- Email: [email protected]
The ultrastructure of the mucilaginous layer on plant roots
Soi Bid. Biochem. Vol. 4, pp. 443-449. Pergamon Press 1972. Printed in Great Britain
THE ULTRASTRUCTURE OF THE MUCILAGINOUS LAYER ON PLANT ROOTS M. P. GREAVESand J. F. DARBYSHIRE The Macaulay Institute for Soil Research, Craigiebuckler,
Aberdeen, Scotland
(Accepted 16 May 1972) Sunnnary-A mucilaginous external layer or mucigel was observed on the roots of all 16 species of common agricultural crop plants examined. On axenic roots, the mucigel appears in the electron microscope as an unevenly distributed layer of granular and fibrillar material covering the outer surface of the root and most root hairs. The greatest quantity of mucigel was found around the root cap. Apart from the presence of soil particles adhering to the external surface of the mucigel on soil-grown plants, structural differences were seldom observed between the mucigels on plants grown in soil, sand or nutrient solution. Some soilgrown axenic peas (Pisum sativum L.) and mustard (Sinapis alba L.) however, had particulate and amorphous soil material distributed throughout the mucigel. When plant roots were colonized by Pseudomonas sp., Cytophaga johnsonii Stanier or a mixture of several micro-organisms, larger quantities of mucigel developed than on axenic roots of the same plant species. The number of root-surface cells, which were either dead or showed changes in the structure of their walls, was also greater when micro-organisms were present. In the root-elongation zone and in other zones where microbial colonization was sparse, the mucigel retained its usual appearance near micro-organisms. In densely colonized regions of the mucigel, however, the region immediately around the micro-organisms could usually be differentiated from the bulk of the mucigel. When micro-organisms were present the mucigel often had a distinct outer boundary.
INTRODUCTION
presence of an outer mucilaginous layer on the roots, especially the root caps, of Angiosperms has been reported by many investigators in the last decade (Dart and Mercer, 1964; De Felipe and Grossenbachet, 1964; Scott, 1965; Juniper and Roberts, 1966; Leiser, 1968; Samtsevich, 1968; Brams, 1969; Floyd and Ohlrogge, 1970, 1971). Jenny and Grossenbacher (1963) showed that this mucilaginous layer or ‘mucigel’ on barley roots, which were grown in bentonite or permutite, sometimes completely filled the space between the root and the surrounding particulate medium. The possibility that micro-organisms may influence the formation and structure of the mucigel has received little consideration in the past. This paper reports the results of a survey of the ultrastructure of the root mucigel of 16 species of agricultural crop plants. It includes the first report dealing with the mucigel on axenic roots grown in soil. THE
MATERIALS
AND METHODS
Rooting media
Three different rooting media, soil, sand and nutrient solution, were used for the plant species listed in Table 1. These plants were grown for 2-4 weeks in a glasshouse with ambient air temperatures between 15 and 25”C, except when peas, Pisum sativum L., were grown in nutrient solution (see below). Glasshouse experiments were done three times. Soil. Soil for axenic soil studies was collected from the surface 15 cm of the Institute 443 SOIL 414-E
M. P. GREAVES
444 TABLE
AND J. F. DARBYSHIRE
1. PLANT SPECIESGROWN IN GLASSHOUSE,WITH ROOTINGMEDIA USED Plant species
Rooting
N, SO
Maize, Zea rnays L. ‘Kelvedon 3 13’ Timothy grass, Phleum prutense L. S50 Winter wheat, Triticum aestivum L. ‘Capelle-Desprez’ Onion, Al&m cepa L. Cultivar is unknown Pea, Pisum sativum L. ‘Tall, sugar white’ Pea, P. sativum L. ‘Meteor’ Pea, P. sativum L. ‘Foremost’ White mustard, Sinapis alba L. Cultivar is unknown Tomato, Lycopersicum esculentum Mill. ‘Market King’ N = nutrient solution, soil.
SA = sand,
SO = non-irradiated
media
SA, SO, SG SA, SO, SG N N, SA, SO, SG N. SA, SO, SG N SA, SO, SG SA, SO, SG soil,
SG = y-irradiated
garden, air-dried, passed through a 3-mm mesh sieve, sealed in nylon packets in 100 g amounts and sterilized by y-irradiation (2.5 Mrad). The contents of these packets of soil were transferred individually to sterile 2 *5 cm dia. boiling tubes plugged with cotton wool. The soil in each tube was remoistened with sterile glass-distilled water and planted with one axenic seedling (Darbyshire and Greaves, 1971). Soon after the plumules had emerged, two thirds of the seedlings were each inoculated with 0.1 ml of a 24-h nutrient broth (Oxoid Ltd, London) culture of either Pseudomonas sp. or Cytophuga johnsonii Stanier. The remaining axenic seedlings each received 0.1 ml of sterile nutrient broth. Pseudomonas sp. and C. johnsonii were isolated from the roots of timothy grass, Phleum prutense L. S50, (Darbyshire and Greaves, 1970) and cocksfoot, Ductylis glomerutu L. S143 (Webley, Duff, Bacon and Farmer, 1965) respectively. A laminar air-flow apparatus (Microflow Limited, Fleet, Hants., England) was used to facilitate the axenic manipulations. Plants were also grown in unsterilized soil from the Institute garden in the manner described above except that the packaging and y-irradiation were omitted. Roots were also sampled on two separate occasions, from plants (Table 2) growing in the Institute garden and from agricultural crops of barley and wheat at Methlick, Aberdeenshire (Thistlyhill and Tarves Series respectively of the Tarves Soil Association). TABLE 2. PLANT SPECIESGROWN
IN
FIELD OR GARDEN SOIL
Spring barley, Hordeum vulgare L. ‘Golden Promise’ Winter wheat, Triticum aestivum L. ‘Champlein’ Winter wheat, T. aestivum L. ‘Jos Cambier’ Potato, Solunurn tuberosum L. ‘Maris Peer’ Potato, S. tuberosum L. ‘Sharpe’s Express’ Onion, Allium cepa L. ‘Ailsa Craig’ Leek, AIlium porrum L. ‘Musselburgh Pea, Pisum sativum L. ‘Admiral Beatty’ Brussels sprouts, Brussica olerucea L. ‘Cambridge Special’ Savoy cabbage, B. oleracea L. ‘Ormskirk, Late Green’ Turnip, Brussicu rupa L. ‘Milan, Extra Early’ Parsnip, Pustinaca sutivu L. ‘Hollow Crown Improved’ Carrot, Daucus curota L. ‘Ideal’ Lettuce, Lactuca sativa L. ‘Webb’s Wonderful’ Broad Bean, Vicia faba L. ‘Bunyard’s Exhibition’
ULTRASTRUCTURE
OF THE MUCIGEL
445
