The Ultrastructure of Plastids in Roots

INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 85

The Ultrastructure of Plastids in Roots JEAN M. WHATLEY Botany School, Oxford University. Oxford, England Introduction . . . . . , . , ......................... ... . ... General Features , . . . . . . , . . , . , . . . . . . . . . . . A . The Root.. . . . . . ............................ .. ..... B. Plastid Development . . . . . . . . . . . . . . . . . . ......................... 111. Nongreen Roots., . . . A. The Forms and Distribution of Plastids in Roots of Seedlings with Particular Reference to P haseolus vulgaris. . . . . . . . . . . . . B. Plastids in Radicles of Embryos.. . . . . . . . . . . . . . . . . ........................ IV. Green Roots.. . . . . . . . . . . . . . . A. A z o h pinnata-the Ultrast Size, and Numbers of Plastids in Different Cell Files . . . . . .................. B. Other Aquatic Species.. . . . . . . . . . . . . . . . . . . . . . . . C. Plastid Dedifferentiation and the Rhizophores of Selaginella

I.

11.

.

V.

VI.

VII. V111.

IX . X.

XI.

The A. B.

................................... .......... ............................

cal Changes (Triticum vulgare. Secale cereale, a Hybrid Triticale, and Lens culinuris) . . . . . . . C. Ultrastructural Changes in Convolvulus arvensis . . D. Ultrastructural Changes in Daucus carota . . . . . . , . . . . . . . . . . . Greening and Plastid Division. . . . . . . . . . . . . . . . . . . . . . . A. The Ultrastructure of Dividing Plastids . . . . . . . . . B. Plastid Numbers and Sites of Plastid Division. . . . ................... C. The Plastid Genome.. . . . , . . . . , . . Plastids in Sieve Elements.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Geotropism and Plastids in Root Caps. . . . . . . . . . . . . . . . . . . . . . . . . A. Amyloplasts as Geoperceptive Organelles . . . . . . . . . . . B. Plastid Distribution in Cells of the C. Plastids in Root Caps of Some Lower Plastid Pigments and Responses to Light. . . A. Protochlorophyllide, Chlorophylls, and ,......... B. Blue Light and Greening.. . . . . . . . . . . . Some Nonphotosynthetic Functions of Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ............................ References . . . . . . . . . .

175 176 176 178 180 180 187 188 188 193 193 196 196

196 200 20 I 202 203 204 205 206 208 208 209 21 I 212 212 213 214 216 217

I. Introduction Most of our information about the ultrastructure, function, and development of plastids has been obtained from work carried out on the leaves of angiosperm I75 Copyright 0 1983 by Academic Press. Inc. All nghts of reprcduclion in any form reserved. ISBN 0 - 1 2 - 3 w n 5 - 2

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seedlings grown either in the dark and subsequently transferred to the light or grown normally with a diurnal cycle of light and darkness (Kirk and TilneyBassett, 1978). The observations made during these investigations have provided a basic model for what is generally accepted as the “normal” plastid. Though the plastids in other plant groups and the plastids in other plant organs (including roots) more or less fit this standard model, their structure and patterns of development nevertheless show subtle differences, the significance of which is far from clear. Though plastids in aerial roots and in the roots of aquatic species often develop into photosynthetically functional chloroplasts indistinguishable from those in leaves, the plastids in roots which penetrate the soil (i.e., most roots) normally lack chlorophyll. The lack of a photosynthetic apparatus in underground roots is not, however, necessarily just the result of growth in darkness, for these roots do not always become green when they are exposed to light. Even when greening does take place, not all cells respond. Fadeel (1962) has estimated that, in the light, proplastids develop into chloroplasts in only 5% of wheat root cells whereas they do so in 80% of leaf cells. When etiolated leaves are exposed to light the etioplasts lose their prolamellar bodies and are rapidly transformed into chloroplasts, sometimes within a matter of hours, but in roots of the same plant the plastids (which lack prolammellar bodies) may take days or even weeks to develop into chloroplasts, if they do so at all. Furthermore the intensity and the wavelengths of light required to promote greening in roots are often different from those required for the greening of leaves. Thus the plastids in nongreen roots clearly behave differently as well as differ in structure from those in nongreen leaves. Neither the structure nor the function of plastids in roots has been rigorously investigated. Ultrastructural studies have been few and then generally limited either to plastids in the apical meristem and the root cap in seedlings or to the changes in plastid structure which take place when the roots (usually the excised roots) become green on exposure to light. Both the small size of most root plastids and their small numbers in most root cells have hampered biochemical investigations using isolated plastids. Thus we have little detailed knowledge of the ways in which root plastids resemble or differ from the “normal” (leaf) plastid and we know even less about how plastids vary in different types of cell within the root or in cells at different distances from the apical meristem. 11. General Features A . THE ROOT A root is usually described as a naked axis of which the only superficial appendages are root hairs; lateral roots and, in some species, root buds both

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originate in tissue deep within the root, most commonly in the pericycle. The apical meristem of a root is subterminal and bidirectional, contributing cells both upward to the root proper and downward to a unique protective structure, the root cap. The main functions of a root are anchorage and the absorption of water and solutes, but roots can also have other physiological functions, e.g., as storage, aerating or supporting organs. The varied morphology of roots reflects these different functions. A young root comprises (1) the epidermis-usually a single layer of cells some of which may be extended to form root hairs; (2) the cortex-concentric files of parenchymatous cells limited toward the interior of the root by the endoderrnis; and (3) the vascular system-a cylinder of parenchymatous cells, within which lie alternating radial strands of phloem and xylem, the system being limited toward the exterior by the pericycle. The epidermis and the cortex may be sloughed off in some older roots. Depending on the physiological function of the root, different types of specialized cell (e.g., sclerenchyma and secretory ducts) may be formed within any of the three main zones. Most dicotyledons and gymnosperms have a root system comprising the primary root and lateral roots which develop acropetally from it. In perennial species these roots can undergo secondary growth. Most monocotyledons have an ephemeral primary root and the functional root system is derived from stem-borne adventitious roots; these do not undergo secondary growth (Esau, 1965). Thus roots may contain many different types of cell and may follow many different patterns of growth. Presumably the plastids show corresponding if less obvious diversity, but, with the exception of some work on sieve elements, the plastids associated with the many specialized types of root cell have never been seriously investigated. It has been postulated that roots evolved from the leafless axes of early land plants, but how this took place is obscure. The nonvascular mosses and liverworts and the most primitive group of vascular plants (the Psilopsida) lack roots and the latter give no evidence in their embryogeny of ever having possessed them. In the Psilopsida the physiological functions of roots are carried out by underground stems and rhizoids (Foster and Gifford, 1974). In other primitive vascular plants the primary root which develops from the radicle is usually short lived; secondary roots commonly arise endogenously near the shoot apex. Among members of the Lycopsida, these roots subsequently branch dichotomously; no lateral roots are formed. In prostrate species of Lycopodiurn the endogenous secondary roots enter the soil more or less directly, but, in upright species, they grow downward through the stem cortex and only emerge near its base. In the Lycopsid genus, Selaginella, secondary roots do not arise endogenously, nor do they grow downward through the stem cortex. Instead cylindrical structures called rhizophores arise superficially in angle meristems at points of branching near the shoot apex. Occasionally these rhizophores give rise to leafy shoots but, usually, they produce no leaves and grow down toward the soil.

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A brief description of the status of rhizophores is included here because the ways in which their plastids develop may provide a clue to the behavior of plastids in roots of less primitive plants. It was formerly believed that the aerial rhizophores of Seluginellu produced no cap and, for this and other reasons, the rhizophores were not considered to be roots. Rather it was believed that true, dichotomously branching roots with root hairs and caps were initiated only at the distal ends of rhizophores when these, after growing aerially for several centimeters, at last came in contact with the soil. Recent anatomical, developmental, and hormonal investigations suggest, however, that rhizophores may indeed be roots able to form both root caps and root hairs and not just the root-bearing organs that their name implies (Grenville and Peterson, 1981; Webster, 1969; Webster and Jagels, 1977; Webster and Steeves, 1963, 1964, 1967; Wochok and Clayton, 1978; Wochok and Sussex, 1974). Though no cap is normally formed in rhizophores of some species of Seluginellu (e.g., S . rnurtensii), a cap does form at a very early stage of rhizophore development in other species (e.g., S. densu, S. kruussiunu, and S . wullucei). In the latter species the apical meristematic cell begins to cut off initials for the cap when the rhizophore is less than 1 mm in length. The first dichotomous branching of the apex which will lead to the formation of future roots can be identified when the rhizophores of S. kruussiunu and S . wullucei are about 2 mm long, though this dichotomy is not visible externally until the rhizophore is some 450 mm in length and is usually just about to enter the soil. It is the series of differential responses by plastids in rhizophores growing (1) above ground, when the contain chlorophyll; (2) below ground, when they lose their chlorophyll; and (3) in moist containers, when they again lose their chlorophyll, which are of particular interest with respect to the behavior of plastids in roots of higher vascular plants, and which will be discussed in some detail below.

B . PLASTIDDEVELOPMENT Ultrastructural investigations, particularly those on leaves, but also those on other plant organs, suggest that in differentiating cells, the development of proplastids into chloroplasts always follows the same basic pathway. Within that pathway five successive stages of development (Fig. 1) can be distinguished (Whatley, 1977). These successive stages can be identified either as a temporal sequence during synchronous development of plastids in tissue becoming green or as a spatial sequence which can be followed along the length of individual cell files from a meristem toward mature green tissue, e.g., from the basal meristem to the tip of a grass leaf (Whatley, 1978; Wellburn, 1982; Wellburn et uf., 1982). This basic pathway of chloroplast development is subject to temporary or permanent modification or blockage. The stage of plastid development at the time of modification or blockage and the nature of the plastid response depend on

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

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J

FIG. I . Stages of plastid development. Eoplast (Stage I t a small more or less spherical plastid containing dense stroma and, usually, a small fragment of thylakoid membrane but no grana. Amyloplast (Stage 2 t r e s e m b l e s an eoplast but contains starch which is not a direct product of photosynthesis. Amoeboid (pleomorphic) plasrid (Stage 3 t w h e n starch is lost from the Stage 2 amyloplast the plastid loses its spherical shape and becomes pleomorphic. Extension of the thylakoid system begins. Pregranal plastid (Stage 4 t t h e plastid usually assumes the discoid shape typical of a mature chloroplast. The thylakoid system becomes much more extensive and appears perforated; bithylakoids (incipient grana) are formed but not true grana. Alternative forms of Stage 4 can be reached in plastids of plants grown either in the light or in darkness. Leaf plastids in plants of Phaseolus vulgaris grown in the light contain chlorophylls a and b in the same ratio as in mature chloroplasts and photosynthesis takes place. Leaf plastids in plants grown in the dark lack chlorophyll and contain crystalline prolamellar bodies in addition to their quite extensive thylakoid system. Mature chloroplast (Stage S t t h e thylakoids lose their performations and true grana are formed. Further development is quantitative and is associated with an increase in extent of the thylakoid system and in the number and depth of stacking of grana.

