Differential Positioning of C4 Mesophyll and Bundle Sheath
Chloroplasts: Aggregative Movement of C4 Mesophyll
Chloroplasts in Response to Environmental Stresses
Regular Paper
Masahiro Yamada1, Michio Kawasaki2, Tatsuo Sugiyama3, Hiroshi Miyake1 and Mitsutaka Taniguchi1,∗
1Graduate
School of Bioagricultural Sciences, Nagoya University, Nagoya, Aichi, 464-8601 Japan
of Agriculture and Life Science, Hirosaki University, Hirosaki, Aomori, 036-8561 Japan
3Research Institute of Life and Health Sciences, Chubu University, Kasugai, Aichi, 487-8501 Japan
2Faculty
In C4 plants, mesophyll (M) chloroplasts are randomly
distributed along the cell walls, while bundle sheath (BS)
chloroplasts are typically located in either a centripetal or
centrifugal position. We investigated whether these
intracellular positions are affected by environmental
stresses. When mature leaves of finger millet (Eleusine
coracana) were exposed to extremely high intensity
light, most M chloroplasts aggregatively re-distributed to
the BS side, whereas the intracellular arrangement of
BS chloroplasts was unaffected. Compared with the
homologous light-avoidance movement of M chloroplasts
in C3 plants, it requires extremely high light (3,000–
4,000 µmol m−2 s−1) and responds more slowly (distinctive
movement observed in 1 h). The high light-induced
movement of M chloroplasts was also observed in maize
(Zea mays), another C4 species, but with a distinct pattern
of redistribution along the sides of anticlinal walls,
analogous to C3 plants. The aggregative movement of M
chloroplasts occurred at normal light intensities (250–
500 µmol m−2 s−1) in response to environmental stresses,
such as drought, salinity and hyperosmosis. Moreover, the
re-arrangement of M chloroplasts was observed in fieldgrown C4 plants when exposed to mid-day sunlight, but
also under midsummer drought conditions. The migration
of M chloroplasts was controlled by actin filaments and
also induced in a light-dependent fashion upon incubation
with ABA, which may be the physiological signal
transducer. Together these results suggest that M and BS
cells of C4 plants have different mechanisms controlling
intracellular chloroplast positioning, and that the
aggregative movement of C4 M chloroplasts is thought to
∗Corresponding
be a protective response under environmental stress
conditions.
Keywords: C4 photosynthesis • Chloroplast • Eleusine
coracana • Environmental stress • Photo-relocation
movement • Zea mays.
Abbreviations: BS, bundle sheath; DAPI, 4′,6-diamidino-2phenylindole; DMSO, dimethylsulfoxide; M, mesophyll; ME,
malic enzyme.
Introduction
Chloroplasts can change their intracellular positions to optimize photosynthetic activity and/or to reduce photodamage in response to light irradiation (Takagi 2003, Wada et al.
2003, Sato and Kadota 2007). Thus, under high intensity
light irradiation, chloroplasts move away from light to minimize photodamage, while under low intensity irradiation
they move toward the light to maximize photosynthesis.
The motility and positioning of chloroplasts appear to be
mediated by actin filaments and/or microtubules (Wada
et al. 2003, Sato and Kadota 2007). A spatial reorganization
of actin filaments occurs during light-dependent redistribution of chloroplasts. Actin filaments not only provide tracks
for chloroplast movement but also anchor the chloroplasts
after photo-orientation (Takagi 2003). These chloroplast
photo-relocation movements are widely observed in a variety of plant species, from green algae to seed plants, although
little attention has been given to C4 plants. There is one
report that some monocotyledonous C4 plants show the
chloroplast photo-relocation movement in response to blue
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Plant Cell Physiol. 50(10): 1736–1749 (2009) doi:10.1093/pcp/pcp116, available online at www.pcp.oxfordjournals.org
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Stress-responsive movement of C4 chloroplasts
light (Inoue and Shibata 1974). However, particular behavior
of the chloroplasts was not described.
C4 plants such as maize and finger millet have two types
of photosynthetic cells, mesophyll (M) and bundle sheath
(BS). Both cell types are arranged into a specialized Kranztype leaf anatomy: BS cells surround the vascular tissues
while M cells encircle the cylinders of the BS cells. The C4
dicarboxylate cycle of photosynthetic carbon assimilation is
distributed between the two cell types and acts as a CO2
pump to concentrate CO2 in the BS chloroplasts (Hatch
1999, Kanai and Edwards 1999). The site of decarboxylation
to feed CO2 to BS chloroplasts is different between C4 subtypes; in NADP-malic enzyme (ME) type species donation of
CO2 from C4 acids occurs in BS chloroplasts while in NAD-ME
type species it occurs in mitochondria in BS cells. M and BS
cells have well developed and numerous chloroplasts. Just as
the M and BS chloroplasts are structurally and functionally
differentiated, so their intracellular orientation is also different: M chloroplasts of all C4 species are randomly distributed
along the cell walls, while BS chloroplasts are typically located
in the centripetal position (close to the vascular tissue, as in
finger millet) or the centrifugal position (close to M cells, as
in maize). The intracellular orientation of BS chloroplasts is
thought to have physiological significance. The centrifugal
position of BS chloroplasts is advantageous to metabolite
exchange between M cells and BS chloroplasts. In contrast,
the centripetal position of BS chloroplasts maximizes the
length of the CO2 diffusion pathway between BS and M cells,
and minimizes CO2 leakage from BS cells to M cells (Hattersley
and Browning 1981, von Caemmerer and Furbank 2003).