Sand. Axenic and inoculated seedlings were grown in acid-washed, coarse sand (No. 52 W, G. Garside, Sand Quarries, Leighton Buzzard, Beds., England) in the manner described for r-irradiated soil, except that the soil and glass distilled water were replaced by approximately 100 g amounts of coarse sand and nutrient solution (Darbyshire and Greaves, 1970). The sand was sterilized in the tubes by autoclaving at 121°C for 20 min after the addition of the nutrient solution. Nutrient solution. Peas were grown as described by Darbyshire and Greaves (1970) with axenic roots or roots inoculated with Pseudomonas sp. Onions, Allium cepa L., were grown in the single-strength medium of Macklon and Higinbotham (1968) and maize, Zea mays L., in DeKock and Hall’s (1962) medium, except that iron was supplied as ferric versenate (2 parts/106). Both solutions were non-axenic. Electron microscopy. Between three and five roots per plant were selected from two plants as representative material for each plant species investigated. After the roots had been gently rinsed in glass distilled water, five I-mm lengths were cut from each of the following zones: root apex, elongation zone, root-hair zone and basal zone. The preparation for electron microscopy was as described by DeKock, Rutherford and Cheshire (1971). The root samples were fixed in glutaraldehyde (6 % v/v in phosphate buffer, pH 6 -9) for 1’ 5 h at 20°C. Post-fixation was in either osmium tetroxide solution (1% w/v in sodium veronal-sodium acetate buffer, pH 7 -9) for 1 h at 4°C followed by 24 h at 20°C or in ruthenium red (10 mg/l., Sorensen’s phosphate buffer, pH 7 *O)for 2 h at 4°C. After dehydration in a graded ethanol series, samples were infiltrated with and embedded in a Durcupan A.C.M. mixture. Sections (50-80 nm thick) were cut on an Ultrotome (L.K.B. Instruments Ltd, South Croydon, Surrey, England) using glass knives. Difficulty was encountered in cutting sections of roots grown in soil owing to the presence of mineral particles on the root surface. The best results were obtained with thicker (80-120 nm) sections. The use of a diamond knife did not improve the results. Some root samples were stained with uranyl acetate (2% w/v) at the 75% ethanol stage of dehydration. Sections were stained with uranyl acetate (1% w/v, aqueous solution), lead citrate (Reynolds, 1963) or both and examined in an electron microscope (EM6, A.E.I.) at 75 kV. The orientation of elongated soil particles in the mucigel was measured on enlarged prints (cu. 60 x 40 cm) of the original electron micrographs.
RESULTS
Many of the roots examined, particularly those colonized by micro-organisms, showed signs of tissue damage reminiscent of ineffective fixation in the epidermis and outer cortex of the root-hair and basal root zones. The apical meristems and elongation zones were undamaged. It was concluded, therefore, that the damage had occurred before the roots were fixed. Light microscopy of unfixed roots also showed that many of the outer root cells were damaged. Mucigel was present on the roots of all the plant species studied and its ultrastructure was little affected by the type of rooting medium used. There were, however, some distinct structural differences between the mucigels found on axenic and non-axenic roots. Accordingly, the observations reported below apply to all plant species studied unless otherwise stated. Axenic roots. The mucigel on axenic roots appears as an unevenly distributed layer of granular and fibrillar material covering the outer surface of the root and most root hairs. The greatest quantity of mucigel was found around the root cap (Fig. 1). The remnants of occasional dead root cells (Fig. 2) and structures which resemble plant organelles in the
446
M. P. GREAVES
AND
J. F. DARBYSHIRE
mucigel, as well as the loss of wall structure from root surface cells (Fig. 3), suggest that the mucigel is partly derived from root surface cells. The middle lamellae between the rootsurface cells and some parts of the mucigel (Fig, 1) were strongly stained with ruthenium red. As this stain is frequently used to detect pectin it is probable that pectic material is common to both regions. The root mucigels on axenic plants grown in soil and other media could only be distinguished by the presence of particulate and amorphous soil material on the former. These soil particles were usually found adhering closely to the outer surface of the mucigel which was also present in some micro-pores between the adhering particles. On some axenic pea and mustard plants, soil particles were distributed throughout the mucigel (Fig. 4). As very long exposures were required to clearly demonstrate these particles in the electron micrographs, structural detail within the cytoplasm has been lost through over-exposure. These over-exposed areas have been darkened on the back of the electron micrograph plates, using a very soft pencil, to reduce their contrast. It was estimated that 67 % of the elongated particles visible in the mucigel in Fig. 4 were inclined at less than 30” to the tangent to the nearest point on the root surface. If the root hair in Fig. 4 was considered alone the corresponding proportion was 88 %. Many of these elongated particles appear to be platy clay minerals. The proportion of similarly orientated particles in the mucigel on axenic mustard roots was only 30%. Non-axe& roots. The thickness of the mucigel on roots colonized with micro-organisms ranged between 0.5 and 8 pm compared with a maximum of about O-5 pm on all parts of axenic roots except the root cap where it was up to 2.5 pm thick. The number of root surface cells which were either dead or had lost some wall structure was also greater when microorganisms were present. The structure of the root mucigel appeared to be very similar irrespective of whether the root surface was colonized with specific micro-organisms (e.g. Pseudomonas sp., C. johnsonii) or a mixture of micro-organisms which developed in nonaxenic nutrient solution. Microbial colonization usually resulted in some electron-transparent areas in the mucigel close to the microbial cells. As the microbial population increased, these areas became more widespread until eventually the matrix of the mucigel was almost entirely electron-transparent (Fig. 6). The observed exceptions to this generalization are described below. Electron-dense material developed around Pseudomonas sp. when these bacteria colonized the root-cap mucigel of peas grown in nutrient solution (Fig. 5). This material became fess electron-dense further away from the root apex and, in the oldest part of the root, the bacteria were embedded in an almost entirely electron-transparent