Microscopy Advance Access published January 23, 2014
Microscopy, 2014, 1–10
doi: 10.1093/jmicro/dft083
A study of the native cell wall structures of the
marine alga Ventricaria ventricosa
(Siphonocladales, Chlorophyceae) using atomic
force microscopy
Downloaded from http://jmicro.oxfordjournals.org/ at University of New South Wales on April 2, 2014
Enid M. Eslick1,3,*, Mary J. Beilby2, and Anthony R. Moon1
1
Department of Physics and Advanced Materials, University of Technology, Sydney, Broadway, NSW
2007, Australia, 2Department of Biophysics, School of Physics, University of New South Wales, Kensington,
NSW 2052, Australia, and 3Present address: Radiation Physics Laboratory, Sydney Medical School,
University of Sydney, Camperdown, NSW 2006, Australia
*To whom correspondence should be addressed. E-mail: enid.eslick@sydney.edu.au
Received 24 November 2013; Accepted 11 December 2013
Abstract
A substantial proportion of the architecture of the plant cell wall remains unknown with a
few cell wall models being proposed. Moreover, even less is known about the green algal
cell wall. Techniques that allow direct visualization of the cell wall in as near to its native
state are of importance in unravelling the spatial arrangement of cell wall structures and
hence in the development of cell wall models. Atomic force microscopy (AFM) was used
to image the native cell wall of living cells of Ventricaria ventricosa (V. ventricosa) at high
resolution under physiological conditions. The cell wall polymers were identified mainly
qualitatively via their structural appearance. The cellulose microfibrils (CMFs) were easily
recognizable and the imaging results indicate that the V. ventricosa cell wall has a
cross-fibrillar structure throughout. We found the native wall to be abundant in matrix
polysaccharides existing in different curing states. The soft phase matrix polysaccharides
susceptible by the AFM scanning tip existed as a glutinous fibrillar meshwork, possibly
incorporating both the pectic- and hemicellulosic-type substances. The hard phase matrix
producing clearer images, revealed coiled fibrillar structures associated with CMFs,
sometimes being resolved as globular structures by the AFM tip. The coiling fibrillar
structures were also seen in the images of isolated cell wall fragments. The mucilaginous
component of the wall was discernible from the gelatinous cell wall matrix as it formed
microstructural domains over the surface. AFM has been successful in imaging the native
cell wall and revealing novel findings such as the ‘coiling fibrillar structures’ and cell wall
components which have previously not been seen, that is, the gelatinous matrix phase.
Key words: atomic force microscopy, cell wall, cellulose microfibril, green algae, mucilage, sulphated
polysaccharides, Valonia, Ventricaria
© Crown copyright 2014
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Introduction
Ventricaria ventricosa, formerly known as Valonia ventricosa [25] has had a long history in cell wall studies dating
back to the early 1900s using spectroscopic and microscopic
techniques [25–28]. The cell wall structures of this alga and
its close relatives in the genus of Valonia are larger in size
compared with related features in land plants; hence intricate polymer associations may be more easily discernible in
such walls. The V. ventricosa cell wall is abundant in cellulose [29], which is in its highest form of crystallinity [25,30].
This has made it a popular sample used in the study of cellulose molecular structure [31–35]. The cellulose molecules
are arranged in chains forming long ribbon-like structures,
the CMFs [30]. The CMFs are arranged parallel to one
another in each lamella of the cell wall and the overall cell
wall of V. ventricosa displays a cross-fibrillar structure,
having three main CMF directions with each lamella containing only one CMF direction [36]. There could be several
layers of parallel CMFs in one lamella [37]. Unlike V. ventricosa the CMFs in the cell wall of land plants is usually in a
helicoidal arrangement with a different arrangement in the
cellulose synthesis enzyme complexes [38].
In this study, we aim to obtain a direct visualization of the
intact native cell wall structures and architecture of a living
cell utilizing the AFM. While there have been several AFM
structural studies on isolated plant cell walls, either dry or
partially hydrated [39–43], there has been only one structural
study on a native plant cell wall [44]. In terms of the application of AFM in imaging algal cell walls, there has been no
previous native cell wall study; however, isolated walls of
Vaucheria terristris sensu Goetz have been imaged [45].
Materials and methods
Algal material
Large spherical cells of V. ventricosa (Siphonocladales,
Chlorophyceae) [24] usually over 2 cm in diameter were collected from Heron Island on the Great Barrier Reef, Queensland, Australia. For experiments, sterile cultures were used
where the aplanospores were propagated artificially and
placed on sterile coral maintained in sterilized f/2 medium
with a pH of 8.0 [46]. Cultures were maintained at tropical
temperatures (22–25°C) and grown in natural light cycles.
The osmotic pressure of the medium was approximately
990 mOsmol kg−1. The osmotic pressure was measured
with a cryoscopic Osmometer (Osmomat 030, Gallay Scientific, North Melbourne, Victoria, Australia).
Preparation for native cell wall imaging (live cells)
Cells with diameters ranging from 2 to 4 mm were selected
for experiments. One week prior to the experiments, corals
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The simplified concept of the complex dynamic plant cell
wall (type I) is that it is a network of cellulose microfibrils
(CMFs) cross-linked with hemicelluloses embedded in a
pectin gel [1]. The mechanical and functional properties of
the cell wall are defined by its detailed molecular architecture [2]. Even though the chemistry of the plant cell wall
polymers is well known [3], there remains a lack of understanding regarding the arrangement of the polymers in the
cell wall and of their functional role [4]. There are a few cell
wall structural models that exist and these models are continually being refined [5,6].