The intracellular arrangement of BS chloroplasts is acquired
during cell maturation (Miyake and Yamamoto 1987). A
mechanism for keeping chloroplasts in the home position
operates after establishment of the intracellular disposition
of chloroplasts, since the original arrangement of chloroplasts can be re-established 1–2 h after disturbance by
centrifugation (Kobayashi et al. 2009). The intracellular positioning of M and BS chloroplasts is dependent on the actomyosin system and cytosolic protein synthesis, but not
tubulin or light (Miyake and Nakamura 1993, Kobayashi
et al. 2009). These findings suggest that M and BS cells in C4
plants have different systems for chloroplast positioning; an
M cell-specific system for dispersing chloroplasts and a BS
cell-specific system for holding chloroplasts in the centripetal or centrifugal disposition. These unique arrangements of
C4 chloroplasts are thought to be caused by the cytoskeletal
network and vacuolar pressure (Kobayashi et al. 2009), but
the molecular mechanism is obscure at present.
A change in the intracellular disposition of C4 chloroplasts
in response to environmental stresses other than light was
initially reported by Lal and Edwards (1996). They described
that the chloroplasts and cytosol in M cells of droughtstressed maize, a monocot NADP-ME type C4 plant,
collapsed inwardly and BS chloroplasts lost their centrifugal
position. The effect of drought stress on chloroplast position
in Amaranthus cruentus, a dicot NAD-ME type C4 plant, is
not as pronounced as for maize. However, the detailed
behavior of chloroplasts and its molecular mechanism were
not mentioned in the report. To gain a better understanding
of chloroplast relocation movement in C4 plants, we closely
investigated the intracellular disposition of chloroplasts in
response to various environmental stresses and plant hormones in this study. When mature leaves of finger millet and
maize were exposed to high intensity light, M chloroplasts
showed aggregative movement but BS chloroplasts did not.
The orientation movement of M chloroplasts was also
observed under natural growing conditions with high sunlight, salinity or drought stress. These findings suggest that
M and BS cells are also differentiated regarding the control
of intracellular chloroplast positioning in response to
environmental changes.
Results
Effect of light on the intracellular positions of M and
BS chloroplasts in finger millet
M cells in leaf blades of finger millet, an NAD-ME type C4
plant, have a great number of chloroplasts dispersed
randomly along the cell walls, while BS cells have larger chloroplasts that are located in the centripetal position. A fiber
illuminator with white light was used to illuminate leaf
blades attached to plants, at light intensities of 250, 2,000,
3,000 or 4,000 µmol quanta m−2 s−1 for 2 h (Fig. 1, Supplementary Fig. S1). Most of the M chloroplasts were disproportionately re-distributed to the BS side in response to the
light, and the centripetal positioning of M chloroplasts was
more distinct at intensities >3,000 µmol m−2 s−1. Although a
fraction of the M chloroplasts did not migrate to the BS side,
there is a possibility that the M chloroplasts gather to circumvent high intensity light. In contrast, the centripetal
arrangement of BS chloroplasts was unchanged, even though
they appeared to swell slightly after strong light illumination
and the degree of chloroplast association was marginally
reduced.
We also investigated the time course of M chloroplast
movement in response to strong light irradiation. Slight
aggregative movement of M chloroplasts was observed 0.5 h
after strong light irradiation, and the one-sided distribution
of M chloroplasts became more remarkable in a timedependent manner (Fig. 2, Supplementary Fig. S2).
Effect of high intensity light on the intracellular
positions of maize chloroplasts
Maize is an NADP-ME type C4 plant. Compared with the
NAD-ME type finger millet, maize has more numerous M
chloroplasts, while its BS chloroplasts are smaller and located
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Fig. 1 Effect of light intensity on the intracellular positions of M and BS chloroplasts in finger millet. Leaf blades of finger millet were continuously
irradiated with white light of intensity 250 (A and B), 2,000 (C and D), 3,000 (E and F) and 4,000 µmol m−2 s−1 (G and H), respectively, at the adaxial
side for 2 h. B, D, F and H are magnified images. In each panel, the upper side of the leaf sections is the adaxial side. B, bundle sheath cell; M,
mesophyll cell; V, vascular bundle. Scale bars = 50 µm.
in the centrifugal position (Fig. 3A, B). Given these differences,
we examined whether maize also shows re-arrangement of
chloroplasts in response to high intensity light. After 2 h of
strong light irradiation, maize M chloroplasts redistributed,
but were found along the sides of anticlinal walls parallel to
the direction of irradiation (Fig. 3C, D). This re-arrangement
pattern was obviously different from the pattern observed in
finger millet in which M chloroplasts mainly migrated
towards the BS side and formed a partial ring around a cylinder of BS cells. The centrifugal positioning of chloroplasts in
maize BS cells was not changed under the high intensity
light.
