Download Arrested Differentiation of Proplastids into Chloroplasts in

Wataru Sakamoto1,∗, Yasuyuki Uno1,3, Quan Zhang2, Eiko Miura1, Yusuke Kato1 and Sodmergen2
1Research
Institute for Bioresources, Okayama University, Kurashiki, Okayama, 710-0046 Japan
2Key Laboratory of Cell Proliferation and Differentiation (Ministry of Education), College of Life Sciences, Peking University, Beijing 100871,
PR China
Leaf variegation is seen in many ornamental plants and is
often caused by a cell-lineage type formation of white
sectors lacking functional chloroplasts. A mutant showing
such leaf variegation is viable and is therefore suitable
for studying chloroplast development. In this study, the
formation of white sectors was temporally investigated in
the Arabidopsis leaf-variegated mutant var2. Green
sectors were found to emerge from white sectors after the
formation of the first true leaf. Transmission electron
microscopic examination of plastid ultrastructures
confirmed that the peripheral zone in the var2 shoot
meristem contained proplastids but lacked developing
chloroplasts that were normally detected in wild type.
These data suggest that chloroplast development proceeds
very slowly in var2 variegated leaves. A notable feature in
var2 is that the plastids in white sectors contain remarkable
globular vacuolated membranes and prolamellar bodylike structures. Although defective plastids were hardly
observed in shoot meristems, they began to accumulate
during early leaf development. Consistent with these
observations, large plastid nucleoids detected in white
sectors by DNA-specific fluorescent dyes were characteristic
of those found in proplastids and were clearly distinguished
from those in chloroplasts. These results strongly imply
that in white sectors, differentiation of plastids into
chloroplasts is arrested at the early stage of thylakoid
development. Interestingly, large plastid nucleoids were
detected in variegated sectors from species other than
Arabidopsis. Thus, plastids in variegated leaves appear
to share a common feature and represent a novel
plastid type.
3Present
Keywords: Arabidopsis thaliana • Chloroplast development •
Electron microscopy • Leaf variegation • Plastid nucleoid •
Thylakoid membrane.
Abbreviations: Col, Columbia; cpDNA, chloroplast DNA;
DAPI, 4′,6-diamidino-2-phenylindolyle; FTL, first true leaf;
MRL, mature rosette leaf; PEND, plastid envelope DNA
binding; ptDNA, plastid DNA; PLB, prolamellar body; PVB,
plastidic vacuolated body; SiR, sulfite reductase; SYBG, SYBR
Green I; TEM, transmission electron microscopy; var1, yellow
variegated 1; var2, yellow variegated 2.
Special Issue – Regular Paper
Arrested Differentiation of Proplastids into Chloroplasts
in Variegated Leaves Characterized by Plastid Ultrastructure
and Nucleoid Morphology
Introduction
Leaf variegation is defined as a patch of different colors in
the same leaf and is often seen in a variety of ornamental
plants. When the variegation pattern is identical in each leaf,
it is likely formed by leaf tissues differentiating into specific
cell types. This type of variegation is termed ‘figurative
(or structural) variegation’ (Kirk and Tilney-Bassett 1978).
On the other hand, non-identical variegation is termed ‘true
variegation’ and is frequently caused by genetic mutations
or by environmental stresses such as nutrient deficiency and
pathogen infection. True variegation caused by genetic mutations sometimes follows cell lineage and is characterized by
splashes of green/white sectors (Kirk and Tilney-Bassett
1978, Rodermel 2002). Other non-cell-lineage types show
unequal variegation patterns such as irregularly shaped marginal sectors. Mutations that result in leaf variegation or
stripes have been known to occur in many plant species
for a long time. Such mutants are invaluable resources to
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∗Corresponding
Plant Cell Physiol. 50(12): 2069–2083 (2009) doi:10.1093/pcp/pcp127, available online at www.pcp.oxfordjournals.org
© The Author 2009. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.
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Plant Cell Physiol. 50(12): 2069–2083 (2009) doi:10.1093/pcp/pcp127 © The Author 2009.
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functionally study chloroplast development (Sakamoto
2003, Aluru et al. 2006, Yu et al. 2007). Unlike albino mutants,
which are completely defective in chloroplast development
and thus do not grow into maturity, the formation of green/
white sectors allows us to simultaneously dissect both
sectors. Chloroplast biogenesis is a complex process and
various plastid types are known (Wise 2006, López-Juez 2007,
Sakamoto et al. 2008). Plastids in white sectors may represent a novel plastid type.
In the last decade, many variegation/stripe mutants have
been characterized at the molecular level in model plant
species including Arabidopsis (Carol and Kuntz 2001,
Rodermel 2002, Sakamoto 2003, Aluru et al. 2006, Yu et al.
2007), maize (Han et al. 1992) and tomato (Keddie et al.
1996, Carol and Kuntz 2001). With the exception of mutations caused by transposons and organelle genomes (leading
to a genetic mosaic), most of the mutants are recessive and
are nuclear encoded. The recessive nature of variegation
indicates that two plastid types separated in green/white
sectors are formed in the same genetic background.
Molecular characterization of corresponding genes in these
mutants revealed that variegation is not caused by identical
genes but rather by various redundant mechanisms associated with chloroplast function. While these studies demonstrate that a threshold of chloroplast factors exists and
determines plastid differentiation into chloroplast in a cellautonomous manner, the precise mechanisms leading to
the formation of variegated sectors still remain unclear.
