Microtubules in root hairs

In contrast to most plant cells, root hairs are supposed to grow at their tip only (Sievers & Schnepf, 1981). At the tip the cellulose microfibrils are deposited in a random pattern, forming the primary cell wall. Behind the tip the microfibrils of the secondary wall are deposited in a regular pattern, giving rise to well-defined wall textures, varying from axial to helical or helicoidal (Sassen, Pluymaekers, Meekes & de Jong-Emons, 1981). In root hairs these specific wall textures have been interpreted as structural adaptations to obtain various mechanical properties, as required by environmental conditions (Sassen et al. 1981).

The deposition of microfibrils is generally thought to be under the control of the cortical cytoskeleton. Support for this concept comes from the observed alignment between the cortical microtubules and newly deposited cellulose microfibrils in cell walls (Heath & Seagull, 1982). Thus, since the orientation of the nascent cellulose microfibrils in helicoidal walls of root hairs changes intermittently, a corresponding change in direction of the cortical microtubules is to be expected. However, in Equisetum hyemale root hairs, microtubules are axially oriented, whereas microfibrils are found in either helicoidal or helical configurations (Emons, 1982; Emons & Wolters-Arts, 1983).

In Raphanus sativus axially oriented microtubules are found in the part of the root hairs where microfibrils are deposited with random orientation (Newcomb & Bonner, 1965). However, it has been suggested that in this species the microtubules control microfibril deposition at a later stage, when microfibrils become oriented axially (Seagull & Heath, 1980; see also: Heath & Seagull, 1982; Robinson & Quader, 1982).

The observations, especially on Equisetum, contradicted present models of microtubular control of microfibril deposition and prompted a further investigation of the cortical microtubules in root hairs. Here we present our results, not only from Raphanus and Equisetum, but also from other species with axial (Lepidium), helicoidal (Limnobium, Allium and Ceratopteris) and helical wall textures (Urtica). Because of the limitation of oblique/serial sectioning, we chose to investigate the cytoskeleton using primarily immunofluorescence techniques (Wickei al. 1981; Wick & Duniec, 1983) in order to visualize the entire cytoskeleton at the light-microscopical level. In addition, we have used the dry-cleaving technique, with special attention to the cortical microtubules, at the electron microscope (EM) level (Traas, 1984; Traas, Braat & Derksen, 1984). The relation between the pattern of microtubules and microfibril orientation and other morphogenetic and physiological properties of the cell is discussed.

Root hairs of the following species were used: Equisetum hyemale (Equisetaceae), Ceratopteris thalictroides (Parkeriaceae) Limnobium stoloniferum (Hydrocharitaceae), Allium sativum (Liliaceae), Raphanus sativus and Lepidium sativum (Brassicaceae) and Urtica dioica (Urticaceae).

Cuttings of Equisetum, Ceratopteris, Limnobium and Urtica were cultured on an aqueous soil extract (H. Meekes, unpublished) under greenhouse conditions. Roots with full-grown hairs developed within about 2 weeks.

Seeds of Allium, Raphanus, Lepidium and Urtica were germinated at room temperature in the dark on wet filter paper. Raphanus and Lepidium developed roots in 24h. Hairs with maximum length were generally present within 48 h. Allium developed roots of about 1 cm within a week. Root hairs appeared after slight drying within 24 h (see also Rosene, 1954). From Urtica only a few seeds germinated. Roots and root hairs developed very irregularly; the first hairs appeared 1 week after germination.

Rootlets with hairs were fixed immediately after the first root hairs obtained their maximum size. From Equisetum, Limnobium and Urtica also older roots with full grown, persistent hairs were used.

Root hairs were prepared for immunofluorescence microscopy essentially as described previously (Wick & Duniec, 1983; Lloyd, 1983). We used 20mM-potassium phosphate buffer (pH6·8), with 5–10 mM-EGTA and 2–5 mM-MgCl₂, but occasionally we used Pipes instead of phosphate buffer.

To degrade the cell wall 5 % cellulase (ONOZUKA R–10; Serva, Heidelberg, FRG) or 1–3 % cellulysin (Calbiochem-Behring, La Jolla, U.S.A.) was used. The primary antibody was a monoclonal anti-tubulin (Mas 077; Sera Labs) and the second antibody was a rabbit, fluorescein isothyocyanate (FITC)-labelled anti-rat immunoglobulin G(IgG) (Nordic Labs. BV; Tilburg, the Netherlands). Preparations were examined under a Leitz Orthoplan microscope with an appropriate filter combination and illumination, using Leitz 50 × and 100 × water-immersion lenses. Photographs were taken with a Leitz Vario Orthomat combination on Agfapan professional film 400 Asa.

