Download FM Dyes Label Sterol-Rich Plasma Membrane

Plant Cell Physiol. 49(10): 1508–1521 (2008)
doi:10.1093/pcp/pcn122, available online at www.pcp.oxfordjournals.org
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FM Dyes Label Sterol-Rich Plasma Membrane Domains and
are Internalized Independently of the Cytoskeleton in Characean
Internodal Cells
Andreas Klima and Ilse Foissner *
Department of Cell Biology, Division of Plant Physiology, University of Salzburg, Hellbrunnerstrasse 34, A-5020 Salzburg, Austria
We applied the endocytic markers FM1-43, FM4-64
and filipin to internodal cells of the green alga Chara
corallina. Both FM dyes stained stable, long-living plasma
membrane patches with a diameter of up to 1 mm. After 5 min,
FM dyes labeled cortical, trembling structures up to 500 nm
in size. After 15 min, FM dyes localized to endoplasmic
organelles up to 1 mm in diameter, which migrated actively
along actin bundles or participated in cytoplasmic mass
streaming. After 30–60 min, FM fluorescence appeared in the
membrane of small, endoplasmic vacuoles but not in that of
the central vacuole. Some of the FM-labeled organelles
were also stained by neutral red and lysotracker yellow,
indicative of acidic compartments. Filipin, a sterol-specific
marker, likewise labeled plasma membrane domains which
co-localized with the FM patches. However, internalization
of filipin could not be observed. KCN, cytochalasin D,
latrunculin B and oryzalin had no effect on size, shape and
distribution of FM- and filipin-labeled plasma membrane
domains. Internalization of FM dyes was inhibited by KCN
but not by drugs which interfere with the actin or microtubule
cytoskeleton. Our data indicate that the plasma membrane
of characean internodal cells contains discrete domains
which are enriched in sterols and probably correspond to
clusters of lipid rafts. The inhibitor experiments suggest that
FM uptake is active but independent of actin filaments,
actin polymerization and microtubules. The possible function
of the sterol-rich, FM labeled plasma membrane areas
and the significance of actin-independent FM internalization (via endocytosis or energy-dependent flippases) are
discussed.
Keywords: Chara — Cytoskeleton — Endocytosis — Lipid
raft — Plasma membrane domain — Sterol.
Abbreviations: AFW, artificial fresh water; BDM, 2,3butanedione monoxime; CD, cytochalasin D; CLSM,
confocal laser scanning microscope/microscopy; DIC,
differential interference contrast; DiOC6(3), 3,30 , dihexyloxacarbocyanine iodide; DMSO, dimethylsulfoxide; FRAP,
fluorescence recovery after photobleaching; LatB, latrunculin B; PBS, phosphate-buffered saline; PIPES, piperazineN,N0 -bis (2-ethanesulfonic acid).
Introduction
Endocytosis is the process by which cells internalize
plasma membrane and extracellular material. It is important for the recycling of plasma membrane components, for
nutrient uptake and for signaling (for recent reviews, see
Geldner 2004, Murphy et al. 2005, Samaj et al. 2005, Polo
and Di Fiore 2006, Samaj et al. 2006a, Samaj et al. 2006b).
A variety of endocytic pathways have been described in
animal and fungal cells which differ mainly in the
participation of proteins and in the size of endocytic
vesicles (Conner and Schmid 2003, Soldati and Schliwa
2006). Until a few years ago, little was known about
endocytosis in plant cells. Although earlier work proved
uptake of electron-opaque markers from the external
medium (Hawes et al. 1991), the existence of endocytosis
in plant cells was even questioned because of the high turgor
pressure (references in Marcote et al. 2000, Aniento and
Robinson 2005). This changed rapidly with the introduction
of FM dyes and filipin as endocytic tracers and with
the development of new molecular biological techniques
(Betz et al. 1996, Grebe et al. 2003, Meckel et al. 2004).
Today, the existence of endocytotic processes in plant cells
is beyond debate, but our understanding of plasma
membrane internalization and uptake of extracellular
material is still very limited. Currently, research focuses
on clathrin-dependent endocytic pathways and clathrinindependent mechanisms via lipid rafts (Grebe et al. 2003,
Geldner 2004, Aniento and Robinson 2005, Murphy et al.
2005, Samaj et al. 2005, Ovecka and Lichtscheidl 2006,
Verma et al. 2006).
In this study, we wanted to investigate constitutive
endocytosis in mature, non-elongating internodal cells of
characean algae which are considered to be close relatives of
higher plants (Graham and Kaneko 1991, Tanabe et al.
2005). We were especially interested in the role of the actin
cytoskeleton which was reported to be involved in plasma
membrane internalization in various types of cells including
those of higher plants (Engqvist-Goldstein and Drubin
2003, Baluska et al. 2004, Ayscough 2005, Ovecka et al.
2005, Voigt et al. 2005, Samaj et al. 2006b, Sun et al. 2006).
The characean internodes are well known for their rapid
*Corresponding author: E-mail, [email protected]; Fax, þ43-662-8044-619.
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FM internalization in characean algae
cytoplasmic streaming which depends on interaction of
myosin-coated organelles with parallel, subcortical actin
bundles (Grolig and Pierson 2000, Shimmen and Yokota
2004) and are a valuable experimental tool for investigating
the plant cytoskeleton (Foissner and Wasteneys 2000).
In our study, we applied FM1-43 and FM4-64 which
are widely used as fluorescent markers of endocytosis in
animal, fungal and plant cells (Betz et al. 1996, FischerParton et al. 2000, Bolte et al. 2004, Aniento and Robinson
2005, Samaj 2006). It has also been suggested that FM
internalization is due to the activity of energy-dependent
flippases (Fischer-Parton et al. 2000) or mechanosensitive
cation channels (Nishikawa and Sasaki 1996), and that the
differential distribution of FM dyes to organelle membranes
is due to a lipid sorting mechanism. An argument against
endocytosis-independent uptake is, however, that FM dyes
injected into growing pollen tubes and other cells became
homogeneously distributed and did not localize to specific
organelles (Parton et al. 2003, Van Gisbergen et al. 2007).
This observation and recent data obtained by fluorescence
resonance energy transfer (FRET) analysis (Johnson and
Griffing 2007) also point to FM uptake via endocytic
vesicles. In plant cells, the FM dyes insert into the plasma
membrane and gradually appear in compartments where
they partially co-localize with endosomal (Ueda et al. 2004),
pre-vacuolar (Tse et al. 2004), trans-Golgi (Dettmer et al.
