde, 50 mM Pipes, pH 6.8, 10 mM
ethylene glycol tetraacetic acid, and 5 mM MgSO4) for 1 h
at room temperature and washed twice with PBS. Instead
of using DMSO, the fixed cells were permeabilized for 30
min at room temperature with 0.1% Triton X-100 in
PBS. For C. patadoxa, cells were further treated with 0.2
mg ml1 lysozyme dissolved in Tris–HCl, pH 7.5, 10 mM
ethylenediaminetetraacetic acid for 10 min at 37°C.
Anti-C. paradoxa FtsZ antibodies were diluted to a concentration of 1:500. Immunofluorescent detection of
FtsZ2-1 in C. merolae was performed as described previously (Miyagishima et al. 2003).
Results
Distribution of the Chloroplast Division Genes of
Cyanobacterial Origin and the Deduced Timing of
Endosymbiotic Gene Transfer
In order to examine and compare the relationship between
the eukaryotic cell cycle and the timing of chloroplast division in the eukaryotic algae that possess primary chloroplasts, in this study, we used the red alga C. merolae, the
green algae C. reinhardtii, and Chlorella vulgaris, the glaucophyte alga Cyanophora paradoxa and the streptophyte
alga Mesostgma viride. C. merolae and C. reinhardtii were
chosen because the whole nuclear and chloroplast genome
sequence data are available and the methods for cell cycle
synchronization by means of a light and dark cycle have
been established (Surzycki 1971; Suzuki et al. 1994). In both
organisms, all the known chloroplast division proteins are
encoded in the nuclear genome (fig. 1B). The other algal
species were chosen because chloroplast genomes have
been fully sequenced and the cyanobacteria-descended
MinD, FtsI, FtsW, and SepF are still encoded in the chloroplast genome (fig. 1B) (it was previously reported that MinE
is encoded in the chloroplast genome of C. vulgaris, but see
below), although the whole nuclear genome sequence data
are not available at present. MinD was shown to regulate
the positioning of the chloroplast FtsZ ring, as does its
cyanobacterial counterpart (Yang et al. 2008; Maple and
Moller 2010; Yoshida et al. 2010; Miyagishima et al.
2011). Although the functions of FtsI, FtsW, and SepF have
not been determined in eukaryotic algae, these proteins
localize at the bacterial division site and have been shown
to be involved in cyanobacterial cell division (Marbouty
et al. 2009a, 2009b).
One of the aims of this study is to examine the relationship between the regulation of chloroplast division gene
expression and the endosymbiotic gene transfer from
the chloroplast to the nuclear genome. Thus, before characterizing each organism separately, we compared the repertoire of the known chloroplast division genes of
cyanobacterial origin and the location of the genes (i.e., nuclear or chloroplast genome) to deduce the timing of endosymbiotic gene transfer (fig. 1B). Among the chloroplast
division genes, FTSZ, ARC6, and MINC (MinC is involved in
the positioning of FtsZ ring in bacteria, but its function in
eukaryotic algae have not been determined) have been
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FIG. 1. Schematic view of the chloroplast division complex and distribution of the chloroplast division proteins of cyanobacterial origin. (A)
Localization of the components of the division complex (modified from Miyagishima et al. 2011). Only the known division site-localized
components and proteins that determine the division site are shown (Yang et al. 2008; Maple and Moller 2010; Yoshida et al. 2010; Miyagishima
et al. 2011). Some lineages possess two types of FtsZ that are involved in chloroplast division. In Viridiplantae, these are called FtsZ1 and FtsZ2. The
division proteins unique to land plants such as PDV1, PDV2, MCD1, and PARC6 are not shown. Although omitted from the diagram, ARC3 is also
involved in the positioning of the FtsZ ring and is encoded in nuclear genomes of land plants and certain lineages of green algae. Because ARC3
was not found in Cyanidioschyzon merolae and Chlamydomonas reinhardtii, in which complete nuclear genome sequences are available, we could
not examine the expression in this study. MinC, FtsW, FtsI, and SepF were shown to be involved in cyanobacterial division (Marbouty et al. 2009a,
2009b), but the function and localization of these proteins in chloroplasts have not been determined. (B) Distribution of the chloroplast division
genes of cyanobacterial origin and the deduced timing of endosymbitotic gene transfer (updated from Miyagishima et al. 2011). The tree topology
is based on Becker and Marin (2009) and Archibald (2009). Branch lengths do not represent phylogenetic distances. Proteins were searched by
Blast against the whole-genome database of C. merolae, Ostreococcus lucimarinus, C. reinhardtii, Physcomitrella patens, and Arabidopsis thaliana. In
other cases, searches were performed in the chloroplast genome databases and EST databases of Cyanophora paradoxa, Guillardia theta,
Nephroselmis olivacea, Chlorella vulgaris, and Mesostigma viride. Because the complete genomic information on the latter five organisms is not
available, there are blank columns. The asterisks indicate the genes that were identified in this study. In the FtsZ column, ‘‘nm’’ indicates that FtsZ
is encoded in the nucleomorph, a relic nucleus that originated from a red algal secondary endosymbiont. ‘‘D,’’ ‘‘E,’’ ‘‘Z,’’ ‘‘6,’’ and ‘‘C’’ indicate the
deduced timing of endosymbiotic gene transfer of minD, minE, ftsZ, arc6 (ftn2 in cyanobacteria), and minC, respectively. C. paradoxa, Cyanophora
paradoxa UTEX555; C. merolae, Cyanidioschyzon merolae 10D; G. theta, Guillardia theta; N. olivacea, Nephroselmis olivacea NIES-484; O.
