SPORE DEVELOPMENT IN THE BROWN ROT FUNGI {SCLEROTINIA SPP.)

Transcription

1 New Fhytol. (1969) 68, SPORE DEVELOPMENT IN THE BROWN ROT FUNGI {SCLEROTINIA SPP.) BY H. J. WILLETTS* AND F. D. CALONGEt Department of Botany, University of Bristol {Received 15 May 1968) SUMMARY Development of macro- and microconidia of several species of Sclerotinia causing brown rot of rosaceous fruits was studied by light and electron microscopy. Macroconidia are produced by budding and develop in acropetal succession. Septa form centripetally across the constricted regions between the spore initials of a chain. A single pore in each septum permits movement of cytoplasm and organelles along the chains of spores and these pores are probably not plugged until just before the spores are liberated. The ultrastructure of the mature macroconidia is similar to that of the vegetative hyphae, indicating the unspecialized nature of these spores. Microconidia are formed in succession by extrusion from the tips of terminal phialides. The developing spore is protected by a thin wall and is finally cut off by formation of a basal septum. An uneven splitting of this septum occurs, the thicker part becoming the basal wall of the spore. The latter remains loosely attached to the phialide until another spore is produced. Extrusion of the spores leaves a collar or rim just behind the apex of the phialide. The mature spore contains a large lipid body, a single large nucleus, a small amount of cjrtoplasm and one or two small mitochondria. It is thus of simpler organization than the macroconidium. The development and structure of the two types of spore are compared and the differences correlated with function. INTRODUCTION The fungi causing brown rot of rosaceous fruits (viz. Sclerotinia fructicola (Wint.) Rehm, S. fructigena Aderh. & Ruhl., S. laxa Aderh. & Ruhl. and S. laxa forma mali (Wormald) Harrison) produce both micro- and macroconidia, the latter being placed in the form genus Monilia. Although both forms of spore have been much studied (e.g. Humphrey, 1893; Woronin, 1900; Heuberger, 1934; Hall, 1963; Honey, 1936; Byrde, 1954) some details of their formation and development remain obscure. The present paper describes the genesis and development of both micro- and macroconidia. It forms part of a general investigation of these fungi by H.J.W. The transmission electron microscopy was done by F.D.C. MATERIALS AND METHODS Isolates and material of Sclerotinia fructicola were obtained from stone fruits grown in Australia; the other species were obtained from various localities in southern England. Isolates were cultured on potato dextrose, malt or V8 juice (350 ml V8 vegetable juice, 20 g agar, 1500 ml water) agar media and also by inoculation on pomaceous and drupaceous fruits. * Permanent address: School of Biological Sciences, University of New South Wales, Australia. t Permanent address: Institxito Botanico 'A. J. Cavanilles' (C.S.I.C), Madrid, Spain. 123