mu&gel (Fig. 6). The mixed micro-flora colonizing the root-cap mucigel of maize grown in non-axenic nutrient solution and the mucigel on decaying root-surface cells of wheat, barley and maize grown in non-axenic soil were sometimes surrounded by electron-dense material in the mucigel. When white mustard was grown in sand and inoculated with C. johnsonii, the mucigel on the flatter areas of the root surface consisted of alternate thick bands of electron-dense material and thin electron-transparent bands (Fig. 7). Each electrondense layer was colonized by myxobacteria, which were orientated so that their long axes lay parallel to the root axis. The bands in the mucigel on white mustard roots were less obvious where the colonization by C. johnsonii was sparse or where the organism accumulated in crevices between root surface cells. This banded appearance was never observed in the root mucigel of other plant species inoculated with C. johnsonii or with white mustard grown in soil and inoculated with this myxobacterium. Another difference between axenic and non-axenic roots was that there was often a distinct outer boundary to the mucigel when micro-organisms were present (Fig. 8). This
FIG. 1. T.S. Root cap of axenic mustard grown in sand showing thick mucigel (m). Densely stained material (arrows) is present throughout the mucigel and interlamellae (1) between root-cap cells. Tissue postfixed with ruthenium red and stained with uranyl acetate. Section stained with lead citrate. ( x 24,000) FIG. 2. T.S. Root-hair
zone of axenic pea (cv. Foremost) grown in nutrient solution. Remains of a sloughed-off cell (cv) are lying in an unevenly distributed mucigel (WI).Unstained. ( x 4920)
FIG. 3. T.S. Root-surface cell wall in root-hair zone of axenic mustard grown in sand. Cell wall (cw) consists of a framework of micro-fibrils (f). Mucigel (m). Tissue post-fixed in ruthenium red and stained with uranyl acetate. Section stained with lead citrate. ( x 80,000)
SBB f.p. 4461
FIG. 4. L.S. Root hair of axenic pea (cv. Meteor) grown contains many soil particIes (arrows) and extends over the electron beam damage is visible as hates in the cytoplasm with uranyl acetate and lead citrate.
in r-irradiated soil. Mucigel (PI). surface of the root hair (v). Some of the root-hair. Section stained ( x 12,000)
Frc. 5, T.S. Root cap of pea(cv. Tall, Sugar White) grown in nutrient solution and inoculated with Pseudomct~as sp. Note the electron-dense material (cl) near the bacteria (b) on the mucigel’ (m). Unstained. (x 15,000) FIG. 6. T.S. Root-hair zone of pea (cv. Tall, Sugar White) grown in nutrient solution and inoculated with Pse~~donronas sp. Electron-transparent areas in the mucigel fm) close to bacteria(6). Unstained. ( x 12,000)
FIG. 7. T.S. Root-hair zone of mustard grown in sand and inoculated with C. johnsonii. Mucigel (nr) has a banded appearance, with the myxobacteria (my) in electron-dense bands. Section stained with uranyl acetate. (X 30,000) FIG. 8. T.S. Root-hair zone of non-axenic onion grown in nutrient solution. Note distinct external boundary (arrow) of the mucigel (m). Section stained with uranyl acetate. ( x 15,000) FIG. 9. T.S. Root-hair zone of pea (cv. Meteor) grown in non-irradiated soil. Soil particles have been washed off to reveal the outer boundary (arrow) of the mucigel (m). Section stained with uranyl acetate. ( x 36,000) Inset shows soil particles (s) adhering to outer boundary (arrows) of mucigel (m) before washing. Unstained. ( x 18,000) 10. T.S. Root-hair zone of mustard grown in sand and inoculated with Pseudomonas sp. External boundary (arrow) of the mucigel (m) has several distinct layers (arrow). Tissue postfixed in ruthenium red and stained with uranyl acetate. Section stained with lead citrate. ( x 24,000) FIG.
ULTRASTRUCTURE
OF THE MUCIGEL
447
boundary was not restricted to the immediate vicinity of microbial cells and it was frequently present in root regions which were apparently free from micro-organisms (Figs. 9 and IO). Usually, the boundary appeared to be a single electron-dense layer (Fig. 9), which was often obscured by adhering particles when the plants were grown in soil (Fig. 9 inset). The boundary was otherwise clearly visible in both stained and unstained preparations. Occasionally, the mucigel outer boundary consisted of several layers (Fig. 10)orwasilldefined (Fig. 7). Micro-organisms were often found on the outer surface of the boundary layer as well as embedded in the mucigel. DISCUSSION
The electron micrograph of soil-grown peas (Fig. 9 inset) shows that the mucigel can sometimes completely fill the space between the root surface and the soil. Similar observations were made by Jenny and Grossenbacher (1963) on barley roots grown in bentonite or permutite. Jenny and Grossenbacher, however, found only isolated mineral particles embedded in the mucigel of non-axenic barley but Fig. 4 in the present study shows that the mucigel can sometimes be intimately mixed with soil particles 0.1-0.5 pm long. These small soil particles have so far only been observed in the mucigel of axenic peas and mustard. Although the buckled appearance of the cells in Fig. 4 is reminiscent of shrinkage that can occur during fixation, this is considered unlikely since the majority of root hairs were undistorted. As described earlier, this type of tissue damage was confined to the older zones of the root. At present it seems more likely that the buckling occurred naturally during growth. Further investigations are required to establish whether or not the non-axenic mucigel is characterized by a smaller number of embedded soil particles than the equivalent mucigel on axenic plants. It is also difficult at present to explain why the majority of the soil particles in the pea root mucigel in Fig. 4 should be orientated with their long axes nearly parallel to adjacent root surfaces, whereas those in the mustard root mucigel should be randomly orientated. Blevins et al. (1970) found, in thin sections of two soils, that 70 % of the elongated mineral grains of more than 30 pm length at tree root-soil interfaces were orientated at less than 30” to the tangent to the root surface. They suggested that this orientation was due to the pressure exerted by root growth or to fluctuations in turgor pressure. When smaller soil particles are involved, as in Fig. 4, it is possible that the ionic and molecular interactions between the mucigel and surrounding particles