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How the deposition of
cellulose microfibrils builds
cell wall architecture
Anne Mie C. Emons and Bela M. Mulder
Cell walls, the extracytoplasmic matrices of plant cells, consist of an ordered
array of cellulose microfibrils embedded in a matrix of polysaccharides and
glyco-proteins. This construction is reminiscent of steel rods in reinforced
concrete. How a cell organizes these ordered textures around itself, creating its
own desirable environment, is a fascinating question. We believe that nature
adopted an economical solution to this design problem: it exploits the
geometrical constraints imposed by the shape of the cell and the limited space
in which microfibrils are deposited, enabling the wall textures essentially to ‘build
themselves’. This does not imply that the cell cannot control its wall texture. On
the contrary, the cell has ample regulatory mechanisms to control wall texture
formation by controlling the insertion of synthases and the distance between
individual microfibrils within a wall lamella.
E
very year, plants produce 180 billion
tons of cellulose1, which has enormous commercial value as wood,
paper and cotton. Cellulose is produced in the
form of cellulose microfibrils, (semi)crystalline aggregates of linear polymers
of D-glucopyranosyl residues2–4, linked in
the b-(1→4) conformation. The pattern
formed by these cellulose microfibrils in
the walls is cell type- and developmental
stage-specific, and is a major determinant of
wall properties.
Hypotheses for cell wall texture
formation
Cortical microtubules direct the individual
cellulose microfibrils during their synthesis:
the textbook hypothesis
In elongating plant cells, the cortical microtubules, as well as the cellulose microfibrils
that are being deposited, are both transverse to
the elongation direction of the cells8. This correlation has led to the hypothesis that cortical
microtubules determine the orientation of
the cellulose microfibrils9 by stiffening the
plasma membrane, forming corridors for the
cellulose synthases locally10. The credibility
of this hypothesis is based on experiments in
which microtubule drugs change the orientation of the cellulose microfibrils11 (reviewed
in Ref. 12). However, at most of the concentrations tested, these drugs also stop unidirectional cell elongation. The conclusion of the
drug experiments should be that cortical
microtubules determine the elongation orientation of anisotropically expanding cells. The
microtubule depolymerizing drugs in many12
(but not all13) cases also change the deposition
orientation of the cellulose microfibrils. From
such data it was concluded that the cortical
microtubule orientation determines the deposition orientation of the cellulose microfibrils.
However, because a second parameter appears
to be changed by the drug application, it could
be concluded that cell elongation orientation
determines microfibril deposition orientation.
Therefore the effect that cortical microtubules
have on nascent cellulose microfibrils should
be studied in fully expanded cells. In the fully
expanded cells that have been studied, cortical microtubules and the most recently
deposited cellulose microfibrils are not generally in parallel alignment14. Mutants disturbed
in either cortical microtubule alignment or cellulose microfibril alignment during deposition
are important for determining the role that the
microtubules play in microfibril deposition.
Membrane flow determines cellulose
microfibril deposition orientation
Plant scientists often take the microtubule–
microfibril paradigm for granted, without even
studying the cellulose microfibrils properly14.
Cellulose synthases in the plasma
membrane
Golgi vesicles carrying wall matrix material
as cargo and cellulose synthases in their membranes, deposit these into the wall and the
plasma membrane, respectively, as a result of
exocytosis. Freeze-fractured plasma membranes show these cellulose synthases, which
consist of six particles arranged in a ring5
(Fig. 1), a so-called particle rosette. These
rosettes spin out the cellulose polymers6,7,
which crystallize as bundles, the cellulose
microfibrils. The polymerization forces cause
the synthases to move in the plasma
membrane. A functional model for cellulose
synthase has been hypothesized (Fig. 1), in
which the particles of the rosette spin out the
cellulose polymers.
Types of cell wall textures
The following cell wall textures are known:
axial, transverse, crossed, helical, helicoidal and
random (Fig. 2). Other types of texture are derived from these basic types by the successive
deposition of different textures.
Fig. 1. (a) Particle rosette, the cellulose synthase, in the plasma membrane of an
Equisetum hyemale root hair. The rosette has a diameter of 25 nm and consists of six particles.
In this cell type, the rosette density is relatively low, 5–15 rosettes/mm22, which agrees
with the large distance between individual microfibrils in the last deposited lamella6. (b)
Hypothesis for a functional model of a particle rosette (adapted from Ref. 6 ). The six proteins constituting the particle rosette (PR) [seen in the plasmatic-fracture (PF) face] form
channels in the plasma membrane. Through these channels UDP-glucose passes while
being polymerized into cellulose molecules. The terminal globule (TG) (sometimes seen
in the exoplasmic (EF)-fracture face) could be the still uncrystallized cellulose polymers.
1360 - 1385/00/$ – see front matter © 2000 Elsevier Science Ltd. All rights reserved. PII: S1360-1385(99)01507-1
January 2000, Vol. 5, No. 1
35
trends in plant science
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are strong counter-arguments against this
hypothesis:
• The cellulose microfibrils have dimensions
(both in linear size and mass) that put them
outside the colloidal domain, thus precluding their equilibration on reasonable
timescales19,20.
• The available space between the plasma
membrane and the already extant primary
cell wall is so limited that this system would be dominated by boundary
effects. In most cases these effects would
disturb the equilibrium configuration of
the particles involved, either by promoting boundary layers of different
structure, or anchoring defects or the
imposition of antagonistic geometrical
constraints21.
• The wall matrix material is probably
more like a dense gel than a thermal solvent and would thus also be an obstacle to
equlibration22.