Effects of environmental stresses on the intracellular
positioning of chloroplasts
The extremely strong light intensity (>3,000 µmol m−2 s−1)
that caused the obvious aggregative movements of M chloroplasts described above is far greater than plants encounter
under normal growing conditions. This strong light irradiance induced strong photoinhibition, because the PSII maximum quantum yield (Fv/Fm) of finger millet leaf blades
declined from 0.8 to near 0.5 after 2 h of the strong light
(4,000 µmol m−2 s−1) treatment. Therefore, there is a possibility
that some stress provoked by photoinhibition acts as a trigger for the aggregative movement of M chloroplasts, in addition to the possibility that strong light itself functions as a
signal. We examined the impact of other environmental
stresses on the intracellular disposition of chloroplasts under
normal intensity light. To induce drought stress, water
supply was withheld from finger millet plants growing under
normal intensity light (500 µmol m−2 s−1). When leaf blades
closed and began to fade after 5–7 d of water shortage, we
observed transverse leaf sections (Fig. 4A, B). Almost all of
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the M chloroplasts were distributed towards the BS cells
(Fig. 4B). We investigated the relationship between M chloroplast movement and water potential in leaf blades of finger
millet after disruption of the water supply (Fig. 4C). When
the water potential was between –0.53 and –0.15 MPa, chloroplast movement was observed in some sections but not in
others. When water potential was below –0.7 MPa, all of the
M chloroplasts showed aggregative movement. The M chloroplast movement in response to drought stress was also
observed in maize (Supplementary Fig. S3).
Next, we observed the intracellular arrangement of chloroplasts in response to salinity or high osmotic stress (Fig. 5).
In finger millet exposed to 3% NaCl (1 osmol kg–1) in normal
intensity light, most of the M chloroplasts migrated towards
the BS cells but the centripetal arrangement of BS chloroplasts was unchanged (Fig. 5B). As no significant difference
was observed in the Fv/Fm values of leaf blades between the
control and NaCl-treated plants (0.75 ± 0.01, n = 4), it suggests that the M chloroplasts in the salinity-treated leaves
responded before the occurrence of photoinhibition. The
aggregative movement of M chloroplasts in salinity-stressed
plants was also observed in semi-thin sections prepared from
resin-embedded leaves (Supplementary Fig. S4). M chloroplasts were distributed toward BS cells but not along the cell
walls directly attached to BS cells. High salinity causes a combined stress due to an imbalance of ions and osmotic homeostasis. We also investigated the effect of osmotic stress on
the intracellular arrangement of chloroplasts in finger millet
by supplying 20% polyethylene glycol (0.52 osmol kg−1) as an
external osmolyte (Fig. 5C). Only the M chloroplasts showed
a change in intracellular positioning in response to high
osmotic stress, similarly to the salinity stress. Therefore,
strong osmotic stress clearly induces aggregative movement
of M chloroplasts. Under these stress conditions, no obvious
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Stress-responsive movement of C4 chloroplasts
Fig. 2 Changes in the intracellular arrangement of chloroplasts in response to high intensity light. Leaf blades of finger millet were continuously
irradiated with the high intensity light (4,000 µmol m−2 s−1). Transverse sections were observed with the light microscope before (A and B) and
after 0.5 (C and D), 1 (E and F), 2 (G and H) and 3 h (I and J) of illumination. Scale bars = 50 µm.
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Fig. 3 Change in the intracellular positions of maize chloroplasts in response to light irradiation. Leaf blades of maize were irradiated for 2 h with
normal intensity (250 µmol m−2 s−1) (A and B) or high intensity (4,000 µmol m−2 s−1) light (C and D), and transverse sections were examined. B and
D are magnified images. B, bundle sheath cell; M, mesophyll cell; V, vascular bundle. Scale bars = 50 µm.
Fig. 4 Change in the intracellular position of chloroplasts in response to drought stress. Finger millet growing under the normal light condition
(500 µmol m−2 s−1 during the light period) was exposed to drought stress by withholding the water supply. When leaf blades began to fade after
5–7 d, leaf sections were examined with a light microscope. (A) Control; (B) drought stress. In each panel, the upper side of the leaf sections is the
adaxial side. Scale bars = 50 µm. (C) Relationship between M chloroplast movement and water potential in leaf blades. Water potential was
measured for 12 d after disruption of the water supply. At the same time, we checked whether M chloroplast movement occurred and the results
were plotted on a graph. The water potentials of non-stressed plants were –0.58 to –0.15 MPa.
plasmolysis was observed. Furthermore, when leaf segments
of finger millet were deaerated in 1 M sorbitol (1 osmol kg–1)
and incubated with the same solution for 4 h in the light,
plasmolysis of M cells was observed but the centripetal
aggregation of M chloroplasts did not occur (data not
shown). Therefore, we conclude that the chloroplast
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movement in response to environmental stresses is not
caused directly by plasmolysis, which hardly occurs in plants
growing under atmospheric conditions.
To examine whether light irradiation is necessary for the
chloroplast movement in response to environmental
stresses, finger millet was subjected to drought or salinity
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Stress-responsive movement of C4 chloroplasts
Effects of natural sunlight on the intracellular
positioning of chloroplasts
To investigate whether the chloroplast aggregative movements occur in C4 plants under natural conditions, we harvested leaf blades of finger millet and maize exposed to
direct mid-day sunlight in midsummer (1,800 µmol m−2 s−1)
and a dry environment, and observed the transverse sections
(Fig. 8). Only the M chloroplasts in finger millet and maize
showed the aggregative movement. In finger millet, M chloroplast movement was more significant on the adaxial side
(upper side of the leaf section) compared with the abaxial
side (Fig. 8A). Maize chloroplasts in M cells that were located
at the adaxial or abaxial side migrated towards the BS cells,
similarly to drought stress (Fig. 8B). The Fv/Fm values of leaf
blades from finger millet and maize were 0.37 ± 0.03 and
0.41 ± 0.03, respectively and, therefore, these plants had
experienced severe photoinhibition. At night-time, M chloroplasts of both plants returned to comparatively random
positions along the plasma membranes (Fig. 8C, D). The
Fv/Fm values were recovered to normal values (0.81 ± 0.01 for
finger millet and 0.75 ± 0.01 for maize) after the end of the
night. These findings suggest that change in the intracellular
arrangement of M chloroplasts is a general phenomenon in
field-growing C4 plants that are exposed to multiple environmental stresses, which cause severe photoinhibition.