For thin sectioning, root hairs of Equisetum, Ceratopteris and Limnobium were fixed in glutaral dehyde/OsO₄, dehydrated and flat-embedded in Spurr’s medium according to standardized procedures (Pluymaekers, 1982; Emons, 1982). Sections were stained with uranyl acetate/lead citrate.

For dry cleaving mostly phosphate, but also cacocylate and Pipes, was used as a buffer. Roots were fixed in glutaraldehyde in buffer, post-fixed with tannic acid, washed and root parts with hairs were attached to poly-L-lysine-coated grids (Traas, 1984). After fixation in OsO₄, the specimens were stained with uranyl acetate, dehydrated and critical-point dried. Finally, preparations were cleaved and examined immediately (Traas, 1984) using a Philips EM300 or EM201 electron microscope.

Micrographs of cleaved cells were printed at a final magnification of about × 25 000, the exact values being established using a grating replica. Microtubules were traced from the micrographs on plastic sheets. Lengths, orientations and relative abundance (expressed as total length/per area (μm/μm²)) were measured with a Kontron Videoplan computer (Traas et al. 1984).

Except for Allium and Urtica all growing roots showed the typical cones of hairs at the tip of the root. The older hairs of Raphanus and Lepidium died some time after reaching their maximum length. Ceratopteris, Limnobium and Urtica cuttings developed root hairs at the tip of the root, which persisted for at least 3 weeks before being degraded, while Equisetum root hairs persisted for over 3 months. On roots of Urtica seeds cone-like groups of hairs were only occasionally formed at the tip. Hairs grew, as in cuttings, over large areas of the root and remained intact for long periods (weeks). Moreover, root hair development was very irregular: young hairs appeared over the entire root between older or even full-grown hairs. Cuttings of Urtica showed more regular root-hair development. On Allium, roots hairs developed not only in a cone at the tip, but also at the transition point of root to hypocotyl and subsequently also over the entire root. As in Urtica, hairs developed between older or even full-grown hairs. Root hairs were regularly observed that had either increased in diameter or decreased in girth during growth, giving rise to slightly cone-shaped hair forms.

If formaldehyde was used as a fixative, especially in Equisetum, Ceratopteris and Urtica, plasmolysis often occurred, resulting in collapse of the cytoplasm and disin-tegration of the cytoskeleton. In order to reduce plasmolysis as much as possible a concentration of 20 mM-phosphate was used in combination with formaldehyde. If buffers other than phosphate were used, i.e. 25 mM-Pipes for immunofluorescence and 0·l M-cacodylate or 50mM-Pipes for dry cleaving, no difference in microtubular organization could be observed. Since plants have considerable amounts of calcium in their walls and in their apoplasts, we always used EGTA in the fixative. The presence of EGTA cannot be expected to induce artificial polymerization of tubulin (see, e.g. Solomon, Magendantz & Salzman, 1977).

We never included GTP in our buffers since this might have induced tubulin polymerization (Solomon et al. 1977; for discussion, see Seagull & Heath, 1980). The susceptibility of the cell wall to degradation appeared to be highly variable in different species. Prolonged treatment in fixative and/or high concentration of enzyme were always effective in degrading the cell wall, but these treatments could also destroy the cytoskeleton. In immunofluorescence preparations of Equisetum and Ceratopteris the cytoskeleton often seemed disturbed, probably due to the fragility of the cells after degradation of the cell wall. For Limnobium, immunofluorescence preparations could only be made from non-enzyme-treated segments cut from the hairs under the dissection microscope.

In all dry-cleaving preparations, and also in sectioned material, both microtubules and endoplasmatic reticulum, mitochondria, coated pits, coated vesicles and other filamentous structures were observed. These results are similar to those described previously for other plant cells after dry cleaving (Traas, 1984; Traas et al. 1984) (see Figs 1–7). Differences in appearance between the preparations of the various species used may depend to a large extent on density and distribution of the cytoplasm; dense and irregularly distributed cytoplasm led to irregular cleavage planes, whereas hairs with a large central vacuole mostly cleaved over the vacuole membrane. Sometimes the membrane-attached cytoplasm had a granular structure and stained darkly (see Figs 1 and 7), possibly due to a strong reaction between membrane-attached cytoplasm and tannic acid. Quantitative measurements were carried out on preparations from Equisetum, Limnobium and Raphanus only, since in other species proper dry-cleave preparations could not be obtained (Ceratopteris and Urtica) or the cytoskeleton varied greatly between individual hairs of the same species (Allium and Urtica).