The permanent appearance of variegation sectors in these
mutants also suggests that plastid differentiation into chloroplast is determined at a particular stage of leaf development and is not an irreversible event.
In Arabidopsis, the yellow variegated2 (var2) mutant shows
a variegation pattern typical of cell-lineage types and is thus
an ideal model for studying chloroplast differentiation. It
was originally reported along with var1 by Martínez-Zapater
(1992) and mapped on chromosome 2. The gene responsible
for variegation encodes FtsH2, one of the isoforms in the
chloroplast FtsH, which is localized in the thylakoid
membrane (Chen et al. 2000, Takechi et al. 2000). FtsH is an
ATP-dependent metalloprotease that belongs to the AAA+
protein family (Neuwald et al. 1999, Ito and Akiyama 2005).
It is one of the major prokaryotic proteases in chloroplasts
and performs processive degradation to maintain functional
chloroplasts (Adam et al. 2006, Sakamoto 2006). Plastidic
FtsH forms a hetero-complex with two types of isoform that
are functionally distinguishable: Type A includes FtsH1 and
FtsH5, and Type B includes FtsH2 and FtsH8 (Yu et al. 2005,
Zaltsman et al. 2005b). FtsH levels are coordinately regulated
between the two types (Sakamoto et al. 2003, Yu et al. 2004),
and we confirmed that leaf variegation in var1 (similar
but not as intense as var2) is caused by the lack of FtsH5
(Sakamoto et al. 2002).
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The results in var1 and var2 appear to imply that a threshold of FtsH levels (or levels of other molecules regulated by
FtsH) is an important functional constituent for proper
chloroplast development. Meanwhile, studies on FtsH in
Arabidopsis and Synechocystis demonstrated that FtsH is
involved in avoiding photoinhibition by contributing to the
light-dependent degradation of the Photosystem II (PSII)
reaction center protein D1 (Nixon et al. 2005, Komenda et al.
2007, Kato and Sakamoto 2009). Thus, FtsH is currently
proposed to play a dual role in chloroplasts; an unrelated
role to variegation in the PSII repair cycle and a variegationrelated role in thylakoid formation. In addition, several transacting recessive mutations that suppressed leaf variegation
in var2 have been identified. A majority of the suppressors
appear to act on chloroplast protein synthesis (Miura et al.
2007, Yu et al. 2008). Based upon these findings, the balance
between protein synthesis and degradation was proposed to
mitigate the variegation phenotype. The accumulating
amount of genetic evidence implied more complexity in the
formation of variegation sectors.
In contrast to the intensive genetic studies, little is known
regarding how these abnormal plastids are formed along
with leaf development. Variegation in var2 is only observed
in true leaves but not in cotyledons. It is more severe in the
first emerging leaves than in laterally developing leaves
(Zaltsman et al. 2005a, Kato et al. 2007). In this study, plastid
ultrastructures and nucleoids in var2 were deeply examined
during the early stage of true leaf development. Our results
demonstrate that white sectors contain a novel type of
plastid that is likely formed as a result of the arrest at the
early stage of chloroplast development.
Results
Characteristic features of plastids in var2 white
sectors in mature leaves
We first prepared ultrathin sections of mature rosette
leaves (MRLs) at 30 d after sowing, and examined plastids in
wild type (ecotype Columbia, Col) and var2 variegated
sectors by transmission electron microscopy (TEM). Green
sectors in var2 MRLs contain chloroplasts that were
almost indistinguishable from those observed in wild type
(Fig. 1A, B). In contrast, plastids in white sectors clearly
lacked developed thylakoid membranes and granal stacks,
but instead contained globular vacuolated membrane structures (termed hereafter as ‘plastidic vacuolated body’, PVB).
They appeared to vary in size but were mostly globular and
constituted of a single membrane (Fig. 1C, D, Supplementary Fig. S1). In addition to PVBs, we observed prolamellar
body (PLB)-like structures in MRL plastids in white sectors
(Fig. 1D, E). Subsequently to TEM analysis, we examined
plastid nucleoids, plastid DNA (ptDNA)–protein complexes
that are detectable with DNA fluorescence dyes such as
Plant Cell Physiol. 50(12): 2069–2083 (2009) doi:10.1093/pcp/pcp127 © The Author 2009.
Plastids in variegated leaf sectors
Fig. 1 Characteristic ultrastructures of plastids and their nucleoids observed in var2 mature leaves (30 d old). Lens-shaped chloroplasts detected
in Col (A) and var2 green sectors (B), and a plastid detected in a var2 white sector (C–E). P, PVB. PLB, PLB-like structure. Plastid nucleoids
detected by SYBG stain in Col (F) and var2 (G). In (G), the image was taken at the border between the green and white sectors. Areas corresponding
to single chloroplasts or a plastid are enclosed by red or white circles, respectively. Signals corresponding to nuclei are indicated by yellow
arrowheads. Bars, 1 µm in (A)–(E); 10 µm in (F) and (G).
4′,6-diamidino-2-phenylindolyle (DAPI) and SYBR Green I
(SYBG) (Kuroiwa 1991) (Fig. 1F, G). The plastid nucleoids in
white sectors were large and resembled those in proplastids
(Miyamura et al. 1986, Miyamura et al. 1990, Fujie et al.
