Download Hyperosmotic stress-induced actin filament reorganization in leaf

Journal of Experimental Botany, Vol. 53, No. 375, pp. 1699±1710, August 2002
DOI: 10.1093/jxb/erf018
Hyperosmotic stress-induced actin ®lament reorganization
in leaf cells of Chlorophyton comosum
G. Komis, P. Apostolakos and B. Galatis1
Faculty of Biology, Department of Botany, University of Athens, Athens 157 84, Greece
Received 3 January 2002; Accepted 4 April 2002
Abstract
Actin ®lament (AF) organization was studied during
the plasmolytic cycle in leaf cells of Chlorophyton
comosum Thunb. In most cells the hyperosmotic
treatment induced convex or concave plasmolysis
and intense reorganization of the AF cytoskeleton.
Thin cortical AFs disappeared and numerous cortical, subcortical and endoplasmic AFs arranged in
thick and well-organized bundles were formed.
Plasmolysed cells displayed a signi®cant increase
in the overall AF content compared with the control cells. Cortical AF bundles were preferentially
localized in the shrunken protoplast areas, lining
the detached plasmalemma regions. The endoplasmic AF bundles were mainly found in the perinuclear cytoplasm and on the tonoplast surface. AFs
also traversed some of the Hechtian strands. AF
disorganization after cytochalasin B (CB) treatment
induced dramatic changes in the pattern of plasmolysis, which lasted for a longer time and led to
a greater decrease of the protoplast volume compared to the untreated cells. In many of the above
cells the protoplasts assumed an `amoeboid' form
and were often subdivided into sub-protoplasts.
Soon after the removal of the plasmolytic solution
both CB-treated and untreated cells were deplasmolysed, while the AF cytoskeleton gradually reassumed the organization observed in the control
cells. The ®ndings of this study revealed for the
®rst time in angiosperm cells that plasmolysis triggers an extensive reorganization of the AF cytoskeleton, which is involved in the regulation of
protoplast shape and volume. The probable mechanism(s) leading to AF reorganization as well as
the function(s) of the atypical AF arrays in plasmolysed cells are discussed.
1
Key words: Actin ®laments, Chlorophyton comosum,
hyperosmotic stress, leaf cells, plasmolytic cycle.
Introduction
During plasmolysis of plant cells the plasmalemma is
detached from the cell wall and the protoplast volume is
signi®cantly reduced, while the intracellular architecture
experiences intense mechanical perturbations due to
compaction (Oparka, 1994). In a previous study it was
demonstrated that in plasmolysed dividing leaf cells of
Chlorophyton comosum all microtubule arrays were disintegrated and free tubulin was incorporated into macrotubules and tubulin paracrystals. These atypical tubulin
polymers are elongated, straight and rigid structures,
which often appeared interconnected into complex networks (Komis et al., 2001). It has been suggested that those
may offer mechanical support to the protoplast to resist
forces exerted on it during plasmolysis (Komis et al.,
2001).
In the present study the effects of the hyperosmotic
stress imposed by mannitol solutions on the organization of the actin ®lament (AF) cytoskeleton in the
leaf cells of C. comosum were investigated. The
disturbance of the AF organization in plasmolysed
cells was expected, considering that, in plant cells, the
osmotic stress induces (a) changes in cytosolic Ca2+
concentration (Busch, 1995; Knight et al., 1997, 1998;
Cessna et al., 1998; Brownlee et al., 1999; Knight
2000); (b) activation of a mitogen-activated protein
kinase cascade (Hirt, 2000); and (c) rapid synthesis of
polyphosphoinositides, which are part of the inositol
signalling system (Munnik et al., 2000; Meijer et al.,
2001). All these phenomena are immediately related to
the mechanism(s) regulating AF organization (Janmey,
1994; Richelme et al., 2000; Sullivan et al., 2000).
To whom correspondence should be addressed. Fax: +30 1 7274702. E-mail: [email protected]
ã Society for Experimental Biology 2002
1700 Komis et al.
As far as is known the effect(s) of the hyperosmotic
stress on AF organization in angiosperms have been
studied only in epidermal cells of Allium cepa (LangPauluzzi and Gunning, 2000) and Tradescantia virginiana
(Cleary, 2001; see also Cleary cited in Gunning and Steer,
1996). In the plasmolysed epidermal cells of Allium cepa
the cortical AFs were disorganized, while the other AF
arrays were not affected detectably. Cleary (2001) studied
the effect of plasmolysis mainly on asymmetrical divisions
involved in stomata formation of Tradescantia virginiana.
Moreover, AF rearrangement is involved in cell volume
regulation in different eukaryotic cell systems that are
often exposed to anisosmotic conditions. The AF cytoskeleton respond to osmotic stress in budding yeast
(Chowdhury et al., 1992; Slaninova et al., 2000; Ooms
et al., 2000), ®lamentus fungi (Bachewich and Heath,
1997), Dictyostelium discoideum (Aizawa et al., 1999;
Zischka et al., 1999) and many different mammalian cell
lines (Rizoli et al., 2000; Kapus et al., 2000). In particular,
the role of AFs in many aspects of cell volume regulation
in animal cells is well documented (Henson, 1999;
Papakonstanti et al., 2000). Considering the above information, the probable involvement of AFs in volume
regulation of the plasmolysed protoplasts of the leaf cells
of C. comosum was also examined in the present work.
This phenomenon has not been investigated so far.
Materials and methods
Plasmolysis
Small leaf segments (approximately 2 mm in length) of C. comosum
Thunb., corresponding to the leaf base, were excised and immersed
in aqueous solutions of 0.3, 0.5 and 1 M mannitol for 30 min and
were subsequently processed for AF localization. For examination of
living material, the thinner leaf segments were chosen due to their
transparency. Those were immersed into a drop of plasmolytic
solution on a microscope slide, rapidly covered under a clean
coverslip and monitored using the DIC optics of a Zeiss Axioplan
microscope. In these specimens epidermal as well as subepidermal
cells can be examined. Photomicrographs were taken on T-MAX
400 ®lm overrated at 1600 ASA.
Cytochalasin-B treatment
Leaf segments were incubated in an aqueous solution of 100 mM
cytochalasin B (CB; Sigma) for 30 min prior to plasmolysis.