In all preparations, using immunofluorescence and/or dry cleaving, ordered patterns of microtubules were seen. In immunofluorescence preparations root hairs often showed no microtubules in the tip, or displayed fuzzy stained tips, probably due to preparation artefacts. There were always some root hair tips that showed more or less clear microtubule patterns. Since striking differences between the various species exist, the cytoskeleton of each of the various species is described individually below.

These closely related species gave identical results. In immunofluorescence preparations net-axial oriented microtubules were seen with no, or almost no, microtubules protruding into the extreme tip, but behind the tip a densely stained zone was often present (Fig. 1A). A net-axial orientation was also observed in dry-cleave preparations (for Raphanus, see Fig. 1B, c; for Lepidium, see Traas, 1984). Microtubules could deviate considerably from the cell axis, up to 30° (see Figs 1c, 8A). In many cases there were even crossing microtubules (Fig. 1c, D). No differences in orientation between various length classes could be observed, though the smaller microtubules showed a larger deviation from the main direction. Lengths and relative abundance are presented in Table 1. They appeared to be highly variable. In growing hairs, however, the relative abundance (average: 4·00μm/μm²) was clearly higher than in full-grown hairs (average: 0·68μm/μm²). Raphanus and Lepidium hairs cleaved very irregularly, often leaving dark patches of cytoplasm on the membrane (Fig. 1c). Axially oriented strands of cytoplasm with mitochondria and bundles of microtubules were regularly observed (Fig. 1c). Often the fine structure was obscured by a heavily stained precipitate (Fig. 1D).

In both immunofluorescence and dry-cleaving preparations from growing and full grown hairs a net-axial direction of microtubules was observed (see Figs 2, 3). In immunofluorescence preparations microtubules were seen protuding into the tip (Fig. 2A). In a few instances we were able to cleave root hair tips. In these preparations short and more or less randomly distributed microtubules were observed (data not shown). Microtubules in growing hairs (average 2·47μm) tended to be shorter than in full-grown hairs (3·60μm; see Table 1). The relative abundance of the microtubules in growing hairs (ranging from l·02 to 2·14μm/μm²) was higher than in full-grown hairs (0·6l–l·33μm/μm²; see Table 1). Microtubules deviated con-siderably (up to 90°) from the main direction in growing hairs and up to 70° in full-grown hairs (see Figs 2A, 3A, C; see also Fig. 8), and crossing microtubules were even observed (Fig. 2c). Differences in the main orientation between the various length classes were not observed, but smaller microtubules showed a larger spreading from the main direction. Growing hairs with a relatively few small vacuoles cleaved at a plane just above the membrane, leaving only traces of cytoplasm on the membrane surface. The microtubules were more or less homogeneously distributed (see Fig. 2B).

Full-grown hairs cleaved over the membrane of the vacuole leaving the cortical cytoplasm with its organelles almost intact (see Fig. 3A). In full-grown hairs strands of cytoplasm with numerous mitochondria and bundles of microtubules were seen (see Fig. 3A, B). Outside the cytoplasmic strands microtubules were not usually present in bundles (compare Figs 3B, c). Occasionally, microtubules were accompanied by a presumptive m?crofiiament with a diameter of about lO?n (Fig. 2c). Sometimes, small bundles of presumptive microfilaments were observed, also in oblique sections (data not shown).

In both immunofluorescence and dry-cleave preparations, microtubules with a net-axial orientation were found (Fig. 4A, B). In immunofluorescence preparations microtubules were found to protrude into the tip. In the few hair tips that we were able to cleave, short, randomly distributed microtubules were observed (data not shown). The lengths of the microtubules (average 2·20 μm) were about the same as in growing hairs of Equisetum (see Table 1). The relative abundance appeared to be very low (from 0·11 to 0·30 μm/μm²) as compared to that inRaphanus and Equisetum (see Table 1). Also in Limnobium microtubules deviated considerably from the main direction (up to 90°; see Figs4B, c, 8). Here, too, crossing microtubules (Fig. 4B) and microtubules accompanied by a presumptive microfilament (diameter, 100 Å) were observed that seemed to split into protofilaments (Fig. 4F). In full-grown hairs the microtubules retained their axial orientation. Plasma strands with mitochondria and with microtubule bundles were seen regularly (not shown), but less frequently than in Raphanus or Equisetum. Huge, mainly axially oriented bundles of putative microfilaments (individual diameters, 100Å) were often observed in dry-cleave preparations (Fig. 4c, E), and also in sectioned material (not shown).