Techniques that allow direct visualization of the cell wall
structures and architecture are of enormous benefit as they
have the potential to reveal how all wall polymers are
related. The methods used in direct visualization studies
have significant limitations in allowing the study of the cell
wall in as near to its native state. Commonly, the cell wall is
isolated from the plant cell and undergoes a variety of
sample preparations necessary for the particular technique
[7]. Unlike conventional electron microscopy techniques,
the rapid freeze deep-etching electron microscopy technique
[8] has been vital in the development of cell wall models,
revealing the existence of the cross-linking nature of wall
polymers such as the hemicellulosic polymer xyloglucan
[9,10]. Histochemical and cytochemical labelling techniques
have been useful in specific localization of some molecules
in the wall but they are unable to provide information on
the three-dimensional arrangement of the molecular
network [8,11]. A relatively recent powerful technique
which can be used in cell wall architectural and structural
studies is atomic force microscopy (AFM).
AFM remains a relatively new form of microscopy in
plant cell wall studies, even though it has been around for
almost three decades [12] ,and has attractive advantages for
its application in biological studies. A major advantage is
that an AFM can be operated in liquid media and thereby
allows the study of structures on living cells in their physiological environment [13,14]. Samples are imaged by a
probing tip either in direct contact with the sample or in
intermittent contact with the sample, allowing subnanometer
resolution on proteins in reconstituted membranes and few
tens of nanometers on live cells, due to their viscoelasticity
[15,16]. Moreover, the AFM can also conduct nanomechanical measurements on cell surfaces and determine the relative
elasticities of the sample [17]. For reviews on the application
of AFM in cell biology studies see [18–20].
Commonly used models for plant cells are large unicellular green algae [21]. They are easy to work with and the
area of disturbance by a probing device, e.g. an AFM tip, is
insignificant in size compared with the cell [22–24].
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E.M. Eslick et al. AFM imaging of Ventricaria cell wall, 2014, Vol. 0, No. 0
containing cells were placed in 35 mm plastic Petri dishes
for acclimatization to the experimental conditions in simplified artificial seawater (ASW), of 990 mOsmol kg−1 and pH
8.0 [47].
Enzyme treatment on native cell wall
Preparation of cell wall fragment specimens
Cells were cut open and small fragments of cell wall were
created and placed in a fixing solution (2.5% glutaraldehyde
in 0.1 M sodium cacodylate buffer of pH 7.4) overnight at
4°C. The samples were washed three times with cacodylate
buffer of pH 7.4 and the fixed cell walls were dehydrated
with 70% ethanol.
Atomic force microscopy
The most commonly used imaging mode for live biological
specimens is the contact mode (CMAFM) [50]. In contact
mode imaging, the tip is in constant contact with the surface
as it scans the sample. This mode usually produces the
highest quality images if the applied force on the sample is
able to be kept to a minimum to minimize sample damage,
e.g. as in low force contact mode (LCMAFM) imaging [51–
53]. We minimized imaging forces on the live cells by
viewing the force curves in the AFM force mode prior to
imaging and setting the imaging set point to correspond to
minimal force. In terms of non-living biological specimens,
tapping mode (TMAFM) [54–56] imaging is usually preferred. In tapping mode imaging, the cantilever is made to
vibrate near its resonant frequency and then brought into
contact with the surface which reduces shear forces as the
tip is in intermittent contact with the surface.
In most AFM imaging studies, the amount of force
applied to the sample is kept constant by a feedback loop
system. Two types of images can be collected each time, one
containing height information, called ‘height’ image, and
the other called the ‘error signal’ image which is obtained by
Native cell wall imaging in ASW
The cells were rigidly immobilized on the coral in ASW and
were placed in the AFM isolation box overnight to acclimatize to the AFM room conditions. Live cell imaging was
carried out using a Dimension 3100 AFM equipped with a
Nanoscope IIIa controller (Veeco Instruments, Santa
Barbara, CA, USA). The Petri dish containing the cells in
ASW was mounted directly on the Dimension 3100 stage
plate and held secure with double-sided tape. The fluid cell
(DTFML, Veeco Instruments, Santa Barbara, CA, USA,)
was used for imaging in liquid. The Dimension 3100 AFM
is equipped with a camera which facilitates positioning the
tip onto the cell.
Cells were imaged in ASW, in contact mode (CMAFM)
using a silicon nitride tip (DNP, Veeco Instruments, Santa
Barbara, CA, USA) with a nominal spring constant of
0.32 N m−1. An imaging scan rate of 0.3 Hz was used.
Cell wall fragments imaging in ambient conditions
Samples of the inner wall were prepared by gluing the wall
fragment onto mica using araldite two-part glue. Excess
ethanol was removed from the sample. Mica was stuck on
an AFM metal sample puck. We used a Multimode™ AFM
(Veeco Instruments™, Santa Barbara, USA) operating with
a NanoScope IIIa™ controller and an E scanner. Tapping
mode imaging (TMAFM) was applied on cell wall fragment
specimens using silicon cantilevers, model tapping mode
etched silicon probe (TESP, Veeco Instruments™, Santa
Barbara, USA) operating at resonant frequencies of
∼300 kHz. The drive frequency was set to ∼8 KHz.
Surface measurements of structures in height images
were made using the section analysis module of the AFM
software (Veeco Instruments, Santa Barbara, CA, USA).
Image flattening and filtering were also done using the AFM
software (Veeco Instruments, Santa Barbara, CA, USA).
Results
Native cell wall
Most living cells contained patchy areas of elevated voluminous coating on their surface of thickness of hundreds of
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The surface cell wall layers including sulphated polysaccharide mucilage was removed by treating the cell surface with
1% Cellulysin (Tricoderma viridae) (Calbiochem-Novabiochem, Sydney, Australia) and 0.5% bovine serum albumin
(BSA) (Sigma, St Louis, MO, USA) in diluted 2-(N-Morpholino)ethanesulfonic acid (MES) (Sigma, St Louis, MO, USA)
buffer solution containing 1:3 ASW/MES buffer (pH 5.8).