2006) and vacuolar marker proteins (Kutsuna and
Hasezawa 2002). Therefore, irrespective of the uptake
mechanism, the FM dyes are at least suited for monitoring
the distribution and dynamics of endocytic organelles. We
also applied filipin which was used to follow steroldependent endocytosis in higher plant cells (Grebe et al.
2003, Ovecka and Lichtscheidl 2006). We reported previously at a meeting that FM dyes label distinct plasma
membrane areas in characean internodal cells and that
FM internalization cannot be inhibited by cytochalasin D
(CD; Foissner and Klima 2008). Here we prove that the
FM-stained plasma membrane areas are enriched in sterols
and that their distribution is independent of the cytoskeleton. Observations in cytoplasmic droplets, additional
inhibitor experiments and quantification of FM internalization in inhibitor-treated and control cells provide further
evidence that constitutive endocytosis, or at least FM
internalization and transport of FM fluorescent organelles
in characean internodes, is not only independent of an
intact actin (and microtubule) cytoskeleton but also
independent of actin polymerization.
Results
The staining results for FM1-43 and FM4-64 were
similar, and co-labeling with both dyes resulted in identical
images (Fig. 1A–C, see below). However, FM1-43 was
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preferred because of its brighter fluorescence and its green
emission which is easier to discriminate from the red
autofluorescence of chloroplasts. Furthermore, FM1-43
could be applied for several days without any obvious
harmful effect, whereas cells treated with the same
concentration of FM4-64 died after 1 d.
Heterogeneous labeling of the plasma membrane by FM dyes
and filipin
The plasma membrane of various cell types have been
reported to be homogeneously labeled by FM dyes (Samaj
et al. 2006a) and filipin (Grebe et al. 2003, Ovecka and
Lichtscheidl 2006). In internodal cells of Chara corallina,
the periphery of the cytoplasm was heterogeneously stained.
Bright patches became visible within 5 min of treatment
with FM dyes and alternated with weakly fluorescent
regions (Fig. 1A–D). Most of these patches were disc-like
(Fig. 1A–C), but elongate (Fig. 1D) or bowl-shaped
structures (not shown) were also observed. Optical crosssections through Z-series revealed that some of these
patches protruded slightly into the cell interior (Fig. 1E).
The FM-stained plasma membrane domains were
present along the whole cytoplasmic surface except for the
cross-walls between adjacent cells (Fig. 1F, G). They were
also largely absent from the neutral line which separates
regions of oppositely streaming endoplasm (Fig. 1H) and
from those regions where the chloroplast surface abutted
the plasma membrane (asterisk in Fig. 1A). The number
and (numerical) density of the FM-stained patches varied
greatly between cells. The highest densities were found in
internodes of the branchlets, with up to 200 disc-shaped
patches per 100 mm2 (Fig. 1A–C). There was also considerable variability in density of FM-stained patches along the
surface of individual internodes (not shown). In contrast to
these spatial variations, both the size and distribution of the
FM-stained patches were rather stable. Although we
investigated several dozens of cells over time periods up to
3 h, detachment, migration or gross shape changes of
patches were never observed.
The distribution of the FM patches was similar in
elongating (not shown) and in mature internodal cells and
therefore independent of the orientation of cortical actin
filaments or microtubules, which are transverse in elongating and random in mature cells (Wasteneys and Williamson
1987). There was also no correlation between the distribution of FM patches and that of the subcortical actin
bundles. For instance, FM patches were absent from the
neutral line which lacks subcortical bundles, but they were
also absent from the cross-walls where the subcortical actin
bundles are in close contact with the plasma membrane (see
above). Neither CD nor oryzalin affected the pattern of FM
fluorescence at the cell surface (Fig. 1I–K), suggesting that
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FM internalization in characean algae
Fig. 1 Heterogeneous labeling of the plasma membrane by FM dyes and filipin, and comparison with the distribution of cortical
organelles in internodes of Chara corallina. (A–C) Double staining of the plasma membrane with FM4-64 (A), FM1-43 (B) and merged
image (C). The asterisk in A indicates the position of an underlying chloroplast. (D) Spot-like and elongate, branched structures after
staining with FM1-43. (E) Optical cross-section through the plasma membrane (stack of four single sections). The FM-stained patches
alternate with weakly fluorescent areas and occasionally project into the cytoplasm (arrow). The plasma membrane appears up to 1 mm
thick due to an optical slice thickness of 550 nm (pinhole size 1.4 Airy units) and imperfect optical cross-sectioning. (F, G) Oblique view on
the cross-wall between a nodal (N) and internodal cell (I). Note the absence of plasma membrane patches (arrows in the fluorescence
image F and the corresponding bright field image G). The asterisk in G indicates a chloroplast. (H) Plasma membrane patches stained by
FM1-43 are less abundant at the neutral line (NL). (I–K) FM1-43-stained patches are stable and are not affected by inhibitors of the
cytoskeleton. Plasma membrane before addition of inhibitors (I), after 30 min in 5 mM cytochalasin C (CD; J) and after another 30 min in
5 mM CD and 10 mM oryzalin (K). (L) FM1-43-stained plasma membrane patches (green) are clearly different from cortical mitochondria
visualized by mitotracker orange (red). (M, N) Secretory vesicles that migrated towards the plasma membrane after local UV irradiation are
seen between and at the surface of cortical chloroplasts at the left side of the DIC image (M, arrow) and are not labeled in the
corresponding FM1-43 image (N, arrow indicates the same area as in M). (O, P) FM patches co-localize with sterol-rich plasma membrane
areas. The internodal cell was simultaneously labeled with FM1-43 (O) and filipin (P). All images were captured within 30 min after pulse
labeling. Bar in E is 4 mm (E), 8 mm (F, G, I–K, M, N), 10 mm (A–D, L, O, P) and 15 mm (H).
the morphology and distribution of the FM patches are
independent of the cytoskeleton.
The plasma membrane remained fluorescent for several
hours even when dyes were washed out after 10 min
incubation. The staining patterns of pulse-labeled cells
were therefore similar to those of cells permanently
immersed in dye-containing artificial fresh water (AFW),
and the patches occasionally remained visible until the day
after pulse labeling. FRAP (fluorescence recovery after
photobleaching) experiments should clarify whether the
FM internalization in characean algae
permanent labeling of patches was due to consistent
recycling of labeled plasma membrane, but laser intensities
sufficient for bleaching induced a wound response, i.e. the
rapid deposition of cell wall material and the deposition of a
new, evenly stained plasma membrane (to be described
elsewhere). Therefore, we can neither prove nor exclude this
possibility. The persistent staining of the plasma membrane
could also be due to the storage of FM dye within the cell
wall, as suggested for stomatal guard cells (Meckel et al.