Fig. 2. Basic wall textures: axial, transverse, crossed, helical, helicoidal and
random. Successive wall lamellae have
been peeled off from top to bottom
showing the random wall texture of the
outermost wall at the base.
However, scientists studying the cell wall
have come up with several alternative
hypotheses for the orientation mechanism of
cellulose mirofibrils. One such alternative is
the membrane flow hypothesis15. According
to this hypothesis the particle rosettes are free
to move in the plane of the viscous medium of
the plasma membrane, and are pushed forward
by synthesis and crystallization forces of the
cellulose microfibril. The first part of this
hypothesis is generally accepted and is also
used by us. But the hypothesis also suggests
that the rotation of the protoplast relative to the
cell wall is the orienting principle. However,
this rotation has never been observed in plants.
Cell wall texture formation is a selfassembly process like liquid crystal
formation
Scientists faced with helicoidal cell walls in
which the microtubules and the cellulose
microfibrils did not match, realized that the
helicoidal walls resemble cholesteric liquid
crystals, which are known to self-assemble
spontaneously16,17. However, this type of equilibrium assembly can only take place when a
sufficient amount of bulk material is present.
There is no evidence that this condition is ever
met because we now know that cellulose
microfibrils are made by plasma membraneembedded synthase complexes in a sequential
growth process18 (reviewed in Ref. 7).
Moreover, from a physics point of view, there
36
January 2000, Vol. 5, No. 1
The geometrical theory
We have proposed a geometrical model
for cellulose microfibril deposition23–26 . The
model quantitatively relates the deposition
angle of the cellulose microfibrils (with
respect to the cell axis) to:
• The density of active synthases in the
plasma membrane.
• The distance between individual microfibrils within a wall lamella.
• The geometry of the cell.
The helicoidal wall texture of root hairs
Inspiration for the geometrical model came
from observations on the secondary wall of
root hairs of Equisetum hyemale. A secondary
wall is deposited in a cell or part of a cell
that does not expand. Thus, at any stage of
Fig. 3. Schematic showing a section
through the consecutive lamellae of a
developing (from the lower to the upper
part of the diagram) helicoidal cell wall.
The already deposited consecutive lamellae are on the left; the most recently
deposited lamella is on the right. The
lengths of the areas with a certain microfibril orientation are much longer than can be
shown in a drawing; therefore, black horizontal lines have been included between
adjacent areas. Both transverse sections
through the wall and surface preparations
show a record of the deposition process in
time. In Equisetum hyemale root hairs, the
length of such an area is ~300 mm (A.M.C.
Emons, unpublished).
Fig. 4. (a) Shadowcast preparation of the helicoidal cell wall texture of an Equisetum hyemale root
hair showing the cellulose microfibrils in three subsequent orientations of the helicoid. A cellulose
microfibril with platinum/carbon shadow deposit is 7–8 nm wide. Scale bar 5 500 nm. (b) Thin section through the cell wall of a root hair of E. hyemale showing the arcs, which are visual artifacts of
the helicoidal texture. A negatively stained cellulose microfibril is 3–4 nm wide. Scale bar 5 500 nm
trends in plant science
Perspectives
its deposition, the secondary wall is a record
of the wall deposition process. The root hair
is a tip-growing cell with primary wall deposition at the growing hair tip and secondary
wall deposition in the shank of the hair.
Root hairs, including those from E. hyemale,
have particle rosettes, the cellulose synthases
(Fig. 1), in the plasma membrane27. The
E. hyemale root hairs have helicoidal cell
walls and the cortical microtubules do not
align with the nascent cellulose microfibrils28–30. A helicoidal cell wall texture (Fig. 2)
consists of one microfibril thick lamellae.
The orientation of the cellulose microfibrils in a lamella makes a constant angle
with the orientation in the subsequent lamella
(Fig. 3). In sections, this gives rise to the optical illusion of arcs (Fig. 4).
Facts and assumptions of the
geometrical model
Cellulose microfibrils are long structures. No
reliable estimate of their length is available
to date because microfibril ends are seen
rarely in wall preparations. An individual
cellulose microfibril can, therefore, contribute to several wall lamellae. Freeze-fracture images show that the cell wall is
appressed against the plasma membrane.
Therefore, wall deposition occurs under
space-limiting conditions.
Consider a cylinder around which several
tapes are being wound, such that the whole
cylinder surface is covered with one single
layer of tape (Fig. 5). The angle that the
tape makes with the long axis of the cylinder
is completely described by the following
formula:
sine a 5
number of tapes N 3 tape width d
p 3 cell diameter D
(1)
where a is the fibril angle with the cell’s long
axis, D denotes the cell diameter, N the
number of active synthases in the plasma
membrane and d is the distance between individual microfibrils within a lamella in a plant
cell (Fig. 5). Therefore, d depends on
the quantity of matrix between cellulose
microfibrils31 and, thus, on the ratio of cellulose to matrix. In an E. hyemale root hair
plasma membrane, this distance is ~150 nm
(Ref. 6).
Because a texture can be described fully by
describing the angle every cellulose microfibril makes with the cell axis during its deposition, the dependencies described in Equation
1 give us the basis of the geometrical theory
of cell wall texture formation.
Now that we can describe how a synthase
moves, we need to consider these questions:
where and at what rate are these synthases
activated? The important assumption of our
model is that this activation takes place in
Fig. 5. (a) The geometrical rule for cellulose microfibril deposition follows from the notion
of winding tapes densely around a cylinder, as is illustrated by unrolling the cylinder into a
planar figure, where D denotes the cell diameter, N the number of active synthases in the
plasma membrane and d is the distance between individual microfibrils within a lamella in a
plant cell. (b) Schematic of a cellulose microfibril surrounded by a wall matrix. The distance
between two microfibrils is the width of one tape, d, used in the model. In Equisetum
hyemale, root hair plasma membrane width is ~150 nm. (c) The influence of changing
cellular parameters on the microfibril winding-angle as predicted by the geometrical theory.