1994), suggesting that plastid development into chloroplasts
is arrested in white sectors. The remarkable inner-membrane
structures and nucleoid structures thus appeared to be
characteristic of var2 white sectors. To examine these two
features during an early stage of true leaf development, we
focused on the first true leaf (FTL) and characterized sector
formation in detail.
Formation of green sectors in FTLs in var2
Previous reports by Chen et al. (1999) showed that the variegation pattern in var2 remained unchanged, when mature
leaves were examined. Consistent with these reports, the
variegation patterns that we observed in MRL remained
unchanged at least for 10 d (Supplementary Fig. S2). When
we observed expanding FTL in var2, however, we found that
some green sectors emerged from the white sector (Fig. 2).
FTL was highly variegated in var2 and often looked
completely white. Photographs taken every day after the
emergence of the FTL demonstrated that green sectors
began to form at 7 d after sowing under our growth conditions. The sectors formed at this stage became clear as the
leaf matured (at 9 d) and it appeared permanent in mature
leaves (after 13 d). These findings indicate that at least in
FTL, the differentiation of proplastids into chloroplasts
occurs at approximately day 7.
Plastid ultrastructure during variegation formation
in the FTL in var2
To examine when abnormal plastids are detectable, ultrathin sections were prepared from 6-day-old seedlings (except
for cotyledons and roots) and 8-day-old FTLs in Col and
var2, and were examined by TEM (Figs. 3, 4). No green sectors were visible at this stage (Fig. 2), although cells that
became green sectors could be included in the preparation.
To follow chloroplast development, we first examined
plastids at the peripheral zones of the shoot meristem. Cells
in Col contained mostly globular chloroplasts of ∼2 µm with
a limited number of thylakoids and no starch granules (Fig. 3A).
Plant Cell Physiol. 50(12): 2069–2083 (2009) doi:10.1093/pcp/pcp127 © The Author 2009.
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Fig. 2 Appearance of the green sectors in TFL of var2 mutants. Photographs of identical plant at 7 d (A, E), 9 d (B, F), 11 d (C, G) and 13 d (D, H)
after sowing. (E)–(H) are close-up images corresponding to (A)–(D), respectively. Bars, 2 mm.
In contrast, cells at the peripheral zone in var2 contained
smaller size (∼1 µm) proplastid-like plastids and only few
detectable thylakoids (Fig. 4A). These results suggest that
the formation of chloroplasts, represented by thylakoid
development, proceeds slowly in var2.
In general, a series of developing chloroplasts can be
found by observing cells from the base to the tip of a single
developing leaf (Leese and Leach 1976). In fact, the FTL in Col
showed that chloroplasts became enlarged (from ∼2 to 8 µm)
and lens shaped, contained an increased number of granal
stacks and finally accumulated starch grains (Fig. 3B–E).
When he FTL in var2 was examined, chloroplast development was found to be remarkably different. In var2, proplastids in shoot meristematic cells appeared to develop
thylakoids along with leaf maturation (Fig. 4A, B). As aforementioned, however, this process was slow in comparison
with that in Col. We also found that the middle to the tip parts
of 6-day-old FTLs accumulated abnormal plastids that were
characterized by irregularly shaped envelopes (Fig. 4C, D).
No apparent PVBs and PLB-like structures were detectable
in 6- and 8-day-old FTLs we observed. Because FTLs in var2
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are dominated by white sectors, most of the cells in FTL
contain abnormal plastids (Fig. 4E). We rarely observed
chloroplast-like plastids that retained thylakoids. These
chloroplast-like plastids were found to coexist with abnormal plastids (not shown).
Formation of abnormal plastids in white sectors
Ultrastructures of plastids in Col and var2 during FTL development are summarized in Fig. 5. In Col, the formation of
leaf primordia around the meristem is followed by initial
thylakoid development. During this stage of development,
chloroplasts are globular and thylakoids begin to develop.
Subsequently, chloroplasts became oval and appeared to be
distributed in proximity to plasma membranes. Afterwards,
thylakoid membranes constituted granal networks and
mature chloroplasts accumulated large starch bodies. In
contrast to the normal chloroplast development as described
above, var2 contained many proplastids within the peripheral zones of the shoot meristem. Overall, we observed distinct plastid development in var2 as schematically presented
in Fig. 6. Three features characteristic of var2 were detected.
Plant Cell Physiol. 50(12): 2069–2083 (2009) doi:10.1093/pcp/pcp127 © The Author 2009.
Plastids in variegated leaf sectors
Fig. 3 TEM analysis of chloroplasts in Col. Chloroplasts in 6-day-old plants at the shoot meristem (A), the basal part of FTL (B), the middle part of
FTL (C) and the tip part of FTL (D). Chloroplasts in 8-day-old plants at the tip part of FTL (E). Bar, 2 µm.
Fig. 4 TEM analysis of plastids in var2. Plastids (indicated by red arrowheads) in 6-day-old plants at the shoot meristem (A), the basal part of
FTL (B), the middle part of FTL (C) and the tip part of FTL (D). Plastids in 8-day-old plants at the tip part of FTL (E). Bar, 2 µm.
Plant Cell Physiol. 50(12): 2069–2083 (2009) doi:10.1093/pcp/pcp127 © The Author 2009.