the type of cell, the plant organ, the species, the physiological state of the plant, and on a number of environmental factors. Such responses are diverse and produce many variations in plastid structure (Whatley, 1977; Klein. 1982). They can result in the synthesis of one normal chloroplast component but not of another, e.g., grana may be formed but not the interconnecting thylakoids or stroma lamellae. Alternatively blockages may result in the accumulation of a

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variety of precursor materials, the form which the accumulation body takes then depending on the nature of the blockage and the period during development at which it occurs. When angiosperms are grown in darkness, for example, the development of leaf proplastids is blocked at a time when some thylakoid extension has already taken place and precursor materials accumulate as a prolamellar body, but in roots, where the blockage apparently takes place slightly earlier, thylakoid extension is minimal and no prolamellar bodies are formed. If the diverse processes of chloroplast development become unbalanced some precursor materials may be produced in excess of requirements and so accumulate for a time before being incorporated into the developing system, e.g., phytoferritin sometimes accumulates in quite massive amounts in eoplasts but later disappears as development proceeds (Whatley, 1978). Though it has in the past been suggested that plastid development is a unidirectional, linear progression leading from proplastids to mature and then to senescent chloroplasts, more recent investigations have shown that this is not necessarily so. Just as proplastids can develop into mature chloroplasts, so can mature chloroplasts dedifferentiate into proplastids, though the latter progression is less common. Depending on the particular aspect of the process which requires emphasis, the basic changes which take place in plastid structure are therefore best represented either as a cycle (Whatley, 1978; Thomson and Whatley, 1980) or as a projection of that cycle representing successive waves of differentiation and dedifferentiation. Diversions from the basic pathway may take place at any point within the cycle. These diversions may be minor or may lead to the development of other distinctive plastid forms such as etioplasts or chromoplasts. These in turn may be further modified to one or another of the five basic stages of development and so join the plastid cycle once again. Senescence, the breakdown of plastid structure often associated with general cellular degeneration, may set in at any point within the plastid cycle; it is not a process restricted only to mature chloroplasts. 111. Nongreen Roots A. THEFORMSAND DISTRIBUTION OF PLASTIDS IN ROOTS OF SEEDLIKGS WITH PARTICULAR REFERENCE TO Phaseolus vulgaris

The basic pathway of plastid development has recently been identified in primary roots of seedlings of Phaseolus vulgaris where it follows two separate spatial sequences (Fig. 2). The first of these sequences extends from the subterminal meristem upward into the root proper and the second downward into the root cap (Whatley, 1983). Thus the bidirectional progress of cellular differentiation within the root is paralleled by the bidirectional progress of plastid develop-

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

-

lo-

-

Root

pro+

-

(2.5mm) -

181

I

I

i

5-

-

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FIG. 2. The sequences of plastid development in a root of Phaseolus vulgaris. 1. Eoplast; 1 ', dedifferentiated plastid = eoplast; 2, amyloplast; 3, amoeboid (plemorphic) plastid; 4, pregranal plastid; +, direction of plastid differentiation;- - 9 , direction of plastid dedifferentiation?; 0 ,level of earliest sieve element plastids (Whatley, 1983).

ment. The basic pathway of plastid development in seedling roots of the grass, Zea mays, follows a similar bidirectional course (Whatley, unpublished), though the root cap of Zea, like that of many other monocotyledons, has separate initials. In roots of Phaseolus vulgaris cv. Canadian Wonder the Stage 1 eoplasts in cells at the tip of the root proper and in adjacent cap cells are spherical or rodshaped (Fig. 3a). The cells which contain spherical eoplasts may more or less coincide with the quiescent center, a feature first identified by Clowes in 1954, but the precise boundaries within the Phaseolus root have not been determined. The eoplasts are sacs containing stroma, phytoferritin (Fig. 4), a few ribosomes, occasional single thylakoid fragments, and aggregations of membranes or tubules (Whatley, 1983). More or less similar aggregations have previously been described from plastids at different stages of development in several organs, including roots, and these have been given a variety of alternative names, e.g., tubular complexes (Newcomb, 1967), thylakoid complexes (Whatley, 1978), and prothylakoid bodies (Wellburn, 1982). Though generally similar to each other in structure these aggregations, which I shall call thylakoid complexes, show some minor variations, viz. those in roots of Phaseofusvulgaris cv Canadi-

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

183

FIG. 4. The distribution of phytofenitin and thylakoid complexes in a root of Phaseolus vulgaris. Small circles, phytofenitin; large circles, thylakoid complexes (Whatley, 1983).

an Wonder are smaller and less well organized and have fewer and less welldeveloped tubules than those observed by Newcomb (1967) in roots of a different cultivar of the same species grown under different environmental conditions. The significance of such subtle variations in structure is not known. What is certain is that thylakoid complexes have some features in common with, but lack the regular paracrystalline organization of, the prolamellar bodies characteristic of leaf etioplasts. Furthermore the thylakoid complexes in root plastids seem to be formed at a slightly earlier stage of development (Fig. 4) than true prolamellar bodies, at a time when no significant thylakoid extension has taken place. ~

FIG. 3. (a) Stage I , possibly dividing, eoplast with a thylakoid complex (t) in the root meristem of Phaseolus vulgaris (Section 1 of the root proper). X24,OOO. (b) Stage 2 amyloplast in the central root cap of Phaseolus vulgaris (Section 3 of the root cap). X20,OOO. (c) Stage 3 pleomorphic plastid at the periphery of the root cap of Phaseolus vulgaris (Section 4 of the root cap): this plastid contains a membrane-bound body (m)as well as starch. X21 ,OOO. (d) All seven of rhe plastid profiles in this micrograph are part of the same Stage 3 pleomorphic plastid in the outer cortex of a root of Phaseolus vulgaris (Section 12 of the root proper). X 10,000. (e) Stage 4 discoid plastid from the outer cortex of a root of Phaseolus vulgaris (Root hair zone): the extent of the thylakoid system is greater in plastids in this part of the root tban in any other zone surveyed. X30,OOO. (0 Stage I ’ eoplast from the lateral root zone: the plastid contains phytoferritin but no thylakoid fragments. X27,OOO.

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..'. .:. .'.

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,

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.

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FIG. 5 . The distribution of starch and membrane-bound bodies in a root of Phaseolus vulgaris. Small circles, starch; large circles, membrane-bound bodies (Whatley, 1983).

The onset of the second stage of plastid development is marked by the accumulation of starch (Figs. 3b and 5), a process which, in the root cap of Phaseolus, reaches its peak in the central cells where the amyloplasts are known to act as statoliths (Fig. 6). Toward the periphery of the cap, in cells which will soon be sloughed off, the plastids begin to lose their starch; a similar loss has also been reported for other species, e.g., Medicago sariva (Maitra and De, 1972)and tomato (Street er al., 1967). In Phaseolus the plastids in the peripheral cells become pleomorphic in shape, i.e., they are entering Stage 3, the maximum stage of development achieved within the cap. Plastids at the periphery of the cap have an electron-dense stroma, some starch, a very restricted system of single thylakoids, and membrane-bound bodies (Figs. 3c and 5 ) . The sometimes crystalline but more commonly granular contents of membrane-bound bodies in plastids of roots and other organs are probably proteinaceous, and may include ribulose 1,5-bisphosphatecarboxylase (Sprey and Lambert, 1977), though other possible components have been proposed, viz. lipids, phenolic compounds, peroxidase, and polyphenoloxidase (reviewed in Hurkman and Kennedy, 1977; Thomson and Whatley, 1980). The pattern of starch accumulation and degradation which forms part of the spatial sequence of plastid development within the root cap (Fig. 6) is displayed as a temporal sequence (Fig. 7) by plastids in the quiescent center of Zea rnays

185

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

Starc area Per lastid profi (urn21

0.8

0.6 0.4 0.2

4 3 2 1

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5 6

Section no. Root cap

8

11

Section number Root proper

---

,---- \---------14 Root hair Lateral m

2

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FIG.6. The distribution of starch in a root of Phaseolus vulgaris. The number of starch grains was counted in successive transverse sections of a root. Starch grains per plastid profile in cells of the inner cortex(& stele (0); root cap The area of starch per plastid profile outer cortex (0); (pm2) in the outer cortex (0); inner cortex (A); root cap (0).

(a).

while a new cap begins to form after the original cap has been removed (Barlow and Grundwag, 1974). Within the root proper of Phaseolus vulgaris, the successive stages of plastid development are found at greater distances from the meristem than they are in the root cap, and, at Stage 2, less starch is accumulated (Whatley, 1983). It is with the beginning of starch accumulation that variations between plastids in different cell files can first be distinguished. Starch appears, for example, nearest to the meristem and in greatest quantity in cells of the outer cortex, and farthest from

,q No. of starch grains

80

40 20 Houn after decapping

FIG.7. Starch accumulation in plastids of the quiescent center following decapping of a root of Zea mays. Compare with root cap in Fig. 6. (From Barlow and Grundwag, 1974.)

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JEAN M. WHATLEY

the meristem and in smallest quantity in cells within the stele (Figs. 5 and 6). Street er al. (1967) found somewhat similar patterns of differential distribution of starch within cultured roots of tomato. The pleomorphic Stage 3 plastids in cortical cells of the root proper are considerably larger and much more highly pleomorphic than the Stage 3 plastids at the periphery of the root cap or in young leaves (Fig. 3c and d). Many Stage 3 plastids in the cortical and epidermal cells contain membrane-bound bodies and the distribution of these at different levels within the root (Fig. 5 ) further illustrate the differential development of plastids in different cell files. However such differential development may in part reflect the fact that cells of the outer cortex are developmentally older than those of the inner cortex (Esau, 1965). In his detailed investigation of root tips of Phaseolus vulgaris, Newcomb (1967) found that the proteinaceous membrane-bound bodies were often connected to subunits of thylakoid complexes and that both of these subsidiary structures and their interconnections sometimes had similar contents. He therefore suggested that the two structures might be developmentally as well as physically linked. It has, inter alia, been proposed that the contents of membrane-bound bodies may later be used during the extension of the thylakoid system, and that both membrane-bound bodies and thylakoid complexes may represent storage deposits which form following temporary (perhaps sequential) blockages or slowing down of the normal pathway of plastid development (reviewed in Thomson and Whatley, 1980). Casodoro and Rascio (1977) followed the development of membrane-bound bodies in roots and shoots of Atropa belladonna. They suggested that vesicles at the periphery of plastids and thylakoid complexes at the center (both of which had electron-dense contents) were the precursors of electron-dense deposits which initially lacked a surrounding membrane. Casodoro and Rascio also observed that in the shoots, but not in the roots, the membranes which later surrounded these bodies were in direct contact with grana. Farther from the tip of the Phuseolus root the plastids lose their pleomorphic form and become discoid (Stage 4 of development-Figs. 2 and 3e). At the same time there seems to be some extension of the thylakoid system, but this is difficult to measure reliably, particularly in the amoeboid plastids. However, the thylakoid system in plastids of Phaseolus roots never approaches the extent or the characteristicessentially spiral conformation of that in Stage 4 chloroplasts or etioplasts in the leaves (Whatley et al., 1982). In addition the single thylakoids in root plastids of Phuseolus (but not in Zea) appear to be less highly perforated than those in leaf plastids and have fewer bithylakoids. In the primary roots of Phuseolus, the most extensive, though still restricted, thylakoid system so far observed is in plastids of cortical cells at the level of root hair maturation. Farther from the tip at the level of the lateral roots, plastids in the cortical cells are spherical sacs containing electron transparent stroma and phytoferritin. These plastids lack even the smallest fragments of thylakoid and

187

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

c

vl

-

Cap

Meristem

\

Tip of root proper zone

zone

Oirections of cellular differentiati;

FIG. 8. vulgaris.