The cells were incubated in the enzyme solution at 30°C for
10 min and then washed several times in ASW. As most commercially available wall-digesting enzymes are not highly
purified, BSA was added to the enzyme digestion recipe to
reduce the activity of proteases [48,49].
using the corrections of the AFM feedback loop system as it
works to maintain the constant specified force set by the
user [57]. The error signal image, referred to as ‘deflection
image’ in CMAFM imaging and ‘amplitude image’ in
TMAFM imaging, consists mainly of high-frequency components that are directly related to the fine topographic
detail in the sample.
Microscopy, 2014, Vol. 0, No. 0
Also revealed at this magnification were coiling fibrillar
structures (Figs. 6 and 7 long open arrowhead). These
coiling structures could possibly represent disorder in the
fibril alignment, or most probably represent solidified fibril
structures caused by features pointed by solid head arrows,
which show glutinous phase meshwork fibrils draping over
CMFs (Fig. 6 dotted solid head arrows). Some cell surfaces
appeared solidified; the effect of the solidified phase matrix
substances created a globular surface appearance (Fig. 8).
Fibril outlines can be made out in the image. Higher magnification of the enzyme treated wall revealed fibril-like structures and cross-linking structures (Fig. 9).
Fig. 3. Error signal image of native cell wall after enzyme treatment.
Imaged in ASW using CMAFM. The CMF network can be seen. Scale
bar, 0.5 µm.
Fig. 1. Error signal image of large area of the native cell wall showing a
voluminous non-transparent coating on its surface. Imaged in ASW
using CMAFM. Contamination with plankton can be seen. Scale bar,
3.5 µm.
Fig. 2. Error signal image of non-transparent coating appearing as a thin
layer over the surface of the wall. Imaged in ASW using CMAFM. Scale
bar, 0.8 µm.
Fig. 4. Image of native cell wall without non-transparent coating imaged
in ASW using CMAFM; height image (above), error signal image
(below). The wall appears disordered in the height image. The
cross-fibrillar wall network can be seen in the error signal image. Scale
bar, 1.5 µm.
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nanometers and best viewed in large scan areas of 20 µm ×
10 µm or more (Fig. 1). In areas of no coating, a flat and
uniform wall surface was observed. Some cells did not
contain voluminous coating but instead a thin layer of a substance with a microstructure of poorly defined domains
(Fig. 2). Enzyme treatment was used to remove the thin
coating substance and the underlying cell wall network was
revealed (Fig. 3). Other cells revealed the cell wall network
immediately, hence lacked a coating (Fig. 4). The height
images were not clear in revealing the alignment of CMFs
network; however, details of fibril alignments could be seen
in the error signal images (Fig. 4). Moreover, image filtering
emphasized the well-known cross-fibrillar cell wall structure
of V. ventricosa (Fig. 5). Two main CMF directions were
discernible and the images showed that the microfibrils were
well aligned and straight.
Imaging at a higher resolution of cell wall surfaces
showed ‘haziness’ in the images (Fig. 6). The haziness was
created by a gelatinous phase substance. Also seen in the
images is a glutinous phase fibrillar meshwork (Fig. 6 solid
head arrows) overlying the CMF network. The meshwork
was found only on areas of the wall with CMFs, voids in the
CMF network corresponding to areas of no meshwork.
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Fig. 5. Filtered image Fig. 4 emphasizing the cross-fibrillar structure
(black arrows). Scale bar, 1.5 µm.
Fig. 9. Error signal image of enzyme-treated native cell wall, imaged in
ASW using CMAFM. Fine details of the thin fibril-like structure (long
arrows) and cross-linking structures (short arrows) can be seen. The
directions of underlying fibrils can also be seen (double headed dashed
arrow). Scale bar, 0.5 µm.
The measurements of the widths of the CMFs were
within a broad range, from 80 to 250 nm. This suggests that
the width of the CMFs may be affected by the amount of
coiling fibrillar structures associated with them. The width
of the thin coiling fibrillar structures ranged between 50 and
140 nm, with the more solidified surface having the smaller
size polymer width.
Cell wall fragments
Fig. 7. Height image of native cell wall in ASW using CMAFM. Meshwork
overlaying CMFs. Coiling bands can be seen (long arrows). Scale bar,
1 µm.
The inner surface of the cell wall imaged in TMAFM
revealed the cross-fibrillar structure of the V. ventricosa wall
(Fig. 10). The CMFs in the surface layers were sparse and
this allowed imaging of the underlying layers of CMFs . In
the deeper layers, the CMFs appeared to be more densely
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Fig. 6. Higher resolution image of the native cell wall without
non-transparent coating in ASW using CMAFM; height image (left),
error signal image (right). The wall appears disordered. A hazy artefact
is seen over areas of the wall, which correspond to areas of the skeletal
network of the surface layer. There is no hazy artefact in areas where
there are voids in the surface skeletal network. Long open head arrows
point to coiling fibrillar structures; short dotted arrows pointing to
‘drapping’ in fibrillar meshwork. The cross-fibrillar network can be seen
clearer in the error signal images. Scale bar, 1 µm.
Fig. 8. Error signal image of native cell wall with a more solidified
surface. Imaged in ASW using CMAFM. Globular fibril outlines can be
seen. Scale bar, 0.2 µm.
Microscopy, 2014, Vol. 0, No. 0
packed. The cell wall lamellae were ∼20 nm thick and the
widths of the CMFs were in the range 40–60 nm. In addition to the CMFs, other cell wall components were visible.