lucimarinus, Ostreococcus lucimarinus CCE9901; C. vulgaris, Chlorella vulgaris C-27; C. reinhardtii, Chlamydomonas reinhardtii; M. virde, Mesostigma
viride NIES-296; P. patens, Physcomitrella patens; A. thaliana, Arabidopsis thaliana. The GenBank accession numbers of the amino acid sequences
are summarized in supplementary table 1 (Supplementary Material online).
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found only in the nuclear genome, suggesting that these
genes were transferred into the nuclear genome before
the emergence of the extant algal lineages (fig. 1B). In contrast, ftsI, ftsW, and sepF have been found only in the chloroplast genome, suggesting that these genes were lost
instead of being transferred into the host nuclear genome.
The distribution patterns of ftsI and ftsW suggest that these
genes were lost multiple times independently in distinct
lineages (fig. 1B).
The distribution patterns of the MIND (minD) and MINE
(minE) genes suggest that these genes were transferred into
the nuclear genome multiple times independently in distinct lineages, as described below. Using Blast searches of
the EST database of C. paradoxa (Reyes-Prieto et al. 2006),
we found the nucleus-encoded MINE (including full length
of the orf; GenBank accession number EC659992) and
MIND (containing 210 bp of the orf; EH035858) cDNA sequences. Their expression was confirmed by RT–PCR, and
we further cloned a longer MIND cDNA (756 bp of the orf).
Although it was previously reported that MinE together
with MinD is encoded in the chloroplast genome of C. vulgaris (Wakasugi et al. 1997) (C-27, the same strain that was
used in this study), the nucleotide sequence and the deduced amino acid sequence (NP_045873) showed no significant similarity to genes or proteins of other organisms,
including minE genes and MinE proteins in our Blast search.
In addition, we found MINE cDNA (CAB42593) in the previously characterized nuclear cDNA clones of Auxenochlorella protothecoides (Hortensteiner et al. 2000), which is
closely related to C. vulgaris. Therefore, we expected that
MinE would be encoded in the nuclear genome of C. vulgaris and, in fact, nuclear MINE cDNA was found by RT–
PCR. MinE and MinD were not found to be encoded in the
C. merolae genome, suggesting that these genes were lost in
the ancestor. However, both proteins are encoded in the
chloroplast genome of the cryptomonad Guillardia theta,
which possesses chloroplasts of a red algal secondary endosymbiotic origin, suggesting that MinD and MinE were
originally encoded in the chloroplast genome of a red algal
ancestor. Given these distribution patterns of the MIND
(minD) and MINE (minE) genes, endosymbiotic gene transfer of minD probably occurred on at least four independent
occasions in the ancestors of glaucophytes, chlorophyceae
(a group in green algae), Mamiellophyceae (a group in
green algae), and land plants (fig. 1B). Endosymbiotic gene
transfer of minE probably occurred independently at least
twice in the ancestors of glaucophytes and Viridiplantae
(green algae and streptophytes) (fig. 1B). Because the times
of endosymbiotic gene transfer are deduced based on parsimonious considerations, it is likely that there may be
more occasions of independent transfer for each gene.