Stages in plastid development in the cap and in the outer cortex of a root of Phaseolus

they contain no starch, membrane-bound bodies, or thylakoid complexes (Fig. 30. If the spatial sequence of plastid development in Phaseolus roots is a true reflection of an earlier temporal sequence, then, after reaching their maximum (albeit limited) state of differentiation (as represented by the Stage 4 plastids in the zone of mature root hairs), the plastids must have undergone dedifferentiation, losing whatever thylakoids they formerly possessed and reverting to a form of eoplast (Fig. 8), as represented by the plastids in the lateral root zone.

B. PLASTIDS IN RADICLESOF EMBRYOS The roots of Phaseolus described above were of dark-grown seedlings 5-7 days old. At this time, the Stage 4 plastids in the root hair zone of the primary roots are very different in ultrastructure from the Stage 4 etioplasts in primary leaves. Several reports in the literature suggest that there are no such major differences between the dedifferentiated plastids in different organs within embryos at the start of germination. Plastids in radicles of developing and mature embryos have been described for several species of angiosperm including Phaseolus lunatus (Klein and Ben-Shaul, 1966; Klein and Pollock, 1968), Pisum sazivum (Bong Yul Yoo, 1970), Secale cereale (Hallam, 1972; Hallam et al., 1972; Sargent and Osborne, 1980), and Zea mays (Deltour and Bronchart, 1971). In ripe seeds, plastids in the radicle contain stroma and osmiophillic globules but few ribosomes and almost no thylakoids or vesicles; phytoferritin is often present and starch gains occasionally so. Plastids in the radicle therefore closely resemble those in other parts of the embryo, including shoot apices and cotyledons. Following germination the numbers of plastid ribosomes in the developing leaves

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increase greatly whether the plants are growing in the light or in the dark, but in the roots the number of plastid ribosomes remains low unless the roots are exposed to light. In the developing embryo of Secale cereale, four zones can be distinguished within the radicle+ap, miristem, procortex, and prostele (Hallam, 1972). At this time the cap of the young radicle is itself protected by an ephermeral coleorhiza. Within I hour of the start of germination starch becomes common in plastids of the coleorhiza (Stage 2) but not in those of the root (Sargent and Osborne, 1980). After 6 hours the coleorhiza plastids are swollen with starch and these amyloplasts also contain polyribosomes. At this time starch is still uncommon in radicle plastids and only monoribosomes are present. Only later, as the root tip breaks through the degenerating coleorhiza, does starch accumulate in the cap of the young root, and at this time the cap takes over its usual role of protecting the root proper.

IV. Green Roots

In roots which penetrate the soil, the normal pathway of chloroplast development is blocked; chlorophyll is not synthesized and thylakoid extension is limited. By contrast, aerial roots and roots of aquatic species often appear green and some plastids at least develop into chloroplasts in much the same way as they do in leaves. A. Azolla pinWtU-THE

ULTRASTRUCTURE, SIZE, AND

PLASTIDSIN DIFFERENT CELLFILES

NUMBERS OF

The only detailed study which has been made of plastid development in complete roots which are normally green is one based on the water fern, Azolla pinnara (Whatley and Gunning, 1981). This investigation made use of a magnificent series of electron micrograph montages of median longitudinal sections of roots of different ages which had previously been assembled and studied by Gunning and his associates (Hardham and Gunning, 1977; Gunning, 1978; Gunning er al., 1978a-c). FIG.9. (a) Small, immature chloroplasts with true grana but with a limited thylakoid system from a cell close to the apical cell in the zone of formative divisions in Azolla pinnara (Root 3; see text). x 12,500. (b) Large, mature chloroplasts with a well-developed system of thylakoids and grana from a cell of the outer cortex in the upper part of Root 3 of Arollu pinnara. X 12,500. (c) Sieve element plastid in a root of Phaseolus vulgaris. X30,OOO. (d) The amyloplasts are concentrated toward the distal wall of this central cap cell of Zea mays (courtesy of Dr.C. R. Hawes). (e) Part of the large single plastid in a root cell of Isoeres lacusrris showing starch grains and many osmiophilic deposits. x5OOO.

190

JEAN M. WHATLEY

The major advantage in using this species of Azollu is that the single apical cell divides in a precise manner to establish a small root within which the lineages of individual cells can easily be determined. The apical cell produces a helical sequence of daughter cells which, in turn, undergo a series of longitudinal formative divisions giving rise to easily distinguished concentric files of cells. The first of the formative divisions gives rise to an outer cell and an inner cell. Following another longitudinal division, the outer cell gives rise to the two outermost cell files of the root, the dermatogen, toward the exterior, and the outer cortex, toward the interior. Longitudinal division of the inner cell produces one cell which is the precursor of both the inner cortex and the endodermis and another cell from which the initials of the pericycle and the inner stele are derived (Gunning et al., 1978~).Subsequent transverse proliferative divisions provide additional cells within each file segment. Gunning uses the term merophyte, for each daughter cell or its derivatives (Gunning et al., 1978a). A complete gyre of the root is made up of three merophytes of which two are visible in longitudinal section. The older the merophyte, the greater its distance from the apical cell and the higher the number it is given in the accompanying figures. In all the roots investigated, the single apical cell contained small pleomorphic plastids which already had some thylakoids and a few true grana, so the state of plastid development in the apical cell was already more “advanced” than in any cells within nongreen roots. In the youngest roots examined and in the stele and the zone of formative divisions in older roots (Fig. 9a), the plastids were more or less discoid in shape but the thylakoid system was scarcely more extensive than in the apical cells. The numbers of plastid profiles present in each cell section suggested that in these cells plastid division had roughly kept pace with cell

G

6

8 10 Gyres

12

FIG.10. The numbers of plastid profiles in cells belonging to different cell files within an older root (root 3) of Arollu pinnaru. 0, Dermatogen; B. outer cortex; 0 ,inner cortex; 0. endodermis; pericycle. (From Whatley and Gunning, 1981.)

A,

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

Fic. 1 I .

..

6

4

8 10 Gyres

191

12

Thylakoid development in plastids of different cell files within root 3 of Azollupinnura. outer cortex; 0 ,inner cortex; 0, endodermis; A,pencycle. (From Whatley and Gunning, 1981.)

0, Dermatogen;

division. Farther from the apex, however, in the outer cell files of the proliferative zone in older roots, the numbers of plastid profiles per cell section, the numbers of bands of thylakoids (Figs. 9b, 10, and 11), and the length of the plastids all increased progressively as the age of the merophyte increased. The apical cell in the Azolla root divides about 55 times (Gunning et al., 1978a). The number of plasmodesmata laid down in the walls between the apical cell and its successive daughter cells declines as the root ages; this suggests that the symplast of the apical cell becomes progressively more isolated from the rest of the root. At the same time, the first sites of xylem thickening, root hair formation, starch deposition, and, within each cell file, particular transverse

t

,

;

,

,

,

,

,

,

,

,

,

,

1 2 3 4 5 6 7 8 9101112 Gyres

FIG. 12. The numbers of plastid profiles in cells of the outer cortex in roots of different ages show acropetal drift. 0 , Root 2 (oldest); Root 4: A, Root 6; Root 8 (youngest). (From Whatley and Gunning, 1981.)

A.

+,

192

JEAN M. WHATLEY

divisions as well as the attainment by plastids of a particular size and state of development all take place progressively closer to the apical cell (Figs. 12 and 13), i.e., these features all show acropetal drift (Gunning, 1978; Whatley and Gunning, 1981). Thus in the green Azollu roots, chloroplast development can be seen as both a spatial sequence extending from the apical cell upward through the root and as a temporal sequence which is reflected by plastids at particular sites within roots of different ages. Though cells of the dermatogen and outer cortex have a common mother cell, their plastids show independent trends in development; cells of the inner cortex and the endodermis also have a common parentage but their plastids similarly follow distinctive pathways. However, cells of the outer and inner cortex, in spite of their different parentage, have plastids which begin to follow a similar course of development soon after these cell files have become established (Figs. 10 and 11). Other cell features, too, show patterns of development characteristic of particular cell files, e.g., the areas occupied by groundplasm and by vacuoles, which may reflect the extent of cytoplasmic protein synthesis, and the area occupied by chromocenters, which may reflect nuclear activity (Barlow er ul., 1982). In the small roots of Azollu pinnara, the differential development within different cell files of plastids and other cell features is easy to observe and the precision with which development appears to be controlled is impressive. We still do not know the basis of this control or how control is exerted but the state of plastid development in a particular cell is clearly related to (1) the age of the root, (2) the file in which the cell lies, and (3) the distance of the cell from the root apex.

8

7

6 5 4 3 2 1 Root no. Increasing age' FIG. 13. Acropetal drift. This is indicated by the several plastid criteria shown; at the same time, the number of plastid profiles in the apical cell declines. 0 ,First site of starch deposition; first achievement by plastids of a mean of 3.0 thylakoid bands; A,first achievement by plastids of a mean length of 1.5 pm; I, the number of plastid profiles per apical cell section. (From Whatley and Gunning, 1981.)