Higher-resolution images revealed that the CMFs had
coiling thinner fibril-like structures along their lengths,
comparable to the native cell wall images (Fig. 11, short
arrows). The widths of these fibril structures were measured
to be in the range 15–30 nm. Occasionally, we found areas
of structureless material in the wall, located between adjacent wall lamellae and covering the underlying CMFs in
that region (Fig. 11 long arrows). The location of the structureless material with reference to the surface layer could be
seen much more clearly in the height images. Unusual swirllike structures were found on the inner wall (Fig. 12) with
diameters of ∼700 nm. In larger area scans, multiple swirllike structures were seen (Fig. 10 arrows).
Discussion
This study highlights the application of AFM in the study of
native cell wall structures and architecture. Cell wall studies
utilizing the AFM have been conducted on fragments of isolated wall, except for [44], using both contact mode and
tapping mode imaging. The results from those studies
reported mainly on the CMF part of the wall with little
information on matrix cell wall components [58]. Studies of
the native cell wall, abundant in matrix polysaccharides, are
likely to provide more insights into cell wall mechanisms.
Native cell wall images revealed that the surfaces of some
cells contained a coating of a substance which was either
present in a voluminous amount or a thin layer, obstructing
Fig. 11. Higher-resolution image of fixed inner wall surface imaged in
ambient conditions using TMAFM; height image (left), error signal
image (right). Thin fibril-like structures can be seen to form coils along
the lengths of the cellulose microfibrils (short white arrows). In some
areas, we observed a thin layer of structureless material between
lamellaes covering the underlying fibrils, but this was rarely seen (long
arrows). Scale bar, 0.2 µm.
Fig. 12. High-resolution image of swirl structures on fixed inner wall
surface imaged in ambient conditions using TMAFM. These structures
are possibly anchoring sites of aplanaspores. Scale bar, 0.2 µm.
the visualization of the cell wall network, while other cells
were coating-free. This coating was attributed to the
sulphated polysaccharide extracellular mucilage that is commonly found on the surface of algal cells including V. ventricose [59,60]. The existence of the mucilaginous substance
over the surface of the cells, in smaller amounts, was identified by its microstructure. Similar sulphated polysaccharide
mucilage microstructures have been observed on diatoms,
also revealed by AFM [61]. Preliminary AFM physical measurements on the mucilaginous surface showed that it is an
adhesive substance (data not shown). Enzymatic treatment
of the wall surface removed this non-transparent coating
and revealed the underlying cell wall network.
The cell wall network was directly observed on cells
which lacked the coating. The height images appeared to
show a complex CMF network, with no particular order,
amongst amorphous soft phase matter. On the other hand,
the error signal images, capable of revealing finer structural
details, showed that the V. ventricosa outer wall is ordered
with a cross-fibrillar structure. This finding concurs with
previous studies on V. ventricosa wall [36,37,62–64].
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Fig. 10. Error signal image of fixed inner wall surface, imaged in ambient
conditions using TMAFM. The cross-fibrillar wall structure is clearer in
TMAFM images. Cytoplasmic contents are seen on the upper left side of
the wall. Scale bar, 0.6 µm. (See Fig. 12 for explanation of arrows in this
figure).
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walls or its Valonia relatives. Interestingly, this fibril structure is also evident in our images of isolated cell wall. The
visualization of these features in our imaging study of V.
ventricosa wall, unlike previous reports, may be due to the
larger cell wall structures of large algal cells compared with
plant cells or the fact that these structures have not been
seen on other Valonia species may suggest that it is native to
V. ventricosa. Of interest, Baker et al. [33] found corrugated
CMF surfaces in their AFM high-resolution study of purified CMFs from Valonia wall, which they attributed to
sample preparation effects. The coiling fibrillar structure
conformational association with CMFs suggests that they
form a major load-bearing network in the wall, and hence
may be a separate matrix polysaccharide from pectin,
similar to the hemicellulosic polymer, xyloglucan, in the cell
wall of land plants [67,68]. Xyloglucan is considered to be
hydrogen bonded to cellulose. Morris et al. [69] have used
AFM to study the desorption energies of xyloglucan from
V. ventricosa cellulose which did not suggest a complex
coiling interaction. However, studies in the conformation of
xyloglucan have suggested that it can form into flexible to
semi-flexible random coils [9,70,71]. There is no information on the hemicellulosic polysaccharides in the cell walls
of V. ventricosa; however, possible candidates for the
coiling fibrillar structures binding to CMFs in the cell wall
of V. ventricosa are xyloglucan-like polysaccharides such as
ulvan [72,73] or mixed linkage xyloglucan [74] ,which have
been found in related algal species in the genus Ulva. While
these polysaccharides are similar to xyloglucan, they differ
in composition, structure and charge. Mannans also have
the capability of bonding with CMFs [75]. Furthermore, a
coiling fibril conformation associated with wall loosening
may raise some questions in relation to the mechanism by
which this change occurs (i.e. ease or difficulty of uncoiling
of the polymer or does this coiling structure change into a
different structure), issues which remain to be elucidated.
Tepfer and Cleland [76] had previously demonstrated that
the wall loosening response in V. ventricosa is less complex
than in land plants, perhaps not involving enzymatic
mechanisms but rather conformational changes in the structures in the cell wall.
Further evidence of the existence of the coiling structures
is reflected in the measurements of the CMFs in our images.
CMFs from green algae are larger in dimension than in land
plants which are ∼5–15 nm wide [3]. CMFs from V. ventricosa have been reported to be both rectangular and square
in shape [27,62]. The majority of studies have found the
width of the V. ventricosa CMFs to be ∼20 nm using
various techniques such as X-ray and electron diffraction
and electron microscopy [27,62,77]. Similar findings have
been reported using AFM [31,33], however, larger CMFs
measurements have been reported (100 nm) and attributed
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Interestingly, our inner wall images of V. ventricosa also
showed a cross-fibrillar ordering and, hence, as suggested
by Preston [36], we conclude that the V. ventricosa wall is
cross-fibrillar ordered throughout and the terms ‘primary
wall’ and ‘secondary wall’ may not apply in this species.
