Download Chloroplast anchoring: its implications for the

Journal of Experimental Botany, Vol. 60, No. 12, pp. 3301–3310, 2009
doi:10.1093/jxb/erp193 Advance Access publication 21 June, 2009
DARWIN REVIEW
Chloroplast anchoring: its implications for the regulation of
intracellular chloroplast distribution
Shingo Takagi*, Hideyasu Takamatsu and Nami Sakurai-Ozato†
Department of Biological Sciences, Graduate School of Science, Osaka University, Machikaneyama 1-1, Toyonaka, Osaka 560-0043,
Japan
Received 8 December 2008; Revised 18 May 2009; Accepted 19 May 2009
Abstract
The intracellular distribution of organelles plays a pivotal role in the maintenance and adaptation of a wide spectrum
of cellular activities in plants. Chloroplasts are a special type of organelle able to photosynthesize, capturing light
energy to fix atmospheric CO2. Consequently, the intracellular positioning of chloroplasts is crucial for plant growth
and development. Knowledge of the photoreceptors and cellular apparatus responsible for chloroplast movement
has gradually accumulated over time, yet recent advances have allowed improved understanding. In this article,
several aspects of research progress into the mechanisms for maintaining the specific intracellular distribution
patterns of chloroplasts, namely, chloroplast anchoring, are summarized, together with a brief consideration of the
future prospects of this subject. Our discussion covers developmental, physiological, ecophysiological, and recent
cell biological research areas.
Key words: Actin cytoskeleton, chloroplast anchoring, chloroplast movement, chloroplast positioning.
What is chloroplast anchoring?
Light-dependent chloroplast movement allows the adaptation of cellular activities to fluctuating environmental light
conditions. Under dim light, chloroplasts move in order to
maximize light interception, while under strong light, chloroplasts move to minimize light interception. It is generally
accepted that the physiological significance of these movements is to increase photosynthetic activity under dim light
(Zurzycki, 1955), while to evade photodamage caused by
excess light (Zurzycki, 1957; Park et al., 1996; Kasahara
et al., 2002). Chloroplasts exhibit species-specific patterns of
movement, for example, the rotation of single ribbon-shaped
chloroplasts in the cylindrical cells of green algae (Wagner
and Klein, 1981), face and profile positioning of multiple
chloroplasts in cylindrical cells of algae, moss, and ferns, or
the noted accumulation and avoidance responses of chloroplasts observed in leaf mesophyll cells of various angiosperms (Haupt and Scheuerlein, 1990).
Conversely, under non-fluctuating light conditions favourable for photosynthesis, chloroplasts actively maintain their
intracellular distribution. Otherwise cytoplasmic streaming
would not allow the chloroplasts to maintain their position
within the cell, resulting in decreased light use efficiency. As
described later, the concept of ‘chloroplast anchoring’ has
arisen from careful analysis of chloroplast behaviour in
various types of plant cells at different developmental stages
and under different physiological conditions. This article
aims to summarize several aspects of chloroplast anchoring
and discuss possible mechanisms underlying this intriguing
phenomenon. Since comprehensive reviews on chloroplast
movement and its motile apparatuses have already been
published (Wada et al., 2003; Lee and Liu, 2004; Shimmen
and Yokota, 2004; Wada and Suetsugu, 2004), this article
does not cover those subjects.
Developmental view
Intracellular movement and distribution of chloroplasts
have been extensively investigated in C3 plants, whereas our
* To whom correspondence should be addressed: E-mail: [email protected]
y
Present address: Department of Anatomy and Molecular Cell Biology, Graduate School of Medicine, Nagoya University, Tsurumai 65, Showa, Nagoya 466-8550,
Japan.
ª The Author [2009]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
For Permissions, please e-mail: [email protected]
3302 | Takagi et al.
knowledge on C4 plants is rather limited. In C4 leaf tissue,
mesophyll cells are the site for the primary fixation of CO2
by phosphoenolpyruvate (PEP) carboxylase into the fourcarbon oxaloacetate, while decarboxylation of malate,
formed from oxaloacetate, and the refixation of CO2 are
carried out in bundle sheath cells (Hatch and Slack, 1970).
Chloroplasts in C4 plant mesophyll cells are more or less
randomly distributed, whereas the distribution pattern of
chloroplasts in bundle sheath cells is unique; chloroplasts
are distributed either centripetally, close to the vascular
tissue, or centrifugally, around the periphery of the bundle
sheath. Different distribution patterns of chloroplasts in
bundle sheath cells are probably advantageous for the
efficient exchange of metabolites between the mesophyll
and the bundle sheath cells, the minimization of CO2
leakage, and so on.
In the NAD-malic-enzyme type C4 plant finger millet
(Eleusine coracana), Miyake and Yamamoto (1987) demonstrated that the distribution pattern of chloroplasts in
bundle sheath cells was established during leaf development,
independent of the light environment, with etioplasts
exhibiting the centripetal chloroplast distribution in mature
leaves developing in the dark. Miyake and Nakamura
(1993) further examined possible factors involved in the
establishment of the centripetal distribution of chloroplasts
using various chemicals. Auxin, cycloheximide, and the
actin-depolymerizing reagent cytochalasin B exhibited substantial inhibitory effects, while other hormones and
colchicine did not. Auxin seemed to affect the early
processes in the establishment of centripetal distribution,
suggesting a role in modulating cell elongation and/or
developmental stage, while cycloheximide and cytochalasin
B appeared to exert more direct effects on the migration of
chloroplasts to the centripetal regions. The inhibitory effects
of cytochalasin B, but not of colchicine, strongly suggest the
involvement of the actin cytoskeleton in chloroplast migration and anchoring. Recently, Kobayashi et al. (2009)
demonstrated that actin filaments, intimately associated
with the surface of chloroplasts in bundle sheath cells, may
play a central role in the re-establishment of the characteristic distribution pattern of chloroplasts after displacement
caused by centrifugation. However, the maintenance of
a specific distribution pattern of chloroplasts did not seem
to be dependent upon the actin cytoskeleton.