(i) Changing the relative number of microfibrils expressed as a fraction of the maximum
Nmax 5 pD/d; (ii) changing the relative effective width of the microfibrils expressed as a
fraction of the maximum possible width dmax 5 pD/N; (iii) changing the relative diameter of
the cell expressed in units of the minimum diameter Dmin 5 Nd/p.
January 2000, Vol. 5, No. 1
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trends in plant science
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Fig. 6. (a) Insertion domain. The assumption of the geometrical model is that synthase activation
takes place in localized mobile insertion domains. An insertion domain consists of a plasma membrane area in which Golgi vesicles, containing the synthases in their membranes, are inserted into
the plasma membrane. (b) This insertion domain itself, or the activation of the synthases in the
domain, moves with a velocity v, while the microfibrils are spun from the synthases in the opposite direction. t 5 time; t† 5 time interval; w 5 the speed with which the synthase moves.
Box 1. The synthase density evolution equation
The equation that describes the evolution of active cellulose microfibril synthase density on
a perfectly cylindrical cell of radius R, is given by
∂N ( z, t )
∂N ( z, t )
wd
−
= w ( N , z, t ) − w † ( N , z, t )
N ( z, t )
∂t
∂z
2pR
where w is the speed with which the synthase moves, d is the effective width of a cellulose
microfibril plus adherent matrix material and N(z,t) is synthase density. w is the local rate of
synthase production for which we choose the following form
w ( N , z, t ) =
N∗
N ( z, t )
1 −
t∗ (1 − g )
N∗
g
if N (z,t),N* and z is
located inside an insertion domain. In all other cases w 5 0. The param-eter g controls the
shape of the synthase production curve and lies between 0 and 1. Synthase production stops
when the maximum density
2pR
N∗ =
d
is reached, which for stationary insertion domains would happen after time t* . The insertion
domains are assumed to have a length l and travel at a speed v. Finally, the local rate of
rosette de-activation w✝ needs to be determined. This rate depends on the full evolution of the
density in a time interval of length t✝ (the synthase lifetime). Fortunately, the resultant equations are of a type that can be readily solved with entirely classical techniques. At first sight
the model appears to have many parameters, but dimensional analysis reveals that there are
only four relevant ones:
l
l =
wt∗
• Length of the insertion domain
•
Speed of the insertion domain
•
Synthase lifetime
n
w
t†
t† =
t∗
b =
g
38
January 2000, Vol. 5, No. 1
Fig. 7. Visualization of the solution of
our geometrical model in a helicoidal
example. For clarity, only a small number of cellulose microfibrils are shown,
and have been drawn at a highly
expanded scale. Four ‘generations’ of
cellulose microfibrils are spun by synthases that were activated at the tip of an
insertion domain at four equally spaced
moments in time. The displacement of
the starting points along the cell is a
consequence of the motion of the insertion domain. Each of these generations
contributes locally to a single lamella.
The inset shows how the angle of deposition between adjacent lamellae is
nearly constant, the hallmark of helicoidal texture.
localized mobile insertion domains. An insertion domain consists of a plasma membrane
area in which Golgi vesicles, containing the
synthases in their membranes32, are inserted
into the plasma membrane (i.e. a domain
where exocytosis occurs; Fig. 6). For insertion domain mobility, at least two experimentally verifiable scenarios are possible:
• The Golgi bodies that produce the Golgi
vesicle-containing synthases move along
the plasma membrane while inserting the
vesicles.
• The Golgi bodies themselves are stationary, but exocytosis of their vesicles is
locally controlled by a time-varying
concentration of an activating–inhibiting
substance.
It is possible that calcium ions fulfil this role.
We believe that the activation of exocytosis in
root hairs involves calcium33,34, and calcium
ion concentration oscillations have been
observed in root hairs upon stimulation35.
trends in plant science
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The mathematical model
We can formulate a model that describes the
whole process of cell wall texture formation
using the elements described for the geometrical model. The theory describes the evolution of active synthase density in the plasma
membrane in space and time. If the number of
active synthases present at a given location in
the plasma membrane is known, the geometrical rule predicts the orientation of the cellulose microfibrils that are deposited. In a sense,
each microfibril ‘records’ the motion of its
synthase. The totality of all these ‘recordings’
creates the cell wall texture. To formulate such
a model, attention needs to be focused on a
small segment of the plasma membrane. How
does the number of active synthases in the portion of plasma membrane belonging to this
segment change in a small interval of time?
There are three sources of change:
• Movement of synthases into and out of the
segment.
• Activation of new synthases by an insertion
domain.
• De-activation of synthases after a finite life
span.
Mathematically, this yields a partial differential equation for synthase density N (z,t ),
which can be solved as a function of position
z along the cell and time t (Box 1).
The Helicoidal solution
The solution for a helicoidal cell wall demonstrates what the geometrical model can do
(Fig. 7). In such a wall, the microfibril orientation changes by a constant angle from one
lamella to another (Fig. 2), producing a staircase-like structure (hence the name helicoid).
To obtain such a structure in which the deposition angle progresses from transversal (a 5
08), through axial (a 5 908) and back to transversal (a 5 1808), after which the cycle can
repeat itself, the parameters of the model need
to be precisely tuned. In our view, the geometrical model is sufficient to explain the
most complicated cell wall texture. However,
its general utility is its ability to describe the
various other wall textures also seen in plant
cells (Fig. 2).