Fig. 5 Change in the intracellular arrangement of chloroplasts in
response to salinity or high osmotic stress. Finger millet was supplied
with 3% NaCl or 20% polyethylene glycol solution to produce salinity
and high osmotic stress, respectively, for 5 d in normal intensity light
(500 µmol m−2 s−1 during the light period), and transverse sections of
leaf blades were examined. (A) Control; (B) salinity stress; (C) high
osmotic stress. Scale bars = 50 µm.
stress under dark conditions. Although the water potential
of leaf blades exposed to drought or salinity stress for 9 d was
decreased to –1.83 ± 0.18 MPa or –0.80 ± 0.17 MPa (n = 4),
respectively, the aggregative movement of M chloroplasts
was not observed (data not shown). Therefore, it was concluded that light is required for the chloroplast movement
in response to environmental stresses.
We also examined the intracellular arrangement of nuclei
in response to salinity stress (Fig. 6). Although BS nuclei
were located close to M cells, M nuclei were distributed
peripherally at the mid position, a little towards BS cells. The
intracellular positions of both types of nuclei were not
changed regardless of salinity stress. We further observed
the intracellular arrangement of mitochondria (Fig. 7). All
BS mitochondria were dominantly located close to vascular
bundles but M mitochondria were randomly distributed
in the cells. The intracellular positions of neither type of
mitochondria were changed regardless of salinity stress.
Involvement of actin filaments in the intracellular
arrangement of chloroplasts in response to strong
light irradiation
We investigated whether actin filaments participate in M
chloroplast movement in response to light irradiation.
Cytochalasin B is a potent inhibitor of actin polymerization,
and we had previously confirmed by immunodetection that
our pre-treatment of leaf segments with cytochalasin B
disrupted actin networks (Kobayashi et al. 2009). Treatment
of finger millet with cytochalasin B showed a prominent
inhibitory effect on the strong light-dependent movement
of M chloroplasts, in contrast to treatment with dimethylsulfoxide (DMSO) as a control (Fig. 9A, B). Cytochalasin B
did not affect the disposition of M chloroplasts under normal
intensity light (Figs. 9C, D). The centripetal position of BS
chloroplasts was unchanged irrespective of cytochalasin B
treatment.
Effect of plant hormones on the intracellular
positioning of chloroplasts
ABA accumulates and functions as a signal transducer in
response to environmental stresses such as drought and soil
salinity (Zhang et al. 2006). To investigate the possibility of
the involvement of ABA in the chloroplast movement in
response to environmental stresses, we allowed leaf
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Fig. 6 Effect of salinity stress on the intracellular positions of nuclei. Transverse sections of leaf blades from control (A and B) or salinity-stressed
(C and D) finger millet were stained with DAPI and observed under a bright-field (A and C) or fluorescence (B and D) microscope. B and D are
merged images of the bright-field and fluorescence images. Nuclei were detected as white particles in cells. Scale bars = 50 µm.
Fig. 7 Effect of salinity stress on the intracellular positions of mitochondria. Transverse sections of leaf blades from control (A–C) or salinitystressed (D–F) plants were stained with rhodamine 123. Mitochondria (yellow) and chloroplasts (red) were imaged using confocal laser scanning
microscopy. C and F are enlarged images of M cells, and the right side in the two panels is the BS side. Scale bars = 50 µm.
segments from non-stressed finger millet to absorb ABA
during incubation for 16 h under low intensity light. This
ABA treatment induced the centripetal assembly of M chloroplasts (Fig. 10). We confirmed that treatment with ABA
above 3 µM was effective in causing this arrangement of
chloroplasts. When the incubation with ABA was conducted
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in the dark, the chloroplast movement did not occur
(data not shown). Incubation with other plant hormones
(IAA, 2,4-D, GA3 and kinetin) in the light had no effect on
the intracellular positioning of chloroplasts (data not
shown). Various concentrations of NaCl (0.3–3%) and H2O2
(1–100 mM) also had no effect (data not shown).
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Stress-responsive movement of C4 chloroplasts
Fig. 8 Aggregative movement of M chloroplasts in field-grown finger millet and maize in midsummer. Leaf blades of finger millet (A and C) and
maize (B and D) growing under natural midsummer conditions with high radiation and a dry environment were sampled in the middle of the day
(14:00 h; atmosphere temperature, 35°C; light intensity, 1,800 µmol m−2 s−1; A and B) or during the night (3:00 h; atmosphere temperature, 26°C;
C and D) of a fair day, and transverse sections were examined. In each panel, the upper side of the leaf sections is the adaxial side. Scale
bars = 50 µm.
Fig. 9 Effect of cytochalasin B on the intracellular arrangement of chloroplasts in response to light irradiation. Leaf segments excised from leaf
blades of finger millet were deaerated in 0.5% (v/v) dimethyl sulfoxide (DMSO) with or without 50 µM cytochalasin B, and floated on the
solution for 2 h under room light (<5 µmol m−2 s−1). Then, the leaf segments were irradiated for 2 h with normal intensity (250 µmol m−2 s−1) or
high intensity (4,000 µmol m−2 s−1) light, and transverse sections were examined. (A) DMSO, high intensity light; (B) DMSO + cytochalasin, high
intensity light; (C) DMSO, normal intensity light; (D) DMSO + cytochalasin, normal intensity light. Scale bars = 50 µm.
