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Cell Biology |
Department of Biology, University of Louisiana at Lafayette, P.O. Box 42451, Lafayette, Louisiana 70504-2451 USA
Received for publication March 19, 2007. Accepted for publication October 9, 2007.
ABSTRACT
To characterize cellular fluidity and mechanical processes, we determined the viscous properties of the cytoplasm of Chara contraria rhizoids in vivo by injecting and displacing superparamagnetic particles. After injection and a 24-h recovery period, the particles were moved to different positions within the rhizoid by an external magnet. The system was calibrated with solutions of known viscosities. The viscosity was determined based on the velocity at which individual beads moved toward the external magnet. The viscosity of the cytoplasm varied with direction of measurement (i.e., was highly anisotropic) and also varied between sites. The highest viscosity was observed near the endogenous statoliths (139 mP·s parallel and 78 mP·s perpendicular to the rhizoid axis). Depolymerization of actin filaments with latrunculin B reduced the viscosity significantly except around the nucleus but did not change the overall viscosity pattern. Microtubule depolymerization with oryzalin reduced viscosity especially between the nucleus and the statolith zone. The data indicate that F-actin but not microtubules affects statolith sedimentation and that cytoplasmic viscosity may be important for the gravisensing system.
Key Words: Chara • Characeae • cytoplasm • cytoskeleton • gravisensing • rheology • rhizoid • viscosity
Dynamic processes within cells, such as vesicle transport, sedimentation of statoliths, or enzymatic reactions, depend on the intracellular organization, cytoskeletal architecture, density, and viscosity of cytoplasm. The viscosity within a cell depends on the quantity and organization of soluble and colloidal matter and the distribution of organelles and cytoskeletal components. Viscosity affects transport of vesicles, flexibility of membranes, and the force and thus energy required to move cellular compounds and structures. Some sensory functions also depend on the viscous properties of fluids, especially the stimulation of hair cells in the vestibular apparatus in animals and the sedimentation of statoliths or equivalents in plants. The sedimentation of otoliths and statoliths generates sensory signals related to accelerations and gravity.
The sedimentation of statoliths in the macro-alga Chara is a finely controlled process that stimulates distinct areas of the plasma membrane (Braun, 2002
). Therefore, changes in viscous properties of the cytoplasm are likely to affect the perception of the gravity signal as well as transport of wall-forming vesicles, the organization and turnover of the cytoskeleton (Bartnik et al., 1990
), and the rate of diffusion or transport of cellular contents.
Gravisensing depends on the translation of positional changes of statoliths into growth signals and reactions that realign the graviresponding structure to the gravitational set point angle (Firn and Digby, 1997
). However, the positioning of the statoliths must not interfere with their ability to move or sediment when the gravity vector changes. Exposure of rhizoids to microgravity in sounding rocket experiments (Buchen et al., 1993
) displaced statoliths temporarily upward. Similarly, the displacement of statoliths toward the rhizoid tip by centrifugation (Braun and Sievers, 1993
) was compensated, indicating that active processes maintain the statoliths in their targeted position. Therefore, statoliths are actively suspended and do not sediment. Longitudinal displacement of endogenous statoliths has been linked to F-actin (Braun and Wasteneys, 1998
; Kuznetsov and Hasenstein, 2001
). However, despite its involvement in the positioning of statoliths, F-actin does not seem to play a direct role in gravisensing as recently suggested for higher plants (Yoder et al., 2001
), but at least in Arabidopsis, it does seem to be involved in the termination of the graviresponse process (Yamamoto and Kiss, 2002
; Hou et al., 2003
).
Sedimentation of statoliths occurs when the forces in the downward direction are greater than those that keep the statoliths suspended, such as buoyancy, Brownian motion, and cytoskeletal interactions. Although viscosity does not represent a force, it greatly affects the movement of cellular components. Using optical tweezers, Leitz et al. (1995)
and Braun (2002)
demonstrated that the repositioning of statoliths in different regions of the rhizoid required different forces. Although optical tweezers provide an assay for mechanical linkages between (statolith) binding proteins and the actin cytoskeleton, these measurements cannot differentiate between active cytoskeleton-derived forces and cytoplasmic viscosity.
The measurement of viscosity in vivo is difficult but provides information about the motility of organelles, vesicles, and molecules, the sedimentation of statoliths and the extent of molecular crowding, which describes enhanced biochemical or enzymatic activity as the viscosity or the concentration of co-solutes increases (Minton, 1998
; Davis-Searles et al., 2001
). Therefore, the interaction between F-actin and statoliths may benefit from (increased) cytoplasmic viscosity.