discussed by Jenny and Grossenbacher (1963) may become more important. The presence of a mucigel on axenic plants suggests that at least part of the mucigel on non-axenic plants has a plant origin. The remains of dead root-surface cells in the axenic mucigels also shows that it is not only the root-cap cells that may contribute to the mucigel. The possession of the same staining reaction to ruthenium red by inter-lamellae between root cells and some parts of the mucigel suggests that pectic material is common to both regions. Using radioautographic techniques, Northcote and Pickett-Heaps (1966) demonstrated the movement of labelled material from the Golgi bodies, tist into the walls of wheat root-cap cells and finally into the external mucilaginous layer. Chemical analysis of this labelled polysaccharide material showed it to consist mainly of glucose and galactose. Northcote and Pickett-Heaps concluded that the labelled polysaccharide was probably pectic in nature. The greater quantity of mucigel generally observed on non-axenic compared with axenic plants is presumably due to bothdirectcontributions and indirect effects of micro-organisms. The greater number of dead root-surface cells, or cells showing loss of cell wall structure,
448
M. P. GREAVES
AND J. F. DARBYSHIRE
observed in non-axenic compared with axenic roots are examples of indirect microbial effects. The several examples described earlier where the mucigel structure immediately adjacent to the microbes is differentiated from the bulk of the mucigel are examples of direct microbial action on the mucigel. The observations in the present study suggest that a distinct external boundary is characteristic of the mucigel of non-axenic roots, although the boundaries in the electron micrographs of non-axenic roots shown by Jenny and Grossenbacher (1963) and Brams (1969) are faint. Dart and Mercer (1964) found that axenic roots of barrel medic Medicugo tribuloides Desr., as well as roots of the same species inoculated with Rhizobium meliloti Dangeard had a “. . . membranous layer 2: 140 L% wide . . .” on the exterior of the mucigel. Dart and Mercer (1964) also described an inner granular matrix layer of the mucigel on the roots of barrel medic where very few Rhizobium cells were observed. No comparable inner layer was observed on the plant species examined in the present study and it is possible that this structure is specifically associated with root nodule bacteria. At present, it is difficult to assess the ecological and physiological significance of either the external mucigel boundary or the entire mucigel. Both structures could influence the movement of chemical compounds and micro-organisms between the root and the surrounding soil. Dart and Mercer (1964) have suggested that the mucigel forms a protected niche for the rapid multiplication of root nodule bacteria and that it provides a possible site for the accumulation of root exudates. The mucigel may also help pioneer microbial species to prevent subsequent colonization of the root surface by other micro-organisms. Darbyshire and Greaves (1971), however, have shown that the mucigel is not an impenetrable barrier against the soil amoeba, Acanthamoeba palestinensis Reich. The present study demonstrates that the mucigel on non-axenic plants is heterogeneous and is derived from micro-organisms and the plant.
REFERENCES BLEVINSR. L., HOLOWAYCHLJK N. and WILDING L. P. (1970) Micromorphology of soil fabric at tree rootsoil interface. Proc. Soil Sci. Sot. Am. 34,460-465. BRAMSE. (1969) The mucilaginous layer of citrus roots-its delineation in the rhizosphere and removal from roots. PI. Soil 30, 105-108. DARBYSHIRE J. F. and GREAVESM. P. (1970) An improved method for the study of the interrelationships of soil microorganisms and plant roots. Soil Biol. Biochem. 2, 63-71. DARBYSHIREJ. F. and GREAVESM. P. (1971) The invasion of nea roots. Pisum sativum L.. by soil microorganisms, Acanthamoeba palestinerwis (Reich) and Pseudomonas sp. Soil Biol. Biochem: 3, 151-155. DART P. J. and MERCERF. V. (1964) The legume rhizosphere. Arch. Mikrobiol. 47, 344-378. DE FELIPER. and GROSSENBACHER K. A. (1964) Investigaciones sobre la naturaleza de la capa mucilaginosa presente en las racines. Farmacognosia 24, 105-127. DEKOCK P. C. and HALL A. (1962) The effect of nutrient concentration upon the growth and mineral composition of mustard plants. Agrochimica 7, 89-95. DEKOCK P. C., RUTHERFORD M. and CHESHIREM. V. (1971) The fine structure of leaf cells of copper deficient oats. Ann. Bot. 35, 193-199. FLOYDR. A. and OHLR~CGEA. J. (1970) Gel formation on nodal root surfaces of Zea muys-I. Investigation of the gel’s composition. PI. Soil 33, 331-343. FLOYDR. A. and OHLROGGEA. J. (1971) Gel formation on nodal root surfaces of Zea maw Some observations relevant to understanding its action at the root-soil interface. PI. Soil 34, 595406. JENNY H. and GROSSENBACHER K. (1963) Root-soil boundary zones as seen in the electron microscope. Proc. Soil. Sot. Am. 27, 273-277.
JUNIPER B. E. and ROBERTSR. M. (1966) Polysaccharide synthesis and the fine structure of root cells. J. R. microsc. Sot. 85, 63-71. LEISERA. T. (1968) A mucilaginous root sheath in Ericaceae. Am. J. Bot. 55, 391-398. MACKLON A. E. S. and HIGIN~OTHAMN. (1968) Potassium and nitrate uptake and cell transmembrane electropotential in excised pea epicotyls. PI. Physiol. Lancaster 43, 888-892.
ULTRASTRUCTURE NORTHCOTE
OF THE MUCIGEL
449
D. H. and PICKET-T-HEAPS J. D. (1966) A function of the Golgi apparatus in polysaccharide synthesis and transport in the root-cap cells of wheat. Biochem. J. 98, 159-167. REYNOLDS E. S. (1963) The use of lead citrate at high pH as an electron-opaque stain for electron microscopy. .I. Cell. Biol. 17,208-212. SAMTSEVICH S. A. (1968) Gel-like excretions of plant roots and their influence upon soil and rhizosphere microflora. In Methods of Productivity Studies in Root Systems and Rhizosphere Organisms. (M. S. Ghilarov, V. A. Korda, L. N. Novichkova-Ivanova, L. E. Rodin and V. M. Sveshnikova, Eds), pp. 200-204, Nauka, Leningrad. SCOTT F. M. (1965) The anatomy of plant roots, In EcoIogy of Soil-borne Plant Pathogens. Prelude to Biological Control (K. F. Baker and W. C. Snyder, Eds), pp. 145-151, Murray, London. WEBLEY D. M., DUFF R. B., BACON J. S. D. and FARMERV. C. (1965) A study of polysaccharide-producing organisms occurring in the root region of certain pasture grasses. J. Soil Sci. 16, 149-157.