Physiological view
Chloroplast movement in C3 angiosperms is regulated by
blue light. In Arabidopsis thaliana, the blue-light receptors
phototropin1 (phot1) and phototropin2 (phot2) have been
identified as being responsible for the accumulation response of chloroplasts, while phot2 alone mediates the
avoidance response (Jarillo et al., 2001; Kagawa et al.,
2001; Sakai et al., 2001). On the other hand, both blue and
red light are effective in inducing chloroplast movement in
several species of cryptogam plants (Wada et al., 2003). The
effects of light in the red region are mediated by the plantspecific photoreceptor phytochrome (Wada and Kadota,
1989). Although molecular species of functional phytochrome for chloroplast movement have not yet been
identified in many cases, in the fern Adiantum capillusveneris, neochrome1, which is a chimeric receptor consisting
of the chromophore-binding domain of phytochrome and
the full-length phototropin (Nozue et al., 1998), is responsible for red-light-induced chloroplast movement
(Kawai et al., 2003).
In protonemal cells of the fern Dryopteris sparsa,
Yatsuhashi and Kobayashi (1993) examined the effects of
red light on light-induced chloroplast movement. After the
completion of the photo-relocation movement of chloroplasts induced by red light, when protonemata were
illuminated with far-red light to convert phytochrome from
its far-red-light-absorbing form (Pfr) back to its red-lightabsorbing form (Pr), the induced specific distribution
pattern of chloroplasts decayed much faster compared with
dark-incubated controls (Yatsuhashi and Kobayashi, 1993).
It appears that phytochrome in this fern is involved not
only in the induction of the photo-relocation movement of
chloroplasts, but also in the maintenance of their specific
distribution pattern, although mechanisms for both processes remain to be elucidated. Whether such roles of
phytochrome are specific to this fern is also an open
question.
In the aquatic hydrocharitaceae species Elodea canadensis
and Vallisneria spiralis, Seitz (1980) extensively investigated
the nature of light-dependent chloroplast movement, developing the concept of ‘chloroplast centrifugability’,
namely, how easily chloroplasts in living cells are displaced
by centrifugal force as an estimate for the motile activity of
cytoplasm. This makes the implicit assumption that chloroplast movement is passive and solely a result of movements
of the cytoplasmic matrix. Putative components involved in
the centrifugability of chloroplasts include the viscosity of
cytoplasm (passive component), the motility of cytoplasm
(active component), the magnitude of adhesion of chloroplasts to the cortical cytoplasm, and various kinds of
interactions with other organelles such as mitochondria and
the vacuole. Even now it is difficult to distinguish and
specify the relative importance of each of these components.
Still, in carefully designed experiments, centrifugation
methods provide useful information on intracellular microconditions associated with organelles, including the mechanical properties of the cytoplasmic matrix (Hiramoto
and Kamitsubo, 1995).
In leaf epidermal cells of V. spiralis, Seitz (1971) noted
dose-dependent effects of illumination; low-fluence-rate
illumination resulted in a decrease in the centrifugability of
chloroplasts, whereas high-fluence-rate illumination increased it. Seitz concluded that reductions in chloroplast
centrifugability reflected lowered cytoplasmic motility under
low-fluence-rates, while it was increased when exposed to
high-fluence-rates. Based on differential wavelength dependence on the effects of blue light of low- and high-fluencerates, Seitz (1979) tentatively concluded that photosynthetic
pigments and flavin function as the photoreceptor systems
under low- and high-fluence-rates, respectively. More
Chloroplast anchoring | 3303
importantly, the effects of blue light of different fluence
rates exhibited a contrasting sensitivity to metabolic inhibitors (Seitz, 1979, 1980). While the inhibitors for photosynthetic electron transport suppressed the blue-light-induced
decrease in chloroplast centrifugability, those for oxidative
phosphorylation suppressed the increase in centrifugability.
This led Seitz (1980) to propose that the cytoplasmic ATP
level, supplied by photosynthesis and respiration, may
change in relation to light intensity, and that a difference in
ATP levels between the periclinal and anticlinal regions is
the first cause for the different distribution patterns of
chloroplasts induced by light of different intensities. Specifically, chloroplasts tend to migrate out of highly motile
cytoplasmic regions with high levels of ATP and accumulate
in quiescent cytoplasmic regions containing less ATP.
However, such a mode of regulation by ATP seems
unlikely, as the cytoplasmic ATP level is now known to be
maintained at a more or less constant concentration,
irrespective of irradiance (Keifer and Spanswick, 1979;
Kikuyama et al., 1979), at concentrations sufficiently high
to account for the observed motile activity of actomyosin interactions (Shimmen, 1978), which seems to be important in
the movement of cytoplasm in higher plant cells (Shimmen
and Yokota, 2004). More recent hypotheses for chloroplast
anchoring based on the roles of the cytoskeleton will be
discussed later.
Ecophysiological view
CO2 diffusion through leaf tissues may be one of the most
important physiological factors influencing chloroplast
movement and positioning (Walczak and Gabryś, 1981).
The area of the chloroplast surface facing the intercellular
airspace (Sc) is important in determining the conductance of
CO2 in leaf tissues. To maximize the diffusion of CO2 from
the intercellular airspace to the site of CO2 fixation, the
chloroplast stroma, chloroplasts tend to be located contiguous with the plasma membrane (PM) (Evans and von
Caemmerer, 1996; Terashima et al., 2006).
Using photoacoustic analysis, Gorton et al. (2003)
examined whether chloroplast movement had any effect on
CO2 diffusion in the leaves of Alocasia brisbanensis. No
supporting result was obtained that was favourable for the
role of the avoidance response of chloroplasts in modulating internal CO2 diffusion in leaves nor the distance
between chloroplasts and the intercellular air space. Rather,
a liming effect of the avoidance response of chloroplasts on
photosynthetic activity was reported recently. Tholen et al.
(2008) analysed the relationship between the distribution
patterns of chloroplasts and Sc in Arabidopsis thaliana.
After induction of the avoidance response of chloroplasts to
high-fluence-rate light, which tends to orientate chloroplasts
along cell–cell boundaries from cell–airspace boundaries, Sc
was decreased significantly together with CO2 conductance.