To investigate the viscous properties of the cytoplasm in Chara rhizoids, we utilized the magnetic particle method (Crick and Hughes, 1949
), modifications of which have been used to determine viscoelastic properties and forces in living macrophages (Bausch et al., 1999
). Our data indicate a strong anisotropy of viscosity in Chara rhizoids, which is likely to contribute to cellular function and organization of this specialized, tip-growing cell.
MATERIALS AND METHODS
Plant material
Chara contraria A. Braun ex Kütz. (Characeae) was grown in 20-L glass tanks containing soil from a local pond under a 3-cm layer of sand. Oogonia were harvested, dried, and stored for 3 mo in the refrigerator (4°C) and germinated on Petri dishes containing nutrient medium solidified with 1.5% agar in a vertical orientation (Scherp and Hasenstein, 2003
). For microinjection experiments, one germinated oogonium with several rhizoids was placed on a microscope slide in liquid nutrient medium. To reduce turgor pressure and facilitate injection, we incubated the entire alga for 10 min in 160 mM sorbitol.
Microinjection
Magnetic particles were microinjected into Chara rhizoids with micropipettes pulled with a P-2000 puller (Sutter Instrument, Novato, California, USA) from borosilicate glass with an outer diameter of 5–7 µm (Scherp and Hasenstein, 2003
). Magnetic beads (Bangs Laboratory, Fishers, Indiana, USA; 1 µm diameter) were washed and suspended in injection buffer (48 mM K2HPO4, 4.5 mM KH2PO4, and 14 mM NaH2PO4, pH 7.2) and back-filled into the pipette by vacuum (30 mm Hg).
The rhizoid was injected with the aide of a holding device (15-µm blunt, solid glass rod) to secure the rhizoid under a Nikon Eclipse E600 FN microscope. The injection pipette was withdrawn within 15–30 min after injection to allow healing of the injection site. The oogonium with the injected rhizoid was then transferred to a Petri dish containing fresh nutrient medium for a 24-h recovery period. Only rhizoids that recovered to growth rates of 60–80 µm·h–1 were used for measurements.
Drug treatment
F-actin was depolymerized by immersing rhizoids in 5 µM latrunculin B (LatB) for 10 min before measurements. Microtubules were depolymerized by incubating rhizoids for 10 min in 5 µM oryzalin. The working solutions were prepared by diluting stock solutions (10 mM LatB and 5 mM oryzalin in dimethyl sulfoxide) with nutrient medium. Longer treatments had no effect on viscosity measurements.
Calibration
Glycerol (4.25–12 M), glucose (0.03–4.3 M), and sucrose solutions (2.07–2.45 M) with a combined range of viscosities from 1 to 147 mP·s (Weast, 1987) were prepared and checked against their refractive index. The magnetic beads were washed three times by suspension in the respective calibration solution. A drop of magnetic bead solution was placed on a hemocytometer and covered with a coverslip. Velocity was measured by placing a cone-shaped magnet (1 mm long, 0.8 mm base diameter, attached to a glass rod) 100 µm from the bead of interest. The time required for beads to traverse 20 or 50 µm toward the tip of the magnet was recorded with a stopwatch and plotted against the known viscosity (Fig. 1). The calibration and the in vivo measurements in the rhizoid were carried out by horizontally moving the magnetic beads. Because the same magnet was used for calibration and measurements, knowledge of the precise strength of the magnetic field was not necessary.
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50 µm above the rhizoid. This arrangement allowed the magnet to be moved to all areas of the rhizoid (Fig. 2). All calibration and in vivo measurements were carried out with the same magnet configuration.
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RESULTS
The viscosity of the cytoplasm in Chara rhizoids fluctuated by almost two orders of magnitude (Fig. 2). In addition to the variation by longitudinal position, viscosity also varied with the direction of bead movement. Typically, displacement of the beads parallel to the rhizoid axis resulted in different viscosity values than displacements perpendicular to the rhizoid axis.
