Mechanical Forces in Plant Growth and Development
Deborah D. Fisher and Richard J. Cyr*
Department of Biology, 208 Mueller Lab, Pennsylvania State University, University Park PA
ABSTRACT
Plant cells perceive forces that arise from the environment
and from the biophysics of plant growth. These forces provide
meaningful cues that can affect the development of the plant.
Seedlings of Arabidopsis thaliana were used to examine the
cytoplasmic tensile character of cells that have been implicated
in the gravitropic response. Laser-trapping technology revealed
that the starch-containing statoliths of the central columella cells
in root caps are held loosely within the cytoplasm. In contrast,
the peripheral cells have starch granules that are relatively
resistant to movement. The role of the actin cytoskeleton in
affecting the tensile character of these cells is discussed. To
explore the role that biophysical forces might play in generating
developmental cues, we have developed an experimental model
system in which protoplasts, embedded in a synthetic agarose
matrix, are subjected to stretching or compression. We have
found that protoplasts subjected to these forces from five
minutes to two hours will subsequently elongate either at right
angles or parallel to the tensive or compressive force vector.
Moreover, the cortical microtubules are found to be organized
either at right angles or parallel to the tensive or compressive
force vector. We discuss these results in terms of an interplay of
information between the extracellular matrix and the underlying
cytoskeleton.
INTRODUCTION
Land plants are relatively sessile organisms, and rigid cell
walls interconnect all the cells within the plant thallus. These
traits have two important consequences. First, plants generally
cannot move quickly in response to a changing environment;
rather they must adapt to where they are. Second, due to the
physical coupling of rigidly bound cells within the plant tissues,
mechanical events in one area of the plant can be transmitted to
other areas. Cells are subjected to a variety of forces during plant
growth and development. Exogenous forces arise from the
environment and endogenous forces stem from the biophysics of
plant growth. The overall objective of this article is to provide
the reader with an appreciation of how the plant can use these
forces as meaningful developmental cues. In the first part of the
article, we will describe how specialized cells perceive
gravitational forces. In the second part, we will discuss how
endogenous forces might be used by plant cells to help monitor
their own growth status—in particular, the relationship between
the microtubule cytoskeleton and the cell wall. We will present
data to support the hypothesis that plants have the ability to
continually use a variety of mechanical cues to monitor and
adjust morphogenetically. These mechanical cues, derived from
the action of exogenous and endogenous forces, are integrated by
the plant to insure that optimal growth occurs under a variety of
conditions.
GRAVITY WORKS ON MOVEABLE OBJECTS
External factors are used as developmental cues by
growing plants (e.g., electromagnetic radiation, gravity, and
temperature; Hangarter, 1997). Undoubtedly, these same
elements have had a profound impact on plant evolution
(Barlow, 1995; Niklas, 1998). Although the entire plant is
continuously subjected to gravitational acceleration, probably
only specialized cells have the ability to perceive this force in
angiosperms (Sack, 1991). Perception appears to be based
largely on cellular rigidity. Basically, if a cellular object is held
tightly within the cytoplasm (i.e., if the tensile strength holding a
particular object is greater than the product of the object’s
gravitational acceleration and its mass) the object remains
stationary. However, if the object is loosely suspended within
the cell, the gravitationally induced force will cause it to fall
downward. This downward movement, in specialized cells, is
transduced to affect the growth of both shoots and roots
(Masson, 1995; Volkmann et al., 1999).
Among the mass types that gravity affects are the heavy
starch grains (statoliths) located within the root caps of
angiosperm cells (Figure 1). Genetic and morphological data
indicate that the starch grains within the root cap are major sites
of gravity perception (Kiss et al., 1989; Kiss and Sack, 1989;
Sack and Kiss, 1989). Amyloplasts likely play a similar role
within the graviresponsive cells of shoots (Kiss et al., 1997;
Weise and Kiss, 1999). It appears that only certain cells within
the root cap are involved in the gravity response. Laser ablation
of the more centrally located columella cells abolishes a root’s
ability to respond to gravity; however,
Figure 1. The Root Tip is Highly Differentiated. A root tip
from Arabidopsis thaliana is shown with outlined central
columella cells, which contain loose statoliths.
*Correspondence to: Richard J. Cyr: fax: 814-865-9131; e-mail:
[email protected]
Gravitational and Space Biology Bulletin 13(2), June 2000
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FORCES AND PLANT DEVELOPMENT
Figure 2. Loose Starch Grains Demonstrated with Laser Trapping . Central columella cells are visualized within an Arabidopsis
thaliana root tip and a starch-containing statolith. (a) The infrared optical trap is turned on, and a starch grain is trapped (arrow). (b) The
steering mirror for the laser trap has been adjusted to reposition the focal point of the laser and the entrapped starch grain to a new
position. Note the upward displacement of the starch grain here, compared with the same grain’s position in a.
ablation of the more laterally located peripheral cells does not
(Blancaflor et al., 1998). The approximate location of these
central columella cells is outlined in Figure 1. These ablation
results correlate well with differences in sedimentation velocities
of amyloplasts in the central cells, versus the peripheral cells, of
the root cap (Sack et al., 1986; Blancaflor et al., 1998).