2004).
In order to prove that the peripheral FM patches were
indeed localized to the plasma membrane and were not due
to staining of cortical organelles or heterogeneous labeling
of the cell wall, we performed a number of experiments. The
cortical cytoplasm of characean internodal cells is dominated by files of stationary chloroplasts but also harbors
other organelles, such as cortical endoplasmic reticulum,
mitochondria, secretory vesicles and peroxisomes. The
reticulate, 3,30 -dihexyloxacarbocyanine iodide [DiOC6(3)]stained cortical endoplasmic reticulum was clearly distinct
from the mostly punctate FM pattern (Supplementary
Fig. S1A). Co-labeling of cells with FM1-43 and mitotracker orange revealed considerable differences between
the distribution of the FM patches and that of mitochondria as long as the observation time did not exceed 2 h
(Fig. 1L). In pollen tubes, FM dyes are rapidly transferred
to secretory vesicles which accumulate at the growing tip
(see, for example, Hörmanseder et al. 2005). In Chara, the
accumulation of Golgi-derived secretory vesicles can be
induced by mechanical damage or local irradiation with
UV. These vesicles are large enough to be visualized in the
light microscope, have characteristic refractile properties
and dynamic behavior, and their involvement in wound wall
deposition has been documented by electron microscopy
and video-enhanced contrast microscopy (Foissner 1988,
Foissner et al. 1996). The comparison of the differential
interference contrast (DIC) with the FM fluorescence image
showed that these organelles were not stained and were
mostly much smaller than the FM patches (Fig. 1M, N).
Immunofluorescence of peroxisomes with an antibody
against catalase revealed roundish organelles with a
diameter of up to 2 mm. Their location and distribution
were clearly different from that of the FM patches (not
shown). Finally, no FM fluorescence was detected in cell
walls isolated from FM-stained cells (not shown).
Obviously, FM dyes accumulated (or fluoresced
stronger) in stable plasma membrane domains, which
differed from their surroundings by an altered composition
of structural components. Additional support for this view
came from the staining pattern obtained with filipin, a dye
supposed to be specific for cholesterol in animal cells and
sterol-like substances in plant cells (Grebe et al. 2003).
Filipin labeled punctate or elongate structures in the
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peripheral cytoplasm of live (Fig. 1P) and fixed cells
(Supplementary Fig. S1C) which had a similar size and
distribution to those of the FM patches. Double staining
with FM1-43 and filipin revealed that both dyes labeled the
same areas (Fig. 1O, P).
Taken together, these data strongly suggest that the
FM patches correspond to sterol-rich plasma membrane
domains.
Time course of FM internalization
In contrast to the rather stationary FM patches
described above, we sometimes observed considerable
dynamics in their periphery (3D environment) and between
them (Fig. 2A–C). Tiny structures appeared to detach from
or to fuse with the immobile FM patches, while others
appeared or disappeared from the focal plane, suggesting
movement from or into deeper regions of the cytoplasm.
These dynamic FM-stained structures were of diffractionlimited size and only occasionally seen because their
visualization requires optimal optical conditions, i.e. superior staining and a thin cell wall free of epiphytes. They may
represent the first detectable stages of plasma membrane
internalization in characean cells (compare Meckel et al.
2004). Large organelles which formed at and pinched off
the plasma membrane as described in root cells or in
protoplasts and guard cells after hyperosmotic treatment
(Baluska et al. 2002, Meckel et al. 2005) were not observed.
A few minutes after dye addition, FM-fluorescent spotlike structures of diffraction-limited size and larger,
vesicular organelles with a diameter of up to 500 nm were
also detected beneath the plasma membrane (Fig. 2D). The
FM-stained organelles were clearly different from cortical
mitochondria and performed oscillating, probably
Brownian motion in the clefts between the chloroplast
files (Fig. 2D–F). Some of them trembled back towards the
plasma membrane, moved along straight or curved tracks
between the stationary FM patches and then returned back
into deeper regions of the cortical cytoplasm (not shown).
In the endoplasm, FM-labeled organelles became visible
about 15 min after dye addition (Fig. 2G). Most of them
participated in cytoplasmic mass streaming which reached
up to 60 mm s–1 in our studies (Fig. 2H, right trajectory).
Those which were close to the subcortical actin bundles
travelled with variable and considerably slower velocity
(Fig. 2H, left trajectory; mean ¼ 11. 2 m 4.9 mm s–1; n ¼ 10).
Double labeling with mitotracker orange excluded the
possibility that the larger FM-stained organelles represent
mitochondria which have a similar dynamic behavior
(Fig. 2G; Kachar 1985, Foissner 2004). The FM-stained
endoplasmic organelles were also different from the tubular
inner endoplasmic reticulum (Supplementary Fig. S1B),
from peroxisomes and secretory vesicles (see above and
section on cytoplasmic droplets below).
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FM internalization in characean algae
Fig. 2 FM-stained organelles in internodal cells of Chara corallina. Time course of FM internalization (A–L) and co-localization with
acidotropic dyes (M–W). (A–C) Dynamics of FM1-43-labeled structures at the plasma membrane. Note dynamic extensions (thick arrows)
of the stable FM patches. Weakly fluorescent, minute structures (thin arrows) appear and disappear between the patches. The time series
was taken 5 min after pulse labeling; the time interval between images is 4 s. (D) At 10 min after pulse labeling, FM1-43-stained organelles
(green) are seen in the cortical cytoplasm underneath the plasma membrane, and are clearly different from mitochondria identified by
mitotracker orange (red). (E, F) FM1-43-stained organelles (arrows in E) between cortical chloroplasts perform trembling motions typical for
Brownian movement. The time interval between positions of organelles indicated by dots in F is 500 ms. (G) At 20 min after pulse labeling,
FM1-43-stained organelles (green) are seen in the endoplasm. Red-labeled organelles are mitochondria. (H) The FM-stained organelles
move slowly and with variable velocity along the subcortical actin bundles (left trajectory in H with 500 ms time interval). In deeper
regions of the endoplasm, FM-stained organelles participate in continuous mass streaming with a considerably faster velocity of about
60 mm s–1 (right trajectory in H with 500 ms time interval). (I–L) Optical sections through internodal cells about 30 min after pulse labeling
with FM1-43. (I, J) The membrane of a small vacuole, which is located at the periphery of the cytoplasm at the chloroplast-free neutral line
and protrudes into the large central vacuole (V), is stained with FM1-43 (white arrows in the fluorescence image I and in the DIC image J;
the cytoplasm is overexposed). The black arrow indicates the cell wall. (K, L) The tonoplast of the large central vacuole (V; arrow in the DIC
image L) is not labeled. The small arrow in the fluorescence image K points to an FM-stained organelle. A file of small chloroplasts typical
of elongating cells is visible in L; the black arrow indicates the cell wall. (M–S) Dual labeling with FM1-43 and neutral red. (M O) Cortical
organelles are stained by FM1-43 (M) or by neutral red (N). Two of them co-localize (arrows and merged image O). (P–S) The membrane of
small vacuoles (asterisks) is stained by FM1-43 (P) and neutral red (Q). Note also organelles at the surface of the vacuoles which are stained
by both dyes. R is the merged image, S is the DIC image. (T–U) Dual labeling with FM4-64 (T) and lysotracker yellow (U). Lysotracker
yellow accumulates in vacuoles (asterisks) the periphery of which is stained with FM or covered by FM-stained organelles. V is the merged
image; W is the DIC image; C ¼ chloroplast. Bar in F is 3 mm (A–C, I–W) and 5 mm (D–H).