Subsequently, they were plasmolysed with 1 M aqueous mannitol
solution further supplemented with 100 mM CB. Some of the leaf
pieces were processed for AF labelling whereas some others were
directly observed by DIC optics and photographed as described
above. Additionally, after plasmolysis some leaf pieces were brie¯y
®xed in 1% OsO4 in 1 M mannitol to allow long-term observations
on protoplast morphology.
Deplasmolysis
Following plasmolysis in 1 M mannitol, both CB-treated and
untreated leaf segments were returned to distilled water.
Subsequently, they were either processed for AF localization or
were monitored as previously described. For the observation of
living material, leaf segments were ®rst allowed to adhere on acid-
washed, poly-L-lysine coated coverslips. Subsequently, they were
placed on top of a drop of plasmolytic solution and were monitored
for a period of 30 min. Afterwards, plain distilled water was allowed
to diffuse from one edge of the coverslip, while the excess of liquid
was removed from the opposite site by the aid of ®lter paper. The
immobilization of leaf segments prevented their dislocation during
liquid exchange, thus allowing the observation of the same cells
during the plasmolytic cycle. To test the viability of the CB-treated
or untreated cells after a plasmolytic cycle, some of the leaf pieces
were re-exposed to hyperosmotic treatment of the same magnitude.
AF localization
For staining of AFs in control and plasmolysed leaf cells, a
¯uorophore-conjugated phalloidin-staining regime was employed.
The staining protocols used were based on previously published ones
with extensive modi®cations and thus will be described in detail. All
chemicals were purchased from Sigma unless stated otherwise.
Control and treated leaf segments were ®xed for 45 min in
formaldehyde freshly prepared from hydrolysis of 4% w/v
paraformaldehyde in either PEMD (50 mM K-PIPES, 5 mM
EGTA, 5 mM MgSO4, 5% v/v DMSO) pH 6.8 or phosphatebuffered saline (PBS) pH 7.4 both supplemented with 1 M glycerol
and 5 IUs of AlexaFluor 568-conjugated phalloidin (Molecular
Probes) per ml of ®xative. Alternatively, prior to ®xation, leaf
segments were pretreated with the bifunctional protein crosslinker
m-maleimidobenzol-N-hydroxysuccinimide ester (MBS; 500 mM in
PEMD plus 0.05% v/v Triton X-100) for 30 min (Panteris et al.,
1992, and references therein). For control experiments, leaf pieces
were immersed in distilled water for 30 min before AF localization.
Afterwards, specimens were thoroughly washed in PEMD for 30
min and they where then subjected to enzymatic digestion of the cell
wall material using a cocktail of 1% cellulase Onozuka R-10, 1%
macerozyme (both from Yakult Honsha) and 3% pectinase (Fluka)
in PEMD, pH 6.8 for 2 h. Leaf segments were then washed in PEMD
and were subsequently forced through a wide-bore Pasteur pipette in
order to release cells. The resulting cell suspension was then ®ltered
through a 200 mm mesh to remove unmacerated tissue. The cell
suspension was mildly centrifuged at 500 rpm for 10 min in a
benchtop clinical centrifuge. The supernatant containing cell debris
was discarded and the loose pellet was resuspended in PEM (PEMD
without DMSO). This procedure was repeated twice. Then 20 ml
aliquots of the cell suspension were spread onto poly-L-lysine coated
coverslips and were allowed to settle for 5 min in a humid chamber
to prevent cells from air-drying. After cells had adhered, they were
permeabilized with 3% v/v Triton X-100 in PBS pH 7.4, supplemented with 5 IUs AlexaFluor 568-conjugated phalloidin per
coverslip for 60 min. Finally, chromatin was counterstained for 60
min with 5 mg ml±1 Hoechst 33258 in PBS with the addition of 5 IUs
per coverslip AlexaFluor 568-phalloidin and specimens were ®nally
mounted in antifade solution (0.1 mg ml±1 of p-phenylenediamine in
PBS, pH 8.0 made in 90% v/v glycerol). For ¯uorescence
microscopy and photomicrography a Zeiss Axioplan microscope
equipped with standard epi¯uorescence ®lters and Neo¯uar objectives was used. Micrographs were taken on Kodak T-MAX 400 ®lm
at 1600 ASA. For the examination of AFs in intact leaf segments the
same protocol was applied with the exception of the enzyme
treatment, which was omitted. In this case photomicrography was
not applicable due to high background generated by the overlapping
cell layers.
Assessment of the AF content
To assess changes in the AF content due to plasmolysis, image
analysis of either photomicrographic negatives or directly of
¯uorophore labelled specimens were used. Although the following
approaches cannot be used to quantify in absolute numbers the
Hyperosmotic stress induces AF reorganization 1701
For digital image analysis the Image-Pro Plus software (Media
Cybernetics) was used. Following background subtraction, at least
®ve areas of prede®ned magnitude were chosen and the number of
AFs that were automatically tracked within were measured,
averaged and expressed as the amount of ¯uorescence per unit
area. Background was measured in cytoplasmic areas devoid of AFs.
Average ¯uorescence intensity per unit area of either plasmolysed or
control cells (arbitrary units) was plotted as a histogram using MS
Excel software (Microsoft Corp.). Digital image analysis to assess
differences in the AF number between different cell populations has
previously been used (Hallows et al., 1996; Schindelholz and Reber,
1999).
Differences in ¯uorescence intensity were also estimated by
measuring the overall ¯uorescence of Alexa-phalloidin labelled
cells. At least 500 cells from plasmolysed and control cells were
compared, using the photometer coupled to the Zeiss Axioplan
microscope, which converts ¯uorescence intensity to exposure time
for photomicrography. The values obtained though this approach
were comparable to those obtained from digital image analysis of the
photomicrographic negatives.
Results
General remarks
Fig. 1. Living ECs as they appear under DIC optics during the ®rst
plasmolytic cycle (A±D, plasmolysis; E, F, deplasmolysis). Numbers
in (A) to (D) indicate time lapsed (in min) after immersion of the
tissue in the plasmolytic solution. Numbers in (E) and (F) indicate
time in seconds following substitution of the plasmolytic solution by
water. Bar 10 mm.
cellular AF content, they are reliable for a comparison of the AF
content between control and plasmolysed cells.
For image analysis photomicrographic negatives of 100 control
and 100 plasmolysed cells were scanned through an Agfa Duoscan
scanner and captured as `tiff' ®les using Agfa FotoLook software.
The hyperosmotic treatment of leaf pieces of C. comosum
with 1 M mannitol induced plasmolysis in every cell type.