The changes in these two parameters were not noted in the
phot2 mutant, which is deficient in the avoidance response
of chloroplasts, and the chup1 mutant, which exhibits no
chloroplast movement. Tholen et al. (2008) concluded that
the avoidance response of chloroplasts may represent
a limiting factor for photosynthetic activity under some
conditions. Under very high light, such responses may be
advantageous in protecting the photosynthetic machinery
from photodamage.
Consequently, there is no doubt that chloroplast anchoring is not only an attractive subject in terms of developmental and cell biology, but is also important in understanding
plant ecophysiology. In the following sections, the possible
mechanisms underlying chloroplast anchoring investigated
to date, in which the actin cytoskeleton has been suggested
to play central roles, will be discussed.
Cytoskeletal involvement
The actin cytoskeleton has been postulated to play an
important role in determining the intracellular distribution
of chloroplasts. Close association of chloroplasts with actin
filaments has been demonstrated in vascular plants as well
as algae (Takagi, 2003). A possible involvement of actin
filaments in chloroplast anchoring was suggested from the
effects of actin-depolymerizing reagents. In A. thaliana
mesophyll cells, disruption of actin filaments in a basketlike configuration surrounding each chloroplast by the
actin-depolymerizing reagents produced a disordered arrangement of chloroplasts (Kandasamy and Meagher,
1999). A similar treatment of leaf epidermal cells of
Vallisneria gigantea rendered the chloroplasts less resistant
to centrifugal force (Dong et al., 1998), although this was
not the case for chloroplasts in the bundle sheath cells of
finger millet as already described (Kobayashi et al., 2009).
Although there are several reports on microtubuledependent chloroplast movement in algae (Mizukami and
Wada, 1981; Maekawa et al., 1986; Menzel and Schliwa,
1986) and a moss (Sato et al., 2001), evidence for the
involvement of microtubules in chloroplast anchoring is
scarce. In the Chenopodiaceae, a unique cell type, in which
the entire C4 photosynthesis cycle is completed within single
chlorenchyma cells, has been reported (Edwards et al.,
2004). In those cells, chloroplasts and other organelles
exhibit specific distribution patterns. Essentially, chloroplasts of two different types occupy distinct intracellular
positions in the single chlorenchyma cell to achieve their
respective roles, namely, primary CO2 fixation by PEP
carboxylase and its subsequent liberation and refixation by
Rubisco. Using cytoskeletal inhibitors and immunofluorescence microscopy, Chuong et al. (2006) suggested that
microtubules play an important role in the maintenance of
chloroplast positioning. Whether the role of microtubules
is specific to chloroplasts or more general in maintaining
the cytoplasmic architecture remains to be investigated.
A role for microtubules was also suggested in the
anchorage of non-green plastids in the hypocotyl
epidermis of dark-grown tobacco (Kwok and Hanson,
2003). Although the motility of plastids is under the control
of the actin cytoskeleton, treatment of the cells with
3304 | Takagi et al.
microtubule-disrupting reagents induced an acceleration of
plastid motility.
Chloroplast motility and actin cytoskeleton
There have been a couple of reports examining a possible
causal relationship between actin organization and the
motility and/or positioning of chloroplasts (Takagi, 2000).
In protonemal cells of a fern, Adiantum capillus-veneris,
circular arrays of actin filaments associated with each
chloroplast appeared after the termination of photorelocation movement and disappeared before the chloroplasts
retrieved their motility in darkness (Kadota and Wada,
1992). The characteristic circular arrays of actin filaments
may function to maintain the specific distribution pattern of
chloroplasts. A brief illumination of leaf epidermal cells of
V. gigantea with low-fluence-rate red light increased chloroplast motility within a few minutes (Izutani et al., 1990;
Dong et al., 1996). Thereafter, the motility drastically
declined and the chloroplasts were almost immobile in less
than an hour. This loss of chloroplast motility appears to be
an important first step in the accumulation response of
chloroplasts along the outer periclinal walls of the cells.
Photobiological dissection of these processes revealed that
the acceleration of chloroplast motility was initiated by the
formation of Pfr, whereas the subsequent deceleration was
regulated by photosynthesis (Dong et al., 1995, 1996). With
the deceleration of chloroplast motility, the configuration of
actin filaments along the outer periclinal walls changed
dynamically from a loose network array to a honeycomb
array, in which individual chloroplasts are tightly surrounded by actin filaments (Dong et al., 1996, 1998).
On the other hand, during the avoidance response of
chloroplasts, immobile chloroplasts must become mobile
before the start of the relocation movement. In the
epidermal cells of V. gigantea, strong blue light initially
induces randomly oriented, short-range movement of chloroplasts within a few minutes (Sakurai et al., 2005). Such
chloroplasts become easily displaced by centrifugation,
suggesting their release from an anchored state (Sakai and
Takagi, 2005). Thereafter the pattern of chloroplast movement becomes increasingly unidirectional. Concomitant
with these changes in the mode of chloroplast movement,
the configuration of actin filaments along the outer periclinal walls changes from being a tightly packed basket-like
array around the chloroplasts to linearly aggregated bundles. By contrast, along the anticlinal walls, the chloroplasts
migrating from the outer periclinal region became surrounded by actin filaments in a basket-like array (Sakai
and Takagi, 2005). These chloroplasts regained their resistance to centrifugal force. These processes may contribute
to an efficient redistribution of chloroplasts in the epidermal
cells exposed to high-fluence-rate illumination.
Importantly, in the presence of an inhibitor of photosynthetic electron transport, neither low-fluence-rate red light
(Dong et al., 1996) nor high-fluence-rate blue light (Sakai
and Takagi, 2005) produced any detectable changes in the
configuration of actin filaments along the outer periclinal
walls or anticlinal walls, respectively. Under these conditions, chloroplasts exhibited a high level of motility and
easily obeyed centrifugal force. Although there was a contradictory report concerning the role of photosynthesis
(Ślesak and Gabryś, 1996), Seitz (1979, 1980) repeatedly
argued that photosynthesis is involved in light-induced
decreases in chloroplast centrifugability. The results
obtained in V. gigantea may support this proposal and,
moreover, strongly suggest the critical involvement of the
actin cytoskeleton in the regulation of light-induced chloroplast anchoring.