THE ULTRASTRUCTURE OF THE MUCILAGINOUS LAYER ON PLANT ROOTS M. P. GREAVESand J. F. DARBYSHIRE The Macaulay Institute for Soil Research, Craigiebuckler,
Aberdeen, Scotland
(Accepted 16 May 1972) Sunnnary-A mucilaginous external layer or mucigel was observed on the roots of all 16 species of common agricultural crop plants examined. On axenic roots, the mucigel appears in the electron microscope as an unevenly distributed layer of granular and fibrillar material covering the outer surface of the root and most root hairs. The greatest quantity of mucigel was found around the root cap. Apart from the presence of soil particles adhering to the external surface of the mucigel on soil-grown plants, structural differences were seldom observed between the mucigels on plants grown in soil, sand or nutrient solution. Some soilgrown axenic peas (Pisum sativum L.) and mustard (Sinapis alba L.) however, had particulate and amorphous soil material distributed throughout the mucigel. When plant roots were colonized by Pseudomonas sp., Cytophaga johnsonii Stanier or a mixture of several micro-organisms, larger quantities of mucigel developed than on axenic roots of the same plant species. The number of root-surface cells, which were either dead or showed changes in the structure of their walls, was also greater when micro-organisms were present. In the root-elongation zone and in other zones where microbial colonization was sparse, the mucigel retained its usual appearance near micro-organisms. In densely colonized regions of the mucigel, however, the region immediately around the micro-organisms could usually be differentiated from the bulk of the mucigel. When micro-organisms were present the mucigel often had a distinct outer boundary.
INTRODUCTION
presence of an outer mucilaginous layer on the roots, especially the root caps, of Angiosperms has been reported by many investigators in the last decade (Dart and Mercer, 1964; De Felipe and Grossenbachet, 1964; Scott, 1965; Juniper and Roberts, 1966; Leiser, 1968; Samtsevich, 1968; Brams, 1969; Floyd and Ohlrogge, 1970, 1971). Jenny and Grossenbacher (1963) showed that this mucilaginous layer or ‘mucigel’ on barley roots, which were grown in bentonite or permutite, sometimes completely filled the space between the root and the surrounding particulate medium. The possibility that micro-organisms may influence the formation and structure of the mucigel has received little consideration in the past. This paper reports the results of a survey of the ultrastructure of the root mucigel of 16 species of agricultural crop plants. It includes the first report dealing with the mucigel on axenic roots grown in soil. THE
MATERIALS
AND METHODS
Rooting media
Three different rooting media, soil, sand and nutrient solution, were used for the plant species listed in Table 1. These plants were grown for 2-4 weeks in a glasshouse with ambient air temperatures between 15 and 25”C, except when peas, Pisum sativum L., were grown in nutrient solution (see below). Glasshouse experiments were done three times. Soil. Soil for axenic soil studies was collected from the surface 15 cm of the Institute 443 SOIL 414-E
M. P. GREAVES
444 TABLE
AND J. F. DARBYSHIRE
1. PLANT SPECIESGROWN IN GLASSHOUSE,WITH ROOTINGMEDIA USED Plant species
Rooting
N, SO
Maize, Zea rnays L. ‘Kelvedon 3 13’ Timothy grass, Phleum prutense L. S50 Winter wheat, Triticum aestivum L. ‘Capelle-Desprez’ Onion, Al&m cepa L. Cultivar is unknown Pea, Pisum sativum L. ‘Tall, sugar white’ Pea, P. sativum L. ‘Meteor’ Pea, P. sativum L. ‘Foremost’ White mustard, Sinapis alba L. Cultivar is unknown Tomato, Lycopersicum esculentum Mill. ‘Market King’ N = nutrient solution, soil.
SA = sand,
SO = non-irradiated
media
SA, SO, SG SA, SO, SG N N, SA, SO, SG N. SA, SO, SG N SA, SO, SG SA, SO, SG soil,
SG = y-irradiated
garden, air-dried, passed through a 3-mm mesh sieve, sealed in nylon packets in 100 g amounts and sterilized by y-irradiation (2.5 Mrad). The contents of these packets of soil were transferred individually to sterile 2 *5 cm dia. boiling tubes plugged with cotton wool. The soil in each tube was remoistened with sterile glass-distilled water and planted with one axenic seedling (Darbyshire and Greaves, 1971). Soon after the plumules had emerged, two thirds of the seedlings were each inoculated with 0.1 ml of a 24-h nutrient broth (Oxoid Ltd, London) culture of either Pseudomonas sp. or Cytophuga johnsonii Stanier. The remaining axenic seedlings each received 0.1 ml of sterile nutrient broth. Pseudomonas sp. and C. johnsonii were isolated from the roots of timothy grass, Phleum prutense L. S50, (Darbyshire and Greaves, 1970) and cocksfoot, Ductylis glomerutu L. S143 (Webley, Duff, Bacon and Farmer, 1965) respectively. A laminar air-flow apparatus (Microflow Limited, Fleet, Hants., England) was used to facilitate the axenic manipulations. Plants were also grown in unsterilized soil from the Institute garden in the manner described above except that the packaging and y-irradiation were omitted. Roots were also sampled on two separate occasions, from plants (Table 2) growing in the Institute garden and from agricultural crops of barley and wheat at Methlick, Aberdeenshire (Thistlyhill and Tarves Series respectively of the Tarves Soil Association). TABLE 2. PLANT SPECIESGROWN
IN
FIELD OR GARDEN SOIL
Spring barley, Hordeum vulgare L. ‘Golden Promise’ Winter wheat, Triticum aestivum L. ‘Champlein’ Winter wheat, T. aestivum L. ‘Jos Cambier’ Potato, Solunurn tuberosum L. ‘Maris Peer’ Potato, S. tuberosum L. ‘Sharpe’s Express’ Onion, Allium cepa L. ‘Ailsa Craig’ Leek, AIlium porrum L. ‘Musselburgh Pea, Pisum sativum L. ‘Admiral Beatty’ Brussels sprouts, Brussica olerucea L. ‘Cambridge Special’ Savoy cabbage, B. oleracea L. ‘Ormskirk, Late Green’ Turnip, Brussicu rupa L. ‘Milan, Extra Early’ Parsnip, Pustinaca sutivu L. ‘Hollow Crown Improved’ Carrot, Daucus curota L. ‘Ideal’ Lettuce, Lactuca sativa L. ‘Webb’s Wonderful’ Broad Bean, Vicia faba L. ‘Bunyard’s Exhibition’
ULTRASTRUCTURE
OF THE MUCIGEL
445