Characean internodal cells contain a huge central
vacuole and numerous smaller vacuoles located at the
periphery of the streaming endoplasm (Fig 2I–L).
Occasionally, these vacuoles are also found in the stagnant
cytoplasm between the chloroplast files where the
imaging conditions are better than in deeper regions
(Fig. 2P–W). At 30–60 min after dye addition, the
membrane of some of these vacuoles became labeled by
FM internalization in characean algae
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Fig. 3 Cytoplasmic droplets prepared from FM-labeled and unstained internodal cells of Chara corallina. (A and B) Spot-like, fluorescent
organelles are present in the cytoplasm isolated from a cell which was stained with FM4-64 for 30 min (A, DIC image; B, fluorescence).
Some of them (arrows) associate with an actin bundle, which is coated with numerous, unlabeled secretory vesicles. (C, D) The higher
magnification shows that fluorescent organelles have a spherical or disk-like shape (left arrow) or an irregular outline (right arrows). The
membrane of the nucleus (N) is not stained (C, DIC image; D, FM4-64 image). (E, F) After 1 d staining, FM dyes also localize to secretory
vesicles (arrows) and other organelles. The membrane of the cytoplasmic droplet is of tonoplast origin and does not fluoresce (arrowheads
in the DIC image E and the FM1-43 image F). (G, H) A cytoplasmic droplet of an unstained cell squeezed into perfusion solution containing
FM4-64 does not internalize the dye (G, DIC; H. fluorescence image taken 15 min after preparation). The asterisk indicates an
autofluorescent chloroplast outside the droplet. Bar in H is 2.5 mm (C, D) and 5 mm (all other images).
FM dyes (Fig. 2I, J). The fluorescence was probably
delivered by punctate FM-labeled organelles which
attached to the surface (membrane) of these vacuoles and
were often unevenly distributed or appeared clumped
(compare Fig. 2P and T). Fusion processes, however,
could not be documented, suggesting that these are longlasting processes. The tonoplast of the central vacuole never
received FM label (Fig. 2K, L) even when cells were
incubated in the dye solution for 24 h (compare Fig. 3E and
F). After 2–3 h, FM dyes gradually appeared in all cortical
and endoplasmic mitochondria, in secretory vesicles and in
various other organelles, except the endoplasmic reticulum
and the tonoplast (see below).
These results suggest that FM dyes can be used for
monitoring endocytosis in Chara internodes as long as the
observation time does not exceed 2 h. In order to prove the
endosomal character of FM-fluorescent organelles, we
applied the acidotropic dyes neutral red and lysotracker
yellow. Neutral red stains the membrane of endosomal
compartments and accumulates in lytic vacuoles (Beilby
and Shepherd 1989, Guttenberger et al. 2000, Dubrovsky
et al 2006). In Chara internodal cells, neutral red labeled
small organelles in the cortex and in the endoplasm and
some of them co-localized with FM1-43-stained particles
(Fig. 2M–O). It also stained the membrane of the FMlabeled vacuoles (Fig. 2P–S). A faint pink color indicated
that neutral red was trapped inside these vacuoles, but the
concentration was not sufficient for documentation. We
also applied lysotracker yellow which is another suitable
probe for lytic compartments (Guttenberger 2000). It
accumulated in vacuoles with FM-stained membranes or
in vacuoles which were surrounded by FM-labeled organelles (Fig. 2T–W). Both neutral red and lysotracker yellow
became trapped in the large central vacuole although the
tonoplast was not labeled by FM (Fig. 2K) or by neutral red
(not shown).
In order to improve the visualization of endoplasmic
membranes and organelles, we used cytoplasmic droplets
squeezed out from internodal cells (Kamiya and Kuroda
1957, Jarosch 1961). They contain cortical and endoplasmic
organelles as well as dynamic actin filaments which
associate into bundles and rotating rings and which have
a strong affinity for secretory vesicles (Fig. 3A–H).
Observations on cytoplasmic droplets from FM-stained
cells confirmed the staining patterns described above. In
droplets prepared from internodes 30 min after pulse
labeling, some of the FM-stained organelles associated
with actin bundles but were clearly different from the
FM internalization in characean algae
majority of secretory vesicles visualized by DIC (Fig. 3A, B)
and from mitochondria identified with mitotracker orange
(not shown). The FM fluorescent structures had either a
punctate, roughly spherical or disc-like appearance, or an
irregular shape. They corresponded to roundish or lobed
organelles which were hardly recognizable with DIC
microscopy (Fig. 3C, D). No fluorescence was detected in
the membrane of the nucleus or that of the endoplasmic
reticulum. If cells were squeezed out 1 d after dye addition,
FM fluorescence also localized to secretory vesicles
(Fig. 3E, F), to mitochondria and to the nuclear membrane
(not shown). The membrane of the cytoplasmic droplets is
of tonoplast origin (Sakano and Tazawa 1986) and did not
fluoresce (Fig. 3E, F), consistent with the observations in
intact cells. Neither the tonoplast-derived membrane nor
the content of droplets became labeled when the cytoplasm
of unstained cells was squeezed into perfusion solution
containing FM dyes (Fig. 3G, H).