2 124 ^- J- WiLLETTS AND F. D. CALONGE Observations were made by light microscopy and with both the Stereoscan and transmission electron microscopes. Material to be viewed with the Stereoscan electron microscope was first examined with a light microscope to confirm the presence of spores, then mounted on an aluminium stub with a proprietary adhesive, and coated under a high vacuum with a thin layer of gold-palladium alloy. Material for examination by transmission electron microscope was fixed in 2% unbuffered potassium permanganate for 30 minutes at room temperature, washed, dehydrated, placed in propylene oxide for 30 minutes, in a i/i mixture (v/v) of propylene oxide and Epon (proprietary embedding material) overnight and finally embedded in Epon. Parallel material was fixed in buffered 6% glutaraldehyde followed by 2% osmium tetroxide, but this method gave less satisfactory fixation. Sections A thick were cut with a glass knife fitted in an L.K.B. Ultramicrotome. These were mounted on copper grids coated with Formvar and stained for 10 minutes in lead citrate (Reynolds, 1963). The sections were examined and photographed with an A.E.I. EM 6B Electron Microscope operating at a potential of approximately 60 kv. RESULTS Macroconidia Development and structure of macroconidia were essentially similar in all the species studied. Thus the following account applies to all of them. Formation of spore chains. Dense tufts (sporodochia) develop on the surfaces of fruits infected with brown rot. Microscopic examination shows that these tufts consist of numerous short conidiophores bearing long chains of conidia (Plate i, Nos. 1-7). The conidiophores are similar in appearance to the vegetative hyphae (Plate i. No. 3) and usually consist of two or three rectangular cells (approximately 13 /i in length). A bud develops at the distal end of a conidiophore and is more ovoid in shape than the mother cell. The bud grows until it is about 10 /x long and then another bud develops at its tip, the original bud continuing to grow until it is the size of a mature spore. This process is continued until ten or more buds are produced. Sometimes, if the environmental conditions are changed, an ordinary vegetative hypha will grow out from the terminal bud (Plate i. No. 5). Such a change in type of growth was observed when Vernon-slide cultures were transferred from a moist to a dry atmosphere. Branched chains of conidia are commonly produced. The branches usually arise from triangular buds which produce two outgrowths (Plate i, Nos. 6 and 7) each of which gives rise to a new branch. The new branches may grow at equal rates but usually one becomes dominant and the growth of the other is restricted. Further branching may take place and many spores then develop from one conidiophore (Plate i, No. 6). When the spore chain breaks, the triangular cells germinate and function as spores. Also a spore some distance from the tip of a chain may bud again and start a sub-apical branch (Plate i. No. 2). Plate I, No. 7 shows the budding off of new cells on one branch, while spores are being liberated on the sister branch. The spore at the distal end of the chain is usually the first to be liberated. When a chain breaks some distance from the free end, each spore in this short chain may germinate. A conidiophore may produce secondary chains after dissemination of the spores of the original ones.

3 Sclerotinia spore development 125 Further details of budding, maturation and release of macroconidia were studied by electron microscopy. Initiation of spores by budding. The ultrastructure of the young conidiophore resembles that of the aerial vegetative hyphae (Plate 2, No. 21) both in the nature and arrangement of the organelles and in wall structure. The wall in both has an outer layer of electron-dense material. Such a layer is absent in submerged hyphae and from this fact and its general appearance the electron-dense material is considered to be derived from mucilaginous material. The first sign of spore initiation is a slight thickening of the wall over the apex of the conidiophore together with the loss of electron density in the outermost layer at this region. The plasmalemma underlying this altered apical section of the wall becomes somewhat wrinkled and contact between it and the wall becomes less close (Plate I, No. 8). The spore bud is blown out by stretching of this altered apical wall and the apical nucleus moves into it from the conidiophore. The electron-dense outer wall layer becomes reconstituted around the spore (Plate i. No. 9) and the whole process is repeated by the development of another spore bud at the apex of the first formed one. This process continues thus producing the characteristic chain of spores. Plate i. No. 9 shows a nucleus passing into the most recently formed spore initial at the apex of a young chain. Delimitation of spores by septum formation. The completion of spore development by the formation of septa separating individual spores does not follow a definite pattern. A septum may develop between the conidiophore and the first bud soon after the bud forms and, as each successive bud is produced, a new septum develops or a whole chain may be formed before septation begins (Plate i. No. 9). When large numbers of chains are being produced in a moist atmosphere, the chains may remain in the aseptate condition for long periods. Byrde (1954) found that a period of exposure to low humidity is an essential preliminary to the differentiation and separation of the macroconidia of Sclerotinia fructigena. Each septum forms centripetally across the constrictions between two spore initials (Plate I, Nos. 13 and 14) and eventually becomes complete except for a simple central pore, through which cytoplasm and organelles continue to move freely along the chain. A nucleus, surrounded by one or more additional membranes is always found at the site of septum formation (Plate i, Nos. 13 and 14). Plate I, No. 12 is a Stereoscan electron micrograph showing a chain of mature spores about to separate. Shrinkage of the spores during processing reveals that they are attached only at the pore area of the septa. Examination of liberated spores suggests that the pores finally become plugged and the last connection between adjacent spores is then readily broken. Ultrastructure of mature liberated macroconidia. The mature spore is more or less ovoid, but with a flattened papilla at each pole. The cell wall is about 0.2 ju thick throughout the later stages of development. The wall consists of the characteristic thin electrondense outer layer, approximately 0.02 jx thick (resembling that of walls of conidiophores and vegetative hyphae described above) and a relatively electron-transparent inner layer (c n thick). The flattened polar papillae can be seen by light microscopy (Plate i. No. 7) and more clearly by electron microscopy (Plate i. No. 11). Plate i. No. 12 shows the external form of these. It is clear that they are more rigid than the lateral walls since they neither collapsed nor shrank noticeably during processing for Stereoscan viewing. Plate I, Nos. 10 and 11 are transmission electron micrographs, the former passing tangentially and the latter medianly through one of these papillae. No. 11 shows the