+,

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

193

B . OTHERAQUATICSPECIES The root tips of several aquatic angiosperms were examined by Kawamatu (1967), Mollenhauer (1967), and Wroblewski (1973). In Lemna, Nvmphoides, Hydrocharis, Trapa, Najas, and Hydrilla the roots appeared slightly green; in Eichornia they were colorless or purple (although the root caps are green). Plastids in the roots of Lemna and Trapa showed the most extensive development of a photosynthetic apparatus; plastids in the roots of other “green” species contained few grana. Kawamatu reported the presence of prolamellar bodies in plastids of some species, but it is not clear whether these were true prolamellar bodies or merely thylakoid complexes of some sort. In Najas roots development of the thylakoid system was preceded by the lining up of vesicles; grana were formed later but interconnecting thylakoids were poorly developed. In Lemna the plastids in different parts of the root were at different stages of development. At the same distance from the apex there were proplastids in the central pith, welldeveloped chloroplasts in the outer pith, and immature to mature chloroplasts in the cortex. The nongreen root proper of Eichornia had plastids which contained vesicles and starch but few thylakoids. These plastids resembled those found near the tips of most nongreen roots but they were very different in structure from the large well-developed chloroplasts present in adjacent cells in that portion of the Eichornia root cap which forms a sheath round the tip of the root proper (Mollenhauer, 1967). Mollenhauer also observed large, mature chloroplasts with many thylakoids and grana in the root caps of the duckweed, Spirodella, and of the epiphytic orchid, Cattleya. In roots of most species, plastid development appears to be bidirectional (Fig. 2), but, in the root of the aquatic fern, Azolla pinnata, there is no downward gradient extending into the cap (Whatley and Gunning, 1981). Instead there is, as in the ensheathing part of the Eichornia cap, a discontinuity in plastid structure. In Azolla the small, amoeboid chloroplasts with a restricted thylakoid system present in the youngest merophytes of the root proper, contrast with the large, fully mature chloroplasts with an extensive thylakoid system present in all cells of the adjacent two-layered root cap. This disjunction may reflect the way in which the root cap in Azolla is initiated. The first division of the apical cell is periclinal. It is from the two products of the daughter cell thus formed that all future cap cells are derived (Gunning er a l . , 1978~).Subsequently the apical cell contributes cells only to the root proper. Thus the root cap is, in effect, the oldest merophyte, and it is perhaps not unexpected that its plastids have reached an advanced state of development similar to that of the plastids in the oldest merophytes of the root proper. C. PLASTIDDEDIFFERENTIATION AND THE RHIZOPHORES OF Selaginella martensii The fact that in Eichornia part of the root cap is green but the adjacent cells in the root proper are nongreen emphasizes the importance of factors other than

194

JEAN M. WHATLEY

light in the control of greening. The rhizophores of the lower vascular plant, Selaginella martensii, may contribute to our understanding of this subject. Unlike the aerial rhizophores of some other species of Selaginella, those of S . martensii normally lack a cap and root hairs. These rhizophores are conspicuously green in the zone of cell maturation, i.e., over much of their length. A second conspicuous green zone extends from 0.2 to 1.4 mm behind the apical initial (Webster and Jagels, 1977). Between these two green zones, in the region of cell elongation, the rhizophore is white or scarcely green. The most obvious components of plastids in the epidermal and subepidermal files of the green tips are osmiophilic deposits, some of which are spherical and some irregular in shape. The plastids also contain a few thylakoids and structures with regularly arranged interconnected tubules. These thylakoid complexes and the thylakoids themselves are both strongly osmiophilic. Indeed the staining properties of the plastids in the electron micrographs published by Webster and Jagels appear similar to the temporary “reversed” staining images shown by thylakoids in immature leaf chloroplasts of some angiosperm species during the early stages of granal formation (e.g., Platt-Aloia and Thomson, 1977). In the cortical cells of the green rhizophore tips the plastids are large and apparently lobed. There may well be only one plastid in each cell. Plastids in cells of the outer cortex contain grana and have a more extensive thylakoid system than plastids in the epidermal and subepidermal cells, but these plastids, too, appear to have a “reversed” staining image. Some of these cortical cell plastids may contain true prolamellar bodies. In cells of the inner cortex, the chloroplasts contain many starch grains as well as a thylakoid system which is more extensive than but lacks the “reversed” staining image of that in plastids of the outer cortex. Webster and Jagels do not describe the ultrastructure of plastids in other parts of the aerial rhizophores. This is a pity as information about plastids in the adjacent white and upper green zones might provide an interesting insight into their behavior during later stages of cellular differentiation. The apparent scarcity of chlorophyll in the white zone may merely be a dilution effect resulting from the dispersal of an unchanged chloroplast complement within an elongating cell or, alternatively, it may reflect a true loss of chlorophyll during a phase of plastid dedifferentiation. Conversely the green of the upper part of the rhizophore may be the result of either (1) redifferentiationof nongreen plastids, or (2) chloroplast growth (perhaps accompanied by division) increasing the total volume of the chloroplast(s) already in each cell. When the aerial rhizophores of Selaginella martensii are kept in the light but placed in moist containers, the tips begin to lose their green color. At the same time root caps and root hairs begin to develop. In these apparently colorless tips, the plastids of epidermal and subepidermal cells cannot be distinguished from those in the same files of the green tips. However plastids in the cortical cells

195

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

become smaller and undergo major structural modification. Grana become swollen and the thylakoids which formerly linked them become inconspicuousor disappear. Later, the now isolated grana are reduced to only a few swollen compartments, though they retain their osmiophilic character. These dedifferentiated plastids now closely resemble those in the epidermal and subepidermal files. Plastids in the inner cortex contain less starch than those in the same cells in green rhizophores. Webster and Jagels state that the modifications in structure of the cortical cell plastids during their loss of chlorophyll are similar to those reported by Thomson (1966) for plastids in ripening oranges during the early stages of their dedifferentiation and transformation into chromoplasts, except that osmiophilic (carotenoid) globules do not accumulate in the rhizophore plastids. Whether the two pathways of dedifferentiation are identical or not, it is clear that in both the rhizophore tips and in the ripening oranges loss of the thylakoid system and of chlorophyll is promoted by factors other than the absence of light. Rhizophores of Sefagineflamarrensii which enter the soil also lose their green color, but they do so much more rapidly than those placed in moist containers. Webster and Jagels make an interesting comparison between their own observations and those of Cormack (1937) on roots of the angiosperm pondweed, Elodea canadensis. In both papers a correlation is made between the absence of chlorophyll and the presence of root hairs, conditions which were promoted in Elodea roots (grown in the light) by the introduction of ethylene (Table I), a hormone produced not only by plants but also by bacteria in the rhizosphere. In rhizophores of Selaginella marrensii which enter the soil, just as in those grown in moist containers, the plastids in the cortical cells become smaller and are modified in structure, but the patterns of modification are quite different under the two environmental conditions. In plastids of rhizophores which penetrate the soil the grana do not swell; structures which seem to be true prolamellar bodies appear within and connected to the thylakoid network. Subsequently some starch is retained but the grana disappear and the prolamellar bodies inTABLE I THEPRODUCTION OF CHLOROPHYLL AND HAIRSI N ROOTSGROWN UNDER DIFFERENT CONDITIONS Species

Growth conditions

S. marrensii Light + air (rhizophores) Light + moist containers Dark + soil Light + water E . canadensis Light + water + (roots) ethylene Dark + soil or water

Chlorophyll Root hairs

+ -

-

+ -

-

-

+ + + +

Source Webster and Jagels (1977) Cormack (1937)

196

JEAN M.WHATLEY

crease in size. A reduced system of single thylakoids radiates from the prolamellar bodies, as it does in leaf etioplasts. This apparently dark-induced pattern of plastid dedifferentiation resembles that described by Cran and Possingham (1973) in ripening fruits of avocado, at a time when light penetration through the blackening skin is severely limited. Both the absence of light and other factors (possibly hormonal) can promote dedifferentiation of chloroplasts. However, each factor may promote its own particular form of dedifferentiation.

V. The Greening of Roots A. GENERAL OBSERVATIONS

In an investigation using light microscopy, Powell (1925) examined the distribution of chlorophyll in intact roots of seedlings of 16 angiosperm species grown for 14 days under continuous illumination. She found chlorophyll in roots of 13 out of the 15 species which grew successfully (Table 11). Longitudinal sections showed that in most species chlorophyll developed throughout the length of the root to within 10-20 mm of the tip, but in Triticum vulgare and Hordeurn vulgare it was limited to the uppermost 15-20 mm. Within the roots chlorophyll was not always present in all cell files and the particular files which contained or lacked chlorophyll were characteristic for the species. Indeed in Ranunculus fiearia the pattern of chlorophyll distribution differed in the two types of root investigated. In two species, Triticum vulgare and Ranunculus fiearia (fibrous roots), the chlorophyll was restricted to a single cortical cell file immediately outside the endodermis. In a similar investigation carried out many years later, Fadeel (1962) found that in roots of Triticum aestivum (T.vulgare) cv. Eroica and in Hordeum vulgare after 7 days exposure to light, chloroplasts were restricted to the two innermost files of the cortex, but that within those cells the chloroplasts were numerous. In roots of Linum usitatissimurn, however, chloroplasts were present in all cortical cells, but each cell contained only a few.

B. ULTRASTRUCTURE AND PHYSIOLOGICAL CHANGES (Triticum vulgare, Secale cereale, A HYBRIDTriticale, AND Lens culinaris) Salema ( 197 1) has investigated chloroplast development during greening of intact roots of the grasses, Triticum vulgare (including cv. ardito), Secale cereale (including cv G23 40) and a hybrid Triticale derived from the two cultivars mentioned. In Triticale the first sign of potential greening in proplastids in root meristematic cells was an increase in the number of vesicles derived from the

TABLE I1

THE DISTRIBUTTON OF CHLOROPHYLL w LIGHT-GROWN INTACTROOTSOF SOMEANGIOSPERM SEEDLINGS~

Species

Single Stele extraendodexmal Inner Middle Outer Pith Rays parenchyma Pericycle Endodermis cell file cortex cortex cortex

+

Acer pseudoplatanus

+

Aesculus hippocastanum R m x sp. Vicia foba Pisum sativum Vicia sativurn Zea mays Bellis perennis Helianthus annuus Hordeurn vulgare Ranunculus ficaria Scilla nutans Triticum vulgare Fagopyrum esculentum Ranunculusficaria Ricinus communis Alliurn cepa

+ + + +

Only in two rows of cells above protoxylem groups

+ +

+

+

+ + + + +

-t

acornpiled from Powell (1925).