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8 days
6 days
Base
Tip
Middle
var2
Col
Meristem
1µm
Fig. 5 Summarized plastid ultrastructures during leaf development in Col and var2 leaves. Representative images of plastids in 6-day-old
(meristem, base, middle, tip parts of FTL) and 8-day-old (tip part of FTL) plants are shown. In var2, two types of plastid representing abnormal
plastids and chloroplasts that likely form green sectors afterwards (after 6 d, middle part) are indicated as separate rows.
First, we found that abnormal plastids started to accumulate
from the tip to the middle part of the FTL at 6 d. These data
suggested that this is the stage where cells are destined to
undergo normal chloroplast differentiation or a possible
arrest of thylakoid development (feature 1, Fig. 6). Secondly,
cells in green sectors form normal chloroplasts, but this process occurs at a much slower rate than the normal chloroplast development in Col (feature 2, Fig. 6). However,
chloroplast development in var2 appears to proceed into
maturity, since green sectors in mature leaves contain fully
developed chloroplasts. Finally, abnormal plastids accumulate PVBs and PLB-like structures instead of thylakoids,
but these bodies were not formed during an early step of
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leaf development and concomitant formation of abnormal
plastids (feature 3, Fig. 6).
Formation of plastid nucleoids during leaf
development
The morphologies of plastid nucleoids, as detected by DAPI
staining of thin sections, are known to change along with
chloroplast development (Lawrence and Possingham 1986,
Miyamura et al. 1990, Rowan et al. 2004). These changes are
also well correlated with thylakoid development and granal
stacking. Because the observation of plastid nucleoids in
var2 MRL indicated that white sectors contained nucleoids
that were larger and less abundant than those observed in
Plant Cell Physiol. 50(12): 2069–2083 (2009) doi:10.1093/pcp/pcp127 © The Author 2009.
Plastids in variegated leaf sectors
Col
(3)
var2
(1)
(2)
Fig. 6 A schematic representation of the green/white sector formation in var2. Temporal chloroplast development in Col FTL is indicated from
left to right. Green bars represent the degree of grana formation. Three features characteristic of var2 are indicated by numbers. Feature 1 is the
point when the abnormal plastids start to accumulate. This point corresponds to 6 day after sowing. Feature 2 is the fact that chloroplast
development, as represented by granal stacking, proceeds very slowly in var2 compared with Col. Feature 3 is the formation of PVBs in plastids.
PVB is predominantly seen in MRL, but not at the early step of leaf development in FTL.
Fig. 7 Morphological change in plastid nucleoids during leaf development as observed by SYBG-stained thin sections of FTL in Col (A–D) and
var2 (E–H). Cells in 8-day-old plants at the basal part of FTL (A, E), the middle part of FTL (B, F) and the tip part of FTL (C, G). Cells in 10-day-old
plants at the tip part of FTL (D, H). Note that fluorescent signals in each image were incorporated by a digital camera at the identical time setting.
Signals corresponding to nuclei are indicated by yellow arrowheads. Areas corresponding to single chloroplasts or plastids are enclosed by red
circles. Bar, 10 µm.
green sectors (Fig. 1G), we characterized plastid nucleoids
during leaf development in Col and var2 FTLs in this study.
We fixed Col and var2 seedlings at 6 or 8 d from which
roots and cotyledons were removed (Supplementary
Fig. S3A, B). Fixed tissues were embedded into Technovit
resin and a cross-section of FTL was stained with SYBG to
visualize plastid nucleoids (Supplementary Fig. S3C).
In Col, developing chloroplasts at the tip and the middle
parts of the FTL contained a few large nucleoids that were
frequently observed as surrounding the envelope (Fig. 7A, B).
As chloroplasts matured and contained granal stacks (tip
part of 8-day-old leaf), nucleoids became smaller in size and
more abundant (Fig. 7C, D). The nucleoids appeared dense
(images in Fig. 7 were recoded by the same time length) and
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were dispersed in the stroma. In some instances, nucleoids
were observed in proximity to the exterior of thylakoid
membranes. In var2, however, it was evident that abnormal
plastids contained nucleoids whose structures differed from
those in Col. Throughout leaf development, plastids retained
nucleoids that appeared at the initial stage. These nucleoids
did not become smaller in size and did not disperse within
plastids and rather remained as large aggregates (Fig. 7E–H).
In some leaf tips at 6 or 8 d, prominent large signals were
occasionally detected. These results are consistent with our
TEM observation, which revealed that thylakoid formation
is strongly impaired in the plastids of white sectors, suggesting that abnormal plastids resulted from an arrest of proper
chloroplast development.
ptDNA levels in the white sectors
Since our cytological observations were not quantitative, we
assumed that the structural change in plastid nucleoids did
not accompany an overall decrease in ptDNA amounts.