The Cell Cycle–Regulated Expression of Chloroplast
Division Genes and Proteins in the Red Alga
C. merolae
The unicellular red alga C. merolae contains a single chloroplast per cell, which divides once before cytokinesis
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(fig. 2). Previous studies using C. merolae synchronized
by a light/dark cycle showed that chloroplast division gene
transcripts and proteins accumulate specifically around the
onset of the dark period when chloroplast division commences (Takahara et al. 2000; Fujiwara et al. 2009; Yoshida
et al. 2010). Because of the use of a light/dark cycle, it was
still unclear whether the expression of the chloroplast division genes and proteins is regulated by the cell cycle
rather than light or circadian rhythms. In addition, the timing of chloroplast division was assigned to the G2 and M
phases (Fujiwara et al. 2009; Yoshida et al. 2010), essentially
based on cell morphology observations, but the exact timing of chloroplast division gene expression and chloroplast
division in relation to the cell cycle has not been reported.
In order to examine whether the timing of chloroplast
division is directly controlled by the cell cycle and, if so, to
define the cell cycle stage in which the chloroplast divides,
we arrested the cells in the synchronous culture in the S or
M phase using inhibitors of cell cycle progression. It has
been shown that cell cycle is regulated by circadian
rhythms, but the circadian clock mechanism oscillates independently of cell cycle progression (Goto and Johnson
1995; Mori and Johnson 2001). To arrest the cells in the
S phase, camptothecin, a specific inhibitor of DNA topoisomerase I, was added to the culture (Nishida et al. 2005).
RT–PCR analyses showed that the level of PCNA (an Sphase marker gene; Pcna is involved in leading strand
synthesis during DNA replication, Morgan 2006) remains
constant and CDC20 (an M-phase marker gene; Cdc20
functions in the metaphase-to-anaphase transition, Morgan
2006) is not transcribed, which is in contrast to the culture
without the inhibitor, in which the PCNA transcript disappears after the S phase and then the CDC20 transcript
accumulates (fig. 2B). These results indicate that cells were
efficiently arrested in the S -phase by camptothecin. To arrest the cells in the M phase, MG-132, a proteasome inhibitor, was added to the culture (Nishida et al. 2005). RT–PCR
analyses showed that the level of CDC20 remains constant,
in contrast to the culture lacking the inhibitor, in which the
CDC20 transcript disappears after the M phase (fig. 2B).
These results indicate that cells were efficiently arrested
in the M phase by MG-132.
When the cells were arrested in the S phase, although
the cells did not divide the chloroplasts continued to divide, producing abnormal cells containing four chloroplasts
per cell (fig. 2A), as described previously (Itoh et al. 1996;
Nishida et al. 2005). RT–PCR analyses showed that the transcript levels of all the known chloroplast division genes remained constant after the emergence of the transcripts at
approximately the 12th h in the synchronous culture (fig.
2B). Immunoblot analyses showed that the levels of FtsZ2-1
and Drp5B proteins also remained constant after the first
appearance at approximately 12th h in the synchronous
culture (fig. 2C). In contrast, when the cells were arrested
in the M phase, chloroplast division stopped after one
round of division during the S phase (fig. 2A). RT–PCR
and immunoblot analyses showed that the transcripts of
the chloroplast division genes and the FtsZ2-1 and Drp5B
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FIG. 2. Expression of the chloroplast division genes and proteins during the cell cycle of the red alga Cyanidioschyzon merolae. C. merolae 10D
was used in this study. (A) Progression of the cell cycle in the synchronous culture of C. merolae. Cells were entrained by a 12-h light and 12-h
dark cycle. The cells in the second cycle are shown. To arrest the cells in the S or M phase, camptothecin (an inhibitor of DNA topoisomerase-I)
or MG-132 (an inhibitor of proteasomes) was added to the culture at the indicated time point. (B) Semiquantitative RT–PCR analyses showing
the change in the mRNA levels of the chloroplast division genes during synchronous culture. The chloroplast division genes are boxed. 18S
rRNA (CMQ305R) was used as the quantitative control. PCNA (CMS101C) and CDC20 (CMA138C) were used as the S- and M-phase markers,
respectively. (C) Immunoblot analyses showing the change in the levels of the chloroplast division proteins during synchronous culture.
(D) Immunofluorescence microscopy showing the timing of FtsZ ring formation. The localization of FtsZ is shown by the green fluorescence
and red is the autofluorescence of chlorophyll. Scale bar 5 5 lm (A), 2 lm (D).
proteins disappeared after the S phase, as was the case in
the culture without the inhibitor (fig. 2B and C). These results in C. merolae suggest that 1) the chloroplast division
occurs only during S phase, 2) this is because the chloroplast division complex is formed specifically during the S
phase (fig. 2D), and 3) this regulation is at least in part
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Miyagishima et al. · doi:10.1093/molbev/mss102
based on the expression and degradation of the components of the chloroplast division complex during the S
phase and M phase, respectively.