This is different from its close relative Valonia-macrophysa
Kütz, which displayed a difference in the fibrillar phase in
the primary and secondary walls [63]. This finding is of
interest as it suggests more differences in their evolutionary
lineages.
The complexity in the native wall network is augmented
by the abundant presence of matrix polysaccharides. The
matrix polysaccharides in the wall appeared in both an
amorphous phase and a fibrillar phase with these two components appearing together, hence making it difficult to
decipher whether this was one cell wall matrix component
or two separate ones. The matrix polysaccharides in the
wall appeared to have gelatinous characteristics appearing
in varying ‘curing’ states, from glutinous fibrillar meshwork
to solidified surface of ill-defined fibrils, having more of a
globular feature. The matrix polysaccharides in its soft
phase created hazy image artefacts due to its susceptibility
to tip scanning. The soft phase matrix was present in
regions that contained CMFs directly under it and not in
areas that contained voids in the CMF network. These
characteristics fit well with the pectin gel matrix which is
known to exist in abundance in the V. ventricosa wall
[28,65]. A similar finding of this behaviour for gelatinous
pectin in the wall was found by Chanliaud and Gidley [66]
in their study of the pectin/cellulose composite, which
showed that pectin bonded with the CMF network, and
voids in the pectin layer were seen to correspond with voids
in the CMF network. Unlike the sulphated polysaccharide
mucilaginous layer, we found this cell wall component not
to have strong adhesive properties (data not shown). It
appears that the gelatinous amorphous matrix component
is susceptible to wall isolation and sample preparation techniques and, unlike the isolated or fixed cell wall, the native
cell wall is abundant in glutinous matrix substances which
provided optimal conditions for their observation.
The solidification of the matrix polysaccharides resulted
in clearer images with no hazy artefacts. The glutinous
fibrillar meshwork seen draping across CMFs solidified into
coiling fibrillar structures. On some cells, these coiling structures are clearly seen and on others the coiling structures
appear as globules as a result of possibly further solidification and tip broading artefact. It is known that cell wall
pectin chemistry can undergo changes such as deesterification [3], which would lead to wall hardening.
There have been no previous reports of the coiling fibrillar
structures in other high-resolution studies of the cell wall,
utilizing AFM or electron microscopy, on V. ventricosa cell
Microscopy, 2014, Vol. 0, No. 0
Conclusions
AFM has been a powerful tool in the study of the native cell
wall allowing direct imaging of the wall’s structures and
architecture. This has been of importance especially in revealing wall structures that are susceptible to wall isolation and
fixation. Fine details of the wall structures were able to be
seen in the error signal mode images. We found that cells
contained different amounts of extracellular mucilage and
that the presence of the extracellular mucilage could be identified by its distinctive microstructure and adhesion properties. We found the wall matrix to be associated with the CMF
network and it existed in different curing states from a glutinous substance of amorphous matter and fibrillar matter to
the solidified fibrillar phase of coiling structures. We surmise
that the larger widths of CMFs found in the native cell wall is
due to the amount of matrix coiling fibrillar structures associated with it. We have found that V. ventricosa has an
ordered wall throughout, unlike its close relative Valonia
macrophysa Kütz. Our images of the native V. ventricosa
wall support the proposed cell wall models of land plants, in
particular the tethered cell wall model [79] and multi-coat
model [80]. The tethered model has cross-linking polymers to
CMFs and the multi-coat model has matrix polysaccharides
coating the CMFs. Further studies involving chemical analyses of the V. ventricosa wall would be of interest to reveal
the identity of the thin fibril-like structures.
Acknowledgements
Enid M. Eslick was awarded an Australian Postgraduate Award
(APA) as part of the work in this study. She also completed an
Atomic Force Microscopy research internship at Veeco Instruments,
Santa Barbara, USA, under the guidance of Dr Peter Harris. Additional Atomic Force Microscopy support from Dr Adam Mechler is
greatly appreciated. We would like to thank Prof. Peter Ralph and Ms
Anthea Harris from UTS for assistance with the making and provision of ASW for these experiments. We would like to thank Dr Lou
De Filippis for his guidance with the enzyme treatment of the cell
wall. Additional support from the Department of Health Sciences,
UTS, for assistance with tissue culture is gratefully acknowledged.
Jenny Norman from the University of New South Wales electron
microscopy unit provided technical support with the isolated cell wall
preparation. We appreciate access to the UTS Microstructural Analysis Unit. We would like to thank the Heron Island Research Station
for collection of the algal samples used in this research.
References
1. Carpita N C, Gibeaut D M (1993) Structural models of primarycell walls in flowering plants - consistency of molecular-structure
with the physical-properties of the walls during growth. Plant J.
3: 1–30.
2. Roberts K (2001) How the cell wall acquired a cellular context.
Plant Physiol. 125: 127–130.
3. Cosgrove D J (1997) Assembly and enlargement of the primary
cell wall in plants. Annu. Rev. Cell Dev. Biol. 13: 171–201.
4. Fry S C (2004) Primary cell wall metabolism: tracking the careers
of wall polymers in living plant cells. New Phytol. 161: 641–675.
5. Cosgrove D J (2001) Wall structure and wall loosening. A look
backwards and forwards. Plant Physiol. 125: 131–134.
6. Brett C, Waldron K (1990) Molecular components of the wall.
In: Black M, Chapman J (eds), Physiology and Biochemistry of
Plant Cell Walls, pp. 4–43 (Unwin Hyman, New York).