The tip (Fig. 2, position 1) of the rhizoid had a "longitudinal" viscosity of 9 ± 0.3 mP·s but a transverse viscosity of 29 ± 1.3 mP·s. The highest viscosity was measured near endogenous statoliths (Fig. 2, position 2). Based on the difference between longitudinal and transverse viscosity, this region had the strongest anisotropy. Longitudinal viscosity was about 139 ± 9 mP·s and transverse viscosity reached 78 ± 8 mP·s. The viscosity in the mid-apical region (between the nucleus and tip; Fig. 2, position 3) was also anisotropic. In contrast to the statoliths region (position 2), beads in the mid-apical region were more readily displaced longitudinally than transversely. The highest viscosity in this zone was measured longitudinally along the cell wall (30 ± 5 mP·s; Fig. 2, position 4).
Compared with the mid-apical and apical region, the cytoplasm around the nucleus (Fig. 2, position 5) had a significantly higher viscosity, similar to that around the vacuole (61 ± 8 mP·s; Fig. 2, position 6). The lowest viscosity in the Chara rhizoid was measured in the zone of cytoplasmic streaming that surrounds the vacuole (3 mP·s; Fig. 2, position 7). This value is comparable to the viscosity of water (1 mP·s).
The application of F-actin depolymerizing drug LatB (Kuznetsov and Hasenstein, 2001
) generally reduced cytoplasmic viscosity (Fig. 2). The depolymerization of the actin network affected viscosity mostly in the statoliths region (Fig. 2, position 2). Viscosity was reduced by 70% to 43 ± 4 mP·s longitudinally and by 50% to 37 ± 3 mP·s in the transverse direction. The viscosity in the apical region (Fig. 2, position 1) was reduced by 50% in the transverse direction and in longitudinal measurements by 33%. After LatB application, the cytoplasm between nucleus and statoliths (Fig. 2, position 3) had no change in the longitudinal component of the viscosity (5 ± 0.3 mP·s) but transverse viscosity decreased from 15 ± 1.3 mP·s to 8 ± 0.7 mP·s. The viscosity near the cell wall (Fig. 2, position 4) did not change. LatB did not affect the viscosity around the nucleus or in the zone of cytoplasmic streaming and surrounding region.
Application of the microtubule-depolymerizing drug oryzalin reduced viscosity less than application of LatB, and the statoliths did not sediment into the apical dome. The viscosity decreased presumably because of the depolymerization of microtubules. The overall viscosity pattern was not changed but oryzalin reduced the viscosity in the longitudinal direction in the area of the statoliths by 10% to 127 ± 5 mP·s (Fig. 2, position 2). The greatest impact of the oryzalin treatment was in the cytoplasm between nucleus and statoliths and in the cytoplasm around the nucleus. Between the nucleus and the tip, oryzalin reduced the viscosity in the longitudinal direction from 6 ± 0.3 mP·s (control) to 4.4 ± 0.1 mP·s (Fig. 2, position 3). The transverse viscosity decreased from 15 ± 1.3 mP·s to 6 ± 0.2 mP·s, and the viscosity near the cell wall (29 ± 5 mP·s) decreased to less than half. After application of oryzalin, the nucleus increased in size so that the space between nucleus and cell wall was reduced. The swelling coincided with an increase in viscosity from 60 ± 6 to 84 ± 5 mP·s. Around the vacuole, the viscosity decreased to half of the control values (62 ± 8 to 35 ± 2 mP·s, Fig. 2, position 6). The viscosity in the tip region of the rhizoid did not change, regardless of direction.
DISCUSSION
The fluid-mechanical properties of the cytoplasm are most important for the dynamics of cellular organization, mixing, transport and stability of intracellular arrangements. Therefore, a careful analysis of the viscous properties of the cytoplasm is likely to enhance our understanding of diverse cellular properties and functions.
The anisotropic viscosity and the influence of F-actin indicate that the cytoplasm is non-Newtonian. Therefore, the measurements describe the viscosity of the cytoplasm for particles 1 µm in diameter, which is the approximate size of many of the individual cellular organelles, most importantly, of the statoliths. Thus, it is important to distinguish microviscosity (important for the movement of small molecules) from intermediate viscosity (important for the movement of proteins; Luby-Phelps, 2000
) and bulk viscosity (important for the movement of organelles; Kamitsubo and Kaneda, 1987
).
In addition, the viscosity may depend on the rate of shear. In thixotropic solutions, the viscosity decreases as the rate of shear increases. These constraints limit the usefulness of a Newtonian calibration for non-Newtonian media, such as the cytoplasm. Nevertheless, the rates of movements of the magnetic beads were similar and responded to the depolymerization of the cytoskeleton so that with a good approximation, the rate of shear primarily depended on the magnetic field moving the beads. Because the same magnet and distance were used in all measurements (replicates in the same rhizoid and repeats in identical regions of several rhizoids), the viscosity values represent a meaningful and reproducible average. The consistency of measurements in each condition indicates that previous shear effects, e.g., dragging beads to the measurement position, did not affect subsequent measurements.