These observations indicate that FM1-43 and FM4-64
are internalized via the plasma membrane and localize to
organelles putatively involved in endocytosis as long as the
observation time does not exceed 2 h. In contrast, filipinlabeled organelles were not observed in intact cells or in
cytoplasmic droplets prepared from filipin-stained internodes, but this is probably due to difficult imaging
conditions (Grebe et al. 2003).
Constitutive FM internalization is not affected by inhibitors
of the cytoskeleton
In most fungal, animal and plant cells investigated so
far endocytosis has been reported to depend on actin–
myosin interaction and/or actin polymerization (see
Introduction). In this study, we applied CD, latrunculin B
(LatB) and 2,3-butanedione monoxime (BDM) in order to
investigate the role of the actin cytoskeleton in FM
internalization in characean algae. We also used oryzalin
to evaluate a possible contribution of microtubules.
CD reversibly reorganizes the cortical actin cytoskeleton of characean internodes into actin rods and rapidly
inhibits cytoplasmic streaming without disassembling the
subcortical actin bundles (Williamson 1978, Collings et al.
1995; Supplementary Fig. S1D, E). We first treated
internodal cells with 5 mM CD for 30 min and subsequently
with a solution containing CD and FM1-43. After another
30 min treatment, cells were washed in AFW, fixed,
mounted and investigated in the confocal laser scanning
microsope (CLSM). FM patches at the plasma membrane
were well preserved, and FM-stained organelles were
present in the cortex and in the endoplasm. The proportion
of fluorescent organelles in the endoplasm of CD-treated
internodes was slightly lower than in control cells, but
means were not significantly different (Fig. 4). In order to
exclude the possibility that fixation caused redistribution of
Fluorescence in endoplasm (vol%)
1514
15
10
5
0
+ CD
− CD
Fixed cells
− CD
+ CD
Live cells
Fig. 4 Effect of cytochalasin D (CD) on FM internalization in
internodal cells of Chara corallina. Cells were treated with 5 mM
CD for 30 min before addition of FM1-43. After another 30 min,
CD-treated and control cells were fixed, mounted and studied
in the CLSM or investigated live. Volume of FM-fluorescent
organelles is given as a percentage of total volume of endoplasm.
Means (SD) are based on Z-series of 16 fixed and four live cells,
respectively, and are not significantly different. For further details,
see text.
plasma membrane-bound FM to cytoplasmic organelles, we
also investigated internodes grown under low light conditions. These cells had thin and weakly autofluorescent
chloroplasts, a prerequisite for live imaging of FM-labeled
organelles in the endoplasm. Internodes treated with 5 mM
CD for 30 min and subsequently with CD and FM1-43
contained abundant fluorescent organelles after 30 min. In
the fast streaming endoplasm of untreated internodes,
individual organelles could not be resolved with conventional CLSM. Therefore, cytoplasmic streaming in control
cells was also arrested with CD but only immediately prior
to observation. Again, we found no significant effect of the
inhibitor on FM internalization in internodal cells (Fig. 4).
Latrunculins bind actin monomers and inhibit their
polymerization into filaments (Spector et al. 1999). At a
concentration of 50 mM, LatB disassembles cortical actin
filaments of Chara internodal cells although subcortical
bundles remain intact and support cytoplasmic streaming
(Supplementary Fig. S1F; compare Foissner and Wasteneys
2007). In combination with CD, latrunculins are most
effective in arresting cytoplasmic streaming and depolymerizing cortical actin in characean internodes (Foissner and
Wasteneys 2007). However, neither LatB alone (Fig. 5A, B)
FM internalization in characean algae
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Fig. 5 Inhibitor effects on FM internalization in internodal cells of Chara corallina. (A, B) Cytoplasmic droplets prepared from internodal
cells which were treated with 50 nM latrunculin B (LatB) for 30 min and stained with LatB/FM1-43 for 20 min contain numerous FMfluorescent organelles. The asterisk indicates an autofluorescent chloroplast. (C, D) Effect of combined treatment with CD and LatB. The
density of fluorescent organelles in the endoplasm of internodes which were treated with 5 mM CD and 50 mM Lat A before and during
staining with FM1-43 (30 min each, C) is similar to that observed in control cells (D). (E) FM1-43-fluorescent organelles are present in
internodal cells treated with 100 mM 2,3-butanedione monoxime (BDM) for 30 min before and during dye addition. (F–H) FM1-43
internalization is completely inhibited by treatment with 1 mM KCN. FM-stained plasma membrane patches are seen at the left and right
side of the image but the endoplasm in the middle of the image is devoid of fluorescent organelles (F). Cytoplasmic droplets from cells
treated with KCN 30 min before and 30 min after dye addition show no FM1-43 fluorescence (G, DIC; H, fluorescence image). The asterisk
indicates an autofluorescent chloroplast outside a droplet. Bar in H is 5 m (A–D, G, H) and 20 m (E, F).
nor the combined treatment with LatB and CD (Fig. 5C, D)
prevented the appearance of FM-stained organelles in intact
internodes and in cytoplasmic droplets prepared from these
cells.
BDM is used as a general myosin inhibitor in spite of
many contradictory data (e.g. McCurdy 1999). In Chara,
concentrations of about 100 mM are required for inhibition
of cytoplasmic streaming and organelle movement, and this
effect is not reversible (McCurdy 1999, Funaki et al. 2004).
Even this concentration did not prevent FM internalization
in internodal cells; it caused, however, clumping of FMfluorescent organelles (Fig. 5E)
Oryzalin at a concentration of 10 mM disassembles
microtubules of characean internodal cells within 30 min
(Supplementary Fig. S1G, H; compare Wasteneys and
Williamson 1987). Experiments with oryzalin-treated
cells ( CD) showed that FM internalization was also
independent of microtubules (Supplementary Fig. S1I).
FM dyes are assumed to enter the cytoplasm via
energy-dependent vesicle-mediated endocytosis (Betz et al.
1996). The unexpected insensitivity of FM internalization
in characean cells against cytoskeletal inhibitors could be
due to passive uptake (diffusion) independent of cellular
metabolism. We therefore applied KCN at a concentration
sufficient to arrest ATP-dependent cytoplasmic streaming
and found that this treatment completely inhibited the
appearance of FM fluorescent organelles in the cytoplasm
(Fig. 5F) and in droplets obtained from such cells (Fig. 5G,
H). Staining of plasma membrane patches was not affected
(Fig. 5F). The same results were obtained when KCN
solutions were supplemented by dimethylsulfoxide (DMSO)
up to a concentration of 1%, i.e. the plasma membrane
became stained but internalization of FM dyes was not
observed.