In fact, our previous semi-quantitative PCR analysis did not
reveal significant changes in ptDNA level between green and
white sectors of var2 (Kato et al. 2007). To examine ptDNA
levels carefully, we separately isolated single green and white
protoplasts from both sectors and performed quantitative
PCR (Fig. 8A). Protoplasts from white and green cells were
clearly distinguishable from each other due to the presence
of chlorophyll auto-fluorescence (Fig. 8B–E). The number of
chloroplasts per protoplast was not significantly different
between Col and var2 green sectors (Supplementary
Fig. S4). Single protoplasts from Col were also subjected to
our PCR analysis. The results using two primer sets, detecting ptDNAs corresponding to psbA or ndhA, showed that
relative ptDNA levels were not significantly different between
Col, and green and white sectors in var2 (Fig. 8F). We also
measured the area size of protoplasts and found that cells
in white sectors are smaller than those in green sectors and
F
psbA
2
Relative cpDNA amount
1
ndhA
2
1
Col
var2
green
var2
white
Fig. 8 Estimation of cpDNA levels in protoplasts derived from green and white cells. (A) Capture of protoplasts with a glass capillary (denoted by
red arrowhead, bar = 200 µm). Protoplasts from green sectors (B, C) and from white sectors (D, E) are shown. Images captured by bright field (B, D)
and by G filter set (C, E, detecting chlorophyll autofluorescence) are shown (bar, 10 µm). (F) CpDNA levels estimated by real-time PCR analysis
using a single protoplast from Col, var2 green and white sectors (n = 10). Results from two gene-specific primers (psbA and ndhA) are shown.
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Plastids in variegated leaf sectors
Ipomoea nil
Oryza sativa
Artemisia princeps Pampan.
Felicia amelloides
Habenaria radiata
Ficus benzyamina
Alternanthera ficoidea
Arabis glabra
Fig. 9 Plastid nucleoids detected by DAPI-stained thin sections from variegated leaves. Images of eight species were shown along with the
photograph of each corresponding variegated leaf. For each species, representative images from green sectors (left) and white sectors (right) are
shown (bars, 10 µm). Note that fluorescent signals in each image were incorporated by a digital camera at the identical time setting. Signals
corresponding to nuclei are indicated by yellow arrowheads. Areas corresponding to single chloroplasts or plastids are enclosed by red and white
circles, respectively.
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Col (Supplementary Fig. S5). As a result, our quantitative
PCR with single protoplasts supported our previous observation that larger plastid nucleoids in white sectors do not
reflect an altered amount of ptDNA per cell.
Enlarged plastid nucleoids are commonly found
in variegated leaves
We assumed that enlarged plastid nucleoids are characteristic of undifferentiated plastids lacking thylakoid membranes
and that white sectors in variegated leaves in species other
than Arabidopsis may contain similar plastid nucleoids. To
test this, mature variegated leaves from eight species were
selected to examine plastid nucleoids with DAPI staining.
The pattern and degree of variegation in these species
were variable. Specifically, Ipomoea, Oryza, Artemisia and
Alternanthera species showed cell-lineage type variegation
and other species showed non-cell-lineage type variegation.
Except for the Oryza stripe mutant, it was unclear whether
leaf variegation is caused by a mutation. Nevertheless,
variegated sectors were permanently observed in each
species under our growth conditions in a greenhouse (Fig. 9,
pictures of leaves).
Leaf tissues containing both green and white sectors were
fixed and embedded in Technovit resin and thin sections
were stained with DAPI to visualize plastid nucleoids. We set
the time period unchanged when recording images with
fluorescence microscopy and the images from both sectors
were compared (Fig. 9). In all species, green sectors were
found to contain small/dense DAPI signals that were dispersed within stroma (Fig. 9, red circles in left panels). In
strong contrast however, white sectors contained enlarged
DAPI signals that were brighter and resembled those
observed in var2 white sectors (Fig. 9, white circles in right
panels). Similar to those in var2, the enlarged nucleoids
detected in cell-lineage type species (e.g. Artemisia and
Ipomoea) were located along with the envelope. Given that
enlarged nucleoids represent plastids with undifferentiated
thylakoids, these results suggest that abnormal plastids in
white sectors of variegated leaves generally contain plastids
with arrested thylakoid formation.
Discussion
Formation of green and white sectors in var2
Although plastids in white sectors of variegated leaves
have been previously observed in many species (Kirk and
Tilney-Bassett 1978), there has not been a systematic analysis to examine at what stage abnormal plastids arise. In this
study, we focused on FTLs of Arabidopsis Col and var2, and
characterized sector formation during leaf development by
observing plastid ultrastructure and nucleoid morphology.
Detailed observations of white sectors demonstrated that
the decision for a leaf cell to undergo normal chloroplast
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differentiation is made at a very early stage of leaf development. The FTL in var2 is dominated by white sectors. In the
shoot meristem, Col contained many small globular chloroplasts with thylakoids and a few granal stacks, whereas the
var2 mutant contained proplastids and a limited number of
thylakoids with no stacking. Thus, one notable feature of
var2 that was revealed in this study was a clear delay in
chloroplast development. Given the observation that the
var2 shoot apical meristem predominantly contained proplastids, we conclude that proplastids are normal in shoot
meristems and have a potential to become functional chloroplasts. It should be noted that prior to the appearance of
green sectors, the FTL appeared yellow at 7 d, an observation
that perhaps indicates that proplastids can initiate chloroplast differentiation at the very initial step (Fig. 2E). The
green sectors became more evident as the yellow color
somewhat disappeared (Fig. 2F). Collectively, all of these
data support our hypothesis that chloroplast development
is initiated but arrested at the initial step.