Downloaded from http://jmicro.oxfordjournals.org/ at University of New South Wales on April 2, 2014
to tip broadening effects [31–33]. In our case, in the native
wall, even larger measurements of the CMFs were obtained.
Although our measurements would have been susceptible to
a certain degree of tip broadening, we suggest that our
larger measurements for CMFs are a consequence of the
association of the coiling fibrillar structures with them.
Peculiar swirl-like structures were observed on the inner
wall. It is unclear as to what these were; we suggest they
could be a site of protoplast (aplanospore) anchoring by
cytoskeletal networks or an unknown cell substance. The
study of Preston and Astbury [26] described modification of
the cell wall into raised circular rims wherever holdfasts (rhizoids) or buds (aplanospores) had developed. Furthermore,
V. ventricosa has a curious ability to eject zoospores, which
form mitotically, directly through the cell wall [26,78]. The
cell wall is perforated enabling direct communication
between the vacuole and the medium, and zoospores are
ejected. However, Preston and Astbury [26] found no
evidence in their polarizing light microscope study of V. ventricosa that the wall structure changes as a result of this
process, the wall appearing to recover the original alignment
of CMFs. The most likely explanation for the swirl structure
is that it represents the region of rhizoid or aplanospore formation, as described by Preston and Astbury [26].
Identification of polymers is difficult using AFM alone
and accompanying methods are necessary. While AFM is a
powerful tool for unravelling structures in both the native
state and fixed state, the identification of the polymers in a
sample specimen is not possible. In cell wall imaging, the
CMFs are obvious structures due to their abundance and
appearance (long and thin) and well-known layout. The
other structures seen in the AFM images of the cell wall have
yet to be identified. AFM force measurements were conducted as an aid to obtain preliminary identification of the
unknown cell wall components seen in our study. In particular, this method was used to aid in distinguishing between
sulphated polysaccharide mucilage and pectin gel in the
native wall (data not shown). Sequential extraction of polymers from the cell wall is often conducted to gain insight into
the structures of cell wall components [40,43,45]. Measurements of CMF sizes before and after sequential extraction of
certain targeted polymers would give an indication of the
polymers’ association with the CMFs.
8
9
E.M. Eslick et al. AFM imaging of Ventricaria cell wall, 2014, Vol. 0, No. 0
25. Sponsler O L (1930) Orientation of cellulose space lattice in the
cell wall. Additional X-ray data from Valonia cell wall.
Protoplasma 12: 241–254.
26. Preston R D, Astbury W T (1937) Structure of cell wall of the
green algae Valonia ventricosa. Proc. R. Soc. Lond. B Biol. Sci.
122: 76–97.
27. Preston R D, Kuyper B (1951) Electron microscopic investigations of the walls of green algae I. A preliminary account of wall
lamellation and deposition in valonia ventricosa. J. Exp. Bot. 2:
247–255.
28. Steward F C, Muhlethaler K (1953) The structure and development of the cell-wall in the valoniaceae as revealed by the electron
microscope. Ann. Bot. (Lond.) 17: 295–316.
29. Cronshaw J, Myers A, Preston R D (1958) A chemical and physical investigation of the cell walls of some marine algae. Biochim.
Biophys. Acta 27: 89–103.
30. Blackwell J, Vasko P D, Koenig J L (1970) Infrared and raman
spectra of cellulose from cell wall of valonia ventricosa. J. Appl.
Phys. 41: 4375–4379.
31. Hanley S J, Giasson J, Revol J F, Gray D G (1992) Atomic force
microscopy of cellulose microfibrils: comparison with transmission electron microscopy. Polymer 33: 4639–4642.
32. Kuutti L, Peltonen J, Pere J, Teleman O (1995) Identification and
surface-structure of crystalline cellulose studied by atomic-force
microscopy. J. Microsc. (Oxford) 178: 1–6.
33. Baker A A, Helbert W, Sugiyama J, Miles M J (1997) Highresolution atomic force microscopy of native Valonia cellulose I
microcrystals. J. Struct. Biol. 119: 129–138.
34. Baker A A, Helbert W, Sugiyama J, Miles M J (2000) New
insight into cellulose structure by atomic force microscopy shows
the I? Crystal phase at near-atomic resolution. Biophys. J. 79:
1139–1145.
35. Hobbs J K, Winkel A K, Mcmasster T J, Humphris A D L, Baker A
A, Blakely S, Aissaoui M, Miles M J (2001) Some recent developments in SPM of crystalline polymers. Macromol. Symposia 167:
1–14.
36. Preston R D (1974) Detailed Structure—Cellulosic Algae.
Physical Biology of Plant Cell Walls, pp. 192–237 (Chapman
and Hall, London).
37. Goto T, Harada H, Saiki H (1973) Cross-sectional view of microfibrils in Valonia (Valonia macrophysa). Mokuzai Gakkaishi 19:
463–468.
38. Tsekos I (1999) The sites of cellulose synthesis in algae:
diversity and evolution of cellulose-synthesizing enzyme complexes. J. Phycol. 35: 635–655.
39. Kirby A R, Gunning A P, Waldron K W, Morris V J, Ng A (1996)
Visualisation of plant cell walls by atomic force microscopy.
Biophys. J. 70: 1138–1143.
40. Davies L M, Harris P J (2003) Atomic force microscopy of microfibrils in primary cell walls. Planta 217: 283–289.
41. Marga F, Grandbois M, Cosgrove D J, Baskin T I (2005) Cell
wall extension results in the coordinate separation of parallel
microfibrils: evidence from scanning electron microscopy and
atomic force microscopy. Plant J. 43: 181–190.
42. Ding S Y, Himmel M E (2006) The maize primary cell wall
microfibril: a new model derived from direct visualization.