Therefore, it appears that there are differences between central
and peripheral columella cells. One difference might be the
tensile strength of the cytoplasm holding the statoliths (Baluska
and Hasenstein, 1997).
To address this possibility, we used laser-trapping
technology to explore the tensile character of the cytoplasm
within the root cap cells. The technology of laser trapping, also
known as laser tweezers, is based on the behavior of highintensity light as it passes through a highly refractive dielectric
sphere. Basically, as light passes into a dielectric sphere with a
higher refractive index than the medium, the light ray is refracted
and reflected, which leads to a change in photonic momentum.
As a result, a major restoring force vector is generated towards
the focal point, which creates a physical trap (Ashkin, 1998).
We grew Arabidopsis seedlings on agarose-coated
coverslips oriented at 45°C. This orientation insures that the tip
of the root will grow along the optical surface of the coverslip,
thereby providing the best possible image. Figure 2a shows
central columella cells within an Arabidopsis thaliana root tip
and statolith (see arrow) that have been trapped by the infrared
laser trap. In Figure 2b, the steering mirror has been moved,
which displaces the trap along with the statolith.
Physically trapping intercellular organelles, such as starch
statoliths, enabled us to determine the relative tensile character
of the cytoplasm in different cells of the root cap. We
systematically scanned across the root cap and determined how
loosely the statoliths were held. The space in which statoliths
freely move within the cells coincides with the central columella
cells outlined in Figure 1. The outline shows that only the central
columella cells have starch grains that are held loosely. Those in
the cap’s periphery are held relatively tightly, and cannot be
moved easily. These results indicate that the starch-containing
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Gravitational and Space Biology Bulletin 13(2), June 2000
statoliths of the central columella cells would predictably move
faster, therefore generating more force in response to gravity, and
thus would be a more sensitive transducer of gravity’s vector.
What cellular component would allow the starch grains to
move more freely in the central columella cells? The actin
moiety of the cytoskeleton is probably the most likely
candidate. In most plant cells, the actin cytoskeleton forms a
tight network that acts to suspend organelles and, in conjunction
with myosin, to affect intracellular motility (Williamson, 1993).
Baluska et al. (1997) report that the central columella cells are
devoid of cytoskeletal elements radiating into the cell interior,
while the peripheral columella cells are rich in these filaments.
Failure to demonstrate cytoskeletal elements in the more interior
locations of central columella cells indicates that the cytoplasm
in these cells is structurally unique. During differentiation of
central columella cells, it is likely that actin is either downregulated and/or that actin-severing/depolymerizing proteins are
up-regulated. Further experimentation will undoubtedly reveal
the molecular basis for this loose state, which appears to be
ideally suited for detecting the movement of heavy organelles in
response to gravity.
ENDOGENOUS FORCES WORK ON THE CELL
Compared to the cells found within a developing animal,
plant cells develop in a somewhat constrained environment and
do not migrate relative to one another (Fosket, 1994). Hence, all
morphogenetic processes must occur within a relatively rigid
milieu. To appreciate how plants grow under these constraints,
consider that osmotic forces cause plant cells to develop high
water pressures. Thus, a turgid plant cell is under pressure, and
this force is exerted isotropically within the cell (Cosgrove,
1993). How does the cell, or a file of cells, elongate in a vectoral
manner when driven by isotropic forces? The strong cellulose
microfibrils within a growing cell (Figure 3a) are not arranged
randomly, rather they are highly oriented at right angles to the
axis of elongation. Cellulose reinforces the wall, much as hoops
strengthen a barrel, allowing little lateral expansion. However,
FORCES AND PLANT DEVELOPMENT
growth is unconstrained in the axis perpendicular to cellulose
alignment (Green and Poethig, 1982). Therefore, as a
consequence of growth, a major strain axis develops along this
axis. This strain axis arises when the isotropic forces of turgor
are permitted to work solely at right angles to the orientation of
cellulose microfibrils (Gertel and Green, 1977; Green and Selker,
1991).