Cell Cycle–Regulated Expression of the Chloroplast
Division Genes and Proteins in the Green Alga
C. reinhardtii
In order to explore the common features and differences in
the regulation of chloroplast division among the eukaryotic
algae, we examined the relationship between the cell cycle
and the expression of the chloroplast division genes and proteins in the green alga C. reinhardtii. C. reinhardtii cell contains a single chloroplast, and this chloroplast divides during
the S/M phase. Cells grow in size by many fold during the G1
phase and then enter into two to four rounds of S/M phases
to produce 4–16 daughter cells (fig. 3A) (Surzycki 1971). Previous studies using C. reinhardtii synchronized by a light and
dark cycle showed that the levels of the FTSZ1, FTSZ2, MIND,
and MINE transcripts oscillate in the early dark period
(Adams et al. 2008; Hu et al. 2008). Because this oscillation
persisted under either a continuous light or a dark condition
(peaking in the subjective-dark period; under the dark condition, cells still divided in a heterotrophic medium) after
previous entrainment by a light and dark cycle, it was suggested that the expression correlates with the circadian
rhythm (Hu et al. 2008). However, the cell cycle of C. reinhardtii is itself also regulated by the circadian rhythm (Goto
and Johnson 1995), and therefore, it is still unclear whether
the expression of the chloroplast division genes are regulated
by the circadian rhythm or the cell cycle.
The RT–PCR analyses performed using the synchronous
culture showed that the transcripts of all the known chloroplast division genes accumulated specifically during the
S/M phases (in addition to FTSZ1, there were also FTSZ2,
MIND, and MINE, ARC6, MINC, and DRP5B) (fig. 3B). This
pattern was also observed under the continuous light condition after entrainment by a light and dark cycle. When
the cells were arrested in the S/M phase by deoxyadenosine, which has been shown to inhibit DNA synthesis in
C. reinhardtii (Harper and John 1986), the transcript levels
of the chloroplast division genes and an S/M-phase marker
in C. reinhardtii, CYCA (Bisova et al. 2005) remained constant after its appearance at around 12th h (fig. 3B). This
result suggests that the expression of chloroplast division
genes is regulated by the cell cycle. In addition, immunoblot
analyses showed that the FtsZ1, MinD, and Drp5B proteins
specifically accumulate during the S/M phases and then are
degraded (fig. 3C). The protein levels remained constant
when the cells were arrested during the S phase (fig. 3C).
Immunofluorescence microscopy showed that the FtsZ ring
forms only during the S/M phases (fig. 3D). These results suggest that the expression of the chloroplast division machinery components during the S/M phase and degradation of
these proteins after chloroplast division restrict the timing of
chloroplast division complex formation in the green alga C.
reinhardtii in a similar manner as in the red alga C. merolae. In
contrast to C. merolae (fig. 2A), chloroplast division appeared
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to stop during constriction of the division site in C reinhardtii
when the cell was arrested in the S/M phases (fig. 3A), but
the reason is not clear at present.
Relationship between the Cell Cycle and Expression
of the Chloroplast Division Genes and Proteins in
the Green Alga C. vulgaris, in Which the MinD
Protein Is Encoded in the Chloroplast Genome
In both C. merolae and C. reinhardtii, all the chloroplast division genes are encoded in the nuclear genome. In contrast,
a few chloroplast division genes of cyanobacterial origin are
still encoded in the chloroplast genome in certain algal lineages (fig. 1B). In order to examine the relationship between
endosymbiotic gene transfer of chloroplast division genes
from the chloroplast to the nuclear genome and also the
regulation of the gene expression by the host cell cycle,
we examined the expression of chloroplast division genes
and proteins in the green alga C. vulgaris, in which the MinD
protein is encoded in the chloroplast genome, as described
above. C. vulgaris cells contain a single chloroplast and the
cell cycle can be synchronized by a light and dark cycle
(Tamiya et al. 1953). Similar to C. reinhardtii, the cells grow
during the G1 phase and then enter into two to four rounds
of the S/M phases so as to produce 4–16 daughter cells
(fig. 4A) (Tamiya et al. 1953).