In the case of A. thaliana mesophyll cells, in which typical
light-dependent accumulation and avoidance responses of
chloroplasts are observed (Trojan and Gabryś, 1996;
Kagawa and Wada, 2000), Krzeszowiec et al. (2007) showed
that blue light never induced any specific reorganization in
the actin cytoskeleton regardless of its fluence rate. However, since the samples appeared to be illuminated with
actinic light immediately after incision into small pieces of
the leaf, the possible effects of physical injury on the
cytoskeleton should be considered. Nevertheless, there is
a possibility that actin reorganization in a very localized,
tiny region is functional in the regulation of chloroplast
movement and positioning in A. thaliana.
Interaction of chloroplasts with the cortical
cytoplasm/plasma membrane
For proper chloroplast anchoring, interaction between
chloroplasts and the cortical cytoplasm and/or PM may be
indispensable. Oikawa et al. (2003) identified a unique
mutant, chloroplast unusual positioning 1 (chup1), in A.
thaliana deficient in chloroplast photo-relocation movement. In palisade mesophyll cells of the chup1 mutant,
chloroplasts constitutively form an aggregate at the bottom
of the cell. The chup1 gene encodes a protein equipped with
several domains exhibiting putative functions, namely, the
N-terminal hydrophobic, coiled-coil, F-actin-binding,
proline-rich, and C-terminal conserved domains. CHUP1
protein translated in vitro was shown to associate with actin
and profilin (Schmidt von Braun and Schleiff, 2008).
However, whether CHUP1 can modulate the polymerization–depolymerization state of actin was not examined.
Oikawa et al. (2008) precisely analysed the functions of
N-terminal hydrophobic and coiled-coil domains in A.
thaliana. In palisade mesophyll cells, CHUP1 is localized
on the outer chloroplast envelope. This localization is
dependent on the presence of its N-terminal hydrophobic
domain, and is impaired by over-expression of GFP fused
to the N-terminal hydrophobic domain, concomitant with
the expression of the chup1 phenotype. On the other hand,
the coiled-coil domain is important in determining the
distribution of chloroplasts along the anticlinal walls of the
cells. Oikawa et al. (2008) proposed that CHUP1 is an
indispensable factor for the association of chloroplasts with
the cortical cytoplasm and the actin cytoskeleton. Further
Chloroplast anchoring | 3305
characterization of CHUP1, especially the modes of regulation of its functions and localization, will provide critical
information on the mechanisms of chloroplast movement
and positioning (Wada and Suetsugu, 2004).
Augustynowicz et al. (2001) examined chloroplast movement in isolated protoplasts from tobacco leaves. They
found that the responsiveness to light was markedly
enhanced when intact leaves were subjected to cold stress
before the isolation of protoplasts. The protoplasts isolated
from cold-stressed leaves exhibited an increased rigidity,
which was greatly antagonized by treatment with an actindepolymerizing reagent. Although cold-induced factors involved in regulation of the rigidity of protoplasts have not
yet been identified, these results may support the idea that
the actin cytoskeleton is an important structural component
of the cortical cytoplasm, and that the interaction with
intact cortical cytoplasm is required for proper chloroplast
movement and positioning.
Dissection of chloroplast anchoring using
spinach
To test the hypothesis that chloroplasts under physiologically stationary conditions are anchored on the cortical
cytoplasm in an actin-dependent manner, cell biological
dissection of the subject may be necessary. For this purpose,
spinach was chosen as the preferred experimental system for
the following reasons; first, blue-light-dependent chloroplast
movement was observed (Inoue and Shibata, 1973; Kumatani
et al., 2006) concomitant with the reorganization of the
actin cytoskeleton (Kumatani et al., 2006; Fig. 1); second,
immunoblotting analyses of biochemically prepared crudemicrosome and PM fractions detected actin as well as phot1
(Fig. 2). In A. thaliana, both phot1 and phot2 are predominantly localized on the PM, and some populations of
them are relocalized to endomembranes after photoperception (Sakamoto and Briggs, 2002; Harada et al., 2003; Kong
et al., 2006; Wan et al., 2008). Third, procedures for the
preparation of protoplasts and isolated chloroplasts are well
established (Walker, 1987). In this section, examination of
the hypothesis described above is summarized using spinach, comparing results obtained in other plant species,
together with future prospects.
As described, the interaction of actin filaments with
chloroplasts in vivo has been suggested in various types of
plant cells (Takagi, 2003), while attempts to detect the
interaction in vitro were assessed for the first time in
spinach. In the final chloroplast fraction isolated from
spinach leaves (Spinacia oleracea L. cv. Torai), endogenous
actin was not detected, although these chloroplasts cosedimented with exogenously added skeletal muscle filamentous actin in vitro (Kumatani et al., 2006). Binding was
strictly dependent on the intactness of the outer envelope of
isolated chloroplasts, and was at least partially light
sensitive (Kumatani et al., 2006). The interaction of
exogenously added actin, regardless of its polymerization
state, with isolated chloroplasts was demonstrated in
Fig. 1. Actin organization in spinach mesophyll cells under
different light conditions. Actin filaments in mesophyll cells either
kept under low-fluence-rate white light (0.9 W m 2) (A) or exposed
to high-fluence-rate blue light (460 nm, 15 W m 2) (C) were
visualized by staining with fluorescent phalloidin. Under dim light,
many fine bundles of actin filaments are associated with chloroplasts (B), whereas under strong blue light, much thicker, linear
actin bundles run in the cytoplasm (D). Bar: 10 lm.
a similar way in A. thaliana (Schmidt von Braun and
Schleiff, 2008). Involvement of actin-binding proteins, such
as CHUP1, in this interaction should be urgently examined.