Most of the epidermal and mesophyll cells (EC and MCs)
examined, displayed convex or concave plasmolysis (Figs
1A±D, 3K, L). In a few plasmolysed cells, interconnected
sub-protoplasts were formed. Many large cells formed an
extensive Hechtian strand network linking the protoplast
with the cell wall. Examination of living ECs and MCs
showed that the protoplast responds instantly to the
hypertonic solution and that plasmolysis is completed
within 1±5 min after immersion of the leaf segments in the
plasmolytic solution. After 5 min the protoplast volume
and shape remain fairly constant (Fig. 1A±D). The same
plasmolysis pattern was induced by 0.5 and 0.3 M
mannitol solutions. However, the number of the plasmolysed cells in these solutions was smaller, while the course
of plasmolysis was prolonged compared with those
induced by 1 M mannitol. A comparative study of living
and isolated plasmolysed cells with DIC optics revealed
that ®xation and separation methods used in this study do
not affect the form of the plasmolysed protoplasts appreciably.
The AF staining protocol followed in this study yielded
reproducible results in plasmolysed dividing, differentiating and mature cells. In the AF staining protocol the MBS
prestabilization of AFs was abandoned, since Triton X-100
used to facilitate penetration of MBS, brought about severe
morphological alterations to the plasmolysed protoplasts.
Fixation in PBS produced slightly better results than
®xation in PEMD, probably due to the stabilizing effects of
inorganic phosphates on AF structure (Rickard and
Sheterline, 1988). Finally, the addition of substoichiometric quantities of Alexa-phalloidin in the ®xative
protected the labile AFs against severing from formalde-
1702 Komis et al.
Fig. 2. AF organization in control cells. Bar 10 mm. (A, B) Cortical AFs in a meristematic (A) and in a differentiating (B) cell. (C) AF-PPB
(arrow). (D) Endoplasmic AFs in a meristematic cell. Those are mostly located around the nucleus. (E) Cortical AFs of a differentiating MC.
(F) Subcortical AFs of a differentiating cell. (G) Endoplasmic AFs of a mature cell. (H) Cortical AFs of a mature cell.
hyde. The higher quantum yield of Alexa 568 ¯uorophore
when compared to rhodamine (Haugland, 2001) provided
the means to compensate for the high levels of auto¯uorescence, and thus to increase the signal-to-noise ratio.
AF organization in control cells
Control leaf cells display all the AF arrays described
elsewhere (Panteris et al., 1992; Cleary, 1995; Cleary and
Mathesius, 1996). In all cell types examined the cortical
cytoplasm was traversed by well-organized arrays of ®ne
AFs or AF bundles (Fig. 2A, B, E, H). In the vast majority
of the interphase meristematic cells, cortical AFs were
arranged perpendicularly to the longitudinal cell axis
(Fig. 2A). In the preprophase±prophase cells, AF preprophase bands (AF-PPBs) of various degrees of organization were routinely observed (Fig. 2C), a feature
commonly observed in other plant species (Mineyuki,
1999). Differentiating and mature cells exhibited cortical
AFs usually arranged in various orientations (Fig. 2B, E,
H).
Thick, mostly longitudinal and sparsely arranged AF
bundles traversed the sub-cortical cytoplasm, which is the
site of cytoplasmic streaming (Fig. 2F). This AF system
was more prominent in differentiating and mature leaf
cells than in the meristematic ones, in which the cortical
AF arrays dominated. In all cell types examined AF
bundles were found in the perinuclear cytoplasm (Fig. 2D,
G), which in many meristematic cells formed distinct
perinuclear AF cages (Fig. 2D). In differentiating and
mature cells AF bundles emerging from the perinuclear
cytoplasm entered the transvacuolar cytoplasmic strands,
often reaching the cell cortex (Fig. 2G).
AF organization in plasmolysed cells
In plasmolysed cells, the AF cytoskeleton is signi®cantly
affected. The changes observed were identical among all
the mannitol solutions used (0.3, 0.5 and 1 M) as well as
among all the epidermal and mesophyll cell types examined. To avoid repetition of the results the AF organization
in cells plasmolysed with 1 M mannitol will be described
only.
A consistent effect of the hyperosmotic stress on the AF
cytoskeleton is the disappearance of the ®ne cortical AFs.
This was most obvious in the interphase plasmolysed
meristematic cells. New AF systems consisting of thick AF
bundles ran through the cortical cytoplasm. Additionally,
subcortical and endoplasmic AF arrays of different organization from those found in control cells were observed
(Fig. 3A±E). Notably, the AF-PPB resisted plasmolysis in
most preprophase cells (Fig. 3F), a phenomenon also
observed in the plasmolysed ECs of Tradescantia (Cleary,
2001). At the PPB site the plasmalemma is not detached
from the cell wall (Kagawa et al., 1992; Cleary, 2001;
Komis et al., 2001).
Hyperosmotic stress induces AF reorganization 1703
Fig. 3. AF organization in cells plasmolysed for 30 min in 1 M aqueous mannitol solution. Bar 10 mm. (A) Meristematic cell displaying convex
plasmolysis. AF bundles underlying the plasmalemma. (B) Numerous well-organized AF bundles underlying the plasmalemma in this
differentiating plasmolysed cell. (C, D) Mature plasmolysed cells displaying a well-organized network of cortical/subcortical AF bundles.
Arrowheads in (D), point to Hechtian strands traversed by AFs. (E) Endoplasmic AF bundles in a meristematic plasmolysed cell. (F) AF-PPB
(arrow) in a plasmolysed cell. (G) Cell displaying concave plasmolysis. Intense actin staining can be observed under the detached regions of the
plasmalemma (arrowheads). (H) Most of the AF bundles in this plasmolysed cell are located in the most shrunken region of the protoplast (arrow).
(I) Endoplasmic AFs in a differentiating plasmolysed cell. Many AFs are localized on the surface of the nucleus (N) and the vacuoles
(arrowheads). (J) The cell displayed in I under DIC optics. (K, L) Differentiating cells displaying concave plasmolysis. Many AF bundles underlie
the detached regions of the plasmalemma (arrows). Arrowheads point to AF bundles in contact with vacuoles.