J. Agric. Food Chem. 54: 597–606.
Downloaded from http://jmicro.oxfordjournals.org/ at University of New South Wales on April 2, 2014
7. Satiatjeunemaitre B, Martin B, Hawes C (1992) Plant-cell wall
architecture is revealed by rapid-freezing and deep-etching.
Protoplasma 167: 33–42.
8. Mccann M C, Wells B, Roberts K (1990) Direct visualization of
cross-links in the primary plant-cell wall. J. Cell Sci. 96:
323–334.
9. Fujino T, Sone Y, Mitsuishi Y, Itoh T (2000) Characterization of
cross-links between cellulose microfibrils, and their occurrence
during elongation growth in pea epicotyl. Plant Cell Physiol. 41:
486–494.
10. Knox J P (1992) Molecular probes for the plant-cell surface.
Protoplasma 167: 1–9.
11. Binnig G, Quate C F, Gerber C (1986) Atomic force microscope.
Phys. Rev. Lett. 56: 930–933.
12. Drake B, Prater C B, Weisenhorn A L, Gould S A C, Albrecht T R,
Quate C F, Cannell D S, Hansma H G, Hansma P K (1989)
Imaging crystals, polymers, and process. Science 243:1586–1589.
13. Hansma H G, Hoh J H (1994) Biomolecular imaging with the
atomic-force microscope. Annu. Rev. Biophys. Biomol. Struct.
23: 115–139.
14. You H, Yu L (1999) Atomic force microscopy imaging of living
cells: progress, problems and prospects. Methods Cell Sci. 21:
1–17.
15. Fotiadis D, Scheuring S, Muller S A, Engel A, Muller D J (2002)
Imaging and manipulation of biological structures with the AFM.
Micron 33: 385–397.
16. Butt H, Cappella B, Kappl M (2005) Force measurements with
the atomic force microscope: technique, interpretation and applications. Surf. Sci. Rep. 59: 1–152.
17. Lal R, John S A (1994) Biological applications of atomic-force
microscopy. Am. J. Physiol. 266: C1–C21.
18. Shao Z, Mou J, Czajkowsky D, Yang J, Yuan J (1996) Biological
atomic force microscopy: what is achieved and what is needed.
Adv. Phys. 45: 1–86.
19. Engel A, Muller D J (2000) Observing single biomolecules at
work with the atomic force microscope. Nat. Struct. Biol. 7:
715–718.
20. Bisson M A, Beilby M J, Shepherd V A (2006) Electrophysiology
of turgor regulation in marine siphonous green algae. J. Membr.
Biol. 211: 1–14.
21. Ryser C, Wang J, Mimietz S, Zimmermann U (1999) Determination of the individual electrical and transport properties of the
plasmalemma and the tonoplast of the giant marine alga Ventricaria ventricosa by means of the integrated perfusion/charge-pulse
technique: evidence for a multifolded tonoplast. J. Membrane Biol.
168: 183–197
22. Hicks G R, Hironaka C M, Dauvillee D, Funke R P, D’hulst C,
Waffenschmidt S, Ball S G (2001) When simpler is better. Unicellular green algae for discovering new genes and functions in
carbohydrate metabolism. Plant Physiol. 127: 1334–1338.
23. Shepherd V A, Beilby M J, Bisson M A (2004) When is a cell not
a cell? A theory relating coenocytic structure to the unusual electrophysiology of Ventricaria ventricosa (Valonia ventricosa).
Protoplasma 223: 79–91.
24. Olsen J L, West J A (1988) Ventricaria (siphonocladales-cladophorales complex, chlorophyta), a new genus for Valonia ventricosa. Phycologia 27: 103–108.
Microscopy, 2014, Vol. 0, No. 0
61.
62.
63.
64.
65.
66.
67.
68.
69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
79.
80.
charophyte alga lamprothamnium papulosum. J. Membr. Biol.
170: 229.
Chiovitti A, Higgins M J, Harper R E, Wetherbee R, Bacic A
(2003) The complex polysaccharides of the raphid diatom pinnularia viridis (bacillariophyceae). J. Phycol. 39: 54.
Revol J F (1982) On the cross sectional shape of cellulose crystallites in valonia ventricosa. Carbohydr. Polym. 2: 123–134.
Itoh T, Brown R M (1984) The assembly of cellulose microfibrils
in valonia-macrophysa kütz. Planta 160: 372–381.
Sugiyama J, Harada H, Fujiyoshi Y, Uyeda N (1985) Lattice
images from ultrathin sections of cellulose microfibrils in the cell
wall of valonia macrophysa kutz. Planta 166: 161.
Long C (1968) The cell walls of algae. In: Rogers H J,
Perkins H R (eds), Cell Walls and Membranes. Spon’s Biochemical Monographs, pp. 114–134 ( E. & F. N. Spon, London).
Chanliaud E, Gidley M J (1999) In vitro synthesis and properties
of pectin/acetobacter xylinus cellulose composites. Plant J. 20:
25–35.
O’Neill M, York W (2003) The composition and structure of
plant primary cell walls. In: Rose J (ed.), The Plant Cell Wall, 1st
edn (Blackwell Publishing/CRC, Oxford/Boca Raton, FL).
Obel N, Neumetzler L, Pauly M (2007) Hemicelluloses and cell
expansion. In: Vissenberg J. P. V. A. K. (ed.), In the Expanding
Cell, pp. 57–88 (Springer, Berlin).
Morris S, Hanna S, Miles M J (2004) The self-assembly of
plant cell wall components by single-molecule force
spectroscopy and Monte Carlo modelling. Nanotechnology 15:
1296–1301.
Rose J K C, Bennett A B (1999) Cooperative disassembly of the
cellulose–xyloglucan network of plant cell walls: parallels
between cell expansion and fruit ripening. Trends Plant Sci. 4:
176–183.