The interaction between chloroplasts and the cortical
cytoplasm and/or PM may be crucial for proper chloroplast
anchoring. The structure and function of plant cortical
cytoplasm have been studied using PM ghosts. PM ghosts
are prepared after rupture or lysis of protoplasts so as to
expose the cortical cytoplasm underlying the PM. Experimental systems using PM ghosts have markedly contributed
to our understanding of the cortical microtubule organization (Cyr, 1991; Sonobe and Takahashi, 1994), actin
organization (Kobayashi, 1996), and the interaction between actin and microtubule cytoskeletons (Collings et al.,
1998).
During the development of tracheary elements in cultured
mesophyll cells of Zinnia elegans, Kobayashi et al. (1987)
observed reticulated arrays of actin filaments between
chloroplasts and the PM. Since those structures disappeared
at the stage of development when chloroplasts are detached
from the PM, Kobayashi et al. (1987) concluded that the
reticulated array of actin filaments functions to anchor
chloroplasts to the cortical cytoplasm and/or PM. Kobayashi
3306 | Takagi et al.
Fig. 2. Immunodetection of phototropins and actin in spinach leaf
cells. The crude-microsome (CM) and plasma-membrane (PM)
fractions prepared from spinach leaves were separated by SDSPAGE (10 lg in each lane). A polypeptide of 120 kDa was
detected with the anti-phot1 antibody, but not with the anti-phot2
antibody. A polypeptide of 43 kDa was detected with the anti-actin
antibody.
(1996) confirmed the presence and disappearance of similar
arrays of actin filaments on PM ghosts prepared from the
mesophyll cells at various developmental stages. He further
demonstrated that such arrays of actin filaments could be
removed from the PM ghosts by washing with a high
concentration of salt, suggesting an involvement of protein–
protein interaction (Kobayashi, 1996).
On PM ghosts prepared from spinach mesophyll protoplasts kept under dim white light, from which most of the
chloroplasts were removed after gentle treatment with
a detergent (Fig. 3A), fine bundles of actin filaments were
seen in configurations suggesting that they had surrounded
individual chloroplasts (Fig. 3B). By contrast, when PM
ghosts were prepared from protoplasts exposed to highfluence-rate blue light (Fig. 3C), much thicker and straighter
bundles of actin filaments were detected (Fig. 3D). These
distinctive configurations of actin filaments are reminiscent
of those observed in living spinach cells under similar light
conditions (Fig. 1). The results suggest that light-induced
reorganization of the actin cytoskeleton maintained some
association with the cortical cytoplasm and/or PM, and
especially, the reticulated array of actin filaments plays an
important role to anchor chloroplasts on the cortical
cytoplasm in spinach as well as Z. elegans. To date, there
has been no indication that suggests an involvement of the
activity of motor proteins in chloroplast anchoring.
Without detergent treatment, more native PM ghosts, to
which chloroplasts remain attached, can be prepared from
spinach mesophyll cells (Fig. 4A). The actin cytoskeleton is
also retained (Fig. 4B), and its organization seems to
Fig. 3. Actin organization on PM ghosts prepared from spinach
mesophyll protoplasts. After chemical fixation, PM ghosts were
treated with a detergent to remove chloroplasts and then labeled
with an anti-actin antibody. On the PM ghosts prepared from
protoplasts kept under dim white light (A), the actin filaments have
surrounded each chloroplast (B), as visualized in living cells (see
Fig. 1B). By contrast, on the PM ghosts prepared from protoplasts
exposed to strong blue light (C), the actin filaments form thicker,
linear bundles (D), very similar to those observed in living cells (see
Fig. 1D). Bar: 10 lm.
surround each chloroplast on the PM ghost. When such
PM ghosts were treated with an actin-depolymerizing
reagent (Fig. 4C) or Ca2+ at physiologically relevant
concentrations (Fig. 4D), detachment of chloroplasts from
the PM ghosts was induced. More intriguingly, the calmodulin antagonist N-(6-aminohexyl)-5-chloro-1-naphthalenesulphonamide (W-7) almost completely blocked the effect
of Ca2+ (Fig. 4E), while its derivative N-(6-aminohexyl)-1naphthalenesulphonamide (W-5) did not (Fig. 4F), suggesting that the mechanism of chloroplast anchoring in spinach
mesophyll cells is under the control of the Ca2+-calmodulin
system. Consequently, the PM ghosts prepared from
spinach mesophyll cells preserve the intactness in terms of
regulation of chloroplast anchoring on the cortical cytoplasm. Although the involvement of calmodulin in bluelight-dependent chloroplast movement was suggested in
Lemna trisulca (Tla1ka and Fricker, 1999), its specific target
process was not clear. The mode of regulation of actin
organization by the Ca2+-calmodulin system and a possible
involvement of actin-binding proteins should be investigated.
Finally, elementary processes are proposed that are
associated with the redistribution of chloroplasts in plant
cells induced by environmental stimuli (Fig. 5). Under
physiologically stable conditions, chloroplasts are anchored
to the cortical cytoplasm and/or PM through their interaction with actin filaments. Although numerous
Chloroplast anchoring | 3307
Fig. 5. A schematic model for the elementary processes associated with the redistribution of chloroplasts in plant cells. Environmental stimuli, such as blue light and mechanical touch, induce
the redistribution of chloroplasts through the modulation of actin
organization in the cortical cytoplasm. For further details, see the
text. ABPs, actin-binding proteins; [Ca2+]cyt, cytoplasmic Ca2+
concentration.
Fig. 4. Bright-field images of PM ghosts prepared from spinach
mesophyll cells. When PM ghosts were prepared in the absence of
detergent, the ghosts were occupied by numerous chloroplasts
(A). Those chloroplasts seem to be tightly associated with actin
filaments (red: chlorophyll autofluorescence, green: actin) (B).