Sand. Axenic and inoculated seedlings were grown in acid-washed, coarse sand (No. 52 W, G. Garside, Sand Quarries, Leighton Buzzard, Beds., England) in the manner described for r-irradiated soil, except that the soil and glass distilled water were replaced by approximately 100 g amounts of coarse sand and nutrient solution (Darbyshire and Greaves, 1970). The sand was sterilized in the tubes by autoclaving at 121°C for 20 min after the addition of the nutrient solution. Nutrient solution. Peas were grown as described by Darbyshire and Greaves (1970) with axenic roots or roots inoculated with Pseudomonas sp. Onions, Allium cepa L., were grown in the single-strength medium of Macklon and Higinbotham (1968) and maize, Zea mays L., in DeKock and Hall’s (1962) medium, except that iron was supplied as ferric versenate (2 parts/106). Both solutions were non-axenic. Electron microscopy. Between three and five roots per plant were selected from two plants as representative material for each plant species investigated. After the roots had been gently rinsed in glass distilled water, five I-mm lengths were cut from each of the following zones: root apex, elongation zone, root-hair zone and basal zone. The preparation for electron microscopy was as described by DeKock, Rutherford and Cheshire (1971). The root samples were fixed in glutaraldehyde (6 % v/v in phosphate buffer, pH 6 -9) for 1’ 5 h at 20°C. Post-fixation was in either osmium tetroxide solution (1% w/v in sodium veronal-sodium acetate buffer, pH 7 -9) for 1 h at 4°C followed by 24 h at 20°C or in ruthenium red (10 mg/l., Sorensen’s phosphate buffer, pH 7 *O)for 2 h at 4°C. After dehydration in a graded ethanol series, samples were infiltrated with and embedded in a Durcupan A.C.M. mixture. Sections (50-80 nm thick) were cut on an Ultrotome (L.K.B. Instruments Ltd, South Croydon, Surrey, England) using glass knives. Difficulty was encountered in cutting sections of roots grown in soil owing to the presence of mineral particles on the root surface. The best results were obtained with thicker (80-120 nm) sections. The use of a diamond knife did not improve the results. Some root samples were stained with uranyl acetate (2% w/v) at the 75% ethanol stage of dehydration. Sections were stained with uranyl acetate (1% w/v, aqueous solution), lead citrate (Reynolds, 1963) or both and examined in an electron microscope (EM6, A.E.I.) at 75 kV. The orientation of elongated soil particles in the mucigel was measured on enlarged prints (cu. 60 x 40 cm) of the original electron micrographs.
RESULTS
Many of the roots examined, particularly those colonized by micro-organisms, showed signs of tissue damage reminiscent of ineffective fixation in the epidermis and outer cortex of the root-hair and basal root zones. The apical meristems and elongation zones were undamaged. It was concluded, therefore, that the damage had occurred before the roots were fixed. Light microscopy of unfixed roots also showed that many of the outer root cells were damaged. Mucigel was present on the roots of all the plant species studied and its ultrastructure was little affected by the type of rooting medium used. There were, however, some distinct structural differences between the mucigels found on axenic and non-axenic roots. Accordingly, the observations reported below apply to all plant species studied unless otherwise stated. Axenic roots. The mucigel on axenic roots appears as an unevenly distributed layer of granular and fibrillar material covering the outer surface of the root and most root hairs. The greatest quantity of mucigel was found around the root cap (Fig. 1). The remnants of occasional dead root cells (Fig. 2) and structures which resemble plant organelles in the
446
M. P. GREAVES
AND
J. F. DARBYSHIRE
mucigel, as well as the loss of wall structure from root surface cells (Fig. 3), suggest that the mucigel is partly derived from root surface cells. The middle lamellae between the rootsurface cells and some parts of the mucigel (Fig, 1) were strongly stained with ruthenium red. As this stain is frequently used to detect pectin it is probable that pectic material is common to both regions. The root mucigels on axenic plants grown in soil and other media could only be distinguished by the presence of particulate and amorphous soil material on the former. These soil particles were usually found adhering closely to the outer surface of the mucigel which was also present in some micro-pores between the adhering particles. On some axenic pea and mustard plants, soil particles were distributed throughout the mucigel (Fig. 4). As very long exposures were required to clearly demonstrate these particles in the electron micrographs, structural detail within the cytoplasm has been lost through over-exposure. These over-exposed areas have been darkened on the back of the electron micrograph plates, using a very soft pencil, to reduce their contrast. It was estimated that 67 % of the elongated particles visible in the mucigel in Fig. 4 were inclined at less than 30” to the tangent to the nearest point on the root surface. If the root hair in Fig. 4 was considered alone the corresponding proportion was 88 %. Many of these elongated particles appear to be platy clay minerals. The proportion of similarly orientated particles in the mucigel on axenic mustard roots was only 30%. Non-axe& roots. The thickness of the mucigel on roots colonized with micro-organisms ranged between 0.5 and 8 pm compared with a maximum of about O-5 pm on all parts of axenic roots except the root cap where it was up to 2.5 pm thick. The number of root surface cells which were either dead or had lost some wall structure was also greater when microorganisms were present. The structure of the root mucigel appeared to be very similar irrespective of whether the root surface was colonized with specific micro-organisms (e.g. Pseudomonas sp., C. johnsonii) or a mixture of micro-organisms which developed in nonaxenic nutrient solution. Microbial colonization usually resulted in some electron-transparent areas in the mucigel close to the microbial cells. As the microbial population increased, these areas became more widespread until eventually the matrix of the mucigel was almost entirely electron-transparent (Fig. 6). The observed exceptions to this generalization are described below. Electron-dense material developed around Pseudomonas sp. when these bacteria colonized the root-cap mucigel of peas grown in nutrient solution (Fig. 5). This material became fess electron-dense further away from the root apex and, in the oldest part of the root, the bacteria were embedded in an almost entirely electron-transparent mu&gel (Fig. 6). The mixed micro-flora colonizing the root-cap mucigel of maize