4 126 H. J. WiLLETTS AND F. D. CALONGE plugged pore of the original interspore septum and both figures show that the clearly defined electron-dense outer wall layer does not extend over the papillae but is broken at the site of the original constriction between the spore and its neighbour. The spore contains several well-developed nuclei which, at least in young spores, show evidence of recent division, numerous globose mitochondria with well-developed cristae, a rather sparse, irregular membrane system, glycogen granules and dark bodies of uncertain nature, which perhaps consist of lipid material, are also present, (Plate i, Nos. 8-11, 13 and 14), and resemble those of the vegetative hypha (Plate 2, No. 21). Microconidia S. fructicola formed microconidia in culture under a greater variety of conditions and in greater numbers than did any of the European species. Of the latter S. laxa f. mali produced microconidia the most readily. Accordingly, the following account refers mainly to S. fructicola. The microconidia of all species were, however, similar in appearance and agreed with Heuberger's (1934) description of those of S. fruticola. Fig, I, Camera lucida drawings and tracings of microconidia produced in culture, (a) Phialides and microconidia of Sclerotinia fructicola. (b) Microconidia of 5, fructicola produced on a germ tube growing from a macroconidium, (c) 5, fructicola: several single phialides arising at rigbt angles from a vegetative hypba growing in agar cultures, (d-h) S. fructigena: pbialides and microspores produced on macroconidia in old agar cultures kept in tbe dark. Production of microconidia. Some isolates of S. fructicola when grown on potato dextrose agar in petri dishes formed many black pycnidium-like masses of microconidia, which were particularly abundant where the margins of the colony were in contact with the side of the dish. The microconidia became dispersed as a cream-coloured suspension in drops of water condensing on the surface of the colony. Plate 2, No. 15 shows a cluster of microconidia and Plate 2, No. 16 illustrates the coalescence of such (h)

5 Sclerotinia spore development 127 clusters to form a single mass. Fig. i shows isolated microconidiophores which occur scattered over the mycelium and also illustrates the formation of microconidia by germ tubes produced by macroconidia of S. fructicola germinating in situ on low-nutrient media. Similar formation of microconidia from germ tubes of macroconidia germinating in situ were seen in old cultures of 5. fructigena, on the surfaces of apricots and plums infected with S. laxa and on apricots with S. laxa f. mali. (a) (d) A PH Fig. 2. The formation of the second and subsequent microconidia of Sclerotinia laxa f. mali. (a and b) Early stages in the extrusion of the tip of the protoplast of a phialide (PH). Note the wall of the phialide (W) which is continuous with the thin spore wall (SW) and the collarette (C); also the layer of mucilage (MUC) over the surface of the phialide and spore walls, (c) The spore wall has thickened, (d) Centripetal growth of a septum (S) across the canal between spore and mother cell; a simple pore (P) remains in septum, (e) Unequal splitting of septum between spore and phialide. The newly formed spore is slightly pear-shaped and has a large oil drop (L) and a comma-shaped nucleus (N). Development of microconidiophores and microconidia. The microconidiophores are dichotomously branched hyphae of diameter )"> i-^- considerably narrower than vegetative hyphae the average diameter of which is 5.8 /i. The degree of branching depends on cultural conditions. The septate branches (Plate 2, No. 19) terminate in bottle-shaped, often asymmetric phialides, from the apices of which the microspores are cut off in succession. After the release of the first spore the phialide shows a collar-like structure at the tip below which it is slightly constricted. The young phialide has an internal structure similar to that of vegetative hyphae and macroconidiophores and is surrounded by a wall resembling that of the vegetative hypha, macroconidiophore and macroconidium already described, i.e. having a thin outer electron-dense layer and a thicker inner one relatively electron transparent. Details I N.P.