Previous investigations of the cytoplasmic properties of the Chara rhizoid have used optical tweezers (i.e., a focused laser beam), which displaced endogenous statoliths (Leitz et al., 1995
; Braun, 2002
). Although this elegant, noninvasive approach can move statoliths within the cell, it is limited to the area between the tip and the nucleus (Leitz et al., 1995
), and the documented interference between endogenous statoliths and the actin network (Braun, 2002
) is likely to affect measurements. The application of high-intensity laser light to the rhizoid damages the cell, reduces growth, and leads to swelling of the tip (Leitz et al., 1995
). Additionally, the heat load is likely to affect the properties of the cytoplasm and cellular functions.
In contrast, microinjection of particles into the rhizoid overcomes these limitations. The measurement system can be calibrated (Fig. 1) and uses inert beads that, unlike statoliths, are not coated with proteins and thus are not affected by direct cytoskeletal interaction. Also, the measurements can be performed for an extended period of time without actin–myosin interactions, and the beads can be positioned anywhere within the rhizoid, which permits detailed mapping of its viscosity.
Because no subcellular viscosity values for plant cells are available, it is difficult to compare our data. Previous studies have estimated the viscosity of maize cytoplasm to be between 1 and 20 mP·s (Sack et al., 1985
). In animal cells, experimental data range from 10 mP·s for sea urchin eggs (Hiramoto, 1969) to 10 000 mP·s for the squid axon (Sato et al., 1984
). Although these data exemplify the wide range of cytoplasmic viscosities in different cells, our data are unique in that they describe the range of viscosities in discrete domains within a single cell.
The lowest viscosity in Chara rhizoids was found in the zone of cytoplasmic streaming, where the energy-dependent streaming process benefits from low friction. In the area around the nucleus, viscosity was considerably higher (Fig. 2). The high viscosity between nucleus and cell wall may assist with the anchoring of the nucleus and contribute to the molecular continuity between plasma membrane and nucleus. This region represented the barrier past which statoliths were not movable by optical tweezers. The elevated viscosity near the nucleus may affect gene expression and enhance molecular crowding (Ingber, 1997
; Minton, 1998
).
Another area of high viscosity was associated with the cytoplasmic ring around the vacuole (62 ± 8 mP·s), which isolates the vacuolar domain from the remainder of the cytoplasm. The high viscosity in this region could stem from the lining of proteins along the cell membrane, which may be necessary for generating the force that produces cytoplasmic streaming. The anchoring of proteins to the plasma membrane is likely to enhance the viscosity for particles that are larger than proteins.
Leitz et al. (1995)
found a pattern of viscosity that was similar to the one described here and concluded that the interaction between statoliths and the actin network is based on an active, acto-myosin-based system, which controls the suspension of statoliths. Because magnetic beads were less likely than endogenous statoliths to interact with F-actin, the anisotropy observed with beads and endogenous statoliths corroborate one another. However, our data also indicate that the rheological properties are not exclusively controlled by the acto-myosin system, as indicated by the high viscosity values after F-actin depolymerization.
Latrunculin B significantly reduced the longitudinal and transverse viscosity in the statolith region but not to the level of the surrounding cytoplasmic domains (center or tip). This reduction may result from the higher fluidity of g-actin monomers (Wilhelm et al., 2003
). The residual viscosity after cytoskeletal depolymerization may depend on other cytoskeletal or cellular compounds. In contrast to its effects in the statolith region, LatB reduced the transverse viscosity in the tip region almost to the value in the streaming zone (see Fig. 2). The disparity of the LatB effects on the transverse and longitudinal directions in the tip and statolith area suggests that the longitudinal viscosity results at least partially from some other, as of yet unidentified, component or condition or that LatB incompletely depolymerizes specific F-actin or differentially degrades different actin isoforms (Foissner and Wasteneys, 2007
). The reduced longitudinal viscosity in the tip domain is likely to facilitate vesicular transport to the growth region (apical dome) of the rhizoid (Bartnik et al., 1990
). Similar to its effects in the statolith region, LatB reduced the viscosity in the tip region but the viscous anisotropy and a relatively high viscosity persisted. The remaining viscosity may be related to the membrane (endoplasmic reticulum) system and vesicle density, both factors that determine non-Newtonian behavior.