grown in non-axenic nutrient solution and the mucigel on decaying root-surface cells of wheat, barley and maize grown in non-axenic soil were sometimes surrounded by electron-dense material in the mucigel. When white mustard was grown in sand and inoculated with C. johnsonii, the mucigel on the flatter areas of the root surface consisted of alternate thick bands of electron-dense material and thin electron-transparent bands (Fig. 7). Each electrondense layer was colonized by myxobacteria, which were orientated so that their long axes lay parallel to the root axis. The bands in the mucigel on white mustard roots were less obvious where the colonization by C. johnsonii was sparse or where the organism accumulated in crevices between root surface cells. This banded appearance was never observed in the root mucigel of other plant species inoculated with C. johnsonii or with white mustard grown in soil and inoculated with this myxobacterium. Another difference between axenic and non-axenic roots was that there was often a distinct outer boundary to the mucigel when micro-organisms were present (Fig. 8). This
FIG. 1. T.S. Root cap of axenic mustard grown in sand showing thick mucigel (m). Densely stained material (arrows) is present throughout the mucigel and interlamellae (1) between root-cap cells. Tissue postfixed with ruthenium red and stained with uranyl acetate. Section stained with lead citrate. ( x 24,000) FIG. 2. T.S. Root-hair
zone of axenic pea (cv. Foremost) grown in nutrient solution. Remains of a sloughed-off cell (cv) are lying in an unevenly distributed mucigel (WI).Unstained. ( x 4920)
FIG. 3. T.S. Root-surface cell wall in root-hair zone of axenic mustard grown in sand. Cell wall (cw) consists of a framework of micro-fibrils (f). Mucigel (m). Tissue post-fixed in ruthenium red and stained with uranyl acetate. Section stained with lead citrate. ( x 80,000)
SBB f.p. 4461
FIG. 4. L.S. Root hair of axenic pea (cv. Meteor) grown contains many soil particIes (arrows) and extends over the electron beam damage is visible as hates in the cytoplasm with uranyl acetate and lead citrate.
in r-irradiated soil. Mucigel (PI). surface of the root hair (v). Some of the root-hair. Section stained ( x 12,000)
Frc. 5, T.S. Root cap of pea(cv. Tall, Sugar White) grown in nutrient solution and inoculated with Pseudomct~as sp. Note the electron-dense material (cl) near the bacteria (b) on the mucigel’ (m). Unstained. (x 15,000) FIG. 6. T.S. Root-hair zone of pea (cv. Tall, Sugar White) grown in nutrient solution and inoculated with Pse~~donronas sp. Electron-transparent areas in the mucigel fm) close to bacteria(6). Unstained. ( x 12,000)
FIG. 7. T.S. Root-hair zone of mustard grown in sand and inoculated with C. johnsonii. Mucigel (nr) has a banded appearance, with the myxobacteria (my) in electron-dense bands. Section stained with uranyl acetate. (X 30,000) FIG. 8. T.S. Root-hair zone of non-axenic onion grown in nutrient solution. Note distinct external boundary (arrow) of the mucigel (m). Section stained with uranyl acetate. ( x 15,000) FIG. 9. T.S. Root-hair zone of pea (cv. Meteor) grown in non-irradiated soil. Soil particles have been washed off to reveal the outer boundary (arrow) of the mucigel (m). Section stained with uranyl acetate. ( x 36,000) Inset shows soil particles (s) adhering to outer boundary (arrows) of mucigel (m) before washing. Unstained. ( x 18,000) 10. T.S. Root-hair zone of mustard grown in sand and inoculated with Pseudomonas sp. External boundary (arrow) of the mucigel (m) has several distinct layers (arrow). Tissue postfixed in ruthenium red and stained with uranyl acetate. Section stained with lead citrate. ( x 24,000) FIG.
ULTRASTRUCTURE
OF THE MUCIGEL
447
boundary was not restricted to the immediate vicinity of microbial cells and it was frequently present in root regions which were apparently free from micro-organisms (Figs. 9 and IO). Usually, the boundary appeared to be a single electron-dense layer (Fig. 9), which was often obscured by adhering particles when the plants were grown in soil (Fig. 9 inset). The boundary was otherwise clearly visible in both stained and unstained preparations. Occasionally, the mucigel outer boundary consisted of several layers (Fig. 10)orwasilldefined (Fig. 7). Micro-organisms were often found on the outer surface of the boundary layer as well as embedded in the mucigel. DISCUSSION
The electron micrograph of soil-grown peas (Fig. 9 inset) shows that the mucigel can sometimes completely fill the space between the root surface and the soil. Similar observations were made by Jenny and Grossenbacher (1963) on barley roots grown in bentonite or permutite. Jenny and Grossenbacher, however, found only isolated mineral particles embedded in the mucigel of non-axenic barley but Fig. 4 in the present study shows that the mucigel can sometimes be intimately mixed with soil particles 0.1-0.5 pm long. These small soil particles have so far only been observed in the mucigel of axenic peas and mustard. Although the buckled appearance of the cells in Fig. 4 is reminiscent of shrinkage that can occur during fixation, this is considered unlikely since the majority of root hairs were undistorted. As described earlier, this type of tissue damage was confined to the older zones of the root. At present it seems more likely that the buckling occurred naturally during growth. Further investigations are required to establish whether or not the non-axenic mucigel is characterized by a smaller number of embedded soil particles than the equivalent mucigel on axenic plants. It is also difficult at present to explain why the majority of the soil particles in the pea root mucigel in Fig. 4 should be orientated with their long axes nearly parallel to adjacent root surfaces, whereas those in the mustard root mucigel should be randomly orientated. Blevins et al. (1970) found, in thin sections of two soils, that 70 % of the elongated mineral grains of more than 30 pm length at tree root-soil interfaces were orientated at less than 30” to the tangent to the root surface. They suggested that this orientation was due to the pressure exerted by root growth or to fluctuations in turgor pressure. When smaller soil particles are involved, as in Fig. 4, it is possible that the ionic and molecular interactions between the mucigel and surrounding particles discussed by Jenny and Grossenbacher (1963) may