The most prominent effect of the hyperosmotic stress is
that it induced a signi®cant increase in the overall AF cell
content. AFs were more numerous in plasmolysed cells
than in the control ones (Fig. 3A±E cf. Fig. 2B, E, F, H)
and formed thick bundles (Figs 3B±D, 4A). The data
obtained using the digital image analysis of the photomicrographic negatives showed that this increase in
plasmolysed differentiating cells is about 120% in relation
to non-plasmolysed cells (Fig. 5A). Estimations made by
the microscope photometer revealed that the increase of
the AF content in differentiating plasmolysed cells is about
150%, while in meristematic cells about 50% compared to
control cells (Fig. 5B). This phenomenon was con®rmed in
ECs and MCs with all the mannitol solutions used.
In plasmolysed cells most of the cortical and subcortical
AF bundles were localized at the areas of intense
protoplast shrinkage, where they formed a network lining
the detached plasmalemma regions (Fig. 3B, G±L).
1704 Komis et al.
Fig. 4. (A) Almost all AFs in this plasmolysed cell are located within
the protoplasmic bridge (arrow) interconnecting neighboring subprotoplasts. (B) Mature plasmolysed cell. Many AFs can be observed
on the surface of the vacuole (V) as well as within the intensely
shrunken region of the protoplast (arrow). Bar 10 mm.
Generally, the AF bundles were signi®cantly enriched
within the most shrunk protoplast regions (Figs 3H, K, 4A,
B). In cells, which had undergone concave plasmolysis, an
intense actin staining was localized underneath the
plasmalemma, at the borders of the detached protoplast
regions (Fig. 3G, K, L), while almost no staining was
observed in the non-detached ones. In many cells, AFs
were also detected within most of the Hechtian strands
(Fig. 3D). The endoplasmic AF bundles ran through the
cytoplasm in various directions (Fig. 3E, I). In the majority
of plasmolysed meristematic cells the endoplasmic AF
bundles were arranged in a cage encircling the nucleus
(Fig. 3E). Differentiating and mature cells displayed AFs
juxtaposed with the tonoplast (Figs 3I±L, 4B). In some
cases AFs completely surrounded the vacuoles (Fig. 4B).
The AF organization in plasmolysed cells in intact tissue
was similar to that described above. In all plasmolysed ECs
examined numerous AF bundles were localized in the
shrunken protoplast areas, lining the detached plasmalemma regions. Therefore, the cell separation procedure does
not affect the AF organization.
CB-treated plasmolysed cells
In plasmolysed cells CB induced AF disorganization and
dramatic changes in the pattern of plasmolysis. In AF
depleted plasmolysed ECs and MCs, the course of
plasmolysis lasted for a longer time (Fig. 6A±D) and led
to a greater decrease of the protoplast volume compared
with the untreated cells (Fig. 6A±D cf. Fig. 1A±D).
Frequently, the protoplast was almost completely detached
from the cell wall (Figs 6D, 7A). As mentioned above,
in cells treated with 1 M mannitol, plasmolysis was
Fig. 5. Control cell (white bars), plasmolysed cell (grey bars). (A, B)
Histograms displaying the average AF ¯uorescence intensity of
control and plasmolysed leaf cells. (A) Histogram of the average AF
¯uorescence intensity measured through digital image analysis.
Sample size: 100 differentiating control cells, 100 differentiating
plasmolysed cells. (B) Histogram of the average AF ¯uorescence
intensity measured through microscopic. Sample size 500 control
cells, 500 plasmolysed cells (a, differentiating cells; b, meristematic
cells).
completed within 1±5 min after immersion in the
hyperosmotic solution, while in those subjected to CB
treatment continued for 30±60 min (Fig. 6A±D cf. Fig. 1A±
D).
In many CB-treated plasmolysed cells the protoplast
assumed an irregular form, usually `amoeboid' with
prominent protrusions (Fig. 7B±D). Frequently, this was
subdivided into numerous subprotoplasts, which were
usually separated from the main protoplast (Fig. 7E, F). On
the surface of many CB-treated protoplasts ®ne and rigid
protoplasmic extensions, shorter and thicker than the
Hechtian strands were observed (Figs 6D, 7A).
The CB-treated plasmolysed cells displayed many
`vesicular elements' exhibiting a rather homogeneous
Hyperosmotic stress induces AF reorganization 1705
Fig. 6. Living CB-treated ECs as they appear under DIC optics during the ®rst plasmolytic cycle (A±D, plasmolysis; E, F, deplasmolysis).
Arrowheads in (D) point to protoplasmic extensions. Numbers in (A) to (D) correspond to time in minutes after immersion of the tissue in 1 M
mannitol solution supplemented with 100 mM CB. Numbers in (E) and (F) indicate time in seconds after replacement of the plasmolytic solution
with water. Bar 10 mm.
content occupying the periplasmic area (Fig. 7C±E).
Besides, OsO4 ®xation of CB-treated cells, as well as
of the untreated ones, revealed the existence of large
osmiophilic bodies in the cytoplasm and in the
vacuoles (Fig. 7F) not found in control cells. The
nature and origin of these bodies remains to be
elucidated.
Deplasmolysis
Within seconds after the replacement of the plasmolytic
solution with water the plasmolysed protoplasts rounded
up and deplasmolysis was rapidly carried out (Fig. 1E, F).
During the protoplast re-expansion the plasmalemma reincorporated the Hechtian strands. Finally, the protoplast is
re-appressed on the cell wall.
Deplasmolysis also takes place in the CB-treated
plasmolysed cells. During this process the protoplast
becomes round and rapidly expands, while the protoplasmic extensions are reincorporated by the protoplast
(Fig. 6E, F). The sub-protoplasts and the `vesicular
elements' located in the periplasmic space were also
fused with the enlarging protoplast. Notably, the severe
plasmolysis, experienced by the cells in the presence of
CB, does not affect their viability even 60 min after the
onset of plasmolysis (Fig. 6A±F).
In deplasmolysed cells the thick AF bundles were
disorganized and ®ne AFs reappeared. They formed the AF
arrays found in control cells, i.e. cortical arrays, endoplasmic AF bundles traversing the perinuclear cytoplasm
as well as others meandering among the vacuoles (Fig. 8A±
D).
Cells that underwent a complete plasmolytic cycle could
be successfully re-plasmolysed suggesting that they still
retain their viability between the two- anisosmotic extreme
treatments. However, during the second plasmolytic cycle,
the protoplast remained plasmolysed for a short time. Then
this is slowly deplasmolysed in the presence of mannitol
solution (Fig. 9A, B). This was not the case for CB-treated
cells though. As soon as the deplasmolysed protoplast reencounters plasmolytic solution containing CB it initially
shrinks but within seconds `explodes' releasing cytoplasmic material.