Oryzalin had no effect on the viscous properties in the rhizoid tip, most likely because the microtubule network does not fully extend into this region but ends close to the area surrounding the endogenous statoliths (Braun and Wasteneys, 1998
). The highest reduction in viscosity occurred in the cytoplasmic region between nucleus and statolith zone, indicative of the high abundance of microtubules in this area. The reduction of viscosity near the cell wall suggests that the depolymerization of cortical microtubules leads to a reduction in viscosity similar to that caused by the depolymerization of F-actin. The enlargement of the nucleus after microtubule depolymerization suggests that the microtubules are involved in the shaping and suspension of the nucleus.
Because depolymerization typically reduced viscous friction, the filamentous organization seems to be necessary for optimal interaction with artificial beads and other similarly sized cellular particles such as vesicles and endogenous statoliths. The exception to the generally decreasing viscosity is the increase in the longitudinal viscosity near the nucleus. The observed swelling of the nucleus inevitably reduces the available cross-sectional area between the nucleus and cell wall, which may reduce fluidity and lead to an apparent increase in viscosity. Instead of increased resistance between magnetic particle and fluid, the enhanced viscosity is likely to result from friction between the particle and static, nonfluid structures such as membrane-bound proteins or membrane-attached cytoskeletal elements. Although understanding cellular responses to a wide variety of mechanical stimuli is important, there is no consensus regarding the best manner in which to measure and interpret mechanical properties in plant or animal cells (Hoffman et al., 2006
). In addition to the viscous drag that affects cellular transport, frequency dependency alters the stiffness of the cytoskeleton (Shafrir and Forgacs, 2002
) and modifies signal transduction and graviresponse (Ma and Hasenstein, 2007
). Because of the many interactions between mechanical properties, a comprehensive mechanical analysis is difficult and individual aspects will have to be tested and integrated into a general framework that illustrates mechanical aspects of cellular functions. The following discussion illustrates the potential effect of viscous properties on the gravisensing system.
The cytoskeleton as dynamic gravisensing system
The statoliths in the Chara rhizoid are composed of barium sulfate (Sievers and Schmitz, 1982
), and embedded in a protein and carbohydrate matrix (Wangcahill and Kiss, 1995
). The statoliths are suspended by prominent, longitudinally oriented F-actin bundles (Braun and Wasteneys, 1998
). These longitudinally oriented actin bundles cause statolith suspension since LatB treatment results in immediate sedimentation of the statoliths into the apical dome (Kuznetsov and Hasenstein, 2001
). The strong reduction of viscosity after LatB application suggests that F-actin contributes to the viscous properties in the statolith domain and to the transverse sedimentation of statoliths. The weaker anisotropy after LatB application (see Fig. 2) may result from incomplete depolymerization of F-actin.
The much lower transverse than longitudinal viscosity is likely to polarize transverse movement and sedimentation upon reorientation to the sensitive site of the plasma membrane (Braun, 2002
). However, the high viscosity in the statolith region is likely to attenuate movements of individually moving statoliths and thus to enhance the concerted activity of statoliths. Bulk movement of the statoliths is likely to produce a stronger signal than movement of individual statoliths. This argument is supported by a similar effect in the tips of negatively gravitropic protonemata of the moss Ceratodon (Kern et al., 2001
), where under microgravity conditions, the number of amyloplasts in the sensitive subapical zone increases, presumably to enhance the overall sensitivity of the gravisensing system. Because the statoliths are actively positioned in the most viscous zone of the rhizoid and the continuous activity of the acto-myosin system prevents sedimentation of the statoliths, the statoliths may provide a continuous signal that constantly verifies the gravity vector. The interaction with the acto-myosin system leads to the suspension of statoliths in the Chara rhizoid and causes saltatory movements of amyloplasts in columella (Sack et al., 1985
) or endodermis cells (Kato et al., 2002
). On the basis of these observations, gravisensing is a dynamic process that depends on the continuous interaction of the acto-myosin system with statoliths, rather than simply on their sedimentation. Viscosity, in addition to gravity and buoyancy, affects the movement of statoliths and amyloplasts and contributes to the organization of rhizoids.
FOOTNOTES
1 This research was supported by NASA grants NAG10–0190 and NAG2–1423. 
2 Corresponding author (e-mail: hasenstein{at}louisiana.edu
)
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