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Fig. 10 Effect of ABA on the intracellular arrangement of chloroplasts.
Leaf segments excised from leaf blades of finger millet were deaerated
in 0.1% ethanol with or without 10 µM ABA and floated on
the solution for 16 h under low intensity light (100 µmol m−2 s−1).
(A) Control; (B) ABA treatment. Scale bars = 50 µm.
Discussion
Stronger light and longer exposure times are
required for aggregative movement of C4 M
chloroplasts compared with C3 M chloroplasts
Photo-relocation movement of chloroplasts is widely
observed in a variety of plant species. In this study, we found
that M chloroplasts of C4 plants showed aggregative movement in response to strong light. Extremely high light intensities >3,000 µmol m−2 s−1 were needed to induce an obvious
movement of M chloroplasts in normally growing C4 plants
(Fig. 1, Supplementary Fig. S1). Inoue and Shibata (1974)
reported that absorbance of leaves from five graminaceous
C4 species decreased in response to blue light (about 86 µmol
quanta m−2 s−1), but the light intensity was much lower than
that necessary for the obvious aggregative movement
induced by white light in our experiment. They used leaves
incubated in darkness for 1 d before measurement and,
therefore, the leaves might become more susceptible to
light. We also confirmed that blue light could induce the
centripetal positioning of M chloroplasts but the extent of
localization was not prominent (data not shown). Inoue and
Shibata did not report the precise migration pattern of chloroplasts, and the wavelength dependency of the aggregative
movement remains to be investigated.
The aggregative movement of M chloroplasts in C4 and C3
plants differs in light intensity and time required. C3 M chloroplasts respond to much lower light intensities than C4 M chloroplasts. For example, the apparent light avoidance movement
of chloroplasts in dark-adapted Arabidopsis thaliana leaf
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occurs upon illumination with blue light at 5 W m−2 (about
19 µmol m−2 s−1) (Trojan and Gabrys 1996). The extent of
chloroplast avoidance movement in A. thaliana increases in
response to the intensity of white light and reaches a maximum at about 500 µmol m−2 s−1 (Kasahara et al. 2002). Similarly, the maximum chloroplast movement in redwood sorrel
occurs upon illumination with blue light at 250 µmol m−2 s−1
(780 µmol m−2 s−1 of daylight) (Brugnoli and Björkman 1992).
The time required for obvious observation of chloroplast
movement is also shorter in C3 plants. For example, chloroplasts of redwood sorrel, sunflower and Arabidopsis start to
move after only a few minutes of light irradiation (Brugnoli
and Björkman 1992, Trojan and Gabrys 1996). In leaf epidermal cells of the aquatic angiosperm Vallisneria gigantea,
about half the chloroplasts move out of the area irradiated
with high intensity blue light within the first 15 min of irradiation, and the percentage increases to 80% after 30 min
(Sakurai et al. 2005). In contrast, the extent of chloroplast
movement was low after 30 min of high intensity light irradiation (Fig. 2, Supplementary Fig. S2) and, therefore, the
response of C4 M chloroplasts to strong light seems to be
slow. C4 plants generally adapt to high intensity light and,
therefore, C4 photosynthetic cells might not be as susceptible to light-inducing stresses in comparison with C3 M cells.
Moreover, growth conditions might be another factor to
yield the differential light responsiveness. C3 plants are generally grown under lower intensity light compared with C4
plants and, therefore, photoinhibition and chloroplast
movement for photoprotection in C3 plants is more likely to
occur at relatively low light intensities.
Aggregative movement of C4 M chloroplasts was
induced in response to environmental stresses
The chloroplast movement in M cells of finger millet
occurred under normal intensity light (500 µmol m−2 s−1)
under stress conditions such as drought, salinity or hyperosmosis (Figs. 4, 5). These abiotic stresses are thought to
reduce the threshold intensity of light at which aggregative
movement of M chloroplasts occurs. Ionic and osmotic
stresses originating from salinity cause damage to metabolic
processes and the ultrastructure of chloroplasts (Yamane et al.
2003, Hasan et al. 2005, Morales et al. 2006, Omoto et al.
2009). Although supplying plant roots with 20% polyethylene glycol solution induced re-arrangement of M chloroplasts (Fig. 5C), incubation of leaf segments with 1 M sorbitol
or 3% NaCl solutions whose osmolality was twice as high as
that of the polyethylene glycol solution had no effect on
chloroplast arrangement. Therefore, it is thought that some
signal associated with the osmotic stress is generated in a
domain outside of leaf tissue and influences M chloroplast
movement. A decrease in water potential during water
shortage is also important in M chloroplast re-arrangement
(Fig. 4C). However, another factor may be involved in the
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Stress-responsive movement of C4 chloroplasts
induction of chloroplast movement, because M chloroplast
movement was occasionally observed in leaves showing high
water potential above –0.53 MPa.
Previously it was reported that water stress induced centripetal re-arrangement of M chloroplasts in leaves of the C4
plant, maize (Lal and Edwards 1996) and the C4 Crassulacean
acid metabolism (CAM) cycling plant, Portulaca grandiflora
(Guralnick et al. 2002). Their intracellular localization is similar to the typical aggregative arrangement of M chloroplasts
which we observed in finger millet, but not in maize, during
high light stress. The chloroplast rearrangement of C4 M cells
is thought to be induced by a combination of light and environmental stresses. In our experiments, most M chloroplasts
in finger millet leaves moved towards the BS, but some M
chloroplasts remained scattered around the opposite side.