1706 Komis et al.
Fig. 8. AF organization in deplasmolysed cells. Bar 10 mm. (A)
Cortical AFs in a meristematic cell. (B±D) Endoplasmic AFs in a
meristematic (B), differentiating (C), and mature (D) cell.
Fig. 7. CB-treated plasmolysed ECs under DIC optics. Bars 10 mm.
(A) In this cell the protoplast is completely detached from the cell
wall. Arrow points to a protoplasmic extension. (B, C) In these cell
groups, the plasmolysed protoplasts have assumed an `amoeboid'
form. Arrows in C mark `vesicular elements'. (D) `Amoeboid'
plasmolysed protoplast exhibiting discrete protoplasmic protrusions
(arrows). Arrowhead shows a `vesicular element'. (E) In this
plasmolysed cell the protoplast (asterisk) has suffered very intense
shrinkage. In the space between the protoplast and the cell wall subprotoplasts (arrows) and `vesicular elements' (arrowheads) can be
observed. (F) Osmiophilic bodies in CB-treated plasmolysed cells
®xed with OsO4. Arrows show sub-protoplasts.
Discussion
The changes in organization of the AF cytoskeleton
occurring during the plasmolytic cycle in leaf cells of
C. comosum are summarized as (a) the hyperosmotic stress
induces the disappearance of the ®ne cortical AFs and the
formation of numerous well-organized cortical, subcortical
Fig. 9. Living ECs under DIC optics during the second plasmolytic
cycle. It is clearly evident that the protoplast volume increases in the
presence of the plasmolytic solution. Numbers indicate time in
minutes after the initiation of the second plasmolytic cycle (starting
immediately as cells completely deplasmolysed and water was
replaced by the plasmolytic solution). Bar 10 mm.
Hyperosmotic stress induces AF reorganization 1707
and endoplasmic AF bundles; (b) the cortical AF bundles
are preferentially localized in the shrunk protoplast areas,
lining the detached plasmalemma regions: the endoplasmic AF bundles are mainly located in the perinuclear
cytoplasm and on the tonoplast surface; (c) the experimental AF disorganization prolongs plasmolysis and
induces dramatic changes in shape and volume of the
plasmolysed protoplast; and (d) during deplasmolysis, the
normal AF cytoskeleton is reinstated quite rapidly.
The ®ndings of the present study show clearly that the
AF cytoskeleton in the angiosperm C. comosum, is
intimately involved in the mechanism by which the cells
regulate the shape and the volume of the protoplast during
plasmolysis. Some of the phenomena mentioned above
will be discussed in the following sections.
AF polymerization in plasmolysed cells
The induction of AF polymerization in plasmolysed cells
observed for the ®rst time here in a plant species, has been
described in other diverse biological systems exposed to
hypertonic conditions (Hoffmann and Pedersen, 1998). For
example, in Erlich ascites tumour cells, a brief exposure to
a hypertonic challenge results in approximately a 25%
elevation of the F-actin content (Pedersen et al., 1999).
Similarly, the overall actin content is elevated in
Dictyostelium amoebae experiencing a hyperosmotic
treatment (Zischka et al., 1999). Therefore, the induction
of AF polymerization is a common response of animal,
fungal and plant cells to hyperosmotic stress. In animal
cells the excellent mechanical properties of the AF
networks seem to relieve the forces imposed on the cell
through volume reduction and compaction (Janmey,
1994).
It is known that the hyperosmotic treatment triggers a
signalling pathway that involves an increase in the
cytosolic Ca2+ concentration (Pedersen et al., 1998;
Reddy, 2001) and the generation of polyphosphoinositides
(Munnik et al., 2000; Meijer et al., 2001; Munnik, 2001).
In animal cells these pathways directly integrate the stress
condition to the AF cytoskeleton by inducing AF
polymerization and/or by affecting the AF physical
properties (Janmey, 1998).
The hyperosmotic stress in plants causes an elevation of
cytosolic Ca2+ concentration by recruiting intra- and/or
extracellular stores (Busch, 1995; Knight et al., 1997,
1998; Cessna et al., 1998; Brownlee et al., 1999; Knight,
2000). This Ca2+ mobilization in the plasmolysed C.
comosum leaf cells, could induce AF formation directly by
increasing the af®nity for polymerization of G-actin
subunits (Strzelecka-Golaszewska, 2001) or indirectly
through a series of Ca2+-dependent signal transduction
events involving the regulation of actin dynamics (Janmey,
1994, 1998).
AFs and plasmalemma protection
During plasmolysis certain plasmalemma areas retain
their attachments to the cell wall, while some others
are retracted following protoplast shrinkage (Oparka,
1994; Lang-Pauluzzi, 2000). As a result shearing forces
are generated, which might injure the plasmalemma
(Oparka, 1994). This work reveals the existence of a
positional relationship between cortical AF bundles and
the plasmalemma at those sites where the most intense
shearing forces are produced. In cells exhibiting
concave plasmolysis, AFs are signi®cantly enriched
underneath the plasmalemma, preferentially near the
boundaries between the detached and non-detached
regions of the latter. Moreover, in plasmolysed cells
displaying convex plasmolysis, AFs line the detached
plasmalemma regions.
The partial persistence of plasmalemma±cell wall
attachments in plasmolysed cells may raise a challenge
against plasmalemma integrity through stretching. It is
thus necessary for the plasmolysed protoplast to develop a
mechanism to compensate for the incoming injury
(Oparka, 1994). In animal cells, being under mechanical
stress, the cytoskeleton provides support to the plasmalemma by reinforcing the cortical framework preferentially at the sites of extensive force application (Ingber,
1997; Ko and McCulloch, 2000). A characteristic example
is the endothelial cells that line the blood vessels, which
are continuously under shear stress (Kano et al., 2000)
imposed by the blood stream. To compensate the stress,
these cells form an extensive cortical network of AF
bundles, the `stress ®bres', which are strongly inducible
upon force application (Galbraith et al., 1998). Often, they
coalign with the direction of ¯uid ¯ow under shear stress
(Galbraith et al., 1998; Frame and Sarelius, 2000). Current
work suggests a role for AF remodelling as a response to
mechanical challenge (Ingber, 1997). Stress ®bres are able
to contract, serving as a compensatory mechanism against
mechanical injury of the plasmalemma (Arora et al., 1999;
Katoh et al., 1998).