become more important. The presence of a mucigel on axenic plants suggests that at least part of the mucigel on non-axenic plants has a plant origin. The remains of dead root-surface cells in the axenic mucigels also shows that it is not only the root-cap cells that may contribute to the mucigel. The possession of the same staining reaction to ruthenium red by inter-lamellae between root cells and some parts of the mucigel suggests that pectic material is common to both regions. Using radioautographic techniques, Northcote and Pickett-Heaps (1966) demonstrated the movement of labelled material from the Golgi bodies, tist into the walls of wheat root-cap cells and finally into the external mucilaginous layer. Chemical analysis of this labelled polysaccharide material showed it to consist mainly of glucose and galactose. Northcote and Pickett-Heaps concluded that the labelled polysaccharide was probably pectic in nature. The greater quantity of mucigel generally observed on non-axenic compared with axenic plants is presumably due to bothdirectcontributions and indirect effects of micro-organisms. The greater number of dead root-surface cells, or cells showing loss of cell wall structure,
448
M. P. GREAVES
AND J. F. DARBYSHIRE
observed in non-axenic compared with axenic roots are examples of indirect microbial effects. The several examples described earlier where the mucigel structure immediately adjacent to the microbes is differentiated from the bulk of the mucigel are examples of direct microbial action on the mucigel. The observations in the present study suggest that a distinct external boundary is characteristic of the mucigel of non-axenic roots, although the boundaries in the electron micrographs of non-axenic roots shown by Jenny and Grossenbacher (1963) and Brams (1969) are faint. Dart and Mercer (1964) found that axenic roots of barrel medic Medicugo tribuloides Desr., as well as roots of the same species inoculated with Rhizobium meliloti Dangeard had a “. . . membranous layer 2: 140 L% wide . . .” on the exterior of the mucigel. Dart and Mercer (1964) also described an inner granular matrix layer of the mucigel on the roots of barrel medic where very few Rhizobium cells were observed. No comparable inner layer was observed on the plant species examined in the present study and it is possible that this structure is specifically associated with root nodule bacteria. At present, it is difficult to assess the ecological and physiological significance of either the external mucigel boundary or the entire mucigel. Both structures could influence the movement of chemical compounds and micro-organisms between the root and the surrounding soil. Dart and Mercer (1964) have suggested that the mucigel forms a protected niche for the rapid multiplication of root nodule bacteria and that it provides a possible site for the accumulation of root exudates. The mucigel may also help pioneer microbial species to prevent subsequent colonization of the root surface by other micro-organisms. Darbyshire and Greaves (1971), however, have shown that the mucigel is not an impenetrable barrier against the soil amoeba, Acanthamoeba palestinensis Reich. The present study demonstrates that the mucigel on non-axenic plants is heterogeneous and is derived from micro-organisms and the plant.
REFERENCES BLEVINSR. L., HOLOWAYCHLJK N. and WILDING L. P. (1970) Micromorphology of soil fabric at tree rootsoil interface. Proc. Soil Sci. Sot. Am. 34,460-465. BRAMSE. (1969) The mucilaginous layer of citrus roots-its delineation in the rhizosphere and removal from roots. PI. Soil 30, 105-108. DARBYSHIRE J. F. and GREAVESM. P. (1970) An improved method for the study of the interrelationships of soil microorganisms and plant roots. Soil Biol. Biochem. 2, 63-71. DARBYSHIREJ. F. and GREAVESM. P. (1971) The invasion of nea roots. Pisum sativum L.. by soil microorganisms, Acanthamoeba palestinerwis (Reich) and Pseudomonas sp. Soil Biol. Biochem: 3, 151-155. DART P. J. and MERCERF. V. (1964) The legume rhizosphere. Arch. Mikrobiol. 47, 344-378. DE FELIPER. and GROSSENBACHER K. A. (1964) Investigaciones sobre la naturaleza de la capa mucilaginosa presente en las racines. Farmacognosia 24, 105-127. DEKOCK P. C. and HALL A. (1962) The effect of nutrient concentration upon the growth and mineral composition of mustard plants. Agrochimica 7, 89-95. DEKOCK P. C., RUTHERFORD M. and CHESHIREM. V. (1971) The fine structure of leaf cells of copper deficient oats. Ann. Bot. 35, 193-199. FLOYDR. A. and OHLR~CGEA. J. (1970) Gel formation on nodal root surfaces of Zea muys-I. Investigation of the gel’s composition. PI. Soil 33, 331-343. FLOYDR. A. and OHLROGGEA. J. (1971) Gel formation on nodal root surfaces of Zea maw Some observations relevant to understanding its action at the root-soil interface. PI. Soil 34, 595406. JENNY H. and GROSSENBACHER K. (1963) Root-soil boundary zones as seen in the electron microscope. Proc. Soil. Sot. Am. 27, 273-277.
JUNIPER B. E. and ROBERTSR. M. (1966) Polysaccharide synthesis and the fine structure of root cells. J. R. microsc. Sot. 85, 63-71. LEISERA. T. (1968) A mucilaginous root sheath in Ericaceae. Am. J. Bot. 55, 391-398. MACKLON A. E. S. and HIGIN~OTHAMN. (1968) Potassium and nitrate uptake and cell transmembrane electropotential in excised pea epicotyls. PI. Physiol. Lancaster 43, 888-892.
ULTRASTRUCTURE NORTHCOTE
OF THE MUCIGEL
449
D. H. and PICKET-T-HEAPS J. D. (1966) A function of the Golgi apparatus in polysaccharide synthesis and transport in the root-cap cells of wheat. Biochem. J. 98, 159-167. REYNOLDS E. S. (1963) The use of lead citrate at high pH as an electron-opaque stain for electron microscopy. .I. Cell. Biol. 17,208-212. SAMTSEVICH S. A. (1968) Gel-like excretions of plant roots and their influence upon soil and rhizosphere microflora. In Methods of Productivity Studies in Root Systems and Rhizosphere Organisms. (M. S. Ghilarov, V. A. Korda, L. N. Novichkova-Ivanova, L. E. Rodin and V. M. Sveshnikova, Eds), pp. 200-204, Nauka, Leningrad. SCOTT F. M. (1965) The anatomy of plant roots, In EcoIogy of Soil-borne Plant Pathogens. Prelude to Biological Control (K. F. Baker and W. C. Snyder, Eds), pp. 145-151, Murray, London. WEBLEY D. M., DUFF R. B., BACON J. S. D. and FARMERV. C. (1965) A study of polysaccharide-producing organisms occurring in the root region of certain pasture grasses. J. Soil Sci. 16, 149-157.