Utility of the model
Numerous cell wall examples that support
the geometrical model have been discussed
before23–26. The important assumption to be
verified is the movement or activation of the
insertion domains. We have proposed two scenarios for local, sequential exocytosis: the
movement of the Golgi vesicle-producing
Golgi bodies along the plasma membrane
while inserting the vesicles, and the local calcium-induced exocytosis of vesicles along the
plasma membrane. Other properties of the
insertion domain that need to be determined
and manipulated include its length and speed.
Furthermore, to date we know nothing about
the synthase life-time. The use of mutants,
GFP-technology and calcium measurements
of high sensitivity along the plasma membrane
of cells with known wall deposition patterns,
should provide an appropriate series of experiments for studying the exocytosis machinery
in living cells in relation to wall deposition.
Acknowledgements
We thank Allex Haasdijk for the artwork in
Figures 1(b), 2, 3 and 6(a), and Ton Bisseling
for critically reading the manuscript.
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Does a light-harvesting
protochlorophyllide a/bbinding protein complex exist?
Gregory A. Armstrong, Klaus Apel and Wolfhart Rüdiger
Recent in vitro studies have led to speculation that a novel light-harvesting
protochlorophyllide a/b-binding protein complex (LHPP) might exist in dark-grown
angiosperms. Structurally, it has been suggested that LHPP consists of a 5:1 ratio
of dark-stable ternary complexes of the light-dependent NADPH: protochlorophyllide
oxidoreductases A and B containing nonphotoactive protochlorophyllide b and
photoactive protochlorophyllide a, respectively. Functionally, LHPP has been
hypothesized to play major roles in establishing the photosynthetic apparatus, in
protecting against photo-oxidative damage during greening, and in determining
etioplast inner membrane architecture. However, the LHPP model is not compatible
with other studies of the pigments and the pigment–protein complexes of dark-grown
angiosperms. Protochlorophyllide b, which is postulated to be the major lightharvesting pigment of LHPP, has, for example, never been detected in etiolated
seedlings. This raises the question: does LHPP exist?
L
ight profoundly influences plant
development and allows photosynthesis to occur, but it also represents a
tremendous risk. Photo-oxidative damage
initiated by excited state photosensitizing
molecules, such as chlorophylls and their
biosynthetic precursors, can be lethal.
Angiosperms that germinate in darkness in
the soil enter the seedling developmental program known as skotomorphogenesis (Fig. 1).
However, such seedlings must be prepared
for a subsequent light-triggered switch to
photomorphogenesis. Upon illumination, the
leaves of etiolated angiosperms synthesize
and accumulate large quantities of chloro40
January 2000, Vol. 5, No. 1
phylls a and b. Seedlings are particularly susceptible to photo-oxidative damage during
this transition to photoautotrophy.
The presence or absence of light dramatically
influences plastid development. Dark-grown
angiosperm seedlings contain an achlorophyllous plastid type known as the etioplast, which
is transformed into a photosynthetically competent chloroplast during photomorphogenesis1.
The etioplast is defined by the presence of two
types of internal membranes, the lattice-like
prolamellar body, which is composed of interconnected tubules, and the unstacked prothylakoids. Etioplasts characteristically accumulate
the chlorophyll precursor protochlorophyllide
Anne Mie C. Emons is at the Laboratory of
Experimental Plant Morphology and Cell
Biology, Dept of Plant Sciences,
Wageningen University, Arboretumlaan 4,
6703 BD Wageningen, The Netherlands
(tel 131 317 484329;
fax 131 317 485005;
e-mail [email protected]);
Bela M. Mulder is at the Condensed Matter
Division of the FOM Institute for Atomic and
Molecular Physics, Kruislaan 407, 1098 SJ
Amsterdam, The Netherlands
(tel 131 20 6081231;
fax 131 20 6684106;
e-mail [email protected]).
(Pchlide), more specifically protochlorophyllide
a (Pchlide a)2–4. Illumination of etioplasts initiates the dispersal of the prolamellar body and
the formation of thylakoid membranes containing the pigment–protein complexes of the
photosynthetic apparatus.
In this context, recent in vitro reconstitution
experiments have been interpreted as providing
evidence for a novel light-harvesting Pchlide
a/b-binding protein complex5, termed LHPP
by analogy to the ubiquitous light-harvesting
chlorophyll a/b-binding proteins (LHCP) of
green plants. LHPP is speculated to:
• Serve as the central structural determinant
of the prolamellar body in etioplasts.
• Be essential for the establishment of the
photosynthetic apparatus.
• Confer photoprotection on greening
seedlings by dissipating excess light
energy, thereby minimizing Pchlideinduced photo-oxidative damage.
On the one hand, if they are correct, these
hypotheses would have a major impact on our
understanding of the seedling transition from
skotomorphogenesis to photomorphogenesis.
On the other hand, to date there are no in vivo
data that directly support the existence of an
LHPP complex6. Here, we critically analyse
the LHPP model in light of the current literature on the properties of etioplast membranes,
pigment–protein complexes and pigments.
Roles of the light-dependent PORA and
PORB proteins in etioplast formation
and photo-oxidative protection
The presence of the prolamellar body and the
accumulation of Pchlide a in etioplasts are
known to correlate with large quantities of the
strictly light-dependent NADPH:protochlorophyllide oxidoreductase (POR; 1.3.1.33)1,7–9.