The delayed emergence of green sectors that we observed
in Fig. 2 has not been previously reported in var2, because
previous studies focused on expanded leaves. In this study,
we showed that expanding FTL contains a patch of the cells
that escape from the arrest and undergo chloroplast development (Fig. 2F–H). Namely, the gradual appearance of
green patches in white sectors indicates that chloroplast
development proceeds very slowly in var2. Variegation is
more severe in the FTL than in the second and subsequent
leaves, partly because other FtsHs than FtsH2 are upregulated in the second and subsequent leaves in var2 (Zaltsman
et al. 2005a). Increased FtsH levels in the second and subsequent leaves may be one of the reasons that FTL only showed
a marked emergence of green sectors. The delay of chloroplast development in var2 can be connected to the suppressor mutations in which leaf variegation was rescued (Park
and Rodermel 2004, Miura et al. 2007, Yu et al. 2008, Zhang
et al. 2009). One possibility is that a suppressor mutation can
delay overall chloroplast development in the var2 background. As a consequence, the threshold level of FtsH (or
other factors regulated by FtsH) was mitigated, thereby
allowing most of the mesophyll cells to contain chloroplasts
of normal appearance. In this scenario, it is possible that any
mutations delaying chloroplast differentiation may also act
as a suppressor. It also implies that the balance between the
leaf maturation process and chloroplast development, rather
than the balance between protein synthesis and degradation in chloroplasts, is important for proper chloroplast
formation.
The other important finding from our TEM analysis pertains to PVBs and PLB-like structures that were found in
abnormal plastids. Because the PVBs were predominantly
detected in MRL white sectors, we predicted that PVBs arise
during leaf development. Unexpectedly, their precursor-like
Plant Cell Physiol. 50(12): 2069–2083 (2009) doi:10.1093/pcp/pcp127 © The Author 2009.
Plastids in variegated leaf sectors
structures were hardly detectable in the early stage of FTL
development, suggesting that PVB formation is very slow or
predominantly proceeds in mature leaves. An alternative
possibility is that apparent PVBs are not formed in FTL. This
possibility seems unlikely but not completely ruled out, since
we did not examine FTL at 30 d. Although the origin of PVB
remains unclear, the co-existence of PLB-like structures
raises a possibility that PVBs are derived from PLB-like structures. The PLB is a particular type of membrane structure in
etioplasts and is thought to represent a novel type of thylakoid under dark conditions (Kirk and Tilney-Bassett 1978,
Wise 2006). It is reasonable to assume that the abnormal
plastids in FTL are unable to stay arrested in development
and show characteristics similar to etioplasts by forming
PLBs (Park et al. 2002, Philippar et al. 2007). Supporting this
observation is the previous study by Chen et al. (2000) that
demonstrated that etioplast formation in cotyledons is
comparable between Col and var2. It is possible that the
white plastids in MRL perhaps have a potential to develop
but they fail to form a functional thylakoid network from
PLB-like structures due to the lack of photosynthetic pigments and other protein components. As a result, these
defective plastids form PVBs instead. Thus, our observations
demonstrate that plastids in the white sectors of variegated
leaves represent a novel plastid type. Membrane structures
like PVBs in a mutant defective in chloroplast development
have been previously reported in other studies (Motohashi
et al. 2001, Abdelkader et al. 2007, Okegawa et al. 2007). On
the other hand, we did not detect invaginated membrane
structures along with the inner envelope, which were
reported in several mutants defective in thylakoid formation
(Kroll et al. 2001, Kobayashi et al. 2007). We do not assume
that PVBs directly derive from inner envelope.
Nucleoid structures in variegated leaves
Similar to our TEM analysis, our initial characterization of
plastid nucleoids was carried out only in MRL. Therefore we
decided to observe plastid nucleoids during FTL development in this study. As previously reported, a series of sections in 6- and 8-day-old FTLs demonstrated that plastid
nucleoids follow the typical morphological alteration
(Kuroiwa 1991, Fujie et al. 1994, Sakamoto et al. 2008). Immature chloroplasts tend to contain nucleoids that are large,
less packed and dispersed along with the inner envelope.
As the chloroplasts mature with many granal stacks, the
nucleoids appeared to move in stroma and they become
small and packed (Fig. 7A–D). It is evident that in var2, the
prominent large SYBG signals remain in abnormal plastids of
white sectors (Fig. 7E–H). These results are consistent with
our previous observation and support our hypothesis that
thylakoid development is arrested in white sectors.
DNA compaction in nucleoids was suggested to affect
gene expression (Sekine et al. 2002). In contrast, our previous
study indicated that expression of plastid genes is not significantly altered between green and white sectors in var2 (Kato
et al. 2007). It thus appears that the altered nucleoid morphologies in var2 white sectors do not affect plastid gene
expression at the transcript level. Some chloroplast proteins
were biochemically identified to interact with ptDNAs and
constitute plastid nucleoids (Sato et al. 2003, Sakai et al.
2004, Sakamoto et al. 2008). Those include, for example,
plastid envelope DNA-binding protein (PEND; Sato et al.
1998) and sulfite reductase (SiR; Sato et al. 2001), MFP1
(Jeong et al. 2003) and CND41 (Murakami et al. 2000) whose
precise roles in nucleoid formation and gene expression
remain unclear. PEND is located in the inner envelope and
was shown to anchor ptDNAs to the plastid envelope.
A study using truncated PEND–green fluorescent protein
showed that nucleoids change their morphology within different tissues (Terasawa and Sato 2005). Large plastid nucleoids detected in white sectors may be associated with PEND
since they are located along with the envelope. DAPI-stained
plastid nucleoids were also studied previously by Fujie et al.