In plasmolysed leaf cells of C. comosum the AF network
formed underneath the plasmalemma might protect against
shearing possibly through an analogous mechanism like
the contraction of stress ®bres. A similar role has been also
attributed to the cortical AF network formed in
Dictyostelium hyperosmotically stressed cells (Aizawa
et al., 1999). Alternatively, it could result in an increased
rigidity of the plasmalemma at the sites of maximal
tension. AF bundles found in close association with the
vacuoles in plasmolysed cells of C. comosum may offer
mechanical support to the tonoplast as well.
In plasmolysed plant cells, numerous multilamellar
vesicles cut off from the tonoplast which at high external
salt concentration became condensed into osmiophilic
bodies (Singh and Johnson-Flanagan, 1987). These struc-
1708 Komis et al.
tures probably reserve membrane material, to be used for
the tonoplast re-expansion during deplasmolysis (Oparka,
1994). Osmiophilic bodies observed in the plasmolysed
cells of C. comosum, but not in the control ones, probably
represent surplus of membrane material.
AFs and protoplast shape regulation
The ®ndings of this work support the hypothesis that in
plasmolysed C. comosum leaf cells the AF network is
somehow involved in the regulation of the protoplast
shape. The effects of CB further support the above
suggestion. AF depletion from the plasmolysed protoplasts
results in anomalous `amoeboid' forms, never observed in
cells plasmolysed in the absence of CB. Protoplasmic
extensions similar to those found in plasmolysed protoplasts isolated from cold acclimated leaves of rye
(Dowgert and Steponkus, 1984; Gordon-Kamm and
Steponkus, 1984) were observed in this material after CB
treatment.
It should be noted here that the regulatory role of the
AFs in the shape of the plasmolysed protoplasts of C.
comosum is expressed in the absence of microtubules. The
latter are disintegrated in hyperosmotically treated
cells (Komis et al., 2001). The tubulin paracrystals and
macrotubules, which are formed, may offer mechanical
support to the protoplast (Komis et al., 2001). Whether the
tubulin paracrystal network underlies the formation of
`amoeboid' protoplast protrusions in the CB-treated
plasmolysed cells remains to be seen.
The sub-protoplasts and `vesicular elements' encountered in the CB-treated plasmolysed cells examined were
not always, if at all, linked to the main protoplast. By
contrast, sub-protoplasts were rarely observed in plasmolysed cells in the absence of CB and were always attached
to the main protoplast through AF-rich cytoplasmic
bridges. AF disruption may result in the excision of subprotoplasts as a result of plasmalemma destabilization and
blebbing.
AFs and protoplast volume regulation
Disruption of AFs in CB-treated plasmolysed cells of C.
comosum uncoupled the ability of the protoplast to
maintain a constant volume under hyperosmotic conditions. Therefore, plasmolysis should trigger a protective
mechanism, to enable the cell to attenuate the loss of water
and the protoplast volume reduction (Lee-Stadelmann and
Stadelmann, 1989). This is achieved by the increase in the
concentration of small organic and chemically inert
osmolytes in the plasmolysed protoplast (Hare et al.,
1998; Tabaeizadeh, 1998). Their synthesis, transport and
accumulation is a relatively slow procedure (Lang et al.,
1998; Tabaeizadeh, 1998). Therefore, osmotic and volume
regulation is initially carried out by the transfer of ions
from the apoplast into the protoplast through plasmalemma
ion channels, which are inducible under hyperosmotic
conditions (Lang et al., 1998). The activation of such a
regulatory mechanism in the plasmolysed cells of C.
comosum is indicated by the fact that when they undergo a
second plasmolytic cycle they are almost completely
deplasmolysed in the presence of the mannitol solution.
In animal cells, cytoskeleton-mediated cell volume
regulation after exposure to anisosmotic conditions is
mostly attributed to the capacity of cytoskeletal elements,
mostly AFs, in controlling ion channels and transporter
activities (Ko and McCulloch, 2000; Papakonstanti et al.,
2000). There are many ways by which the cytoskeleton is
implicated in ion channel function (Janmey, 1998;
Khurana, 2000; Szaszi et al., 2000). In hyperosmotically
treated animal cells the cell volume is reconstituted though
a process termed `regulatory volume increase' (RVI). As
described above, exposure to hypertonic solutions generally induces AF polymerization. Excessive AFs are
required, among others, for the activation of ion channels
(Lang et al., 1998; Pedersen et al., 1999; Szaszi et al.,
2000), which in turn trigger the RVI process.
In hyperosmotically C. comosum treated cells, the
volume regulatory response is expressed differentially
during two successive plasmolytic cycles. At the ®rst
plasmolytic cycle, where no net volume increase is
observed, it was documented as the ability of the
plasmolysed protoplast to maintain a constant volume.
This observation suggests the accumulation of osmolytes
or ions in the protoplast to counterbalance the extracellular
hypertonicity. During the second plasmolytic cycle the
accumulation of the osmolytes should be high enough to
drive a net volume increase. The ability of the protoplast to
withstand continuous shrinkage during the ®rst plasmolytic cycle and the capacity to undergo volume increase
during the second, are strongly inhibited by CB. This
implies that an AF-dependent mechanism, probably controlling the plasmalemma ion channel and aquaporin
regulation, might be responsible for protoplast volume
maintenance.
It should be noted that the implication of the AFs in
plant cell volume regulation, is not well documented.
Reversible changes in cortical AF organization related to
changes of cell volume have been observed only in stomata
of Vicia faba and Commelina communis. In open stomata,
guard cells display well-organized radial cortical AF
arrays, which disintegrate when the stomata close (Kim
et al., 1995; Eun and Lee, 1997), changes probably
mediated by cytosolic Ca2+ levels and by protein kinase
and phosphatase activities (Hwang and Lee, 2001). It has
been suggested that AFs are involved in the regulation of
the guard cell volume, by modulation of the activity of the
plasmalemma ion channels (Hwang et al., 1997), as it
probably happens in the case of plasmolysed cells of C.
comosum.
Hyperosmotic stress induces AF reorganization 1709
Acknowledgements
We thank Dr M Issidorides for allowing access facilities to the
digital image analysis of the Cozzica Foundation. Thanks are also
extended to Mr S Yietos for developing the suitable macro for
automation of actin ®lament tracking and ¯uorescence intensity
measurements. G Komis was awarded a scholarship by the State
Scholarship Foundation.