In maize leaves irradiated by high intensity light, M chloroplasts were distributed along the sides of the anticlinal walls
(Fig. 3C, D), but the direction of chloroplast movement in
field-growing water-stressed maize was rather towards the
BS (Fig. 8B), similar to the observation of Lal and Edwards
(1996) under drought stress. Therefore, C4 M chloroplasts
might show light avoidance movement similar to C3 M chloroplasts, but prominent aggregation of M chloroplasts
occurs in C4 plants that receive severe stresses for long periods of time.
C4 plants attain higher rates of photosynthesis in full sunlight and are also more efficient in water use compared with
C3 plants (Hatch 1992). As a result, C4 plants are said to be
more tolerant to environmental stresses. We found aggregative movement of M chloroplasts of finger millet and maize
growing in a field in midsummer (Fig. 8). The leaf surface at
that time was exposed to a light intensity of about 1,800 µmol
m−2 s−1, which was not high enough to induce chloroplast
movement in the laboratory. The field-grown plants can be
subject to other stresses in addition to high intensity light.
Under the mid-day field condition, plants were exposed to
strong light and high temperature for several hours. Although
plants were well watered, a high transpiration rate may
nonetheless cause low leaf water potential (Hirasawa and
Hsiao 1999). Indeed, the field-growing plants that we measured showed severe photoinhibition at mid-day. Thus, a
combination of stresses may induce chloroplast movement
in C4 plants in the field. The intracellular disposition of M
chloroplasts changes diurnally as the aggregative arrangement is partially eliminated at night-time when plants
recover from photoinhibition.
Possible physiological roles of the aggregative
M chloroplast movement
A study with Arabidopsis mutants revealed that chloroplast
avoidance movement decreases the amount of light absorption by chloroplasts, and therefore protects plants from
photodamage under high light (Kasahara et al. 2002). C4
plants growing under environmental stresses are exposed to
an excess of light energy and are subjected to photoinhibition (Lal and Edwards 1996, Jia and Lu 2003, Xu et al. 2008).
Under these conditions, the assemblage of M chloroplasts is
thought to provide photoprotection through mutual shading of the chloroplasts, similarly to C3 chloroplasts. Actually,
we observed an increase in light transmittance through leaf
blades in response to high intensity light (Supplementary
Figs. S1, S2). Although the degree of PSII photoinhibition by
high intensity light is similar between M and BS thylakoids of
maize (Pokorska and Romanowska 2007), M chloroplasts are
more sensitive to the damaging effect of salinity than are BS
chloroplasts (Hasan et al. 2005, Omoto et al. 2009). Previously, we found that salinity-induced damage in M chloroplasts of maize and rice is light dependent, and not due to
direct effects of excessive accumulation of sodium in the leaf
tissues (Mitsuya et al. 2003, Hasan et al. 2005). We therefore
assumed that reactive oxygen species are involved in the
chloroplast damage induced by salinity (Mitsuya et al. 2003,
Yamane et al. 2004a, Yamane et al. 2004b, Hasan et al. 2005).
Moreover, the distribution of antioxidant enzymes is
reported to be different between M and BS cells in maize
(Doulis et al. 1997, Foyer et al. 2002). It is presumed that antioxidant status could be different between the photosynthetic cell types under stress conditions. Under the salinity
stress that caused aggregative movement of M chloroplasts
(Fig. 5B), symptoms of photoinhibition were not observed.
The C4 M chloroplast movement may be one means of photoprotection which occurs prior to photoinhibition.
Another possible role of C4 M chloroplast movement is
maintenance of photosynthetic activity under stress conditions. Most M chloroplasts in finger millet moved toward
the BS, unlike C3 chloroplasts that migrate to the cell walls
parallel to strong light. The centripetal aggregation of C4 M
chloroplasts might be to enable communication with BS
cells. The centripetal position of M chloroplasts shortens the
diffusion pathway of metabolites between M and BS cells,
and may contribute to keeping C4 photosynthesis active.
Moreover, leakiness of CO2 from BS cells is increased in
stressed C4 plants (Ghannoum 2009). M chloroplasts and
cytosol might move towards the BS to refix CO2 released
from BS cells more efficiently. However, the centripetal
aggregation of M chloroplasts towards the BS side could
increase the diffusion distance between the intercellular air
space and the primary carboxylation step (cytosolic phosphoenolpyruvate carboxylase and M chloroplast) and, therefore, decrease the production of C4 dicarboxylates (Lal and
Edwards 1996, Tholen et al. 2008). Indeed, the chloroplast
avoidance response in A. thaliana leaves results in a smaller
chloroplast surface area adjacent to intercellular airspaces
and decreases internal conductance to CO2 diffusion (Tholen
et al. 2008). Attempts to characterize the relationship between
chloroplast disposition and photosynthetic parameters are
Plant Cell Physiol. 50(10): 1736–1749 (2009) doi:10.1093/pcp/pcp116 © The Author 2009.
1745
M. Yamada et al.
currently in progress to determine whether C4 plants adapt
to refix released CO2 under environmental stress conditions
which lead to stomatal closure.