RT–PCR analyses of C. vulgaris synchronized by a light
and dark cycle showed that the nucleus-encoded FTSZ1,
DRP5B, and MINE transcripts accumulated around the
time of chloroplast division (fig. 4B). The transcript level
of the chloroplast-encoded minD also oscillated, peaking
around the light/dark transition, which corresponds to
the time of chloroplast division. However, under continuous light after entrainment by a light and dark cycle, the
minD transcript level continued to increase, in contrast to
FTSZ1, DRP5B, and MINE, which decreased after chloroplast division (fig. 4B). These results suggest that the transcript level of the chloroplast-encoded minD changes in
accordance with the light condition rather than the cell
cycle and timing of chloroplast division. Immunoblot
analyses showed that the nucleus-encoded FtsZ1, Drp5B,
and MinE proteins accumulated prior to (MinE) or around
(FtsZ1 and Drp5B) the time of chloroplast division,
whereas the level of the chloroplast-encoded MinD remained constant throughout the cell cycle. This pattern
of chloroplast-encoded MinD protein contrasts with the
nucleus-encoded MinD protein in C. reinhardtii, suggesting that the regulatory system of MinD expression by the
host cell cycle was established after the endosymbiotic
gene transfer of minD from the chloroplast to the nuclear
genome, at least in this lineage of green algae. We could
not observe the timing of the chloroplast division complex formation by immunofluorescence microscopy because of the very thick cell walls of C. vulgaris.
However, the Drp5B protein is expressed exclusively at
the time of chloroplast division, suggesting that the mature division complex forms only during a specific period
of the cell cycle in C. vulgaris.
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FIG. 3. Expression of the chloroplast division genes and proteins during the cell cycle of the green alga Chlamydomonas reinhardtii. C. reinhardtii
137c (Chlorophyceae) was used in this study. (A) Progression of the cell cycle in the synchronous culture of C. reinhardtii. Cells were entrained by
a 12-h light and 12-h dark cycle. The cells in the second cycle are shown. To arrest the cells in the S phase, deoxyadenosine, which blocks DNA
synthesis, was added to the culture at the indicated time point. (B) Semiquantitative RT–PCR analyses showing the change in the mRNA levels of
the chloroplast division genes during synchronous culture. The chloroplast division genes are boxed. EF-1a (XM_001696516) was used as the
quantitative control. CYCA (XM_001693115) was used as the S-phase marker. (C) Immunoblot analyses showing the change in the levels of
the chloroplast division proteins during synchronous culture. (D) Immunofluorescence microscopy showing the timing of FtsZ ring formation.
The localization of FtsZ is shown by the green fluorescence and red is the autofluorescence of chlorophyll. Scale bar 5 10 lm (A), 2 lm (D).
Relationship between the Cell Cycle and Expression
of the Chloroplast Division Genes and Proteins in
the Glaucophyte Alga C. paradoxa
In order to determine whether chloroplast division is regulated by the host cell cycle also in glaucophytes, we examined the expression of the chloroplast division genes
and proteins in the glaucophyte C. paradoxa. Glaucophytes
have retained a peptidoglycan layer between the two chloroplast envelope membranes, unlike the chloroplasts in
other lineages, and are suggested to have branched off earliest in the evolution of the Plantae (Reyes-Prieto and
Bhattacharya 2007). The number of chloroplasts (called cyanelles) per C. paradoxa cell varies from one to six, but the
majority of the cells in our culture contained two or four
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Miyagishima et al. · doi:10.1093/molbev/mss102
FIG. 4. Expression of the chloroplast division genes and proteins during the cell cycle of the green alga Chlorella vulgaris. C. vulgaris C-27
(Trebouxiophyceae) was used in this study. (A) Progression of cell cycle in the synchronous culture of Chlamydomonas reinhardtii. Cells were
entrained by a 16-h light and 8-h dark cycle. The cells in the third cycle are shown. (B) Semiquantitative RT–PCR analyses showing the change
in the mRNA levels of the chloroplast division genes during synchronous culture. The chloroplast division genes are boxed. EF-1a and
chloroplast-encoded rpoA (NC_001865) were used as the quantitative controls. ‘‘N’’ and ‘‘CP’’ indicate genes that are encoded in the nuclear
and chloroplast genome, respectively. (C) Immunoblot analyses showing the change in the levels of the chloroplast division proteins during
synchronous culture. N and CP indicate the proteins that are encoded in the nuclear and chloroplast genome, respectively. Note that the cells
were cultured for 4 more hours in the dark in (B) and (C). Scale bar 5 5 lm.
chloroplasts per cell, as reported previously (Pickett-Heaps
1972; Sato et al. 2007). We tested several conditions in an
effort to synchronize the cell cycle with a light and dark
cycle, but we did not achieve synchronization. Therefore,
we synchronized the cells by arresting the cells in the S
phase with aphidicolin and restarting the cell cycle by removing aphidicolin under a continuous light condition.