References
Aizawa H, Katadae M, Maruya M, Sameshima M, MurakamiMurofushi K, Yahara I. 1999. Hyperosmotic stress-induced
reorganization of actin bundles in Dictyostelium cells overexpressing co®lin. Genes to Cells 4, 311±324.
Arora PD, Janmey PA, McCulloch CA. 1999. A role for gelsolin
in stress ®ber-dependent cell contraction. Experimental Cell
Research 250, 155±167.
Bachewich CL, Heath IB. 1997. Differential cytoplasm±plasma
membrane±cell wall adhesion patterns and their relationships to
hyphal tip growth and organelle motility. Protoplasma 200, 71±
86.
Brownlee C, Goddard H, Hetherington AM, Peake L-A. 1999.
Speci®city and integration of responses: Ca2+ as a signal in
polarity and osmotic regulation. Journal of Experimental Botany
50, 1001±1011.
Busch DS. 1995. Calcium regulation in plant cells and its role in
signaling. Annual Review of Plant Physiology and Plant
Molecular Biology 46, 95±122.
Cessna SG, Chandra S, Low PS. 1998. Hypo-osmotic shock of
tobacco cells stimulates Ca2+ ¯uxes deriving ®rst from external
and then internal Ca2+ stores. Journal of Biological Chemistry
273, 27286±27291.
Chowdhury S, Smith KW, Gustin MC. 1992. Osmotic stress and
the yeast cytoskeleton: phenotype-speci®c suppression of an actin
mutation. Journal of Cell Biology 118, 561±571.
Cleary AL. 1995. F-actin redistributions at the division site in
living Tradescantia stomatal complexes as revealed by
microinjection of rhodamine-phalloidin. Protoplasma 185, 152±
165.
Cleary AL. 2001. Plasma membrane±cell wall connections: roles in
mitosis and cytokinesis revealed by plasmolysis of Tradescantia
virginiana leaf epidermal cells. Protoplasma 215, 21±34.
Cleary AL, Mathesius U. 1996. Rearrangements of F-actin during
stomatogenesis visualized by confocal microscopy in ®xed and
permeabilized Tradescantia leaf epidermis. Botanica Acta 109,
15±24.
Dowgert MF, Steponkus PL. 1984. Behavior of the plasma
membrane of isolated protoplasts during a freeze±thaw cycle.
Plant Physiology 75, 1139±1151.
Eun SO, Lee Y. 1997. Actin ®laments of guard cells are
reorganized in response to light and abscisic acid. Plant
Physiology 115, 1491±1498.
Frame MD, Sarelius IH. 2000. Flow-induced cytoskeletal changes
in endothelial cells growing on curved surfaces. Microcirculation
7, 419±427.
Galbraith CG, Skalak R, Chien S. 1998. Shear stress induces
spatial reorganization of the endothelial cell cytoskeleton. Cell
Motility and the Cytoskeleton 40, 317±330.
Gordon-Kamm WJ, Steponkus PL. 1984. The in¯uence of cold
acclimation on the behavior of the plasma membrane following
osmotic contraction of isolated protoplasts. Protoplasma 123,
161±173.
Gunning BES, Steer MW. 1996. Plant cell biology: structure and
function. Boston, MA: Jones and Bartlett.
Hallows KR, Law F-Y, Packman CH, Knauf PA. 1996. Changes
in cytoskeletal actin content, F-actin distribution, and surface
morphology during HL-60 cell volume regulation. Journal of Cell
Physiology 167, 60±71.
Hare PD, Cress WA, Van Staden J. 1998. Dissecting the roles of
osmolyte accumulation during stress. Plant, Cell and
Environment 21, 535±553.
Haugland RP. 2001. Handbook of ¯uorescent probes and research
chemicals. Oregon: Molecular Probes.
Henson JH. 1999. Relationships between the actin cytoskeleton
and cell volume regulation. Microscopy Research and Techniques
47, 155±162.
Hirt H. 2000. MAP kinases in plant signal transduction. Results
and Problems in Cell Differentiation 27, 1±9.
Hoffmann EK, Pedersen SF. 1998. Sensors and signal
transduction in the activation of cell volume regulatory ion
transport systems. Contributions in Nephrology 123, 50±78.
Hwang J-U, Lee Y. 2001. Abscisic acid-induced actin
reorganization in guard cells of day¯ower is mediated by
cytosolic calcium levels and by protein kinase and protein
phosphatase activities. Plant Physiology 125, 2120±2128.
Hwang J-U, Suh S, Yi H, Kim J, Lee Y. 1997. Actin ®laments
modulate both stomatal opening and inward K+-channel activities
in guard cells of Vicia faba L. Plant Physiology 115, 335±342.
Ingber DE. 1997. Tensegrity: the architectural basis of
mechanotransduction. Annual Review of Physiology 59, 575±599.
Janmey PA. 1994. Phosphoinositides and calcium as regulators of
cellular actin. Annual Review of Physiology 56, 169±191.
Janmey PA. 1998. The cytoskeleton and cell signaling: component
localization and mechanical coupling. Physiological Reviews 78,
763±781.
Kagawa T, Kadota A, Wada M. 1992. The junction between the
plasma membrane and the cell wall in fern protonemal cells, as
visualized after plasmolysis, and its dependence on arrays of
cortical microtubules. Protoplasma 170, 186±190.
Kano Y, Katoh K, Fujiwara K. 2000. Lateral zone of cell±cell
adhesion as the major ¯uid shear stress-related signal transduction
site. Circulation Research 86, 425±433.
Kapus A, Di Ciano C, Sun J, Zhan X, Kim L, Wong TW,
Rotstein OD. 2000. Cell volume-dependent phosphorylation of
proteins of the cortical cytoskeleton and cell-cell contact sites.
The role of Fyn and FER kinases. Journal of Biological
Chemistry 275, 32289±32298.
Katoh K, Kano Y, Masuda M, Onishi H, Fujiwara K. 1998.
Isolation and contraction of the stress ®ber. Molecular Biology of
the Cell 9, 1919±1938.
Khurana S. 2000. Role of actin cytoskeleton in regulation of ion
transport: examples from epithelial cells. Journal of Membrane
Biology 178, 73±87.
Kim M, Hepler PK, Eun SO, Ha KS, Lee Y. 1995. Actin
®laments in mature guard cells are radically distributed and
involved in stomatal movement. Plant Physiology 109, 1077±
1087.