6 128 H. J. WiLLETTS AND F. D. CALONGE of the formation and release of the first spore formed by a phialide have not been completely worked out but the available evidence suggests that the outer layer of the wall over the apex of the phialide is ruptured and that the inner wall is either ruptured or rendered plastic, thus allowing the extrusion of the microspore as a bubble-like structure. The ruptured rim of the original wall forms the collar through which later-formed microconidia extrude. Plate 2, No. 17 is believed to show the extrusion of the first microconidium formed by a particular phialide since the collar is not yet fully formed. Development of the second and subsequent spores by the phialide can be more easily followed (Fig. 2). These are formed by the protrusion of the tips of the protoplast of the phialide beyond the collar. At first it is difficult to distinguish a definite wall around the apex of the phialide but the blown-out spore initial is surrounded by a thin electron-transparent zone (Fig. 2a) which is continuous with the innermost layer of the phialide wall. An electron-dense layer is soon developed on the outer side of the spore wall and is continuous with the phialide wall within the collar (Fig. 2b). As the spore develops the wall becomes thicker (Fig. 2c) and the centripetal growth of a septum across the canal linking spore and phialide seals off the spore except for a central septal pore (Fig. 2, d and e; Plate 2, Nos. 18 and 20). Finally the septum splits unequally leaving a thin layer over the tip of the phialide and a thicker one forming the basal wall of the spore (Plate 2, Nos. 18 and 20). The newly formed microconidium is slightly pear-shaped (Fig. 2e) and usually remains loosely connected to the phialide until another spore begins to form or even longer. Short chains of up to six microconidia, probably held together by mucilage, have been seen. Ultrastructure of the mature microconidium. The mature spore wall (approximately O.I^ fi thick) consists of the electron-transparent inner layer and electron-dense outer one (Plate 2, No. 20) characteristic of vegetative hyphae and conidiophores of these fungi. In young spores there are usually two or three bodies which coalesce to form a large body in the mature spore. These bodies probably contain lipid material as they show black with glutaraldehyde and osmic fixation and were seen as blank white spaces in comparable material fixed with permanganate. Each microconidium also contains several small, globose or ellipsoidal mitochondria, with obvious cristae, and one clearly defined nucleus which often fills much of the space left between the lipid body and the cell wall (Plate 2, No. 20). In immature spores there is a loose and irregular reticulate membrane system (Plate 2, No. 17) but in spores completely separated from the phialide (Plate 2, No. 20), only a few membranes are seen and the large lipid body and the nucleus occupy most of the available space. DISCUSSION Light microscope studies support Hall's (1963) suggestion that macroconidia of the brown rot fungi are blastospores, i.e. produced by budding, and the development is acropetal. However, the buds have broad bases, and septa are laid down across the constrictions. These features tend to emphasize the gradation that exists from blastospores to arthrospores. Turian (1966) described conidium formation in Monilia fructigena (Fries) Westendorp as arthrosporic, but the present writers consider that these spores should be interpreted as blastospores. The macroconidia of the form genus Monilia are usually considered to be unspecialized. This is supported by the similarity in ultrastructure of spores and vegetative hyphae demonstrated in the present paper and by the readiness with which spore formation