Cells during cytokinesis were predominantly observed
around 6 h after aphidicolin removal (fig. 5A), suggesting
that the cells had entered into M phase synchronously. RT–
PCR analyses of an S-phase marker, PCNA, and an M-phase
marker, CYCB (encoding ann M-phase cyclin), also showed
that the culture was synchronized (fig. 5B).
RT–PCR showed that throughout the cell cycle, the transcript levels of the nucleus-encoded MIND and MINE
changed, peaking during the S phase (fig. 5B), whereas
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the transcript levels of the nucleus-encoded FTSZ and
the chloroplast-encoded ftsW and sepF stayed constant
(fig. 5B). Immunoblot analyses showed that levels of the
MinD and MinE proteins change, peaking during S phase,
whereas the FtsZ level is constant throughout the cell cycle
(fig. 5C), as are levels of the corresponding transcripts
(fig. 5B). In contrast to the constant expression of sepF transcript (fig. 5B), the SepF protein level changed, peaking during the S phase (fig. 5C). Although FtsZ level remained
constant during the cell cycle, the FtsZ rings were predominantly observed in cells during the S phase (fig. 5D). These
results indicate that the transcription of chloroplastencoded division genes is not regulated by the host cell
cycle as in the case of C. vulgaris minD. The results also
indicate that the host cell is able to determine the timing
of chloroplast division by the cell cycle–based regulation of
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FIG. 5. Expression of the chloroplast division genes and proteins during the cell cycle of the glaucophyte alga C. paradoxa. C. paradoxa UTEX555
was used in this study. (A) Progression of the cell cycle in the synchronous culture of Cyanophora paradoxa. Cells were arrested in the S phase
with aphidicolin (an inhibitor of DNA polymerase) for 24 h under continuous light and then the cell cycle was restarted under continuous light
along with the removal of aphidicolin. (B) Semiquantitative RT–PCR analyses showing the change in the mRNA levels of the chloroplast
division genes during synchronous culture. The chloroplast division genes are boxed. EF-1a was used as the quantitative control. Pcna and cycB
were used as the S- and M-phase markers. ‘‘N’’ and ‘‘CP’’ indicate the genes that are encoded in the nuclear and chloroplast genome,
respectively. (C) Immunoblot analyses showing the change in the levels of the chloroplast division proteins during synchronous culture. N and
CP indicate the proteins that are encoded in the nuclear and chloroplast genome, respectively. (D) Immunofluorescence microscopy showing
the timing of FtsZ ring formation. The localization of FtsZ is shown by the green fluorescence and red is the autofluorescence of chlorophyll.
Scale bars 5 10 lm (A and D).
not all but only some of the nucleus-encoded chloroplast
division genes/proteins.
Relationship between the Cell Cycle and Expression
of the Chloroplast Division Genes and Proteins in
the Streptophyte Alga M. viride
We have examined the regulation of the timing of chloroplast division in three major groups of eukaryotic algae,
which possess primary chloroplasts (glaucophytes, as well
as red and green algae). Finally, we examined the relationship in the streptophyte alga M. viride, one of the earliest
diverging members of streptophytes, which include certain
algal lineages and land plants (Becker and Marin 2009;
Delwiche and Timme 2011). M. viride contains a single
cup-shaped chloroplast per cell. Because we could not synchronize M. viride by a variety of light and dark cycles or
drug treatments, we fractionated the nonsynchronous proliferating cells under continuous light using a microfluidic
device into two fractions containing smaller and larger
cells, respectively (fig. 6A). The fractionation yielded a sufficient number of cells for RT–PCR analysis but not immunoblotting. RT–PCR confirmed that cells in the S phase
(expressing PCNA) and M phase (expressing CYCB) are enriched in the fraction of larger cells (fig. 6B). The analyses
showed that the transcripts of the nucleus-encoded FTSZ
and DRP5B are also enriched in the fraction of larger cells,
whereas the transcripts of the chloroplast-encoded minD,
ftsI, and ftsW are evenly distributed in the smaller and larger
cell fractions (fig. 6B). Immunofluorescence microscopy revealed that the FtsZ ring exists only in the larger cells (fig.