When the prepared PM ghosts were treated with the actindepolymerizing reagent latrunculin B (C, +LatB) or Ca2+ at 10 6 M
(D, +Ca), many of the chloroplasts were detached from the PM
ghosts. The effect of Ca2+ was antagonized by the calmodulin
antagonist W-7 (E) but not by its derivative W-5 (F). Bar: 10 lm.
functional actin-binding proteins have been identified and
characterized in plant cells (Staiger, 2000; Higaki et al.,
2007), at present, the only known factor involved in
chloroplast anchoring is CHUP1. Essentially, there is no
information about protein factors mediating the interaction
of the actin cytoskeleton with the cortical cytoplasm and/or
PM. Various kinds of environmental stimuli, including blue
light (Harada and Shimazaki, 2007) and mechanical touch
(Knight, 2000), which is also known to induce chloroplast
movement (Makita and Shihira-Ishikawa, 1997; Sato et al.,
1999, 2003), trigger a transient increase in the cytoplasmic
Ca2+ concentration. Ca2+-sensitive actin-binding proteins
may modulate the actin organization in the cortical
cytoplasm, inducing a release of chloroplasts from the
anchored state (de-anchoring). According to signals produced from the receptors, for example, phototropins and
mechanosensory systems (Nakagawa et al., 2007), chloroplasts migrate to their intracellular destinations depending
on the interaction of cytoskeletal components with specific
motor proteins such as actomyosin systems (Shimmen and
Yokota, 2004). Little is known about the identity of the
various signals regulating chloroplast movements, namely,
the direction and velocity of movement, to date. In their
proper positions, chloroplasts are re-anchored through the
reorganization of the actin cytoskeleton. Although an
involvement of photosynthetic activity in the regulation has
been suggested in several plant species, the factors involved
in chloroplast re-anchoring are also elusive. Further multidisciplinary approaches to these elementary processes
should shed light on the mechanism of actin-dependent
chloroplast anchoring and positioning in plant cells and its
significance for plant life.
Acknowledgements
We are grateful to the anonymous reviewers for numerous
invaluable comments to improve this article. Thanks also go
to Louis John Irving of Tohoku University for his critical
reading of the manuscript.
References
Augustynowicz J, Lekka M, Burda K, Gabryś H. 2001. Correlation
between chloroplast motility and elastic properties of tobacco mesophyll protoplasts. Acta Physiologiae Plantarum 23, 291–302.
Chuong SDX, Franceschi VR, Edwards GE. 2006. The cytoskeleton maintains organelle partitioning required for single-cell C4 photosynthesis in Chenopodiaceae species. The Plant Cell 18, 2207–2223.
Collings DA, Asada T, Allen NS, Shibaoka H. 1998. Plasma
membrane-associated actin in bright yellow 2 tobacco cells: evidence
for interaction with microtubules. Plant Physiology 118, 917–928.
3308 | Takagi et al.
Cyr RJ. 1991. Calcium/calmodulin affects microtuble stability in lysed
protoplasts. Journal of Cell Science 100, 311–317.
Dong XJ, Nagai R, Takagi S. 1998. Microfilaments anchor
chloroplasts along the outer periclinal wall in Vallisneria epidermal cells
through cooperation of Pfr and photosynthesis. Plant and Cell
Physiology 39, 1299–1306.
Dong XJ, Ryu JH, Takagi S, Nagai R. 1996. Dynamic changes in
the organization of microfilaments associated with the photocontrolled
motility of chloroplasts in epidermal cells of Vallisneria. Protoplasma
195, 18–24.
Dong XJ, Takagi S, Nagai R. 1995. Regulation of the orientation
movement of chloroplasts in epidermal cells of Vallisneria: cooperation of phytochrome with photosynthetic pigment under lowfluence-rate light. Planta 197, 257–263.
Edwards GE, Franceschi VR, Voznesenskaya EV. 2004. Single
cell C4 photosynthesis versus the dual-cell (Kranz) paradigm. Annual
Review of Plant Physiology and Plant Molecular Biology 55,
173–196.
Kagawa T, Wada M. 2000. Blue light-induced chloroplast relocation
in Arabidopsis thaliana as analyzed by microbeam irradiation. Plant
and Cell Physiology 41, 84–93.
Kagawa T, Sakai T, Suetsugu N, Oikawa K, Ishiguro S, Kato T,
Tabata S, Okada K, Wada M. 2001. Arabidopsis NPL1: a phototropin homolog controlling the chloroplast high-light avoidance response. Science 291, 2035–2262.
Kandasamy MK, Meagher RB. 1999. Actin–organelle interaction:
association with chloroplast in Arabidopsis leaf mesophyll cells. Cell
Motility and the Cytoskeleton 44, 110–118.
Kasahara M, Kagawa T, Oikawa K, Suetsugu N, Miyao M,
Wada M. 2002. Chloroplast avoidance movement reduces photodamage in plants. Nature 420, 829–832.
Kawai H, Kanegae T, Christensen S, Kiyosue T, Sato Y,
Imaizumi T, Kadota A, Wada M. 2003. Responses of ferns to red
light are mediated by an unconventional photoreceptor. Nature 421,
287–290.
Evans JR, von Caemmerer S. 1996. Carbon dioxide diffusion inside
leaves. Plant Physiology 110, 339–346.
Keifer DW, Spanswick RM. 1979. Correlation of adenosine triphosphate levels in Chara corallina with the activity of the electrogenic
pump. Plant Physiology 64, 165–168.
Gorton HL, Herbert SK, Vogelmann TC. 2003. Photoacoustic
analysis indicates that chloroplast movement does not alter liquidphase CO2 diffusion in leaves of Alocasia brisbanensis. Plant
Physiology 132, 1529–1539.
Kikuyama M, Hayama T, Fujii S, Tazawa M. 1979. Relationship
between light-induced potential change and internal ATP concentration in tonoplast-free Chara cells. Plant and Cell Physiology 20,
993–1002.
Harada A, Shimazaki K. 2007. Phototropins and blue light-dependent calcium signaling in higher plants. Photochemistry and
Photobiology 83, 102–111.
Knight H. 2000. Calcium signaling during abiotic stress in plants.
International Review of Cytology 195, 269–324.
Harada A, Sakai T, Okada K. 2003. Phot1 and phot2 mediate blue
light-induced transient increases in cytosolic Ca2+ differently in
Arabidopsis leaves. Proceedings of the National Academy of Sciences,
USA 100, 8583–8588.