Molecular mechanism of M chloroplast movement
A potent inhibitor of actin polymerization, cytochalasin B,
inhibited the aggregative movement of M chloroplasts in
response to high intensity light (Fig. 9). Therefore, actin filaments are considered to participate in the M chloroplast
movement. Actin filaments encircle M and BS chloroplasts
of finger millet and maize, and seem to be involved in their
positioning and anchorage (Kobayashi et al. 2009). The actomyosin system is necessary for arrangement of both chloroplasts during cell maturation and rearrangement of
chloroplasts after disturbance by centrifugal force (Miyake
and Nakamura 1993, Kobayashi et al. 2009). The involvement of actin filaments as a track in chloroplast photorelocation movement has been confirmed in several C3 plant
species by pharmacological studies (Wada et al. 2003). A
basket structure of microfilaments surrounding Arabidopsis
M chloroplasts was observed with immunofluorescent labeling (Kandasamy and Meagher 1999). Actin filaments change
their organization before and after chloroplast movement,
and also function in anchoring chloroplasts to the site
(Takagi 2003). As C4 M chloroplasts move toward the BS
side, the M cells might possess a system for determining cell
polarity and machinery for polarized motility. Whether a
similar motility system for chloroplast movement works in
both C3 and C4 plant cells remains to be investigated.
Even though M chloroplasts show aggregative movement
in response to salinity stress, nuclei and mitochondria did
not change their positions (Figs. 6, 7). In contrast, lightdependent nuclear positioning was reported in leaf cells of
A. thaliana and prothallial cells of Adiantum capillus-veneris
(Iwabuchi et al. 2007, Tsuboi et al. 2007). While both the
nuclear and chloroplast photo-relocation movements share
photoreceptors and cytoskeletons, some components
involved in the moving machinery are thought to be specific
to each organelle (Iwabuchi et al. 2007). Recently, blue lightinduced co-localization of mitochondria with chloroplasts
was shown in Arabidopsis palisade M cells (Islam et al. 2009).
The authors presumed a relationship of the co-localization
with their mutual metabolic interactions. The nuclear and
mitochondrial movement in C3 leaves is speculated to be an
adaptive response for light as well as chloroplast photorelocation movement, while the aggregative movement of
C4 M chloroplasts independent of nuclei and mitochondria
may be induced for a special physiological requirement association with C4 photosynthesis.
Treatment of finger millet leaf segments with ABA
induced the centripetal assembly of M chloroplasts in a
light-dependent manner (Fig. 10). Because ABA was vacuum
infiltrated into the leaf segments, M chloroplast movement
1746
is thought to be caused by a direct effect of ABA on M cells
and not by secondary effects such as stomatal closure. Participation of ABA in chloroplast movement has also been
reported in succulent plants (Kondo et al. 2004). Clumping
of chloroplasts in response to water stress was first found in
cortical cells of P. grandiflora stems (Guralnick et al. 2002).
After that, Kondo et al. (2004) showed that chloroplasts in a
variety of succulent CAM plants become densely clumped
under combined light and water stress. The chloroplast
clumping induced by ABA is dependent on light. ABA, which
is a signal transducer in response to environmental stresses,
is proposed to function as a trigger for the chloroplast movements in C4 and CAM plants. Because M chloroplast movement occurred in the leaf segments irradiated with high
intensity light (Fig. 9A), ABA may be synthesized in the
leaves and initiate chloroplast movement, as well as ABA
which is synthesized in roots and transported to leaves. Light
is essential to chloroplast movement induced by ABA, and it
is also required for the aggregative movement of C4 M chloroplasts in response to drought or salinity stress. Under environmental stress conditions, a decrease in consumption of
reducing equivalents can result in accumulation of electrons
in the photosynthetic electron transport chain, that produces harmful reactive oxygen species. Thus, reactive oxygen
species are another potential trigger for chloroplast movement. Indeed, it was reported that hydrogen peroxide is generated by high fluence blue light in Arabidopsis M cells and
was suggested to promote chloroplast avoidance movement
in the presence of blue light (Wen et al. 2008). However, the
incubation of leaf segments of finger millet with various concentrations of hydrogen peroxide had no effect on the intracellular arrangement of chloroplasts. This indicates that
hydrogen peroxide itself cannot induce chloroplast movement in C4 plants, but further work is required to determine
whether other reactive oxygen species affect chloroplast
movement in C4 M cells.
In summary, the present study has shown the aggregative
movement of C4 M chloroplasts in response to environmental stresses. The movement is light dependent, and evidence
is provided that it is mediated by ABA. At present, the
physiological significance and molecular mechanism of the
chloroplast response are unknown and need further study.
Materials and Methods
Plant materials and growth conditions
Finger millet (Eleusine coracana L. Gaertn. cv. Yukijirushi)
and maize (Zea mays L. cv. Golden Cross Bantam T51) were
grown in vermiculite in a growth chamber with 14 h of illumination (500 µmol m−2 s−1) at 28°C and 10 h of darkness at
20°C per day. Plants were fertilized regularly with Arnon and
Hoagland solution (Arnon and Hoagland 1940) during
growth. The middle regions of the fourth leaf blades from
Plant Cell Physiol. 50(10): 1736–1749 (2009) doi:10.1093/pcp/pcp116 © The Author 2009.
Stress-responsive movement of C4 chloroplasts
plants of about 4 weeks old were normally used for
experiments.
The experiments with field-growing plants were conducted in August 2008 at the University Farm of Nagoya
University. Plants were grown in well-watered and periodically fertilized soil for 10 weeks, and fully-matured leaves
were used for experiments.