6C). These results indicate that the formation of the chloroplast division complex and transcription of the nucleusencoded but not chloroplast-encoded chloroplast division
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FIG. 6. Expression of the chloroplast division genes during the cell
cycle of Mesostigma viride. M. viride NIES-296 (Streptophyta,
Mesostigmatophyceae) was used in this study. (A) Smaller cells
and larger dividing cells were separated by a microfluidic device
from log-phase culture under continuous light. (B) Semiquantitative
RT–PCR analyses comparing the mRNA levels of the chloroplast
division genes between the smaller cells (S) and the larger dividing
cells (L). Chloroplast division genes are boxed. EF-1a (DQ394295)
was used as the quantitative control. PCNA (DN262911) and CYCB
(EC728922) were used as the S- and M-phase markers. ‘‘N’’ and ‘‘CP’’
indicate the genes that are encoded in the nuclear and chloroplast
genome, respectively. (C) Immunofluorescence microscopy showing
the timing of FtsZ ring formation. The localization of FtsZ is shown
by green fluorescence. The differential interference contrast images
and fluorescent images are merged. Scale bar 5 20 lm (A) and
10 lm (C).
genes are regulated by the host cell cycle in M. viride, as had
been observed in the green alga C. vulgaris and the glaucophyte alga C. paradoxa.
Discussion
Regulation of the Chloroplast Division Genes by the
Algal Cell Cycle
Because cells in many algal lineages contain just one or only
a few chloroplasts per cell, chloroplasts have to be duplicated during a specific phase of the cell cycle before cytokinesis. Thus, there is a correlation between the timing of
chloroplast division and the cell cycle. Previous studies
showed that the transcripts (and proteins in the red alga
C. merolae) of certain chloroplast division genes accumulate during a period of chloroplast division in 24-h light/
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dark synchronous culture of a few species of algae (Takahara et al. 2000; Adams et al. 2008; Gillard et al. 2008; Hu
et al. 2008; Fujiwara et al. 2009; Yoshida et al. 2010). However, it had remained unclear whether this gene expression
is directly controlled by the cell cycle or is regulated by light
and/or circadian rhythms. In this study, we showed that the
expression of the chloroplast division genes and proteins is
controlled by the cell cycle by examining cells arrested in
the S phase or M phase. This regulation was observed in
glaucophytes as well as red, green, and streptophyte algae,
which include all the lineages of algae possessing primary
chloroplasts. In addition, the results showed that the expression of all the known nucleus-encoded chloroplast division genes and proteins (i.e., components of the
chloroplast division complex and its direct regulators) of
both cyanobacterial and eukaryotic host origin, except
for FtsZ in C. paradoxa, are regulated by the host cell cycle.
Based on these results, there is probably a mechanism
that controls the transcription of chloroplast division genes
in the nuclear genome specifically during the S phase,
which results in the expression of these proteins. In addition, nucleus-encoded chloroplast division proteins (except
for C. paradoxa FtsZ and chloroplast-encoded SepF proteins) disappeared in the late M phase, suggesting that degradation of the chloroplast division proteins also likely plays
a role in the regulation of chloroplast division. At present, it
is not clear whether this degradation is just because of the
instability of the proteins or involves proteases or other
systems for proteolysis specific to the chloroplast division
proteins. If the latter is the case, there should be at least two
kinds of proteases or mechanisms, which are responsible
for the degradation of the stromal (FtsZ, MinD, MinE,
and SepF) and cytosolic (Drp5B) chloroplast division proteins, respectively. Because Drp5B disappeared during the
M phase in C. merolae even when cells were arrested by the
proteasome inhibitor MG-132, it is unlikely that proteasomes are responsible for the degradation of Drp5B.
In contrast to the nucleus-encoded division genes and
proteins, the expression of the chloroplast-encoded division genes and proteins was found to not be regulated
by the host cell cycle. In addition, nucleus-encoded FtsZ
in C. paradoxa was expressed throughout the cell cycle,
whereas FtsZ ring formation was restricted to the S phase.
Thus, the regulation of the expression of some but not all
the chloroplast division genes and proteins are apparently
sufficient to restrict the timing of chloroplast division to
a period between the S phase and late M phase. It is possible that posttranslational modification of the chloroplast
division proteins is also involved in the regulation of the
timing of chloroplast division, such as phosphorylation
or processing that is observed in mitochondrial division
proteins (Nishida et al. 2007; Taguchi et al. 2007; Yoshida
et al. 2009). Nevertheless, the regulation of both expression
and the degradation of certain chloroplast division proteins
is apparently sufficient to inhibit chloroplast division in
cells during the G1 phase and late M phase because the
cells do not express the set of components that is required
to form a functional chloroplast division complex.