These data suggest that FM internalization in
unwounded cells of characean algae is active but independent of the cytoskeleton and not mediated by up to 1%
DMSO which was used as solvent for dyes and inhibitors in
this study. They do, however, not prove that internalization
occurs via vesicular transport.
Discussion
Sterol-rich plasma membrane domains in Chara—clusters of
lipid rafts?
To our knowledge, characean internodes are the only
plant cells in which the plasma membrane is heterogeneously labeled by FM1-43 and FM4-64. Both dyes
accumulated or fluoresced more brightly in stable, mostly
circular patches with a diameter of up to 1 mm. In this study,
we prove that these plasma membrane patches are
also labeled by filipin, which is specific for cholesterol and
1516
FM internalization in characean algae
sterol-like substances in plants (Robinson and Karnovsky
1980, Grebe et al. 2003), suggesting that the FM-labeled
membrane domains are enriched in sterols. In yeast, filipin
used at high concentrations was shown to cause clustering
of sterols into plasma membrane domains which also colocalized with FM4-64 for unknown reasons (ValdezTaubas and Pelham 2003; see also Robinson and
Karnovsky 1980, Miller 1984 for filipin-induced artifacts).
In Chara, FM dyes labeled plasma membrane patches in the
absence of filipin and, on the other hand, plasma membrane
patches were stained by filipin after fixation of cells, which
excludes the possibility of a staining artifact.
Sterol-like substances that could be stained by filipin
(Grebe et al. 2003) have been isolated from various
characean species (Patterson et al. 1991). Such lipids are
enriched in detergent-resistant plasma membrane microdomains, called lipid rafts (Simons and Van Meer 1988).
Typical lipid rafts are small (in the nm range), quite
unstable (Pralle et al. 2000, Pike 2006), and can therefore
not be resolved in a laser scanning microscope (Grebe et al.
2003). In the Characeae, lipid rafts appear to be clustered
into larger, stable aggregates as reported from
Saccharomyces and other fungi (Malinska et al. 2004,
Alvarez et al. 2007) and from plant cells during pathogen
attack (Bhat and Panstruga 2005). Our observations thus
prove that sterol-rich domains are indeed present in intact
plant cells, consistent with biochemical data (Mongrand
et al. 2004, Shahollari et al. 2004, Borner et al. 2005, see also
Kirkham and Parton 2005, Martin et al. 2005).
Possible function of the plasma membrane domains
The fact that the plasma membrane domains of
characean internodal cells are labeled by endocytic markers
as well as reports about the involvement of lipid rafts in
sterol-dependent endocytosis suggests a participation in
plasma membrane recycling (Nichols and LippincottSchwartz 2001, Cheng et al. 2006). Indeed, minute FMstained particles have been observed at the periphery of the
FM patches, suggesting detachment or fusion of tiny
organelles, but these observations may simply reflect the
overlap of signals from closely associated, separate organelles or structures. Filipin can be used to follow
constitutive recycling of sterols in plant cells (Grebe et al.
2003, Geldner 2004, Murphy et al. 2005, Ovecka and
Lichtscheidl 2006). In our study, filipin labeled the plasma
membrane domains, but uptake of filipin into the cytoplasm
was never observed. Whether this is due to insufficient
imaging conditions, to the presence of quenching substances
in the cytoplasm or to sterol recycling which excludes the
fluorescent marker remains to be investigated.
Lipid rafts and sterol-rich plasma membrane domains
have also been implicated in actin cytoskeleton organization (Bhat and Panstruga 2005, Alvarez et al. 2007).
The comparison of the actin cytoarchitecture with the
distribution of the FM patches as well as the inhibitor
experiments clearly argue against such a possibility in
characean internodal cells.
It is possible that the FM- and filipin-stained patches
in Chara internodal cells correspond to complex
plasma membrane invaginations, known as charasomes.
Charasomes also show a non-uniform distribution, are
absent from cross-walls and the neutral line and may
protrude deeply into the cytoplasm (Barton 1965, Crawley
1965, Franceschi and Lucas 1980, Lucas and Franceschi
1981). They are probably involved in enhanced transport of
ions or inorganic carbon (Franceschi and Lucas 1982, Price
and Whitecross 1983, Price et al. 1985, Bisson et al. 1991,
Chau et al. 1994). Charasomes develop from coated pits,
and coated vesicles appear to pinch off their surface (Lucas
and Franceschi 1981), which corroborates our findings
about a possible detachment of FM-stained organelles from
these plasma membrane domains. However, FM patches,
although at lower densities, are also present in internodes of
the genus Nitella, which lack charasomes (own unpublished
observations). Therefore, further research is needed to
clarify the identity of FM-labeled, sterol-enriched plasma
membrane domains in characean algae. Whatever their
function is, the possibility of visualizing these domains in
the living state by the rather non-toxic FM dyes offers new
perspectives for in vivo observation of plasma membrane
organization and metabolism.
Actin-independent FM internalization and transport of
putative endosomes
There is a large body of evidence from yeast and
mammalian cells implicating the actin cytoskeleton in
several endocytic pathways (see reviews by EngqvistGoldstein and Drubin 2003, Ayscough 2005). In plant
cells as well, both actin nucleation and myosin motor
activity along actin filaments have been described to be
required for plasma membrane internalization (for references, see Baluska et al. 2002, Grebe et al. 2003, Meckel
et al. 2004, Ovecka et al. 2005, Voigt et al. 2005, Ovecka and
Lichtscheidl 2006, Samaj et al. 2006b). In Chara internodal
cells, neither CD, BDM nor LatB arrested uptake of FM
dyes and transport of FM-stained organelles, suggesting
that both processes are largely independent of an intact
actin cytoskeleton, independent of acto-myosin interaction
and independent of actin polymerization (actin comets).
Experiments with oryzalin excluded a possible participation
of microtubules.
We cannot fully exclude the possibility that the
insensitivity of FM uptake against cytoskeleton inhibitors
is due to non-vesicular internalization of the dyes. Our
KCN experiments prove that FM1-43 and FM4-64 were
actively internalized and did not enter the cytoplasm
FM internalization in characean algae
by diffusion. However, FM uptake in characean internodes
could be mediated by the activity of energy-dependent
flippases (Fischer-Parton et al. 2000; see also Introduction).