Comments

+

+ + + +

Seven varieties tested: all gave same results Outer stele only

+ + Fibrous roots Thick contractile mots

Tuberous roots Experiment failed-no

growth

198

JEAN M. WHATLEY

inner plastid envelope. Salema reported that these vesicles assembled into “prolamellar bodies” after 4-5 days of exposure to light, but the low magnification of the published micrographs makes it difficult to determine whether or not these are prolamellar bodies in the now accepted sense. However micrographs in a more recent publication by Oliveira (1982) on roots of Secale suggest that they resemble the thylakoid complexes formed in the light by plastids in developing leaves rather than true prolamellar bodies. Nevertheless, in Triricale root plastids, the extending system of single, perforated thylakoids radiates from the periphery of these complexes as it does from prolamellar bodies in leaf etioplasts. During these early stages of plastid development in the light there is an increase in number of both the stromal and the membrane-bound plastid ribosomes (Oliveira, 1975). After a short time, grana are formed and by the twelfth day of illumination, the root chloroplasts are said to be indistinguishable from leaf chloroplasts. Plastid development during greening of roots of Secale cereale followed a similar course to that in the hybrid Triticale (Salema, 1971). However, in Triticum vulgare, the other parent, no plastids were observed which contained “prolamellar bodies.” Oliveira (1982) calls particular attention to the possibility of genetic control of prolamellar body formation which is suggested by these observations. Salema and Oliveira both consider that extension of the thylakoid

Days

FIG. 14. Chloroplast development during greening of mots of Secale cereule. (A) Hill reaction (02pmol mg-1 chl h-1); (B) chlorophyll content (Fg g - ’ dry wt.); (C) protochlorophyll content (pg g-1 dry wt.); (D) stromal ribosomes; (E) membrane-bound ribosomes. (From Oliveira, 1982.)

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

24

I

48

1

72

199

I

I

Hours

FIG.15. Chloroplast development during greening of roots of Lens culinaris. (A) Hill reaction

I chl h- 1); (B) chlorophyll content (pg g - I dry wt.); (C) RUBC-ase (COz fixed pmo1/25 roots h-1). (From Lance-Nougakde and Pilet, 1965; Nato and Deleens, 1975a.)

(02pmol mg-

system may follow either one of two distinct morphogenetic pathways, the first, as in Tritium, depending entirely on vesicles derived from the inner plastid envelope where synthesis of new membrane takes place, and the other, as in Secale and Triticale, at the expense of the prolamellar bodies. A more detailed investigation of greening in roots of Secale cereale, grown at 25°C and exposed to light of 20,000 lux, has been carried out by Oliveira (1982). This provides data on features such as chlorophyll synthesis and the Hill reaction as well as ultrastructural information (Fig. 14). Somewhat similar information is available for the greening roots of the legume, Lens culinaris (Fig. 15), also grown at 25°C but exposed to light at only 4500 lux (Caporali, 1959; LanceNougarkde and Pilet, 1965; Nato and Deleens, 1975a,b). An interesting comparison between these two species is thus possible. Chloroplast development and chlorophyll synthesis took place much more rapidly, and the Hill reaction began earlier and worked faster in Lens than in Secale (Figs. 14 and 15). However, in both species the relative timings of the biochemical activities measured and particular states of plastid development appear to be quite similar. The more rapid synthesis of chlorophyll in Lens may be the result of the lower light intensities to which the roots were exposed. This is perhaps borne out by the equally rapid synthesis of chlorophyll at low light intensity in the roots of another legume, Lupinus albus (Mesquita, 1971), and by

200

JEAN M. WHATLEY

an investigation on excised roots of Convolvulus arvensis which showed that the chlorophyll content during greening was inversely proportional to light intensities within the range 3000 to 30,000lux (Heltne and Bonnett, 1970).

C. ULTRASTRUCTURAL CHANGESIN Convolvulus arvensis A detailed description of the ultrastructure of the quiescent center and the root cap of Convolvulus arvensis has been given by Phillips and Torrey (1974a,b). The structure of the plastids is essentially similar to that already described for Phaseolus vulgaris and, in the root cap, the characteristic distal positioning of amyloplasts in the elongated cells of the central columella is clearly illustrated. In excised roots of Convolvulus, cultured at 3000 lux, the rate of chloroplast development in the root proper differs in different types of cell (Fig. 16). Heltne and Bonnett found that though cells were fully differentiated within 10 mm of the root tip, it was only at a distance of about 40 mm that chloroplasts with grana first appeared. Chloroplasts were then found only in the cortex, the rate of development of the chloroplasts being the same within all cortical cell files. The mature chloroplasts which developed in the light were similar in structure to leaf chloroplasts. Plastids in dark-grown roots had a dense stroma and contained starch and membrane-bound bodies but no phytoferritin, the accumulation of which, in this species, is apparently light induced. Neither prolamellar bodies nor thylakoid complexes were observed in plastids of either light- or dark-grown roots. In Convolvulus, as in many roots exposed to light, greening takes place first in

e + d a t

E

2

c

0) >

''cO b

p a + lA

60 80 100 150 200 m m from root tip FIG. 16. Chlorophyll content and stages of plastid development in different types of cell during greening of cultured mots of Convolvulus arvensis. (a) Proplastids with few thylakoids; (b) proplastids with many irregular thylakoid configurations; (c) phytoferritin present; (d) early stages of granal development (Stage 4); (e) mature chloroplasts with true grana (Stage 5). 0 ,Inner and outer cortex; xylem parenchyma; M, pericycle; 0, chlorophyll content (pglg fresh weight) of roots grown in the light at 3000 lux. (From Heltne and Bonnett, 1970.)

0

A,

20

LO

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

20 1

fully differentiated cells at some distance from the root tip and subsequently progresses in the direction of the apical meristem (Gautheret, 1932; LanceNougarkde and Pilet, 1965; Salema, 1971). A similar acropetal pattern of development is found in dark-grown leaves of Hordeum vufgarefollowing their transfer to light, when chloroplast maturation proceeds from the most fully differentiated etioplasts in the upper leaf toward the proplastids of the basal meristem (Robertson and Laetsch, 1974). CHANGES IN Daucus carota D. ULTRASTRUCTURAL

Orange roots of the domesticated carrot, Daucus carota, contain chromoplasts rather than colorless proplastids, but these chromoplasts, too, are transformed into chloroplasts on exposure to light. An early description of the ultrastructural changes which took place during chromoplast development in carrot roots was given by Frey-Wyssling and Schwegler (1965). They found that, as the roots developed, starch accumulated in the eoplasts, but that the starch later disappeared as carotenoid crystals were formed and the roots changed color. FreyWyssling and Schwegler suggested that “vacuoles” formed within the plastids as the stroma broke down, and that carotenoid crystals later formed in these large “vacuoles.” Later, Ben-Shaul e? a f . (1968) carried out a more detailed survey using newer and better fixation techniques. They reported that, for the first 2 or 3 weeks of growth, the root remained colorless; the small proplastids were circular in section and contained some thylakoids. During the next few weeks the roots expanded and became yellow to orange in color; in the plastids the thylakoid system became reduced in extent and the remaining thylakoids “broke down” to become swollen vesicles. These thylakoids seemed thicker, more electron dense, and apparently “rigid. ” The electron transparent “spaces” within the thylakoid sacs often appeared angular. It was concluded from isolation, recrystallization, and electron diffraction pattern studies that the carotenoid crystals were formed within the swollen thylakoid sacs rather than in “vacuoles” derived from the stroma. This conclusion has been supported by subsequent investigations on chromoplast development in carrot roots and in other tissues. Ben-Shad and Klein (1965) have also measured the increase in content of the a and p carotenes in carrot roots during the 20 weeks following germination. They found that initially a carotene was synthesized most rapidly but after about 6 weeks synthesis of p carotene became more rapid and the final concentration of p carotene was more than twice that of ci carotene. The ultrastructural changes undergone by the chromoplasts when carrot roots are exposed to light were investigated by Gronegress (1971). He found that all the chromoplasts in cells of the cortex were rapidly transformed into chloroplasts, but that this transformation took place at different rates in different cell files. After 48 hours exposure to light (8000 lux) the plastids in cells of the

202

JEAN M. WHATLEY

innermost cortex retained much of their chromoplast character, but in cells of the outer cortex the plastids contained carotenoid crystals which had become smaller and had acquired occasional wide irregular grana but only a few single thylakoids. In the subepidermal parenchyma the chromoplasts had lost their carotenoid crystals and had become transformed into chloroplasts with a somewhat limited thylakoid system, which nevertheless contained true grana interconnected by stroma lamellae. The gradient across the carrot root represented by these differential states of chromoplast transformation is further illustrated by differences in the chlorophyll content in these three zones; innermost cortex, 0.5; outer cortex, 10.5; subepidermal parenchyma, 48.0 pg/g dry weight.

VI. Greening and Plastid Division The amount of chlorophyll in a plant organ depends not only on the quantity of chlorophyll present in each plastid but also on the number of plastids in each cell. In the mesophyll cells in expanding leaves of most plants grown in the light, plastid division initially seems to keep pace with cell division but later exceeds it, so that the plastid population in each cell increases to a number which tends to be characteristic of the species (Whatley, 1980); mature leaf palisade cells of Phuseolus vulgaris contain about 40 plastids, whereas those of Spinacea oleracea contain 200-250. In most leaves the total amount of chlorophyll usually increases most during this period of rapid chloroplast division and for a short time afterward. There are, however, a few species, mostly tropical, in which the leaves become visibly green only after leaf expansion is virtually complete. In Theobromacacao, for example, the expanding leaves do not look green although their chloroplasts contain a normal complement of thylakoids and grana. However, each cell contains only three small chloroplasts and it is for this reason that the total chlorophyll content of the leaf is low (Baker et al., 1975). In roots which have become green following exposure to light the chlorophyll content as measured per gram dry or fresh weight is significantly lower than it is in leaves (Fig. 17). Electron micrographs of root chloroplasts show that they often have similar numbers of grana to leaf chloroplasts, though the extent of the stromal thylakoids and the numbers of granal compartments are frequently less than in leaf plastids. Many roots may resemble young leaves of Theobroma in that the low chlorophyll content reflects a low chloroplast population as much as a scarcity of chlorophyll in the individual plastids. Certainly when the large, highly vacuolated cortical cells of nongreen Phaseolus vulgaris roots are examined in the electron microscope many grid squares must usually be scanned before a single plastid profile can be found. Though counts of plastid profiles in sections from the immediate root tip of Zea mays indicate that plastid division takes place within this zone, and indeed that the rate of plastid division within

1liIN t

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

203

3

8c

2

8

4

a -

b

1

C -

FIG. 17. A comparison between chloroplasts in green roots and leaves of 13 day old seedlings of Secale cereale. (a) Grana per cell; (b) number of compartments per granum; (c) photosynthetic efficiency (mg 0 2 mg- I chl.); (d) chlorophyll (mglg fresh weight); (e) total carotenoids. (From Oliveira, 1982.)