This nuclear-encoded but plastid-localized
protein is unusual in that it mediates the only
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Perspectives
How the deposition of
cellulose microfibrils builds
cell wall architecture
Anne Mie C. Emons and Bela M. Mulder
Cell walls, the extracytoplasmic matrices of plant cells, consist of an ordered
array of cellulose microfibrils embedded in a matrix of polysaccharides and
glyco-proteins. This construction is reminiscent of steel rods in reinforced
concrete. How a cell organizes these ordered textures around itself, creating its
own desirable environment, is a fascinating question. We believe that nature
adopted an economical solution to this design problem: it exploits the
geometrical constraints imposed by the shape of the cell and the limited space
in which microfibrils are deposited, enabling the wall textures essentially to ‘build
themselves’. This does not imply that the cell cannot control its wall texture. On
the contrary, the cell has ample regulatory mechanisms to control wall texture
formation by controlling the insertion of synthases and the distance between
individual microfibrils within a wall lamella.
E
very year, plants produce 180 billion
tons of cellulose1, which has enormous commercial value as wood,
paper and cotton. Cellulose is produced in the
form of cellulose microfibrils, (semi)crystalline aggregates of linear polymers
of D-glucopyranosyl residues2–4, linked in
the b-(1→4) conformation. The pattern
formed by these cellulose microfibrils in
the walls is cell type- and developmental
stage-specific, and is a major determinant of
wall properties.
Hypotheses for cell wall texture
formation
Cortical microtubules direct the individual
cellulose microfibrils during their synthesis:
the textbook hypothesis
In elongating plant cells, the cortical microtubules, as well as the cellulose microfibrils
that are being deposited, are both transverse to
the elongation direction of the cells8. This correlation has led to the hypothesis that cortical
microtubules determine the orientation of
the cellulose microfibrils9 by stiffening the
plasma membrane, forming corridors for the
cellulose synthases locally10. The credibility
of this hypothesis is based on experiments in
which microtubule drugs change the orientation of the cellulose microfibrils11 (reviewed
in Ref. 12). However, at most of the concentrations tested, these drugs also stop unidirectional cell elongation. The conclusion of the
drug experiments should be that cortical
microtubules determine the elongation orientation of anisotropically expanding cells. The
microtubule depolymerizing drugs in many12
(but not all13) cases also change the deposition
orientation of the cellulose microfibrils. From
such data it was concluded that the cortical
microtubule orientation determines the deposition orientation of the cellulose microfibrils.
However, because a second parameter appears
to be changed by the drug application, it could
be concluded that cell elongation orientation
determines microfibril deposition orientation.
Therefore the effect that cortical microtubules
have on nascent cellulose microfibrils should
be studied in fully expanded cells. In the fully
expanded cells that have been studied, cortical microtubules and the most recently
deposited cellulose microfibrils are not generally in parallel alignment14. Mutants disturbed
in either cortical microtubule alignment or cellulose microfibril alignment during deposition
are important for determining the role that the
microtubules play in microfibril deposition.
Membrane flow determines cellulose
microfibril deposition orientation
Plant scientists often take the microtubule–
microfibril paradigm for granted, without even
studying the cellulose microfibrils properly14.
Cellulose synthases in the plasma
membrane
Golgi vesicles carrying wall matrix material
as cargo and cellulose synthases in their membranes, deposit these into the wall and the
plasma membrane, respectively, as a result of
exocytosis. Freeze-fractured plasma membranes show these cellulose synthases, which
consist of six particles arranged in a ring5
(Fig. 1), a so-called particle rosette. These
rosettes spin out the cellulose polymers6,7,
which crystallize as bundles, the cellulose
microfibrils. The polymerization forces cause
the synthases to move in the plasma
membrane. A functional model for cellulose
synthase has been hypothesized (Fig. 1), in
which the particles of the rosette spin out the
cellulose polymers.
Types of cell wall textures
The following cell wall textures are known:
axial, transverse, crossed, helical, helicoidal and
random (Fig. 2). Other types of texture are derived from these basic types by the successive
deposition of different textures.
Fig. 1. (a) Particle rosette, the cellulose synthase, in the plasma membrane of an
Equisetum hyemale root hair. The rosette has a diameter of 25 nm and consists of six particles.
In this cell type, the rosette density is relatively low, 5–15 rosettes/mm22, which agrees
with the large distance between individual microfibrils in the last deposited lamella6. (b)
Hypothesis for a functional model of a particle rosette (adapted from Ref. 6 ). The six proteins constituting the particle rosette (PR) [seen in the plasmatic-fracture (PF) face] form
channels in the plasma membrane. Through these channels UDP-glucose passes while
being polymerized into cellulose molecules. The terminal globule (TG) (sometimes seen
in the exoplasmic (EF)-fracture face) could be the still uncrystallized cellulose polymers.
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January 2000, Vol. 5, No. 1
35
trends in plant science
Perspectives
are strong counter-arguments against this
hypothesis:
• The cellulose microfibrils have dimensions
(both in linear size and mass) that put them
outside the colloidal domain, thus precluding their equilibration on reasonable
timescales19,20.
• The available space between the plasma
membrane and the already extant primary
cell wall is so limited that this system would be dominated by boundary
effects. In most cases these effects would
disturb the equilibrium configuration of
the particles involved, either by promoting boundary layers of different
structure, or anchoring defects or the
imposition of antagonistic geometrical
constraints21.
• The wall matrix material is probably
more like a dense gel than a thermal solvent and would thus also be an obstacle to
equlibration22.