The orderly deposition of cellulose occurs during its
synthesis. Cellulose synthase complexes are found in complexes
within the plasma membrane (Delmer and Amor, 1995). The
synthesis of cellulose microfibrils can be minimally described as
a two-step process: glucose is first assembled into a β1,4
polyglucan chain, which then combines with 30-100 other
polyglucan chains to crystallize into a microfibril (Brett and
Waldron, 1990). Crystallization is an exergonic reaction that can
work to move the entire cellulose synthase complex into the
plane of the fluid membrane. However, the movement of these
complexes is not random. Somehow, the cortical microtubules,
which are attached to the inner face of the plasma membrane,
restrict the gliding cellulose synthase complexes, as well as
nascent microfibrils, along a particular path (Cyr and Palevitz,
1995). Cortical microtubules flanking the cellulose synthase
complexes act like railroad tracks to guide the complex (Figure
3b; Giddings and Staehelin, 1991; Cyr, 1994). One weakness of
this so-called “microtubule/microfibril paradigm” is that the
molecular nature of how microtubules interact with the plasma
membrane is not understood, nor is it clear whether the cellulose
synthase complexes can interact directly with the underlying
microtubules.
Although it is clear that microtubules influence the
deposition of microfibrils in many, if not most, elongating plant
cells, how microtubules become organized remains uncertain. It
has been proposed that biophysical forces orient microtubules
(Green et al., 1970). The evidence that supports this hypothesis
is largely circumstantial (Williamson, 1990; Williamson, 1991;
Cyr, 1994) and somewhat controversial (Nick, 1999). We have
taken an experimental approach to verify or refute this
hypothesis.
To understand the possible role that mechanical forces
play in orienting microtubules, Nagata et al. (1992) used a
cultured tobacco cell line, designated BY-2. This cell line is easy
to culture and, depending upon the culture conditions, will grow
either as rapidly dividing cells or as elongating cells (Hasezawa
and Syono, 1983). As with many plant cells, it is relatively easy
to remove the cell wall via enzymatic digestion. This is
important, because the rigid nature of the cell wall makes it
difficult to know what forces are perceived when an external
force is applied. By removing the wall, we can directly measure
the effect of the force as a deformation of the protoplast. Once
the cell wall is removed, a spherical protoplast is released that
contains cortical microtubules that usually are very randomly
oriented. Upon culturing, protoplasts regenerate a wall,
reorganize their cortical microtubules, and within 24-48 hours
begin to elongate (Figure 4). Our experiments address the
following questions:
Figure 3. Organization of Cellulose in Elong ating Cells .
(a) Turgor pressure, coupled with wall relaxation, drives cellular
expansion that is limited to one major axis by the organized
cellulose microfibrils. (b) The organized deposition of cellulose is
affected by cortical microtubules, which are attached to the
underside of the plasma membrane.
•
•
Can newly isolated protoplasts have the future
axis of elongation manipulated?
Simultaneously, do the same manipulations that
provide an axial “cue” also affect the organizational
status of the cortical microtubules?
Wymer et al. previously showed that a brief centrifugation
of newly isolated protoplasts could cue the future axis of
elongation (1996). However, in those studies, it was difficult to
know precisely what forces the cells were experiencing. To
further extend the studies, we used a modified, commercially
available apparatus marketed by Flexcell Corporation (Banes et
al., 1985) to manipulate protoplasts embedded in agarose on a
flexible sheet of silastic. The agarose served as an artificial elastic
matrix that allowed us to study the response of the cell to a
given amount of applied force. We poured a protoplast/ agarose
suspension onto the membrane. After the agarose solidified, we
placed the silicon membrane (held tightly in a sterile chamber)
over a vacuum manifold containing an immobile post.
Gravitational and Space Biology Bulletin 13(2), June 2000
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FORCES AND PLANT DEVELOPMENT
Figure 4. Experimental System for Studying Elongation.
Top row: Cultured tobacco cells grow in elongate cell files, and
each cell contains an organized cortical microtubule (MTs) array
that can be visualized with anti-tubulin antibodies. Middle row:
Upon enzymatic removal of the wall, spherical protoplasts with
disorganized cortical microtubules are produced.. Bottom row:
Upon reculturing in a medium that supports elongation, the cells
reorganize their microtubules and elongate, as seen at 7 days
post-protoplasting.
When we drew a vacuum on the manifold, the silicon membrane
deformed around the post and the attached protoplasts/agarose
matrix was stretched or compressed. By changing the geometry
of the loading post, we exposed the cells to different forces.
Note that we set up this apparatus for microscopic examination,
which permitted visualization of the cells as a force was applied.
Also, by adding fluorescent reference beads, we could quantify
the nature of applied force on the matrix and embedded
protoplasts.
Immediately after isolation, cells appeared mostly
spherical (if slightly aspherical, there was no preferential axis of
asymmetry). During the application of force, the cells appeared
either deformed at right angles to the compression force vector or
parallel to the tension vector (data not shown). Significantly,
after the stretching period, the cells returned to a spherical shape
(if non-spherical, there was no preferential axis of asymmetry).