I

this selected area (which includes the quiescent center) is greater than the overall rate of cell division (Whatley, unpublished), we do not know how far upward through the root proper plastid division continues in dark-grown roots. (The presence in electron micrographs of constricted plastids is not adequate evidence that plastid division is being completed.) Light is known to promote plastid division in leaves. It is therefore probable that when roots are induced to green on exposure to light, plastid numbers increase [and indeed this has been shown by Heltne and Bonnett (1970) for Convolvulus arvensis], but neither the lag period likely to be involved nor the extent to which plastid division is promoted has ever been investigated. A. THEULTRASTRUCTUREOF DIVIDING PLASTIDS

It has long been assumed, and has now been established (Chaly et al., 1980), that mature chloroplasts in higher plants replicate following constriction. A similar division mechanism appears to operate in proplastids (Chaly and Possingham, 1980; Whatley, 1980). It is probable that, as division proceeds, the constriction between the two plastid halves becomes progressively narrower until, finally, the two daughter plastids become separated. During the final stages of division of mature chloroplasts, small osmiophilic deposits or plaques are commonly found associated with that part of the plastid envelope which lies within the constriction (Suzuki and Ueda, 1975; Leech er al., 1981). A recent survey of root apices of nine species of angiosperm and one species of conifer shows that dividing proplastids, too, have osmiophilic plaques and confirms that these plaques are part of an annulus which encircles the constriction (Chaly and Possingham, 1981). In most of the species examined the annuli in the dividing root proplastids were 15-40 nm wide but those in proplastids of tomato roots

204

JEAN M. WHATLEY

were 40-70 nm wide. The constrictions themselves varied in width from 50 to 400 nm within each sample, suggesting that the root proplastids do not divide in synchrony. In thin sections of mature leaf chloroplasts the constriction associated with division is usually central; in the root proplastids examined by Chaly and Possingham and in proplastids and immature chloroplasts in leaves of Phaseolus vulgaris (Whatley, 1980; Whatley ef al., 1982) the constriction often appears to be off center. It was suggested for Phaseolus that asymmetrical division might take place when polarized elongation growth of the plastid was out of phase with the rate of plastid division.

B. PLASTIDNUMBERS AND SITESOF PLASTID DIVISION Chaly and Possingham found that constricted plastids were most common in the root cap and common in the cortex and stele near the tip of the root proper. Following serial sectioning, no constrictions suggesting division were observed in plastids in epidermal cells, in amoeboid plastids, or in amyloplasts of the central root cap. In dark-grown roots of Phaseolus vulgaris constricted plastids have been seen in the apical meristem and throughout thecortex of the root up to the level of the root hair zone, but not in cortical cells of the lateral root zone or in the cells at the center of the root cap (Whatley, unpublished). Almost no information is available about the numbers of plastids in different types of cells in dark-grown roots. An early investigation of plastid numbers in the root cap and root tip of Zea mays was made from montages of electron micrographs (Clowes and Juniper, 1964; Juniper and Clowes, 1965) but no further work of this type seems to have been carried out. Using median longitudinal sections of resin embedded roots of Zea mays generously given to me by Dr. C. Hawes, I was able to prepare new montages of electron micrographs which I used to count the number of plastid profiles in cell sections within five adjacent files extending through the center of the root from 12 tiers above (i.e., toward the stele) to 12 tiers below the cap junction (Fig. 18). Counts were limited to these 24 tiers for, within them, the plastids were more or less spheroidal, or slightly rod-shaped or just beginning to become amoeboid, and so not subject to the complication of overestimation of plastid number which results when several profiles of the same highly pleomorphic plastid are included within one cell section. Furthermore, within the selected tiers, the cells of the root proper and of the upper part of the cap were of approximately the same size; cells of the central cap were larger but the distally concentrated plastids occupied a volume of cytoplasm not markedly different from that throughout which the plastids were dispersed in the other cells. It is therefore reasonable to assume that the average counts of plastid profiles per cell section within the five files provide a rough but direct correlation with the total number of plastids present in each cell (between two and three times the number of plastid profiles). The results obtained in this

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

205

No. plastid profiles

I

D :I C

'D

I

4 Stages of development

I D

:

Plastid distribution

120

_______ ________ 1st -Cycle?

--;) 4-

- - - -- - - Basic plastid 4

12

8

8 4 Cell tiers

Cell tiers

COP

Stele

12 I

complement

survey are in general agreement with the earlier estimates of plastid numbers by Juniper and Clowes for root tips of the same species. Figure 18 suggests that plastid division takes place in both the cap initials and in the root proper. It also suggests that within the 12 tiers of the root proper included in the survey, the plastids undergo two cycles of division whereas plastids undergo only one division cycle within the 12 tiers of the cap. No division takes place among the displaced statoliths of the central cap, but both the counts of plastid profiles and the presence of constricted plastids in the electron micrographs point to plastid division taking place in cells of the quiescent center. However, none of the observations made here provides any information about comparative rates of plastid division in different parts of the root tip. Furthermore it should be noted that though Fig. 18 indicates that the average numbers of plastid profiles from the five central cell file sections show sharp increases which suggest cycles of plastid division, the plastid populations within these cells do not divide in synchrony. It is also apparent that the phasing of plastid division, the stage of plastid development, and the distribution of plastids within the cell are all features which are independent of each other.

C. THE PLASTIDGENOME Meristematic cells at the base of spinach leaves 2 cm long contain 10-15 plastids, each with about 200 copies of their genome (Scott and Possingham, 1980). These plastids divide but there is no significant synthesis of plastid DNA

206

JEAN M.WHATLEY

after the time when the young leaf cells contain approximately 25 chloroplasts which together have about 5000 genome copies. As these chloroplasts continue to divide the available DNA is partitioned between successive pairs of daughter plastids. By the time the leaves are 10 cm long each cell contains 150-200 chloroplasts each with about 30 copies of their genome. The DNA in root proplastids initially appears to be much less than in leaf proplastids. In the tips of spinach roots in cells containing about 10 proplastids, each plastid has been estimated to contain only about 10 genome copies. When spinach roots are fed with [3H]thymidine,one or two silver grains were found in each plastid profile within the cortex and stele after 60 minutes and three or more grains after 24 hours (Possingham er al., 1983). In these proplastids 75% of the grains were close to the plastid envelope; few were near the infrequent thylakoids. By contrast in mature leaf chloroplasts most of the silver grains are associated with the thylakoid system and few are seen near the plastid envelope. In the root cap, amyloplasts were labeled after 24 hours but not after 60 minutes. The silver grains in the cap amyloplasts were associated with small areas of stroma or with the few thylakoids but not with the starch grains or with the plastid envelope.

VII. Plastids in Sieve Elements Of the publications concerned with the ultrastmcture of plastids in roots, most restrict their observations to plastids in the cortex of the root proper: very few indeed consider plastids in other types of cell. Indeed only sieve elements and the root cap have received any significant attention. Plastids in mature sieve elements of angiosperms are diverse in structure and these plastids are unusual in that their structural differences are taxonomically distinctive (Behnke, 1972). Though plastids in all organs and in all types of cell generally follow the same basic pathway of development, the proplastids and mature chloroplasts in roots, for example, are usually not identical in structure to those in the leaves of the same plant. However, in any one species, sieve element plastids in all organs seem to develop in the same way from the eoplast stage add, at maturity, have the same distinctive characteristics. Behnke divides sieve element plastids into two main groups, S-type plastids, containing at maturity a sieve element starch which often appears coarsely granular (Fig. 9c), and P-type plastids, which may contain starch but, in addition, contain proteinaceous filaments or cuneate or crystalline inclusions that are not membrane bound. Both S-type and P-type plastids are found among dicotyledons and in gymnosperms (Evert, 1977); S-type plastids have not been reported in any monocotyledon. During their development, sieve element plastids become spheroidal in shape;

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

207

any thylakoids which were present disappear; the plastids accumulate the appropriate starch and proteinaceous deposits within the finely granular stroma; the stroma itself becomes progressively less electron dense until, by maturity, it seems to have disappeared; the plastid envelope becomes wavy in outline. Ultrastructural investigations of sieve element differentiation in roots which describe plastid development include those on the dicotyledons Gossypium hirsurum (Thorsch and Esau, 1981), Nicoriana rabaccum (Esau and Gill, 1972), and Phaseolus vulgaris (Esau and Gill, 1971) and on the monocotyledons Allium cepa (Esau and Gill, 1973), Lemna minor (Melaragno and Walsh, 1976), Zea mays (Walsh, 1980), and several palms (Parthasarathy, 1974a-c; Parthasarathy and Klotz, 1976). In roots, the sequential development of sieve elements and their plastids can be followed along the length of selected cell files. The crystalloid inclusions (subunit spacing 80A) appear in the sieve element plastids of Lemna minor roots at a very early stage of differentiation (Melaragno and Walsh, 1976). Indeed in Allium cepa the appearance of crystalline inclusions is the first visible indication that a cell will develop into a sieve element (Esau and Gill, 1973). Esau and her associates have reported on the distribution of plastids within individual sieve elements. In immature sieve elements of Gossypium hirsurum (Thorsch and Esau, 1981) the plastids are dispersed throughout the cytoplasm but in older cells the plastids move to the periphery of the cells often near the end walls; by contrast, mitochondria in these cells tend to move toward the lateral walls. Later some mitochondria, endoplasmic reticulum, and plastids appear to be aggregated around the disintegrating nucleus. In roots of Nicoriana rabaccum the distribution of sieve element plastids seems to be particularly precisely controlled (Esau and Gill, 1972). Even in young sieve elements most plastids lie near the end wall adjacent to the next older cell, i.e., the plastids occupy a site which is the mirror image of that occupied by the amyloplasts in the central cells of the root cap. This polarized distribution of sieve element plastids is very clearly illustrated in Figs. 1, 3, and 4 of the publication of Esau and Gill (1972). No such polarized distribution is shown by plastids in adjacent files of other types of cell in this species, nor does a polarized distribution appear to be shown by sieve element plastids in the other species investigated. In Allium cepa, for example, the published electron micrographs show sieve element plastids adjacent to the side walls and to both end walls (Esau and Gill, 1973). The means by which the distribution of plastids within cells is controlled is not known. Most investigations on plastids in the sieve elements of roots concentrate on the ways in which they develop or on their state when they reach maturity. Parthasarathy (1974~)observed that, in some palms, sieve elements could be long lived, that their mitochondria could persist for 3-6 years, depending on the species, and that plastids could remain for an even longer period. In old sieve elements, the plastids appear to have degenerated, the plastid envelope often

208

JEAN M.WHATLEY

appearing dilated and the stroma being electron transparent. The crystalline and other inclusions also broke down or disappeared. Parthasarathy points out that though some of these responses may be due to fixation injury, light microscopic evidence also suggests that the plastids do, indeed, degenerate.