Fig. 2. Basic wall textures: axial, transverse, crossed, helical, helicoidal and
random. Successive wall lamellae have
been peeled off from top to bottom
showing the random wall texture of the
outermost wall at the base.
However, scientists studying the cell wall
have come up with several alternative
hypotheses for the orientation mechanism of
cellulose mirofibrils. One such alternative is
the membrane flow hypothesis15. According
to this hypothesis the particle rosettes are free
to move in the plane of the viscous medium of
the plasma membrane, and are pushed forward
by synthesis and crystallization forces of the
cellulose microfibril. The first part of this
hypothesis is generally accepted and is also
used by us. But the hypothesis also suggests
that the rotation of the protoplast relative to the
cell wall is the orienting principle. However,
this rotation has never been observed in plants.
Cell wall texture formation is a selfassembly process like liquid crystal
formation
Scientists faced with helicoidal cell walls in
which the microtubules and the cellulose
microfibrils did not match, realized that the
helicoidal walls resemble cholesteric liquid
crystals, which are known to self-assemble
spontaneously16,17. However, this type of equilibrium assembly can only take place when a
sufficient amount of bulk material is present.
There is no evidence that this condition is ever
met because we now know that cellulose
microfibrils are made by plasma membraneembedded synthase complexes in a sequential
growth process18 (reviewed in Ref. 7).
Moreover, from a physics point of view, there
36
January 2000, Vol. 5, No. 1
The geometrical theory
We have proposed a geometrical model
for cellulose microfibril deposition23–26 . The
model quantitatively relates the deposition
angle of the cellulose microfibrils (with
respect to the cell axis) to:
• The density of active synthases in the
plasma membrane.
• The distance between individual microfibrils within a wall lamella.
• The geometry of the cell.
The helicoidal wall texture of root hairs
Inspiration for the geometrical model came
from observations on the secondary wall of
root hairs of Equisetum hyemale. A secondary
wall is deposited in a cell or part of a cell
that does not expand. Thus, at any stage of
Fig. 3. Schematic showing a section
through the consecutive lamellae of a
developing (from the lower to the upper
part of the diagram) helicoidal cell wall.
The already deposited consecutive lamellae are on the left; the most recently
deposited lamella is on the right. The
lengths of the areas with a certain microfibril orientation are much longer than can be
shown in a drawing; therefore, black horizontal lines have been included between
adjacent areas. Both transverse sections
through the wall and surface preparations
show a record of the deposition process in
time. In Equisetum hyemale root hairs, the
length of such an area is ~300 mm (A.M.C.
Emons, unpublished).
Fig. 4. (a) Shadowcast preparation of the helicoidal cell wall texture of an Equisetum hyemale root
hair showing the cellulose microfibrils in three subsequent orientations of the helicoid. A cellulose
microfibril with platinum/carbon shadow deposit is 7–8 nm wide. Scale bar 5 500 nm. (b) Thin section through the cell wall of a root hair of E. hyemale showing the arcs, which are visual artifacts of
the helicoidal texture. A negatively stained cellulose microfibril is 3–4 nm wide. Scale bar 5 500 nm
trends in plant science
Perspectives
its deposition, the secondary wall is a record
of the wall deposition process. The root hair
is a tip-growing cell with primary wall deposition at the growing hair tip and secondary
wall deposition in the shank of the hair.
Root hairs, including those from E. hyemale,
have particle rosettes, the cellulose synthases
(Fig. 1), in the plasma membrane27. The
E. hyemale root hairs have helicoidal cell
walls and the cortical microtubules do not
align with the nascent cellulose microfibrils28–30. A helicoidal cell wall texture (Fig. 2)
consists of one microfibril thick lamellae.
The orientation of the cellulose microfibrils in a lamella makes a constant angle
with the orientation in the subsequent lamella
(Fig. 3). In sections, this gives rise to the optical illusion of arcs (Fig. 4).
Facts and assumptions of the
geometrical model
Cellulose microfibrils are long structures. No
reliable estimate of their length is available
to date because microfibril ends are seen
rarely in wall preparations. An individual
cellulose microfibril can, therefore, contribute to several wall lamellae. Freeze-fracture images show that the cell wall is
appressed against the plasma membrane.
Therefore, wall deposition occurs under
space-limiting conditions.
Consider a cylinder around which several
tapes are being wound, such that the whole
cylinder surface is covered with one single
layer of tape (Fig. 5). The angle that the
tape makes with the long axis of the cylinder
is completely described by the following
formula:
sine a 5
number of tapes N 3 tape width d
p 3 cell diameter D
(1)
where a is the fibril angle with the cell’s long
axis, D denotes the cell diameter, N the
number of active synthases in the plasma
membrane and d is the distance between individual microfibrils within a lamella in a plant
cell (Fig. 5). Therefore, d depends on
the quantity of matrix between cellulose
microfibrils31 and, thus, on the ratio of cellulose to matrix. In an E. hyemale root hair
plasma membrane, this distance is ~150 nm
(Ref. 6).
Because a texture can be described fully by
describing the angle every cellulose microfibril makes with the cell axis during its deposition, the dependencies described in Equation
1 give us the basis of the geometrical theory
of cell wall texture formation.
Now that we can describe how a synthase
moves, we need to consider these questions:
where and at what rate are these synthases
activated? The important assumption of our
model is that this activation takes place in
Fig. 5. (a) The geometrical rule for cellulose microfibril deposition follows from the notion
of winding tapes densely around a cylinder, as is illustrated by unrolling the cylinder into a
planar figure, where D denotes the cell diameter, N the number of active synthases in the
plasma membrane and d is the distance between individual microfibrils within a lamella in a
plant cell. (b) Schematic of a cellulose microfibril surrounded by a wall matrix. The distance
between two microfibrils is the width of one tape, d, used in the model. In Equisetum
hyemale, root hair plasma membrane width is ~150 nm. (c) The influence of changing
cellular parameters on the microfibril winding-angle as predicted by the geometrical theory.