Irrespective of the mechanism for FM internalization, the
inhibitor experiments definitely show that transport of FMstained organelles, putative endosomes, from the cortex to
the endoplasm requires neither intact actin filaments nor
actin polymerization (actin comets). These findings are
consistent with the special cytoarchitecture of the nonelongating, mature internodes where cortical and subcortical actin cytoskeleton are widely separated from each
other and rarely linked by actin filaments which traverse
the chloroplast layer (Foissner and Wasteneys 2000).
Consistent with actin-independent motility, FM-stained
organelles performed trembling movements within the
clefts between the chloroplast files even in non-treated
cells, suggesting that Brownian movement is sufficient to
account for the transport of putative endosomes from the
cortex towards the endoplasm. In the subcortex, however,
fluorescent organelles migrated slowly and with variable
velocity along tracks that corresponded to the orientation of
subcortical actin bundles. These observations indicate that
at least some of the FM-stained organelles carry myosin at
their surface and are able to move actively along actin
filaments if possible and required (compare Kachar 1985,
Foissner et al. 1996 for a similar dynamic behavior of other
organelles).
Actin-independent plasma membrane internalization
and transport of FM-stained organelles may be regarded
as an adaptation to the special cytoarchitecture of
characean internodes. However, there exist several reports
about actin-independent endocytosis and transport of
endosomes in yeast and mammalian cells (Fujimoto et al.
2000, Gachet and Hyams 2005, Boucrot et al. 2006). The
current evidence in support of a role for the actin
cytoskeleton in plant endocytosis is circumstantial and is
mainly based on drug studies (Samaj et al. 2006b).
Endocytosis is usually coupled to exocytosis in order to
maintain plasma membrane tension and the surface area of
the cell (Murphy et al. 2005, Samaj et al. 2005). In plant
cells, transport of exocytotic vesicles towards the plasma
membrane is usually actin based. The frequently observed
arrest of endocytosis by inhibitors of the actin cytoskeleton
may therefore be a secondary effect of disturbed exocytosis.
The results of our study and the discovery that in maize root
cells PIN proteins may be retrieved from the plasma
membrane via a non-actin dependent process (Boutte
et al. 2006) suggest that actin-independent plasma membrane internalization is more widespread than thought.
However, we cannot exclude the possibility that FM
internalization in Chara (and other cells) is not due to
endocytosis but to the activity of flippases and ion channels
(see Introduction).
1517
FM dyes as endocytic markers in characean algae
In characean algae, FM dyes stained the plasma
membrane, cytoplasmic organelles and vacuoles with
kinetics similar to those reported from various plant and
fungal cells (Samaj et al. 2006a). Double staining with other
markers and the comparison with electron microscopic
images makes it likely that the FM-labeled organelles
correspond to various types of endosomes (see below).
Preliminary data indicate further that FM-stained organelles can be labeled with antibodies against various
endocytic markers (own unpublished data). We found that
FM1-43 and FM4-64 are both suited for studying
endocytosis, or at least the distribution of endocytic
organelles in characean algae (compare Ovecka et al.
2005). After pulse labeling, FM dyes gradually disappear
from the plasma membrane of most plant cells and are
transported to the tonoplast, where the majority of the dye
molecules remain (Parton et al. 2001, Kutsuna and
Hasezawa 2002, Ovecka et al. 2005). In the characean
algae, the plasma membrane remained fluorescent up to
24 h after pulse labeling just as in stomatal guard cells
(Meckel et al. 2004). So far it is not clear whether this is due
to rapid recycling of labeled plasma membrane or to dye
storage in the cell wall.
FM dyes are widely accepted as markers for non-sterol,
clathrin-mediated endocytosis, although FM internalization
via coated vesicles has not yet been demonstrated due to
their small size (Holstein 2002, Bolte et al. 2004, Meckel
et al. 2004). On electron micrographs of characean algae,
coated pits and coated vesicles are distributed over the
plasma membrane of unwounded cells (Pickett-Heaps 1967,
Lucas and Franceschi 1981, Pesacreta and Lucas 1984,
McLean and Juniper 1986, own unpublished results) and
probably contribute to constitutive endocytosis. The
identity of the cortical FM-stained organelles, which are
obviously larger than coated vesicles, remains to be
established.
Both clathrin- and lipid raft-mediated pathways in
plants are reported to meet at the partially coated reticulum,
the equivalent of early endosomes in other cell types
(Geldner 2004, Murphy et al. 2005, Samaj et al. 2005). In
mature, unwounded characean internodal cells, the partially
coated reticulum and organelles involved in late stages of
endocytosis, i.e. multivesicular bodies and Golgi bodies, are
located within the streaming endoplasm, up to several
micrometers away from the plasma membrane where
internalization occurs (Pickett-Heaps 1967, Pesacreta and
Lucas 1984, Foissner 1988). They may correspond to the
lobed and roundish FM-fluorescent organelles described
from cytoplasmic droplets.
Characean internodal cells contain various vacuoles
which differ in size, shape and content (compare Neuhaus
and Paris 2006 for vacuoles in higher plants). This diversity
1518
FM internalization in characean algae
was also reflected by the staining results. The membrane of
small vacuoles located at the inner periphery of the
endoplasm and occasionally between the chloroplast files
became labeled with FM dyes 30–60 min after dye addition.
These vacuoles sequestered the acidotropic dyes neutral red
and lysotracker yellow, and probably have lytic functions.
Our staining results indicate that the big central vacuole is
likewise an acidic compartment, but its membrane, the
tonoplast, was never labeled by FM dyes, suggesting that
this huge compartment has mainly storage function.
Consistent with this observation, Schulte et al. (1994)
found that the central vacuole of characean internodes
accumulates huge amounts of sugar.
Materials and Methods
Plant material and culture conditions
Shoots of C. corallina Klein ex Willd., em. R.D.W. were
grown in a substrate of soil, peat and sand in 10–50 liter aquaria
filled with distilled water. The temperature was about 208C, and
fluorescent lamps provided a 16/8 h light/dark cycle. Internodal
cells of the main axis or the branchlets were harvested 1 d prior to
experiments, trimmed of neighboring internodal cells and left
overnight in AFW (10–3 M NaCl, 10–4 M KCl, 10–4 M CaCl2). In
this study we used mainly mature, non-elongating internodes.
Younger, elongating cells with small and weakly fluorescent
chloroplasts were preferred for the study of endoplasmic organelles
and vacuolar membranes.
In order to improve the visualization of FM-stained
membranes and organelles, we also prepared cytoplasmic droplets
as described in Kamiya and Kuroda (1957) and Jarosch (1961).