VIII. Geotropism and Plastids in Root Caps It is more than 80 years since it was first suggested that plants perceive gravity by means of heavy sedimenting bodies, the statoliths. However the mechanism for geotropism is known only for the unicellular rhizoids of the green alga, Chara (Schroter er al., 1975). In Chara the statoliths are large vesicles containing barium sulfate crystals. These inert statoliths provide a gravity-dependent barrier to the movement of Golgi vesicles toward the growing tip of the rhizoid. The direction of growth of the rhizoid is controlled by the symmetry or asymmetry of release of the contents of the Golgi vesicles at the tip. In land plants many different organs, including roots, show geotropic sensitivity but the means by which geotropism is accomplished is uncertain, though it is assuredly different from that in Chara. The subject of geotropism in land plants has been discussed at length in the literature; only those aspects of it relating to plastids in the root cap will be considered here. A. AMYLOPLASTS AS GEOPERCEFTIVE ORGANELLES

The sensing organelles for geotropic response in land plants are generally believed to be amyloplasts and these are thought to function only in conjunction with other organelles. Displaced dictyosomes (McNitt and Shen-Miller, 1978), endoplasmic reticulum (Sievers and Volkmann, 1977; Hensel and Sievers, 1981; Olsen and Iversen, 1980), or endoplasmic reticulum and plasmodesmata operating together to form a multiple valve system (Juniper, 1976, 1977) are among the organelles which have been proposed to operate with the amyloplasts. The difficulty in finding out how the geotropic response is effected in land plants is accentuated by the short time (a few seconds) required for presentation to the stimulus and by the fact that the zone of perception can be many cells re:moved from the zone of response. In the roots of many land plants the main zone of gravity perception is the columella, a core of cells situated in the center of the root cap and acting together. This core of cells has been estimated to be 2000 in Zea muvs (Clowes, 1976), 250-600 in Pisum sativum (Olsen and Iversen, 1980), but only 96 in lateral roots of Nasturrium amphidium (Haberlandt, 1914); in lateral roots generally, geotropic sensitivity is low or absent. The sensitive cells contain more or less spheroidal amyloplasts (Stage 2 of development) (Figs. 2, 3b, 5 . 6, and 9d)

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

209

which are usually packed with starch grains. This starch is unusual in that it is highly resistant to digestion by artificial means (Audus, 1962); nevertheless there is rapid turnover of starch within these amyloplasts (Northcote and PickettHeaps, 1966). When starch is removed from the amyloplasts of the central cap of Lepidium sativum as a result of treatment with gibberellic acid and kinetin, the roots are no longer able to respond to geotropic stimulus (Iversen, 1969). Although plastids in peripheral cells of the root cap also commonly contain considerable amounts of starch, the quantity is less than in plastids in the central cap (Figs. 3b and c and 6) and the earlier capacity of these peripheral cells (formerly central cells) for geoperception has been lost. When the cap of a maize root is removed a new cap forms, following reactivation of mitotic activity in the quiescent center (Juniper et al., 1966). The eoplasts in the quiescent center undergo a temporal sequence of developmental changes, including starch accumulation and loss (Barlow and Grundwag, 1974; Barlow and Sargent, 1978), which parallel those in the spatial sequence of plastid development in the normal root cap (Figs. 6 and 7). In maize, starch accumulation begins within 3 hours of decapping, geotropic sensitivity is first apparent after 14 hours, but a new cap is not regenerated for 3 to 4 days (Barlow, 1974). Thus the presence of a morphologically identifiable cap is not a prerequisite of geotropic response in rootshdeed Haberlandt ( 1914) found that the starch-containing elongation zone of the cortex in roots of Viciafaba, Phaseolus multi’orus, and Lupinus albus all showed some geotropic sensitivity. B. PLASTIDDISTRIBUTION IN CELLSOF THE ROOT CAP In geotropically sensitive roots the amyloplasts in the central cells of the root cap are not randomly distributed but are generally concentrated toward the distal transverse walls (Juniper and French, 1970; Phillips and Torrey, 1974b). When roots are turned from the vertical to the horizontal, these amyloplasts are displaced and eventually become concentrated toward the former longitudinal wall which now forms the base of each cell. Both the Stage 1 eoplasts in the cap initials and the Stage 3 amoeboid plastids toward the periphery of the cap (Fig. 2) remain dispersed throughout their cells, as do the Stage 2 amyloplasts in the subapical cortex of the roots of most angiosperm species. Though the distal displacement of amyloplasts is a regular feature of geoperceptive cells, this displacement alone does not ensure a geotropic response. In Convolvulus arvensis (Tepfer and Bonnett, 1972) and in Zea mays (Shen-Miller, 1978), for example, the geotropic behavior of the roots is influenced by light (a phytochrome reaction). Roots show a positive orthogeotropic response only after exposure to red light even though the statoliths are distally displaced in roots grown both in the light and in the dark. Tischler ( 1905) investigated several angiosperm species with permanently

GEDTROPISM AND

Species Selaginella martensii (aerial rhizophore) Selaginelka martensii (aerial rhizophore in moist container) Sekaginella kraussiana (aerial rhizophore) Selaginelka kraussiana (underground root) Isoetes macrospora Equisem arvense Ophioglossum petiolarum %a., not applicable.

TABLE Ill STARCH IN SOME LOWER VASCULAR h N T S

Single Cap Starch apical Cap Dichotomous Starch starch in Geotropic cell present branching in cap displaced cortex

Cortical starch displaced

Numberof plastids per cell

+ +

+ +

-

+

-

n.a.O

+

+

1

+

+

-

n.a.

+

?

1

+ + +

+ + + + +

+ + + + +

+ + +

-

n.a.

+

1

-

n.a.

+ +

?

1

+ + +

-

?

? ? ?

JO

+? +?

-

+ +

? ?

1

=

Principal souxe Webster and Jagels (1977) (Figs. 7, 10, and 11) Webster and Jagels (1977) Grenville and Peterson (1981) (Fig. 4) Grenville and Peterson (1981) Peterson er al. (1979) Foster and Gifford (1974) Peterson and Brisson (1977)

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

21 1

ageotropic roots. These included some orchids and climbers with aerial roots, some aquatic species, including Eichorniu, and some species with subterranean roots. In these ageotropic species the cells of the central root cap either had plastids which lacked starch or had amyloplasts which were distributed throughout the cell. Tischler also found that in roots of Festucu ovinu and Pou spp. which were initially ageotropic, the appearance of starch in the caps coincided with the acquisition of geotropic sensitivity. In aroids the geotropic “nutritional” aerial roots were well provided with amyloplasts but the only slightly geotropic “grasping” roots had a reduced statolith system. The negatively geotropic “breathing” roots of some species had a normal statolith system. The distal concentration or displacement of amyloplasts within the central cells of the root cap is characteristic of many angiosperm species and of the few gymnosperm species which have recently been examined (Hestnes and Iversen, 1978; Peterson and Vermeer, 1980). However some lower vascular plants show some interesting variations from this pattern (Table 111).

C. PLASTIDS IN ROOTCAPSOF SOMELOWERVASCULARPLANTS Haberlandt (19 14) refers to the absence of starch from root caps of Seluginellu murrensii but to its presence in distally displaced amyloplasts in cells of the inner cortex of the root proper. More recent investigations have shown that some other species of Seluginellu also have root or rhizophore caps which lack starch (Table 111). The thickened, transverse, distal walls and osmiophilic globules have both been suggested as possible alternative sites but as earlier suggested by Haberlandt, the distally displaced amyloplasts of the cortex seem the more probable sites of geoperception. No species of Lycopodium seems to have been examined, but in another lycopod, Zsoeres macrosporu (Fig. 9e), the large, usually single plastid in each central cell of the cap contains starch. These amyloplasts are, however, amoeboid rather than spheroidal in shape and are positioned close to and often curving round the nucleus rather than adjacent to the distal transverse wall (Peterson et al., 1979). The investigation of Isoeres was not extended to the root proper and so information about the presence or otherwise of a possible geoperceptive starch zone in the cortex is not available. Among ferns, only the root cap of Ophioglossum petiolutum has been the subject of recent investigation (Peterson and Brisson, 1977). In Ophioglossum even the plastids in the single apical cell contain starch. Each plastid commonly contains a single, unusually elongated grain, a characteristic feature also of plastids in young fronds of the species. The root amyloplasts are distally displaced even in the proximal cells of the columella and in these cells also the plastids become amoeboid. One of the electron micrographs published by Peterson and Brisson suggests that toward the outer edge of the columella the plastids may contain a few grana, but this restricted thylakoid system, like the starch,

’

212

JEAN M. WHATLEY

appears to be lost toward the periphery of the cap and at the same time osmiophilic globules increase in number. These structural changes suggest plastid development in the root cap of Ophioglossum petiolatum proceeds not only to the mature Stage 5 chloroplast as it does in aquatic species like Eichornia (Mollenhauer, 1967) and Azolla pinnata (Whatley and Gunning, 1981), but that toward the outer edge of the cap the plastids may, uniquely, undergo dedifferentiation.

IX. Plastid Pigments and Responses to Light A. PROTOCHLOROPHYLLIDE, CHLOROPHYLLS, AND CAROTENOIDS

Using the characteristic red fluorescence as an indicator, Hejnowicz (1958) found protochlorophyllideto be present in the root meristems of all 16 species of plants (ranging from ferns to angiosperms) which he investigated. Just as Powell (1925) had found that during greening of seedling roots, the distribution of chlorophyll in different types of cell was characteristic of the species, so Hejnowicz found some species differences in the distribution of protochlorophyllide, though in most species the greatest red fluorescence response was shown by cells in the cortex. The state of protochlorophyllide in dark-grown roots seems to differ from that in etioplasts of dark-grown leaves (Bjom, 1976, 1980). In primary seedling roots of Zea mays the protochlorophyllideis mostly in the esterified form, though there is some unesterified protochlorophyllide at the extreme tip. In these roots the protochlorophyllide is slowly converted into chlorophyllide in red or blue light but, unless the intensity of the light is very low, the chlorophyllide is then destroyed (Bjom, 1963, 1976). In seedling roots of wheat and in the adventitious roots of maize (but not in the primary roots) synthesis of protochlorophyllide and chloroplast development are both triggered by light. The greening of intact roots generally takes place after a considerably longer lag period than it does in etiolated leaves. Once the roots are green, however, they are capable of normal photosynthesis. The low chlorophyll content of root plastids is reflected in the low rates of photosynthesis in roots (Fadeel, 1963; Oliveira, 1982). However, the photosynthetic assimilation number in roots is very high compared with that in green leaves but is similar to that found in yellow leaves of “golden” varieties of some species. The high photosynthetic efficiency in root plastids (Fig. 17) may also be increased by the presence of CO, concentrations in the intracellular spaces, which are significantly higher than those in leaves. These observations of course indicate that the complete photosynthetic apparatus has been synthesized in green root plastids. The chlorophyll a:b ratio appears to be similar to that in leaf plastids (Fadeel, 1962; Oliveira, 1982) but the carotenoids show some differences. Fadeel (1962) found that in