Hatch MD, Slack CR. 1970. Photosynthetic CO2-fixation pathways.
Annual Review of Plant Physiology 21, 141–163.
Haupt W, Scheuerlein R. 1990. Chloroplast movement. Plant, Cell
and Environment 13, 595–614.
Higaki T, Sano T, Hasezawa S. 2007. Actin microfilament dynamics
and actin side-binding proteins in plants. Current Opinion in Plant
Biology 10, 549–556.
Hiramoto H, Kamitsubo E. 1995. Centrifuge microscope as a tool in
the study of cell motility. International Review of Cytology 157, 99–128.
Inoue Y, Shibata K. 1973. Light-induced chloroplast rearrangements
and their action spectra as measured by absorption spectrophotometry. Planta 114, 341–358.
Izutani Y, Takagi S, Nagai R. 1990. Orientation movements of
chloroplasts in Vallisneria epidermal cells: different effects of light at
low- and high-fluence rate. Photochemistry and Photobiology 51,
105–111.
Jarillo JA, Gabryś H, Capel J, Alonso JM, Ecker JR,
Cashmore AR. 2001. Phototropin-related NPL1 controls chloroplast
relocation induced by blue light. Nature 410, 952–954.
Kadota A, Wada M. 1992. Photoinduction of formation of circular
structures by microfilaments on chloroplasts during intracellular
orientation in protonemal cells of the fern Adiantum capillus-veneris.
Protoplasma 167, 97–107.
Kobayashi H. 1996. Changes in the relationship between actin
filaments and the plasma membrane in cultured Zinnia cells during
tracheary elements differentiation investigated by using plasma membrane ghosts. Journal of Plant Research 109, 61–65.
Kobayashi H, Fukuda H, Shibaoka H. 1987. Reorganization of actin
filaments associated with the differentiation of tracheary elements in
Zinnia mesophyll cells. Protoplasma 138, 69–71.
Kobayashi H, Yamada M, Taniguchi M, Kawasaki M,
Sugiyama T, Miyake H. 2009. Differential positioning of C4 mesophyll
and bundle sheath chloroplasts: recovery of chloroplast positioning
requires the actomyosin system. Plant and Cell Physiology 50,
129–140.
Kong S-G, Suzuki T, Tamura K, Mochizuki N, Hara-Nishimura I,
Nagatani A. 2006. Blue light-induced association of phototropin 2
with the Golgi apparatus. The Plant Journal 45, 994–1005.
Krzeszowiec W, Rajwa B, Dobrucki J, Gabryś H. 2007. Actin
cytoskeleton in Arabidopsis thaliana under blue and red light. Biology
of the Cell 99, 251–260.
Kumatani T, Sakurai-Ozato N, Miyawaki N, Yokota E,
Shimmen T, Terashima I, Takagi S. 2006. Possible association of
actin filaments with chloroplasts of spinach mesophyll cells in vivo and
in vitro. Protoplasma 229, 45–52.
Kwok EY, Hanson MR. 2003. Microfilaments and microtubules
control the morphology and movement of non-green plastids and
stromules in Nicotiana tabacum. The Plant Journal 35, 16–26.
Lee Y-RJ, Liu B. 2004. Cytoskeletal motors in Arabidopsis. Sixtyone kinesins and seventeen myosins. Plant Physiology 136,
3877–3883.
Chloroplast anchoring | 3309
Maekawa T, Tsutsui I, Nagai R. 1986. Light-regulated translocation
of cytoplasm in green alga Dichotomosiphon. Plant and Cell Physiology 27, 837–851.
Sato Y, Kadota A, Wada M. 1999. Mechanically induced avoidance
response of chloroplasts in fern protonemal cells. Plant Physiology
121, 37–44.
Makita N, Shihira-Ishikawa I. 1997. Chloroplast assemblage by
mechanical stimulation and its intercellular transmission in diatom
cells. Protoplasma 197, 86–95.
Sato Y, Wada M, Kadota A. 2001. Choice of tracks, microtubules
and/or actin filaments for chloroplast photo-movement is differentially
controlled by phytochrome and a blue light receptor. Journal of Cell
Science 114, 269–279.
Menzel D, Schliwa M. 1986. Motility in the siphonous green alga
Bryopsis. II. Chloroplast movement requires organized arrays of both
microtubules and actin filaments. European Journal of Cell Biology 40,
286–295.
Sato Y, Wada M, Kadota A. 2003. Accumulation response of
chloroplasts induced by mechanical stimulation in bryophyte cells.
Planta 216, 772–777.
Miyake H, Nakamura M. 1993. Some factors concerning the
centripetal disposition of bundle sheath chloroplasts during the leaf
development of Eleusine coracana. Annals of Botany 72, 205–211.
Schmidt von Braun S, Schleiff E. 2008. The chloroplast outer
envelope protein CHUP1 interacts with actin and profilin. Planta 227,
1151–1159.
Miyake H, Yamamoto Y. 1987. Centripetal disposition of bundle
sheath chloroplasts during the leaf development of Eleusine coracana.
Annals of Botany 60, 641–647.
Seitz K. 1971. Die Ursache der Phototaxis der Chloroplasten: ein
ATP-Gradient? Zeitschrift für Pflanzenphysiologie 64, 241–256.
Mizukami M, Wada S. 1981. Action spectrum for light-induced
chloroplast accumulation in a marine coenocytic green alga, Bryopsis
plumosa. Plant and Cell Physiology 22, 1245–1255.
Seitz K. 1979. Light induced changes in the centrifugability of
chloroplasts: different action spectra and differnt influence of inhibitors
in the low and hight intensity range. Zeitschrift für Pflanzenphysiologie
95, 1–12.
Nakagawa Y, Katagiri T, Shinozaki K, et al. 2007. Arabidopsis
plasma membrane protein crucial for Ca2+ influx and touch sensing in
roots. Proceedings of the National Academy of Sciences, USA 104,
3639–3644.
Seitz K. 1980. Light induced changes in the centrifugability of
chloroplasts mediated by an irradiance dependent interaction of
respiratory and photosynthetic processes. In: Senger H, ed. The blue
light syndrome. Berlin, Heidelberg: Springer-Verlag, 637–642.