Immunofluorescence preparations invariably showed net-axial orientations of microtubules (see Fig. 5A). In oblique sections axial orientations were also observed (Fig. 5B). In immunofluorescence preparations microtubules were observed that had protruded into the tip of the hair (Fig. 5A).

Urtica mainly showed net-axially oriented microtubules, but helical patterns were also present in the same preparations (see Fig. 6). No differences were found between hairs of seeds and cuttings. In older roots with persistent hairs, both axial and helical patterns were observed. No obvious difference in relative abundance between hairs with helical and axial patterns could be seen. Microtubules with axial orientations protruded into the tip of the hair, whereas microtubules with helical orientations passed uninterrupted through the tip. Though slight variations occurred, the orientations generally remained the same in the entire root hair.

The microtubules of Allium root hairs showed various patterns: in both immunofluorescence and dry-cleave preparations helical patterns with an angle of about 45 º were regularly seen, but axial and transverse patterns were also observed (Fig. 7). Similar observations were made also in older roots with persistent hairs.

The axial patterns were very similar to those in the other species studies here. The helical pattern was the same as that in Urtica. Sometimes, the direction of the microtubules differed considerably within one root hair (see Fig. 7c).

In hairs with axial patterns the density of the microtubules was conspicuously higher than in hairs with other orientations (compare Fig. 7B and c, and D and E). Axial microtubules protruded into the tip of the hair, whereas helically oriented microtubules passed uninterrupted through the tip. In dry-cleave preparations very often a precipitate covers the membrane-attached structures, often even obscuring the tubular structure of the microtubules (see Fig. 7E) and increasing their apparent diameter.

Irrespective of their orientation, microtubules could form bundles and, though no quantitative estimations were made so far, no clear indication of an important difference in length as compared to other root hairs was found.

Using immunofluorescence and dry-cleaving we were able to show the presence of highly ordered microtubule arrays in root hairs of various species. In dry-cleave preparations, microtubules and regularly distributed endoplasmic reticulum, coated pits, coated vesicles and thin filaments attached to the plasmalemma were always observed. So far such observations on large areas of the cytoplasmic side of the plasmalemma can be made only by dry cleaving methods. Similar observations have been made in various root cell types (Traas, 1984; Traasetal. 1984). The density and distribution of all these structures differ between cells or even within one cell, depending on the species and physiological conditions. In the present investigation we studied mainly the microtubule system because of its presumptive role in microfibril orientation.

The lengths of the microtubules were within the range of those measured previously for Raphanus root hairs (Seagull & Heath, 1980). The lengths of the microtubules reported here for growing root hairs of Raphanus and Equisetum must be considered minimum values, since microtubules broken off at the edge of the preparation and microtubules partly covered by cell organelles were included in our measurements. The dry-cleaving procedure may have caused some breakage of microtubules, especially when the cells cleaved just over the membrane, as in Equisetum. The frequencies (microtubules/μm as computed from our data) of the microtubules in growing hairs of Raphanus (from less than 1 /μm to 7/μm) showed a larger range than those in a previous report (2/μm to 4/μm) (Seagull & Heath, 1980). Still, the average frequency calculated from our data (5/μm) is higher than reported by Seagull & Heath (1980) (3/μm).

The root hairs studied show strong differences in cell wall texture: axial in Raphanus and Lepidium (Sassen et al. 1981); helicoidal in growing hairs of Equisetum, Ceratopteris, Limnobium (Sassen et al. 1981) and Allium (Sassen et al. unpublished data). Helical textures are found in full-grown hairs of Equisetum, with an angle of about 25 ⁰ to the cell axis (Emons & Wolters-Arts, 1983) and in Urtica with an angle of about 20 º to the cell axis (Sassen et al. 1981). In the root hairs that we have used the microfibril depositing zone was always included. The microtubules in the hairs are axial, with the exception of Allium and Urtica. Our results are consistent with previous reports (Seagull & Heath, 1980; Emons, 1982). A comparison of microtubule and microfibril orientation is given in Table 2. The results show that there is not necessarily a co-alignment between microtubules and microfibrils in hairs depositing helicoidal or helical walls. Since all models involving microtubules in microfibril orientation require, at least temporarily, alignment between microtubules and microfibrils, our results cannot be accepted as support for these models (for reviews see: Heath & Seagull, 1982; Robinson & Quader, 1982; Lloyd, 1984; see also Emons, 1982; Emons & Wolters-Arts, 1983). It cannot be excluded, however, that microtubules play some role in microfibril orientation. It may well be that microfibrils are orientated by microtubules in combination with another membrane adherent cytoplasmic component. It is suggested that microtubules act together with microfilaments and thus give rise to the complicated textures observed in cell walls.