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

213

flax and wheat and Oliveira (1982) found that in rye (Fig. 17) the ratio of carotenoids to total chlorophyll was lower in leaf than in root plastids, the carotenoids being enriched with xanthophylls. Absorption spectra also showed peaks for lycopene and lycophyll in wheat roots but not in wheat leaves. Continuous illumination with white light of low intensity is the regime most commonly used to promote greening in roots. No evaluation seems to have been made of the effects on greening in intact roots by light of different wavelengths or of different intensities, though work of this type has been carried out on excised cultured roots of several species, inter alia, by Heltne and Bonnett (1970) and by Bajaj and McAllan (1969), but principally by Bjorn and by Richter and Dirks. B. BLUELIGHTAND GREENING An absolute requirement for blue light for chlorophyll synthesis and chloroplast development has been shown for excised roots of wheat,cucumber and pea (Bjorn, 1963, 1965, 1967, 1980; Bjorn and Odhelius, 1966; Dirks and Richter, 1975; Richter and Dirks, 1978). However, blue light alone does not bring about chloroplast development. Bjorn (1965, 1980) has suggested a scheme for chloroplast differentiation in roots which involves a series of reactions: (1) the first “red light” reaction brought about by either red or blue light, (2) a dark reaction, (3) a reaction requiring blue light alone, and (4) a second “red light” reaction, brought about by red light alone. The initial response to blue light appears to be the synthesis of specific types of plastid RNA; this response takes place without any lag period. Other slower responses to blue light include the synthesis of chlorophyll and of the chloroplast enzyme, D-glyceraldehyde 3-phosphate: NADP oxidoreductase, which take place only after a lag phase. From electron micrographs of Viciafaba, Dyer et al. (1971) calculated that the ratio of cytoplasmic to plastid ribosomes was low in both the roots and the shoot apices but high in both etiolated and green leaves. Though they give no absolute counts of plastid ribosomes per unit of stromal area it appears that even the most fully differentiated plastids in dark-grown roots usually contain many fewer ribosomes and have a much less extensive thylakoid system than do etioplasts in young leaves kept in the dark. When roots are exposed to light plastid ribsomes very quickly begin to increase in numbers (Fig. 14) but chlorophyll synthesis takes place only after a significant lag period (Oliveira, 1982). Plastids in dark-grown roots generally lack prolamellar bodies. Stetler (1973) examined dark-grown tobacco tissue cultures developed from the pith and equated an absence of prolamellar bodies with the failure (over a short term) of protochlorophyllide to be converted to chlorophyllide. Sundqvist et al. (1980) state that prolamellar bodies generally appear to be formed (as they may be during greening of some roots) only when a blue-mediated process, possibly protein synthesis, runs ahead of a red-mediated one such as the pro-

214

JEAN M. WHATLEY

tochlorophyllide conversion or a phytochrome-dependent reaction. However the results of other investigations (reviewed by Klein, 1982) show that the properties of prolamellar bodies are far from fully understood and their role in thylakoid development is by no means as certain as was formerly believed. The action spectra of the blue light effects in roots resemble the absorption spectra for various carotenoids and flavoproteins, and agree well with that for blue-light-requiring chlorophyll formation in glucose-bleached Chlorellu. It should, however, be noted that an absolute requirement for blue light during greening has only been shown for excised roots and other dark-adapted tissues cultured in media which, like the Chlorellu cultures, are rich in glucose or sucrose (Bjom, 1980). This response to blue light is reminiscent of the curious switch from carbohydrate to amino acid metabolism directed by blue light observed in many organisms (Voskresenskaya, 1972).

X. Some Nonphotosynthetic Functions of Plastids in Roots Little is known about the functions of plastids in roots and it is not the purpose of this article to consider them in any detail. Nevertheless it is perhaps appropriate to refer briefly to some of the few nonphotosynthetic functions which have been ascribed to root plastids, particularly as their isolation for biochemical investigation is now becoming more practicable and one can, perhaps, look forward to future research projects which compare their structure and function. Although it is some years since plastids were first isolated from roots (Thomson et al., 1972), most functional studies have, until recently, made use of' other methods. Autoradiography has been used by Northcote and Pickett-Heaps (1966) among others to trace the synthesis and transport by way of Golgi vesicles of polysaccharides in cells of the root cap. In the summary scheme proposed by Northcote and Pickett-Heaps, plastids have an obvious role as a site of starch storage and a source of the soluble pool of hexose phosphates in the cytoplasm. Plastids and Golgi vesicles have also been implicated in the formation and movement of phenolics, whose widespread occurrence and role, inter alia, in defense against pathogens, make them of particular interest. Mueller and Beckman have tried to determine the origin of phenolics in specialized xylem parenchyma cells in banana roots (1 974) and in endodermal cells in cotton roots (1976). In the endodermal cells of cotton, accumulation of phenols apparently takes place very rapidly (calculated from growth rate to occur within 30 minutes), and can first be detected in vacuoles of cells within 1 mm of the root tip. Mueller and Beckman had previously found a correlation in glandular hairs of tomato, between the accumulation of phenolics and the disappearance of starch from the plastids and they believe that their work on cotton roots shows a similar correlation. However, in the stele of banana roots, the large plastids in phenol-

ULTRASTRUCTURE OF PLASTIDS IN ROOTS

215

accumulating cells have a homogeneous stroma without starch; they also lack thylakoids and invaginations of the plastid envelope, although these plastids do occasionally contain osmiophilic globules which the authors suggest may represent phenolic material. These plastids are closely surrounded by endoplasmic reticulum (ER), but the contents of the ER cisternae are not osmiophilic. The membrane-bound bodies of plastids in some other plant organs and in cell cultures (Gifford and Stewart, 1968; Israel et al., 1969) and osmiophilic deposits in plastids in the coralloid roots of the cycad, Cycas revoluta (Obukowicz et al., 1981) have also been proposed as sites of phenolic accumulation. However, at present no good ultrastructural or histochemical evidence exists which clearly implicates plastids in roots or any other organs of land plants as the site of origin of phenolic material. Nato and Deleens ( 1975a,b) have identified ribulose-bis-phosphate carboxylase (RUBP-carboxylase) inside the plastids of roots of Lens culinaris. Though the amount of RUBP-carboxylase present in dark-grown roots is small there is a significant increase when the roots are exposed to light. The increase in RUBPcarboxylase parallels the increase in chlorophyll synthesis and both processes have the same initial lag period (Fig. 15). When CO, is fixed via PEP-carboxylase, similar results are obtained in the light and in the dark. The amount of CO, fixed in light-grown roots by RUBP-carboxylase is similar to that fixed via PEPcarboxylase, though the latter is the main pathway of CO, fixation in dark-grown roots. Nato and Deleens confirmed (1975b) that the rates of synthesis by Lens roots of C , compounds is the same in the light as in the dark, but that the rates for C , compounds are markedly increased in light-grown roots. More recently there has been increased interest in the role of plastids in nitrite reduction in roots. Emes and Fowler (1979a,b), for example, using isolated amyloplasts from the apices of pea roots, have investigated the intracellular location of the enzymes associated with nitrate assimilation and have found that nitrite reductase, glutamate synthase, and glutamine synthetase are all located in plastids, though there is glutamine synthetase activity also in the cytoplasm. Emes and Fowler also found that all the enzymes of the pentose phosphate pathway were present in small amounts in plastids, though their main site was the cell cytoplasm. They have linked these and other observations together in a scheme which suggests how the pathways of carbohydrate oxidation and nitrate assimilation may interact within the plastids and with reactions taking place in the surrounding cytoplasm. Observations on the reduction of nitrite to ammonia in isolated amyloplasts from root tips of pea and wheat have been further extended to include examination of their responses to waterlogging or anaerobiosis. Dry et al. (198 1) found that under both these conditions nitrite is accumulated and not reduced; they propose that the inhibition of nitrite reduction results from an interruption in the supply of glucose 6-phosphate to the plastids, although ATP appears normally to regulate nitrite reduction. These few examples suggest

216

JEAN M.WHATLEY

that further biochemical work may well provide a much greater insight in the near future into the metabolic functions of plastids in roots.

XI. Conclusions Few land plants can synthesise chlorophyll in the dark. In evolutionary terms the last remnants of this earlier capacity seem to be found in the embryonic cotyledons of some gymnosperms (Kirk and Tilney-Bassett, 1978). Most roots grow below ground in darkness and lack chlorophyll. However, the absence of chlorophyll from these roots is not merely the result of absence of light. Plastid development in most roots (green, nongreen, or becoming green) seems to follow a bidirectional course equivalent to that of cell differentiation. The plastids in roots undergo the same basic sequence of developmental changes (Stages 1-5) as do the plastids in leaves and other organs, though development in roots is often brought to a halt before the plastids differentiate into mature (Stage 5 ) chloroplasts. Although the plastids in roots and, say, leaves follow the same basic pathway of development, the plastids in the two organs differ quantitatively from each other at every stage of development. The extent of the thylakoid system, for example, is usually considerably less in root plastids than in leaf plastids at the same developmental stage and under the same environmental conditions, and, when mature chloroplasts are formed in roots, they generally have fewer thylakoids linking the grana, though they may have as many grana as leaf chloroplasts (Azolla pinnata may be an exception here). In addition, the ancillary structures (containing precursor materials) which are formed in response to the temporary or permanent blockage of the developing thylakoid system differ in character and in the stages at which they develop in different organs. In dark-grown Phaseolus vulgaris seedlings, for example, development of the plastid thylakoid system is initially blocked earlier in the roots and hypocotyls than it is in the primary leaves and prolamellar bodies are formed later in the hypocotyls than in the leaves and not at all in the roots (Whatley, 1978, 1983). When dark-grown plants are transferred into the light, greening takes place much more slowly (if at all) in the roots than in the leaves and different intensities and possibly wavelengths of light are most effective. An early sign of future differences between roots and leaves in the precise patterns of development of their plastids is the smaller number of ribosomes in root eoplasts. The blue light effect shown for excised, cultured roots of several species (Bjorn, 1980) may be associated with reducing this disparity. When roots become green on exposure to light, mature chloroplasts develop in a much smaller proportion of cells than in the leaves; an extreme example of this, observed in the roots of several species, is the restriction of photosynthetically functional chloroplasts to

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a single cell file of the inner cortex (Powell, 1925). In the rhizophores and roots of a Selaginella, loss of chlorophyll can be promoted either by growth in darkness or by growth in the light at high relative humidity (Webster and Jagels, 1977). However, in this Selaginella, each of these two factors promotes a different pattern of chloroplast dedifferentiation. Clearly, then, factors other than light are of major importance in controlling greening; these factors, and light, too, may all influence plastid ultrastructure in different ways, although the basic pattern of plastid development persists. The factors other than light appear to be of particular importance in controlling plastid development in roots, but, unfortunately, it is about these very factors that we have the least information.

ACKNOWLELXMENTS 1 would like to thank Drs. Barlow, Oliveira, Possingham, and Wellburn for sending me prepnnts of their recent publications; Dr. C. R. Hawes for providing me with sections of Zea roots and for an electron micrograph; and Miss T. Scaysbrook and Mr. I. D. A. Kerr for their help in printing the electron micrographs.

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