(i) Changing the relative number of microfibrils expressed as a fraction of the maximum
Nmax 5 pD/d; (ii) changing the relative effective width of the microfibrils expressed as a
fraction of the maximum possible width dmax 5 pD/N; (iii) changing the relative diameter of
the cell expressed in units of the minimum diameter Dmin 5 Nd/p.
January 2000, Vol. 5, No. 1
37
trends in plant science
Perspectives
Fig. 6. (a) Insertion domain. The assumption of the geometrical model is that synthase activation
takes place in localized mobile insertion domains. An insertion domain consists of a plasma membrane area in which Golgi vesicles, containing the synthases in their membranes, are inserted into
the plasma membrane. (b) This insertion domain itself, or the activation of the synthases in the
domain, moves with a velocity v, while the microfibrils are spun from the synthases in the opposite direction. t 5 time; t† 5 time interval; w 5 the speed with which the synthase moves.
Box 1. The synthase density evolution equation
The equation that describes the evolution of active cellulose microfibril synthase density on
a perfectly cylindrical cell of radius R, is given by
∂N ( z, t )
∂N ( z, t )
wd
−
= w ( N , z, t ) − w † ( N , z, t )
N ( z, t )
∂t
∂z
2pR
where w is the speed with which the synthase moves, d is the effective width of a cellulose
microfibril plus adherent matrix material and N(z,t) is synthase density. w is the local rate of
synthase production for which we choose the following form
w ( N , z, t ) =
N∗
N ( z, t )
1 −
t∗ (1 − g )
N∗
g
if N (z,t),N* and z is
located inside an insertion domain. In all other cases w 5 0. The param-eter g controls the
shape of the synthase production curve and lies between 0 and 1. Synthase production stops
when the maximum density
2pR
N∗ =
d
is reached, which for stationary insertion domains would happen after time t* . The insertion
domains are assumed to have a length l and travel at a speed v. Finally, the local rate of
rosette de-activation w✝ needs to be determined. This rate depends on the full evolution of the
density in a time interval of length t✝ (the synthase lifetime). Fortunately, the resultant equations are of a type that can be readily solved with entirely classical techniques. At first sight
the model appears to have many parameters, but dimensional analysis reveals that there are
only four relevant ones:
l
l =
wt∗
• Length of the insertion domain
•
Speed of the insertion domain
•
Synthase lifetime
n
w
t†
t† =
t∗
b =
g
38
January 2000, Vol. 5, No. 1
Fig. 7. Visualization of the solution of
our geometrical model in a helicoidal
example. For clarity, only a small number of cellulose microfibrils are shown,
and have been drawn at a highly
expanded scale. Four ‘generations’ of
cellulose microfibrils are spun by synthases that were activated at the tip of an
insertion domain at four equally spaced
moments in time. The displacement of
the starting points along the cell is a
consequence of the motion of the insertion domain. Each of these generations
contributes locally to a single lamella.
The inset shows how the angle of deposition between adjacent lamellae is
nearly constant, the hallmark of helicoidal texture.
localized mobile insertion domains. An insertion domain consists of a plasma membrane
area in which Golgi vesicles, containing the
synthases in their membranes32, are inserted
into the plasma membrane (i.e. a domain
where exocytosis occurs; Fig. 6). For insertion domain mobility, at least two experimentally verifiable scenarios are possible:
• The Golgi bodies that produce the Golgi
vesicle-containing synthases move along
the plasma membrane while inserting the
vesicles.
• The Golgi bodies themselves are stationary, but exocytosis of their vesicles is
locally controlled by a time-varying
concentration of an activating–inhibiting
substance.
It is possible that calcium ions fulfil this role.
We believe that the activation of exocytosis in
root hairs involves calcium33,34, and calcium
ion concentration oscillations have been
observed in root hairs upon stimulation35.
trends in plant science
Perspectives
The mathematical model
We can formulate a model that describes the
whole process of cell wall texture formation
using the elements described for the geometrical model. The theory describes the evolution of active synthase density in the plasma
membrane in space and time. If the number of
active synthases present at a given location in
the plasma membrane is known, the geometrical rule predicts the orientation of the cellulose microfibrils that are deposited. In a sense,
each microfibril ‘records’ the motion of its
synthase. The totality of all these ‘recordings’
creates the cell wall texture. To formulate such
a model, attention needs to be focused on a
small segment of the plasma membrane. How
does the number of active synthases in the portion of plasma membrane belonging to this
segment change in a small interval of time?
There are three sources of change:
• Movement of synthases into and out of the
segment.
• Activation of new synthases by an insertion
domain.
• De-activation of synthases after a finite life
span.
Mathematically, this yields a partial differential equation for synthase density N (z,t ),
which can be solved as a function of position
z along the cell and time t (Box 1).
The Helicoidal solution
The solution for a helicoidal cell wall demonstrates what the geometrical model can do
(Fig. 7). In such a wall, the microfibril orientation changes by a constant angle from one
lamella to another (Fig. 2), producing a staircase-like structure (hence the name helicoid).
To obtain such a structure in which the deposition angle progresses from transversal (a 5
08), through axial (a 5 908) and back to transversal (a 5 1808), after which the cycle can
repeat itself, the parameters of the model need
to be precisely tuned. In our view, the geometrical model is sufficient to explain the
most complicated cell wall texture. However,
its general utility is its ability to describe the
various other wall textures also seen in plant
cells (Fig. 2).