Nozue K, Kanegae T, Imaizumi T, Fukuda S, Okamoto H,
Yeh KC, Lagarias JC, Wada M. 1998. A phytochrome from the fern
Adiantum with features of the putative photoreceptor NPH1. Proceedings of the National Academy of Sciences, USA 95,
15826–15830.
Shimmen T. 1978. Dependency of cytoplasmic streaming on intracellular ATP and Mg2+ concentrations. Cell Structure and Function
3, 113–121.
Oikawa K, Kasahara M, Kiyosue T, Kagawa T, Suetsugu N,
Takahashi F, Kanegae T, Niwa Y, Kadota A, Wada M. 2003.
CHLOROPLAST UNUSUAL POSITIONING1 is essential for proper
chloroplast positioning. The Plant Cell 15, 2805–2815.
Ślesak I, Gabryś H. 1996. Role of photosynthesis in the control of
blue light induced chloroplast movement. Inhibitor study. Acta
Physiologiae Plantarum 18, 135–145.
Oikawa K, Yamasato A, Kong S-G, Kasahara M, Nakai M,
Takahashi F, Ogura Y, Kagawa T, Wada M. 2008. Chloroplast
outer envelope protein CHUP1 is essential for chloroplast anchorage
to the plasma membrane and chloroplast movement. Plant Physiology
148, 829–842.
Park Y-I, Chow WS, Anderson JM. 1996. Chloroplast movement in
the shade plant Tradescantia albiflora helps protect photosystem II
against light stress. Plant Physiology 111, 867–875.
Sakai T, Kagawa T, Kasahara M, Swartz TE, Christie JM,
Briggs WR, Wada M, Okada K. 2001. Arabidopsis nph1 and npl1:
blue light receptors that mediate both phototropism and chloroplast
relocation. Proceedings of the National Academy of Sciences, USA
98, 6969–6974.
Sakai Y, Takagi S. 2005. Reorganized actin filaments anchor
chloroplasts along the anticlinal walls of Vallisneria epidermal cells
under high-intensity blue light. Planta 221, 823–830.
Sakamoto K, Briggs WR. 2002. Cellular and subcellular localization
of phototropin 1. The Plant Cell 14, 1723–1735.
Sakurai N, Domoto K, Takagi S. 2005. Blue-light-induced reorganization of the actin cytoskeleton and the avoidance response of
chloroplasts in epidermal cells of Vallisneria gigantea. Planta 221,
66–74.
Shimmen T, Yokota E. 2004. Cytoplasmic streaming in plants.
Current Opinion in Cell Biology 16, 68–72.
Sonobe S, Takahashi S. 1994. Association of microtubules with the
plasma membrane of tobacco BY-2 cells in vitro. Plant and Cell
Physiology 35, 451–460.
Staiger CJ. 2000. Signaling to the actin cytoskeleton in plants. Annual
Review of Plant Physiology and Plant Molecular Biology 51,
257–288.
Takagi S. 2000. Roles for actin filaments in chloroplast motility and
anchoring. In: Staiger CJ, Baluška F, Volkmann D, Barlow PW, eds.
Actin: a dynamic framework for multiple plant cell functions. Dordrecht:
Kluwer Academic Publishers, 203–212.
Takagi S. 2003. Actin-based photo-orientation movement of chloroplasts in plant cells. Journal of Experimental Botany 206, 1963–1969.
Terashima I, Hanba YT, Tazoe Y, Vyas P, Yano S. 2006.
Irradiance and phenotype: comparative eco-development of sun and
shade leaves in relation to photosynthetic CO2 diffusion. Journal of
Experimental Botany 57, 343–354.
Tholen D, Boom C, Noguchi K, Ueda S, Katase T, Terashima I.
2008. The chloroplast avoidance response decreases internal conductance to CO2 diffusion in Arabidopsis thaliana leaves. Plant, Cell
and Environment 31, 1688–1700.
Tla1ka M, Fricker M. 1999. The role of calcium in blue-lightdependent chloroplast movement in Lemna trisulca L. The Plant
Journal 20, 461–473.
3310 | Takagi et al.
Trojan A, Gabryś H. 1996. Chloroplast distribution in Arabidopsis
thaliana (L.) depends on light conditions during growth. Plant
Physiology 111, 419–425.
Walker D. 1987. The use of oxygen electrode and fluorescence
probes in simple measurements of photosynthesis. Sheffield: Produced Oxgraphics Ltd.
Wada M, Kadota A. 1989. Photomorphogenesis in lower green
plants. Annual Review of Plant Physiology and Plant Molecular Biology
40, 169–191.
Wan Y-L, Eisinger W, Ehrhardt D, Kubitscheck U, Baluska F,
Briggs W. 2008. The subcellular localization and blue-light-induced
movement of phototropin 1-GFP in etiolated seedlings of Arabidopsis
thaliana. Molecular Plant 1, 103–117.
Wada M, Kagawa T, Sato Y. 2003. Chloroplast movement. Annual
Review of Plant Physiology and Plant Molecular Biology 54, 455–468.
Wada M, Suetsugu N. 2004. Plant organelle positioning. Current
Opinion in Plant Biology 7, 626–631.
Wagner G, Klein K. 1981. Mechanism of chloroplast movement in
Mougeotia. Protoplasma 109, 169–185.
Walczak T, Gabryś H. 1981. The carbon dioxide (CO2) effect on
light-induced chloroplast translocations in higher plant leaves.
Zeitschrift für Pflanzenphysiologie 101, 367–375.
Yatsuhashi H, Kobayashi H. 1993. Dual involvement of phytochrome in light-oriented chloroplast movement in Dryopteris sparsa
protonemata. Journal of Photochemistry and Photobiology B: Biology
19, 25–31.
Zurzycki J. 1955. Chloroplasts arrangement as a factor in photosynthesis. Acta Societa Botanica Poland 24, 27–63.
Zurzycki J. 1957. The destructive effect of light on the photosynthetic
apparatus. Acta Societa Botanica Poland 26, 157–175.