In Urtica and Allium root hairs, not only was the orientation of the microtubules different from that in other species but also large differences between individual hairs, and in Allium even within one hair, were observed. The microtubules always ran through the tip of the hair. The same observations have also been made in expanding root cortex cells from Raphanus, and also Urtica and Allium (Traas et al. 1984; see also Lloyd, 1984), suggesting that the root hairs of Allium and Urtica, at least temporarily, are not solely tip-growing but might also stretch to some extent. The variable microtubule helices in Urtica hardly align with the 20° helix of the microfibrils (Sassen et al. 1981), but at least a temporary alignment between microtubules and microfibrils cannot be excluded. Also in Allium an alignment between microtubules and microfibrils cannot be excluded. Unfortunately, no discrimination between growing and full-grown hairs could be made, since they are found intermingled on the rootlets. Such root-hair formation seems to be rather atypical (compare: Cormack, 1949, 1962).

The patterns of microtubules observed in this study may relate to various other cell properties, such as speed of growth (Lloyd, 1983, 1984), distribution and transport of cytoplasm within the cell (e.g., see Heath & Heath, 1978; Mizukami & Wade, 1983; Traas et al. 1984), cell expansion (Gunning, 1981; Marchant, 1974; Traas et al. 1984) and cell polarity (Sievers & Schnepf, 1981; Mizukami & Wada, 1983).

A relation between growth speed and microtubule orientation can neither be excluded nor corroborated, since no exact data concerning growth speed of the species described here are available.

In Raphanus, Lepidium, Limnobium and Equisetum axially oriented plasma strands with many mitochondria can be seen. These strands contained bundles of microtubules, whereas in the strictly cortical cytoplasm outside the plasma strands fewer and more homogeneously distributed microtubules were seen. Such a relationship between plasma strands and microtubule distribution has also been described for Uromyces (Heath & Heath, 1978) and Bryopsis (Mizukami & Wada, 1983). A cooperative motile function for microtubules and microfilaments has been proposed in several reports (e.g., see Neuhaus-Uri & Kiermayer, 1982). A relation between microtubules and microfilaments is also suggested by our own observations showing that microfilaments often coalign with microtubules. Further-more, in Limnobium large filament bundles occur in combination with low microtubule densities. This could mean that these bundles have taken over part of the microtubule function.

In hairs of all species microtubules may protrude into the tip. In the tips of Equisetum and Limnobium relatively small and randomly distributed microtubules were observed. Randomly distributed microtubules have also been found in other globular cells, such as protoplasts (Valk, Rennie, Conolly & Fowke, 1980; Lloyd, Slabas, Powell & Lowe, 1980). In expanding cells microtubules orientate transversely to the axis of expansion (Lloyd et al. 1980; Simmonds, Setter-field & Brown, 1983; Gunning, 1981). It may well be that a similar relationship exists in many root hairs; in the extreme tip of the hairs the expansion may be the same in all directions, whereas in the lower part of the dome the expansion may be transverse (circumferential) to the cell axis with, accordingly, axial orientation of the microtubules.

Such ordered patterns of microtubules are probably involved in the maintenance of cell polarity. Indeed, in Bryopsis colchicine treatment leads to a loss of polarity, to be restored only after removal of colchicine from the medium and restoration of the microtubules (Mizukami & Wada, 1983).

It must be emphasized that the different possible functions of the microtubules are not mutually exclusive; their expression may depend on the demands of the moment. The observations on Allium and Urtica, especially, indicate that root hairs are much more versatile than is generally assumed.

We thank Dr M. M. A. Sassen for support in various ways, Dr G. Egberink for her help with the videoplan and Dr G. Barendse for critical reading of the text. We also thank Mrs H. Uyen-de Vaan and Mrs J. Broekmans for typing the manuscript.

J,D. is indebted to Dr C. Lloyd (Norwich) for help with immunofluorescence studies. This work was supported by the Dutch Organization for Advancement of Pure Research, ZWO (grant no. 14.25.007, to J.A.T.).