High-light treatment
A fiber illuminator illuminated the middle regions of the
fourth leaf blades with a halogen lamp (MHF-150L, Moritex,
Tokyo, Japan or PICL-NEX, Nippon P-I Co. Ltd., Tokyo, Japan)
at a distance of 2.5 cm. The photosynthetic photon flux density at the leaf surface was checked with a quantum meter
(LI-250, LI-COR, Lincoln, NE, USA). Small segments (5 × 5 mm
square) were excised from the treated leaf blades and vacuum
infiltrated for 10 min with fixation buffer [50 mM PIPESNaOH, pH 6.9, 4 mM MgSO4, 10 mM EGTA, 0.1% (w/v) Triton
X-100, 200 µM phenylmethylsulfonyl fluoride, 5% (v/v) formaldehyde and 1% (v/v) glutaraldehyde]. After incubation at
4°C overnight, the fixed segments were embedded in 5%
(w/v) agar and sectioned at 70–80 µm with a micro-slicer
(DTK-3000W, Dosaka EM, Kyoto, Japan). Transverse sections
were observed with a light microscope (BX51, Olympus,
Tokyo, Japan) equipped with a CCD camera (DP70, Olympus).
Chlorophyll fluorescence was measured with a portable chlorophyll fluorometer PAM-2100 (Walz, Effeltrich, Germany).
Stress treatment
Three- to four-week-old plants were exposed to drought
stress by withholding water supply until the appearance of
the first sign of wilting. Leaf segments were then excised
from the upper developed leaf blades and fixed as described
above. Transverse sections were observed with the light
microscope. Water potential in leaves was measured with a
WP4 Dewpoint Meter (Decagon Devices, Pullman, WA,
USA).
Three plants per pot were grown in a 300 ml plastic pot
filled with vermiculite in the growth chamber. High salinity
treatment was achieved by supplying 30 ml per day of Arnon
and Hoagland solution containing 3% (w/v) NaCl for 5 d. For
high osmotic stress, 15 ml d–1 of 20% (w/v) polyethylene
glycol 6,000 solution was supplied for 5 d. Transverse
sections of the fixed leaf segments were observed with
the light microscope. The osmolality values of the solutions
were determined by the freezing point method in an
Osmotoron-5 (Orion Riken Inc., Tokyo, Japan).
For microscopic observation of semi-thin sections, leaf
segments were fixed as previously reported (Omoto et al.
2009). Semi-thin sections (1 µm thickness) were cut with
glass knives on an ultramicrotome. Then, the sections were
stained with toluidine blue O and observed with the light
microscope.
Nuclear and mitochondrial staining
For nuclear staining, leaf segments from the salinity-stressed
plants were fixed as described above and transverse sections
were stained with 1 mg ml−1 4′,6-diamidino-2-phenylindole
(DAPI) for 1 h. After washing with distilled water for 10 min
twice, the sections were imaged with a light microscope
(BX51, Olympus) equipped with an epifluorescence system
(U-LH100HG, Olympus).
For mitochondrial staining, non-fixed leaf segments from
the salinity-stressed plants were tucked into carrot blocks
and sectioned with a microslicer. Transverse sections were
stained in PME buffer (50 mM PIPES-NaOH, pH 6.9, 5 mM
MgSO4, 5 mM EGTA and 0.15 M NaCl) containing 1 µM rhodamine 123 for 4 min. After washing with PME buffer for
10 min twice, the sections were imaged with a confocal laser
scanning microscope (LSM5 PASCAL, Carl Zeiss, Germany).
Rhodamine 123 was excited with the 488 nm wavelength of
an ArKr laser and the images were collected using a BP505–
530 bandpass filter. Autofluorescence of chloroplasts was
excited with the 543 nm wavelength of a HeNe laser and
imaged using an LP560 longpass filter. Serial confocal optical
images at 0.50 µm intervals were collected, and projections
of 20–40 µm thickness were created with LSM Imaging
Browser software.
Chemical treatment
For cytochalasin treatment, small leaf segments (5 × 5 mm
square) were excised and vacuum infiltrated for 10 min with
0.5% (v/v) DMSO with or without 50 µM cytochalasin B (MP
Biomedicals, Irvine, CA, USA). After floating on the same
solution for 2 h, the leaf segments were exposed to normal
(250 µmol m−2 s−1) or high light (4,000 µmol m−2 s−1) for 2 h.
Then, the leaf segments were fixed and transverse sections
were observed with a light microscope.
For ABA treatment, small leaf segments were excised and
vacuum infiltrated for 10 min with 0.1% (v/v) ethanol with
or without 10 µM ABA. After floating on the same solution
for 16 h under low light (100 µmol m−2 s−1), the leaf segments
were fixed and transverse sections were observed with a light
microscope.
For other chemical treatments, small leaf segments were
excised and vacuum infiltrated for 10 min with 10 mM MESKOH (pH 6.9) containing IAA (0.3, 1 or 3 µM), 2,4-D (3, 10 or
30 µM), GA3 (15, 50 or 150 µM), kinetin (30, 100 or 300 µM),
ABA (1, 3, 10 or 30 µM), NaCl [0.3, 1 or 3% (w/v)] or H2O2
(1, 5, 10, 20 or 100 mM). After floating on the same solution
for 16 h under low light (100 µmol m−2 s−1), the leaf segments
were hand-sectioned with a razor blade and transverse
sections were observed with a light microscope.
Supplementary data
Supplementary data are available at PCP online.
Plant Cell Physiol. 50(10): 1736–1749 (2009) doi:10.1093/pcp/pcp116 © The Author 2009.
1747
M. Yamada et al.
Funding
The Ministry of Education, Culture, Sports, Science and
Technology (Grants-in-Aid for Scientific Research No.
21380014).
Acknowledgments
We thank Mr. Yasuki Tahara, University Farm of Nagoya
University, for growing the plants.
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(Received June 6, 2009; Accepted August 2, 2009)
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