Utility of the model
Numerous cell wall examples that support
the geometrical model have been discussed
before23–26. The important assumption to be
verified is the movement or activation of the
insertion domains. We have proposed two scenarios for local, sequential exocytosis: the
movement of the Golgi vesicle-producing
Golgi bodies along the plasma membrane
while inserting the vesicles, and the local calcium-induced exocytosis of vesicles along the
plasma membrane. Other properties of the
insertion domain that need to be determined
and manipulated include its length and speed.
Furthermore, to date we know nothing about
the synthase life-time. The use of mutants,
GFP-technology and calcium measurements
of high sensitivity along the plasma membrane
of cells with known wall deposition patterns,
should provide an appropriate series of experiments for studying the exocytosis machinery
in living cells in relation to wall deposition.
Acknowledgements
We thank Allex Haasdijk for the artwork in
Figures 1(b), 2, 3 and 6(a), and Ton Bisseling
for critically reading the manuscript.
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Does a light-harvesting
protochlorophyllide a/bbinding protein complex exist?
Gregory A. Armstrong, Klaus Apel and Wolfhart Rüdiger
Recent in vitro studies have led to speculation that a novel light-harvesting
protochlorophyllide a/b-binding protein complex (LHPP) might exist in dark-grown
angiosperms. Structurally, it has been suggested that LHPP consists of a 5:1 ratio
of dark-stable ternary complexes of the light-dependent NADPH: protochlorophyllide
oxidoreductases A and B containing nonphotoactive protochlorophyllide b and
photoactive protochlorophyllide a, respectively. Functionally, LHPP has been
hypothesized to play major roles in establishing the photosynthetic apparatus, in
protecting against photo-oxidative damage during greening, and in determining
etioplast inner membrane architecture. However, the LHPP model is not compatible
with other studies of the pigments and the pigment–protein complexes of dark-grown
angiosperms. Protochlorophyllide b, which is postulated to be the major lightharvesting pigment of LHPP, has, for example, never been detected in etiolated
seedlings. This raises the question: does LHPP exist?
L
ight profoundly influences plant
development and allows photosynthesis to occur, but it also represents a
tremendous risk. Photo-oxidative damage
initiated by excited state photosensitizing
molecules, such as chlorophylls and their
biosynthetic precursors, can be lethal.
Angiosperms that germinate in darkness in
the soil enter the seedling developmental program known as skotomorphogenesis (Fig. 1).
However, such seedlings must be prepared
for a subsequent light-triggered switch to
photomorphogenesis. Upon illumination, the
leaves of etiolated angiosperms synthesize
and accumulate large quantities of chloro40
January 2000, Vol. 5, No. 1
phylls a and b. Seedlings are particularly susceptible to photo-oxidative damage during
this transition to photoautotrophy.
The presence or absence of light dramatically
influences plastid development. Dark-grown
angiosperm seedlings contain an achlorophyllous plastid type known as the etioplast, which
is transformed into a photosynthetically competent chloroplast during photomorphogenesis1.
The etioplast is defined by the presence of two
types of internal membranes, the lattice-like
prolamellar body, which is composed of interconnected tubules, and the unstacked prothylakoids. Etioplasts characteristically accumulate
the chlorophyll precursor protochlorophyllide
Anne Mie C. Emons is at the Laboratory of
Experimental Plant Morphology and Cell
Biology, Dept of Plant Sciences,
Wageningen University, Arboretumlaan 4,
6703 BD Wageningen, The Netherlands
(tel 131 317 484329;
fax 131 317 485005;
e-mail [email protected]);
Bela M. Mulder is at the Condensed Matter
Division of the FOM Institute for Atomic and
Molecular Physics, Kruislaan 407, 1098 SJ
Amsterdam, The Netherlands
(tel 131 20 6081231;
fax 131 20 6684106;
e-mail [email protected]).
(Pchlide), more specifically protochlorophyllide
a (Pchlide a)2–4. Illumination of etioplasts initiates the dispersal of the prolamellar body and
the formation of thylakoid membranes containing the pigment–protein complexes of the
photosynthetic apparatus.
In this context, recent in vitro reconstitution
experiments have been interpreted as providing
evidence for a novel light-harvesting Pchlide
a/b-binding protein complex5, termed LHPP
by analogy to the ubiquitous light-harvesting
chlorophyll a/b-binding proteins (LHCP) of
green plants. LHPP is speculated to:
• Serve as the central structural determinant
of the prolamellar body in etioplasts.
• Be essential for the establishment of the
photosynthetic apparatus.
• Confer photoprotection on greening
seedlings by dissipating excess light
energy, thereby minimizing Pchlideinduced photo-oxidative damage.
On the one hand, if they are correct, these
hypotheses would have a major impact on our
understanding of the seedling transition from
skotomorphogenesis to photomorphogenesis.
On the other hand, to date there are no in vivo
data that directly support the existence of an
LHPP complex6. Here, we critically analyse
the LHPP model in light of the current literature on the properties of etioplast membranes,
pigment–protein complexes and pigments.
Roles of the light-dependent PORA and
PORB proteins in etioplast formation
and photo-oxidative protection
The presence of the prolamellar body and the
accumulation of Pchlide a in etioplasts are
known to correlate with large quantities of the
strictly light-dependent NADPH:protochlorophyllide oxidoreductase (POR; 1.3.1.33)1,7–9.
This nuclear-encoded but plastid-localized
protein is unusual in that it mediates the only
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