Long internodal cells were blotted dry, and cut open with fine
scissors after loss of turgor. The cell sap and the cytoplasm were
extruded with forceps, collected on a slide and covered by a
coverslip. For some experiments, the cytoplasm was squeezed into
isotonic perfusion solution containing 200 mM sucrose, 70 mM
KCl, 4.5 mM MgCl2, 5 mM EGTA, 1.48 mM CaCl2, 10 mM
piperazine-N,N0 -bis(2-ethanesulfonic acid) (PIPES; pH 7) and
10 mM FM1-43 or FM4-64 (see below).
In vivo staining and inhibitor treatments
FM4-64 [N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino)phenyl)hexatrienyl)-pyridinium dibromide] and the fixable
analog of FM1-43 [N-(3-triethylammoniumpropyl)-4-(4-(dibutylamino)styryl)pyridinium dibromide], which retains its fluorescence
after fixation with glutaraldehyde, were purchased from Invitrogen
(Carlsbad, CA, USA) and used at 10 mM diluted from 10 mM stock
solutions in DMSO or from 500 mM stock solutions in distilled
water. The stock solution of filipin III (Sigma, St Louis, MO,
USA; 25 mg ml–1) was prepared in DMSO. Aliquots were frozen
at 708C, thawed immediately before use and diluted 1: 4,000 in
AFW for labeling live cells or in phosphate-buffered saline (PBS:
140 mM NaCl, 3 mM KCl, 10 mM phosphate buffer, pH 7.4,
0.12% NaN3) for labeling cells fixed in 1% (v/v) glutaraldehyde/
PBS. Lysotracker yellow HCK-123 (Invitrogen) was used at 10 mM
diluted from a 1 mM stock solution in DMSO, and neutral red
(Invitrogen) was used at 10–4% diluted from a 10–1% stock
solution in distilled water. Cells were loaded with lysotracker
yellow and neutral red for at least 2 h before addition of FM dyes.
Mitochondria were stained with 1 mM mitotracker orange
(Invitrogen; 1 mM stock solution in DMSO) and the endoplasmic
reticulum was labeled with 1 mM DiOC6(3) (Invitrogen, 1 mM
stock solution in DMSO).
Cells were exposed to 5 mM CD (Sigma; stock solution 10 mM
in DMSO), 50 mM LatB (Calbiochem, San Diego, CA, USA; stock
solution 10 mM in DMSO), 50 mM BDM (Sigma), 10 mM oryzalin
(Riedel-de Haen, Seelze, Germany; stock solution 10 mM in
acetone) and 1 mM KCN (Sigma).
All dyes and inhibitors were diluted with AFW which has a
pH of about 6. The pH of the AFW used for dissolving neutral red
and lysotracker yellow was adjusted to pH 8 with sodium
bicarbonate in order to achieve loading into acidic compartments
(Dubrovsky et al. 2006). We used internodal cells of the same
culture and of similar age as controls and treated them with the
corresponding amounts of solvent, which never exceeded 1%
DMSO or 0.1% acetone, respectively. These concentrations had
no visible effect on cytoplasmic streaming or internalization of
FM dyes.
Actin filaments and microtubules were visualized by perfusion of cells with fluorescent phalloidin (Alexa 488 phalloidin;
Invitrogen) and fluorescent taxol (Flutax-2; Calbiochem) as
described in Foissner (2004).
Confocal scanning microscopy
The CLSMs used in this study were a Zeiss LSM 510 and a
Zeiss LSM Meta coupled to Zeiss Axiovert inverted microscopes
(Jena, Germany). Images produced by the software were further
processed with Adobe Photoshop. Organelle dynamics were
studied by analyzing time series taken at minimum laser intensity
and pixel time in order to avoid photobleaching and stress
response.
For co-localization of FM dyes, FM1-43 was excited with the
488 nm argon laser line at an intensity insufficient to excite FM464, and the resulting emission wavelengths were collected between
505 and 530 nm. FM4-64 was excited with the 543 nm helium–neon
laser line, and emission wavelengths were recorded between 560
and 615 nm. The same settings were used for the dual labeling with
FM1-43 and mitotracker orange and for the dual labeling with
FM1-43 and neutral red. Lysotracker yellow and FM4-64 were
excited with 488 and 543 nm, respectively, and their emission
wavelengths were collected between 505 and 550 nm and between
560 and 615 nm. Filipin was excited with 351 nm, and the emission
was collected between 470 and 490 nm.
All images presented in this study are single sections. Images
of cells are positioned with vertical sides parallel to the long axes of
the internodes.
Quantification of FM internalization
The effect of CD on FM internalization was quantified in
living and in fixed cells. Internodes were treated with 5 mM CD
30 min before and 30 min after addition of FM1-43, whereas
control cells were incubated in FM1-43 alone for 30 min. In order
to obtain comparable images of the stagnant endoplasm,
cytoplasmic streaming in control cells was also arrested with
5 mM CD but only immediately prior to observation in the CLSM.
The fluorescence of endoplasmic organelles in intact cells is
attenuated by the cortical chloroplasts. Therefore, we also
quantified FM internalization in fixed cells. Internodes were
treated with 5 mM CD 30 min before and 30 min after addition of
FM1-43 as described above. CD-treated cells were fixed for 30 min
FM internalization in characean algae
in a solution containing 1% glutaraldehyde and 5 mM CD
dissolved in PBS. Control cells were fixed in the absence of CD.
Fixed cells were washed in PBS and cut into cylinders which were
split longitudinally. Cell fragments were mounted in PBS/glycerol
(1:1) with the cytoplasmic side facing the coverslip so that the
endoplasm could be studied without the disturbing chloroplast
autofluorescence. Endoplasmic volume fractions occupied by
FM1-43-fluorescent organelles (volume of fluorescent organelles
as a percentage of total cytoplasmic volume) were calculated from
Z-series of at least four cells using the Zeiss 3D-software.
Differences between means were analysed by t-test and considered
to be significant if P 0.01 (Sachs 1984).
Supplementary material
Supplementary material are available at PCP Online.
Acknowledgments
We are grateful to Richard Trelease (Arizona State
University), to Ted Farmer (University of Lausanne) and to
Takashi Ueda (University of Tokyo) for antibodies. We thank
Beatrice Satiat-Jeunemaitre (CNRS, Gif-sur-Yvette) and Markus
Grebe (Umea Plant Science Centre) for stimulating discussions.
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(Received July 21, 2008; Accepted August 14, 2008)