(1994). Plastid nucleoids detected in 6- and 8-day-old
Col FTL in this study were similar to those reported in the
previous studies. Plastid nucleoids detected in white tissues
resemble those from non-photosynthetic tissues such as
roots and petals (Fujie et al. 1993). Once again, these data
indicate that nucleoid morphologies are well correlated with
the development of thylakoid membranes.
Although our focus was on nucleoid morphology between
the green and white sectors in var2, it appeared to accompany quantitative differences in DNA signals. To clarify this
question, we carefully estimated ptDNA levels using single
protoplasts. However, no significant differences were
detected between Col, white and green sectors. This result
was consistent with our previous semi-quantitative PCR
analysis (Kato et al. 2007) and confirms that alteration of
nucleoid morphologies do not represent ptDNA amounts.
Regarding chloroplast DNA (cpDNA) amounts during leaf
development, contradictory results were reported from different laboratories (Rowan and Bendich 2009). Chloroplasts
in mature leaves from Arabidopsis were shown to contain a
substantially reduced level of ptDNAs based on fluorescence
detection and quantitative RT-PCR (Rowan et al. 2004). On
the other hand, cpDNA levels were reported to remain
unchanged based on Southern blot analysis (Li et al. 2006,
Zoschke et al. 2007). Given that the var2 white sectors contain plastids whose development into chloroplasts is arrested
at the initial step, our data indirectly indicate that ptDNA
amounts do not significantly change throughout chloroplast
development. It is possible, however, that ptDNAs were
unchanged between the two sectors because the number of
plastids decreased in white sectors. These data also suggest
that each plastid in the white sectors indeed contained
a larger amount of DNA than chloroplasts in green sectors.
Plant Cell Physiol. 50(12): 2069–2083 (2009) doi:10.1093/pcp/pcp127 © The Author 2009.
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W. Sakamoto et al.
We were unable to exclude this possibility since the plastid
number in white cells was not determined.
Besides the quantitative change of ptDNA amounts, we
raised a possibility in this study that many variegated sectors, other than var2 in Arabidopsis, display large plastid
nucleoids due to the impairment of normal thylakoid development. The examination of variegated leaves from different species, including both cell-lineage and non-cell-lineage
types, demonstrated that white sectors contain large nucleoids as expected. It is unclear whether variegated leaves
examined in this study were due to a genetic deficiency
(except for rice) or from physiological effects. In addition,
cell area size, number of chloroplasts per cell and the size of
chloroplasts varied among these species. Nevertheless, white
sectors always tended to be abundant with large nucleoids
that were clearly distinguishable from those in the green
sectors. Our results indicate that despite various patterns
found among different variegated leaves, white sectors have
common features that represent an arrest in thylakoid
differentiation.
Materials and Methods
Plant materials and growth conditions
Arabidopsis thaliana ecotype Columbia (Col) was used as
a wild-type plant in this study. For observing leaf variegation
in var2, the var2-1 allele was used (with the exception that
photos shown in Supplementary Fig. S2 used var2-6).
This var2-1 allele possesses a nonsense mutation at Gln597
(Sakamoto et al. 2004). Sterilized seeds were germinated
and grown on 0.7% (w/v) agar plates containing Murashige
and Skoog medium supplemented with Gamborg’s vitamins
(Sigma-Aldrich) and 1.5% (w/v) sucrose. After stratification
for 2 or 3 d at 4°C under darkness, plates were maintained
under 12 h light (approximately 100 µ mol/m2/s at a constant temperature at 22°C. For further analysis, plants were
transferred to soil after 4–5 weeks. Additional sources for
variegated plants used in Fig. 9 (with the exception of the
rice stripe mutant) were purchased from a local nursery and
subsequently grown in a greenhouse at the Research Institute for Bioresources, Okayama University. The rice stripe
mutant (st-5, mutation mapped on chromosome 4) was
generously provided by Dr Masahiko Maekawa (Okayama
University).
Transmission electron microscopy
Leaf samples for TEM were prepared as follows. For 6-day-old
FTLs, cotyledons and roots were removed from plate-grown
seedlings and the whole remaining leaf tissues were fixed. For
8-day-old FTLs, only middle and tip regions from the FTLs
were excised and fixed for microscopic observation. For
30-day-old MRLs, a 1 × 1 mm region of leaves was excised and
fixed. Leaf samples were fixed in 4% (v/v) glutaraldehyde in
2080
cacodylate buffer (pH 7.4) for at least 24 h at room temperature. After four separate rinses with cacodylate buffer, the
samples were treated with 2% (w/v) osmium tetraoxide for
4–6 h at room temperature. After four separate rinses with
cacodylate buffer, the samples were stained by 2% (w/v)
uranyl acetate in 50% (v/v) ethanol for 1 h at room temperature. Samples were further dehydrated with a graded ethanol series [50%, 70%, 85%, 95%, 100% (v/v)]. Ethanol was
subsequently replaced by a series of Spurr’s resin dilutions
[25%, 50%, 75%, 100% (v/v)] and the resin was hardened
for 16 h at 65°C. Ultrathin sections (60–70 nm) were made
with an Ultracut N (Reichert-Nissei). Thin sections were
stained with 0.25% (w/v) lead citrate and examined with
a JEM-1010 JEOL transmission electron microscope (at the
College of Life Sciences, Peking University.