We directly quantified the magnitude of applied force by
measuring the displacement of fluorescent microbeads, which
confirmed the major compressive or tensive axis. It is
noteworthy that, once the applied force was released, the
reference beads returned to their original positions, thereby
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Gravitational and Space Biology Bulletin 13(2), June 2000
indicating that the agarose matrix behaved as a completely elastic
membrane and that the applied force had no demonstrable
plastic-deforming effects (data not shown). Freshly isolated
protoplasts were deformed for two hours, then released and
cultured to favor elongation. Typically, during the first 24 hours
after force application they remained somewhat spherical (if
asymmetrical, there was no dominant asymmetric axis).
However, within 72 hours, the majority of cells showed some
anisodiametric growth; within seven days, their length-to-width
ratio was typically between three and six. In areas of the agarose
sheet where cells were exposed to biaxial forces (i.e., the center
of the round loading post), the elongative axes were random.
However, in areas where the cells were exposed to a uniaxial
tensive force, the majority had a nonrandom elongative axis
parallel to the tension vector (Figure 5). Similarly, in areas that
were under uniaxial compression, we observed a nonrandom
elongative axis—but it was at right angles to the compression
vector (data not shown).
Finding that a brief uniaxial tensive or compressive force
can cue elongative axes raises questions about how these forces
are perceived and about the nature of the memory once
perception has occurred. Recall that force application occurs 2244 hours before any morphological change is observed. Cortical
microtubules are involved in the deposition of cellulose
microfibrils and, because the ordered deposition of cellulose is
required for elongation in these cells, the microtubules are
candidates for both force perception and the memory
component. That is, random microtubules might perceive the
force, become aligned, and commence orienting cellulose
deposition, which consequently affects the axis of elongation. To
test this hypothesis, we subjected cells to compressive and
tensive forces, then fixed and processed them for
immunolocalization of microtubules (Figure 6). The cortical
microtubules in this cell are not random; rather they show a net
organization in the horizontal direction similar to the tension
force vector (Figure 6, arrow). Also, when protoplasts were first
subjected to compressive forces, then fixed and processed for
cytoskeletal visualization, the microtubules in many cells
showed alignment. However, this alignment was at right angles
to the compressive force vector (data not shown).
CONCLUSION
The above data support the hypothesis that physical forces can
influence axis determination in a plant cell. The finding that both
compressive and tensive forces can elicit this response is
consistent with observations reported in whole-plant studies.
For example, it has been reported that physically bending maize
coleoptiles induces microtubule alignment at right angles to the
tension vector (Zandomeni and Schopfer, 1994). The role of
compression in affecting microtubule alignment might also
explain the change in microtubule orientation observed in
graviresponding roots. In vertically grown roots, microtubules
within the epidermis and cortical cells are aligned at right angles
to the root axis. When roots are tilted horizontally, the cells
within the upper epithelium show the typical transverse
alignment of microtubules regularly observed in rapidly growing
FORCES AND PLANT DEVELOPMENT
Figure 5 . Stretching Cues the Axis of Elongation. Tobacco
protoplasts that were stretched for 2 hours were visualized after
7 days of culture. The majority of cells have elongative axes
parallel to the direction of stretching.
compressed, and high-resolution growth studies have reported a
negative growth rate in this region (Ishikawa et al., 1991). If the
tissue is shrinking, then it is also compressing; and the lower
epidermal cells could perceive this force, which would explain
their transverse alignment.
We suggest that the relationship between microtubules,
microfibrils, and elongative growth is functionally interrelated.
This relationship can be summarized as an extension of the
microtubule/ microfibril paradigm, in which cortical microtubules
serve as templates to affect the deposition of cellulose
microfibrils. The microfibril’s high tensile strength then restricts
isotropic turgor forces to one axis, and the resulting biophysical
forces act as cues that the cells use to affect microtubule
alignment. The alignment cue has previously been inferred from
work with cellulose synthesis inhibitors (Fisher and Cyr, 1998).
Here, we provide data indicating that both tensive and
compressive forces can orient microtubules. The molecular
details of how microtubules relay information to the cellulose
synthase are currently speculative. Therefore, we do not know
the sequence of events by which biophysical forces might be
perceived and transmitted to the microtubules. It has been
proposed that this process may be indirect and that it may
involve the molecular alignment of linking microtubule
nucleators, which then serve to orient microtubules (Williamson,
1990). Conversely, it has been proposed that microtubules
themselves may be the strain sensors (Green et al., 1970). All
models will remain speculative until we determine the molecular
nature of microtubule interaction with the plasma membrane and
the cellulose synthase. The challenge for future research will be
to identify these components and study their behavior under
conditions that are known to affect microtubule alignment and
the resulting elongative growth.
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