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The Relationship between the Expression of the
Chloroplast Division Genes and Endosymbiotic
Gene Transfer
Unlike the nucleus-encoded division genes and proteins, it
was shown that the chloroplast-encoded division genes
and proteins do not correlate with the host cell cycle.
One exception was the SepF protein in C. paradoxa, which
predominantly accumulates during S phase, but the level of
the sepF transcript remained constant through the cell cycle. Thus, it is suggested that the system for the transcriptional regulation of each chloroplast division gene by the
host cell cycle was established after endosymbiotic gene
transfer into the host nuclear genome. In addition, despite
the fact that MIND was independently transferred into the
nuclear genome at least four times and MINE was transferred at least twice in distinct lineages, the expression
of these genes came to be regulated by the host cell cycle
in each case. These observations indicate that there is an
evolutionary trend, such that a division gene comes under
the regulatory control of the cell cycle after endosymbiotic
gene transfer, even though there are exceptions, such as
FTSZ in C. paradoxa. It should be noted that the parallel
independent endosymbiotic gene transfers predicted for
minD and minE are not a rare occurrence. A previous study
indicated that both independent gene loss and transfer
have occurred frequently in the chloroplast genome (Martin
et al. 1998).
In general, a gene transferred from an organelle to the
nuclear genome must acquire a eukaryotic promoter in order to be transcribed by the eukaryotic transcription machinery. In addition, the gene also needs to acquire a transit
peptide-coding sequence in order to be imported into the
organelle (Timmis et al. 2004). In the case of chloroplast
division genes, each gene most likely acquired a promoter
that is regulated by the cell cycle. In this regard, the immediately upstream genomic region of FTSZ and DRP5B in
C. merolae possesses sequences that are similar to an
E2F-biding site, TTTCGCGC, which is known to be recognized
by the S-phase transcription factor E2F (Morgan 2006). Characterization of the promoter region and transcription factors
of the chloroplast division genes are expected to yield important insights into this matter in the near future.
This study suggests that endosymbiotic gene transfer
and subsequent acquisition of transcriptional regulation
by the host cell cycle have contributed to the synchronization of the host cell and primary chloroplast division. An
additional question is whether this scheme is applicable to
other endosymbiotic systems, such as mitochondria, secondary chloroplasts, and the chromatophore of Paulinella
chromatophore (Marin et al. 2005). In C. merolae, the transcripts of the nucleus-encoded mitochondrial division
genes DRP3, MDA1, ZED, and a-proteobacteria-descended
FTSZ along with the corresponding proteins accumulate in
a specific period of the 24-h light–dark cycle when the mitochondria divide (Nishida et al. 2007; Fujiwara et al. 2009;
Yoshida et al. 2009). In the diatom (a stramenopile)
Seminavis robusta, a single cell of which contains two
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chloroplasts of a red algal secondary endosymbiotic origin,
the nucleus-encoded FTSZ transcript accumulates in a specific period of the 24-h light–dark cycle in which the chloroplasts divide (Gillard et al. 2008). Thus, the scheme
presented in this study for primary chloroplasts appears
to be applicable to secondary chloroplasts and mitochondria. In addition, a similar scheme might also be applicable
to the regulation of eukaryotic cell cycle of an endosymbiont by a eukaryotic host cell. Chlorarachniophyte algae
possess a relic nucleus (nucleomorph) derived from a green
algal endosymbiont. In Bigelowiella natans, nuceomorph division occurs during a specific period before mitosis
(Moestrup and Sengco 2001). A recent study showed that
endosymbiont-derived histone proteins are encoded in the
host nuclear genome and are targeted to the nucleomorph.
Transcripts of these histone genes were shown to accumulate during the time corresponding to S phase in a culture,
which was partially synchronized by a 24-h light–dark cycle
(Hirakawa et al. 2011). Further study of the mechanism of
cell cycle–based expression of chloroplast division genes
and other endosymbiotic systems will yield important insights into the common features that underlie the establishment and evolution of permanent endosymbiotic
relationships.
Supplementary Material
Supplementary tables 1 and 2 are available at Molecular
Biology and Evolution online (http://www.mbe.
oxfordjournals.org/).
Acknowledgments
We thank Y. Ono and A. Yamashita for technical support
and BSI’s Research Resources Center of RIKEN for production of antibodies. We are also grateful to National Institute
for Environmental Studies (NIES) for providing some algal
strains. This study was supported by grant-in-aid for Scientific Research from Japan Society for the Promotion of Science (no. 19870033 to S.M.) and by Core Research for
Evolutional Science and Technology (CREST) Program of
Japan Science and Technology Agency (JST) (to S.M.).
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