Technovit thin sections and staining with SYBG
and DAPI
For preparation of Arabidopsis sections, cotyledons and
roots were removed from 6- and 8-day-old seedlings under
a dissection microscope (as illustrated in Supplementary
Fig. S3). The leaf tissues were fixed in FAA solution [5% (v/v)
Formalin, 5% (v/v), acetic acid, 45% (v/v) ethanol] for at
least 3 h or stored at 4°C. For other plant species, a 1 × 2 mm
region of leaves containing green and white sectors was
excised prior to fixation. Fixed tissues were pre-stained by
2.5% (w/v) safranin and washed in 0.1 M phosphate-buffered
saline (PBS pH 7.4). The tissues were dehydrated by a graded
series of ethanol dilutions (50%, 70%, 80%, 90%, 95%,
99.5%, 100% (v/v). Ethanol was replaced by a series of
Technovit7100 resin dilutions [25%, 50%, 74%, 100% (v/v)].
The samples were kept overnight at room temperature and
the resin was finally hardened overnight at 4°C. Thin sections (0.9 µm) were stained with 1 µg/ml DAPI (Invitrogen)
or 1 µg/ml SYBRG (Invitrogen) and examined by an Olympus
AX-84-FLBD fluorescent microscope (U-MWU2 filter set for
DAPI and U-MW13 for SYBG). Images were recorded with
a DP71 digital camera (Olympus). Images from green and
white sectors were recorded with identical time settings.
Protoplast preparation and manipulation
Mesophyll protoplasts were prepared as previously
described in Kato et al. (2007). Leaves from Col and var2-1
plants were cut into small pieces and were gently suspended
in enzyme solution [0.1% (w/v) cellulose Onozuka R10
(Yakult), 0.05% (w/v) pectolyase Y23 (Kyowa Chemical
Products), 400 mM mannitol, 10 mM CaCl2, 20 mM KCl,
5 mM EGTA, 20 mM MES, pH 5.7]. After incubation at room
temperature for 1 h, protoplasts were collected by centrifugation at 60 × g for 1 min. The isolated protoplasts were then
resuspended in wash buffer (154 mM NaCl, 125 mM CaCl2,
5 mM KCl, 2 mM MES, pH 5.7). For PCR analysis, protoplasts
were placed on a slide glass and were detected via inverse
Plant Cell Physiol. 50(12): 2069–2083 (2009) doi:10.1093/pcp/pcp127 © The Author 2009.
Plastids in variegated leaf sectors
microscopy (Olympus CKX41N-31PH). Single protoplasts
were collected into a glass capillary (pore size 100 µm) by
using a pressure-controlled PicoPipet device (Alatair Corp).
Protoplasts from green or white sectors in var2 were easily
distinguishable due to the presence or absence of chloroplasts, respectively. Protoplasts from white sectors contained
abnormal plastids that were detectable under bright-field
microscopy, and were separated by protoplasts derived from
epidermal cells. Protoplasts were carefully transferred into
a PCR tube, suspended in 9.2 µl of sterilized water, sonicated
in a water bath for 1 min (Type UR-20P; Tomy Seiko) and
stored at 4°C until further use. For estimating the area of
protoplasts, they were suspended in wash buffer and
observed by fluorescence microscopy. Images of protoplasts
from Col, green and white sectors in var2 were recorded by
digital camera and the area for each protoplast was calculated with Metamorph software (Molecular Devices). For
counting chloroplast numbers, protoplasts were fixed in
0.02% (v/v) glutaraldehyde and immediately squashed on
a slide glass by gently pressing down on the cover glass.
Images of broken protoplasts with intact chloroplasts were
recorded by a digital camera and were then used to count
chloroplasts.
Real-time PCR analysis
The entire protoplast suspension in a PCR tube was used for
RT-PCR reactions (SYBR Premix ExTaq; Takara). RT-PCR was
performed in 20 µl of a reaction mixture as recommended
by the supplier. PCR cycles were set to 50 under the follow
conditions: 94°C for 13 s, 55°C for 15 s and 72°C for 20 s.
LightCycler Software Version 4.0 (Roche Diagnostics) was
used to quantify DNA levels. For detecting cpDNA, two sets
of primers, detecting the region corresponding to psbA
and ndhA were used: psbA-F (5′-TTGCGGTCAATAAGGT
AGGG-3′), psbA-R (5′-TAGAGAATTTGTGCGCTTGG-3′),
ndhA-F (5′-TGAGATCCGCTAAAACAAGG-3′), ndhA-R (5′CTAGCCGATGGGACAAAA-3′)
Supplementary data
Supplementary data are available at PCP online.
Funding
The Ministry of Education, Culture, Sports, Science
and Technology Grant-in-Aid for Scientific Research
(No. 16085207 to W.S.), the Oohara Foundation (to W.S.),
the Asahi Glass Foundation (to W.S.), the Novartis
Foundation (to W.S.) and the National Natural Science
Foundation of China for a Creative Research Group Program
(No. 30421004 to Sod.). Eiko Miura is a postdoctoral fellow
supported by the Japan Society for the Promotion of
Science.
Acknowledgments
We thank Dr Ryo Matsushima for microscopic analysis, Dr
Masahiko Maekawa for providing rice stripe mutants, Dr
Kazuhide Rikiishi for RT-PCR analyses. We also thank Rie
Hijiya for her technical assistance.
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(Received July 17, 2009; Accepted September 8, 2009)
Plant Cell Physiol. 50(12): 2069–2083 (2009) doi:10.1093/pcp/pcp127 © The Author 2009.
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