Knight H. 2000. Calcium signaling during abiotic stress in plants.
International Review of Cytology 195, 269±324.
Knight H, Trewavas AJ, Knight MR. 1997. Calcium signalling in
Arabidopsis thaliana responding to drought and salinity. The
Plant Journal 12, 1067±1078.
Knight H, Brandt S, Knight MR. 1998. A history of stress alters
drought calcium signalling pathways in Arabidopsis. The Plant
Journal 16, 681±687.
Ko KS, McCulloch CA. 2000. Partners in protection:
interdependence of cytoskeleton and plasma membrane in
adaptations to applied forces. Journal of Membrane Biology
174, 85±95.
Komis G, Apostolakos P, Galatis B. 2001. Altered patterns of
1710 Komis et al.
tubulin polymerization in dividing leaf cells of Chlorophyton
comosum after a hyperosmotic treatment. New Phytologist 149,
193±207.
Lang F, Busch GL, Ritter M, Volkl H, Waldegger S, Gulbins E,
Haussinger D. 1998. Functional signi®cance of cell volume
regulatory mechanisms. Physiological Reviews 78, 247±306.
Lang-Pauluzzi I. 2000. The behaviour of the plasma membrane
during plasmolysis: a study by UV microscopy. Journal of
Microscopy 198, 188±198.
Lang-Pauluzzi I, Gunning BES. 2000. A plasmolytic cycle: the
fate of cytoskeletal elements. Protoplasma 212, 174±185.
Lee-Stadelmann OY, Stadelmann EJ. 1989. Plasmolysis and
deplasmolysis. Methods in Enzymology 174, 225±247.
Meijer HJ, Arisz SA, Van Himbergen JA, Musgrave A, Munnik
T. 2001. Hyperosmotic stress rapidly generates lyso-phosphatidic
acid in Chlamydomonas. The Plant Journal 25, 541±548.
Mineyuki Y. 1999. The preprophase band of microtubules: its
function as a cytokinetic apparatus in higher plants. International
Review of Cytology 187, 1±49.
Munnik T. 2001. Phosphatidic acid: an emerging plant lipid second
messenger. Trends in Plant Science 6, 227±233.
Munnik T, Meijer HJ, Ter Riet B, Hirt H, Frank W, Bartels D,
Musgrave A. 2000. Hyperosmotic stress stimulates
phospholipase D activity and elevates the levels of phosphatidic
acid and diacylglycerol pyrophosphate. The Plant Journal 22,
147±154.
Ooms LM, McColl BK, Wiradjaja F, Wijayaratnam AP,
Gleeson P, Gething MJ, Sambrook J, Mitchell CA. 2000.
The yeast inositol polyphosphate 5-phosphatases inp52p and
inp53p translocate to actin patches following hyperosmotic stress:
mechanism for regulating phosphatidylinositol 4,5-bisphosphate
at plasma membrane invaginations. Molecular and Cellular
Biology 20, 9376±9390.
Oparka KJ. 1994. Plasmolysis: new insights into an old process.
New Phytologist 126, 571±591.
Panteris E, Apostolakos P, Galatis B. 1992. The organization of
F-actin in root tip cells of Adiantum capillus veneris throughout
the cell cycle. A double label ¯uorescence microscopy study.
Protoplasma 170, 128±137.
Papakonstanti EA, Vardaki EA, Stournaras C. 2000. Actin
cytoskeleton: a signaling sensor in cell volume regulation.
Cellular Physiology and Biochemistry 10, 257±264.
Pedersen SF, Jorgensen NK, Hoffmann EK. 1998. Dynamics of
Ca2+i and pHi in Ehrlich ascites tumor cells after Ca2+-mobilizing
agonists or exposure to hypertonic solution. P¯ugers Archives
436, 199±210.
Pedersen SF, Mills JW, Hoffmann EK. 1999. Role of the F-actin
cytoskeleton in the RVD and RVI processes in Ehrlich ascites
tumor cells. Experimental Cell Research 252, 63±74.
Reddy AS. 2001. Calcium: silver bullet in signaling. Plant Science
160, 381±404.
Richelme F, Benoliel AM, Bongrand P. 2000. Dynamic study of
cell mechanical and structural responses to rapid changes of
calcium level. Cell Motility and the Cytoskeleton 45, 93±105.
Rickard JE, Sheterline P. 1988. Effect of ATP removal and
inorganic phosphate on length redistribution of sheared actin
®lament populations. Evidence for a mechanism of end-to-end
annealing. Journal of Molecular Biology 201, 675±681.
Rizoli SB, Rotstein OD, Parodo J, Phillips MJ, Kapus A. 2000.
Hypertonic inhibition of exocytosis in neutrophils: central role for
osmotic actin skeleton remodeling. American Journal of
Physiology 279, C619±633.
Schindelholz B, Reber BF. 1999. Quantitative estimation of Factin in single growth cones. Methods 18, 487±492.
Singh J, Johnson-Flanagan AM. 1987. Membrane alterations in
winter rye and Brassica napus cells during lethal freezing and
plasmolysis. Plant, Cell and Environment 10, 163±168.
Slaninova I, Sestak S, Svoboda A, Farkas V. 2000. Cell wall and
cytoskeleton reorganization as the response to hyperosmotic
shock in Saccharomyces cerevisiae. Archives of Microbiology
173, 245±252.
Strzelecka-Golaszewska H. 2001. Divalent cations, nucleotides,
and actin structure. Results and Problems in Cell Differentiation
32, 23±41.
Sullivan R, Burnham M, ToÈroÈk K, Koffer A. 2000. Calmodulin
regulates the disassembly of cortical F-actin in mast cells but is
not required for secretion. Cell Calcium 28, 33±46.
Szaszi K, Grinstein S, Orlowski J, Kapus A. 2000. Regulation of
the epithelial Na(+)/H(+) exchanger isoform by the cytoskeleton.
Cellular Physiology and Biochemistry 10, 265±272.
Tabaeizadeh Z. 1998. Drought-induced responses in plant cells.
International Review of Cytology 182, 193±247.
Zischka H, Oehme F, Pintsch T, Ott A, Keller H, Kellermann J,
Schuster SC. 1999. Rearrangement of cortex proteins constitutes
an osmoprotective mechanism in Dictyostelium. EMBO Journal
18, 4241±4249.