Zhou Q, Rutland M W, Teeri T T, Brumer H (2007) Xyloglucan
in cellulose modification. Cellulose 14: 625–641.
Lahaye M, Jegou D, Buleon A (1994) Chemical characteristics
of insoluble glucans from the cell-wall of the marine green-alga
ulva-lactuca (l) thuret. Carbohydrate Res. 262: 115–125.
Lahaye M, Ray B (1996) Cell-wall polysaccharides from the
marine green alga ulva ‘‘rigida’’ (ulvales, chlorophyta) - NMR
analysis of ulvan oligosaccharides. Carbohydrate Res. 283: 16.
Popper Z A, Fry S C (2003) Primary cell wall composition of
bryophytes and charophytes. Ann. Bot. (Lond.) 91: 1–12.
Whitney S E C, Brigham J E, Darke A H, Reid J S G, Gidley M J
(1998) Structural aspects of the interaction of mannan-based
polysaccharides with bacterial cellulose. Carbohydrate Res. 307:
299–309.
Tepfer M, Cleland R E (1979) Comparison of acid-induced cellwall loosening in Valonia ventricosa and in oat coleoptiles. Plant
Physiol. 63: 898–902.
Gardner K H, Blackwell J (1971) The substructure of the cellulose microfibrils from the cell walls of the algae valonia ventricosa. J. Ultrastruct. Res. 36: 725–731.
Fritsch F E (1945) The Structure and Reproduction of the Algae,
791 pp (Cambridge University Press, Cambridge, England).
Hayashi T (1989) Xyloglucans in the primary-cell wall. Annu.
Rev. Plant Physiol. Plant Mol. Biol. 40: 139–168.
Fry S C (1989) Cellulases, hemicelluloses and auxin-stimulated
growth—a possible relationship Physiol. Plant 75: 532–536.
Downloaded from http://jmicro.oxfordjournals.org/ at University of New South Wales on April 2, 2014
43. Kirby A R, Ng A, Waldron K W, Morris V J (2006) AFM investigations of cellulose fibers in bintje potato (Solanum tuberosum
L.) cell wall fragments. Food Biophys. 1: 163–167.
44. Thimm J C, Burritt D J, Ducker W A, Melton L D (2000) Celery
(Apium graveolens L.) parenchyma cell walls examined by atomic
force microscopy: effect of dehydration on cellulose microfibrills.
Planta 212: 25–32.
45. Mine I, Okuda K (2007) Fine structure of cell wall surfaces in the
giant-cellular xanthophycean alga Vaucheria terrestris. Planta
225: 1135–1146.
46. Berges J A, Franklin D J, Harrison P J (2001) Evolution of an artificial seawater medium: improvements in enriched seawater, artificial water over the last two decades. J. Phycol. 37: 1138–1145.
47. Bisson M A, Beilby M J (2002) The transport systems of Ventricaria ventricosa: hypotonic and hypertonic tugor regulation.
J. Membr. Biol. 190: 43–56.
48. Heidecker M, Wegner L H, Zimmermann U (1999) A patchclamp study of ion channels in protoplasts prepared from the
marine alga Valonia utricularis. J. Membr. Biol. 172: 235–247.
49. Kennedy B F, De Filippis L F (2004) Tissue degradation and
enzymatic activity observed during protoplast isolation in two
ornamental Grevillea species. In Vitro Cell Dev. Biol. Plant 40:
119–125.
50. Haberle W, Horber J K H, Ohnesorge F, Smith D P E, Binnig G
(1992) In situ investigations of single living cells infected by
viruses. Ultramicroscopy 42: 1161–1167.
51. Rotsch C, Braet F, Wisse E, Radmacher M (1997) AFM imaging
and elasticity measurements on living rat liver macrophages. Cell
Biol. Int. 21: 685–696.
52. Schaus S S, Henderson E R (1997) Cell viability and probe-cell
membrane interactions of xr1 glial cells imaged by atomic force
microscopy. Biophys. J. 73: 1205–1214.
53. Le Grimellec C, Lesniewska E, Giocondi M C, Finot E, Vie V,
Goudonnet J P (1998) Imaging of the surface of living cells by
low-force contact-mode atomic force microscopy. Biophys. J. 75:
695–703.
54. Zhong Q, Inniss D, Kjoller K, Elings V B (1993) Fractured
polymer silica fiber surface studied by tapping mode atomic-force
microscopy. Surf. Sci. 290: L688–LL92.
55. Putman C A J, Vanderwerf K O, Degrooth B G, Vanhulst N F,
Greve J (1994) Viscoelasticity of living cells allows highresolution imaging by tapping mode atomic-force microscopy.
Biophys. J. 67: 1749–1753.
56. Davis J J, Hill H A O, Powell T (2001) High resolution scanning
force microscopy of cardiac myocytes. Cell Biol. Int. 25:
1271–1277.
57. Putman C A J, Vanderwerf K O, Degrooth B G, Vanhulst N F,
Greve J, Hansma P K (1992) A new imaging mode in atomic
force microscopy based on the error signal. In: Proceedings of the
SPIE—International Society of Optical Engineering, on Scanning
Probe Microscopies, Bellingham, pp. 198–204.
58. Morris V J, Ring S G, Macdougall A J, Wilson R H (2003) Biophysical characterization of plant cell walls. In: Rose J (ed.), The
Plant Cell Wall, 1st edn (Blackwell Publishing, USA).
59. Kirst G O (1989) Salinity tolerance of eukaryotic marine algae.
Annu. Rev. Plant Physiol. Plant Mol. Biol. 40: 21–53.
60. Shepherd V A, Beilby M J (1999) The effect of an extracellular mucilage on the response to osmotic shock in the
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