7 Sclerotinia spore development 129 changes to vegetative growth. Apart from the regular constrictions, a chain of spores is very similar to a septate hypha and there is an unobstructed route for the movement of cell contents along the chain to the developing apical bud. This movement takes place through the simple pores in the septa and the development of new buds will continue until one of the septal pores is plugged or environmental conditions change the type of growth. The lateral walls of separated macroconidia were found to have an outer electrondense layer which did not continue over the ends of the spores (Plate i, Nos. 10 and 11). This layer may be the remains of the primary wall that dries out and ruptures just before the chain breaks up, disseminating the spores. Woronin (1900), working with Sclerotinia fructigena and S. laxa, presented very careful drawings to show the process of separation of conidia in the chains and Honey (1936) also described how conidia of the brown rot fungi are set free by the continued growth in size of each conidium in the chain, leading to a final rupturing of the outer primary wall. The absence of an outer electron dense layer on hyphae growing in an agar medium (Willetts and Calonge, unpublished observation), where desiccation of tissues is greatly reduced, may indicate that the electron-dense layer is formed by collapse and drying out of an outer, probably mucilaginous layer. Turian (1966) reported a similar thin layer on the outside of conidia of Neurospora crassa which seemed to be related to their hydrophobic property. He suggested that the layer might be mainly phospholipid associated with protein. The mode of formation of the microconidia of the brown rot fungi differs considerably from that of the macroconidia. The present observations support Heuberger's (1934) account of microspore structure and formation and are similar to those of Brierley (1918) for microspores of Botrytis cinerea. All microspores apart from the first are formed by the extrusion of the tip of the phialide through an opening at the apex of the cell. The constricted neck of the tube, with the surrounding collar may be of importance in the build up of the internal pressures required to blow out the microspore. A spore formed in this way agrees with Tubaki's (1958) description of a phialospore. The microspores of these fungi are thus radically different from the macroconidia of the same species, which are interpreted as blastospores. It is unfortunate that the method of formation of the first microconidium by a phialide and the initiation of the collar, which is such a conspicuous feature at later stages, could not be ascertained with certainty. Such evidence as is available suggests that the first spore of a phialide emerges through rupturing of the apex of the phialide, so leaving the typical collar through which later spores emerge. Such a process has been described in the budding of yeast cells (Nickerson, Falcone and Kessler, 1961; Marchant and Smith, 1967)- As long ago as 1851, Tulasne referred to the microspores of Discomycetes as either spermatia or conidiola. Drayton (1934) produced conclusive experimental evidence that the microconidia of Sclerotinia gladioli are functional spermatia, and similar results have since been obtained with several related species (Drayton, 1937; Gregory, 1938, 1941; Groves and Drayton, 1939). It has been suggested (Griffiths, 1959), but without clear experimental proof, that the microconidia of the brown rot Sclerotinias may also function as spermatia. The relatively simple internal structure of the microconidia is thus of particular interest. The microconidia described in the present paper possess all the essentials to allow them to function as spermatia, viz. a nucleus large in proportion to the size of the cell, a relatively small amount of cytoplasm and a protective cell wall. In addition these spores contain a few small mitochondria and a large lipid body. Thus

8 130 H. J. WiLLETTS AND F. D. CALONGE the mechanisms and materials for some aerobic energy production are present. While this might be expected to suffice for the functioning of a spermatium (i.e. fusion with a receptive hypha or trichogyne and subsequent nuclear migration into the latter) it is unlikely that germination of such a spore would lead to the development of an active germ tube except under unusually favourable conditions. Although Humphrey (1893) and Jehle (1914) claimed that normal germination can take place, later investigators were unable to verify this and the present writers have seen no example of germinating microconidia. In contrast to the simple organization of the microconidia, the internal structure of macroconidia resembles that of active vegetative hyphae and is correlated with their capacity for rapid germination leading, under suitable environmental conditions, to the establishment of a mycelium. The fact that both the mode of formation and the internal structure of macro- and microconidia are so strikingly different supports the view that their functions are also different. The macroconidia are an efficient type of dispersal spore. If microconidia ever function similarly they must do so with exceedingly low efficiency and ail the evidence suggests their originally spermatial nature, whether or not they are now functional. ACKNOWLEDGMENTS The authors wish to express their gratitude to Professor L. E. Hawker for her encouragement and advice in planning and writing up the work; to Dr R. J. W. Byrde and Dr M. F. Madelin for their interest and suggestions during the investigation and the preparation of the manuscript; to Professor H. E. Hinton, E.R.S., for granting permission to use the Stereoscan electron microscope provided for him by the Scientific Research Council, and to the authorities of the Universities of Bristol and New South Wales for providing facilities for this work. Thanks are due to the Agricultural Research Council for a grant to one of us (F.D.C.). REFERENCES BRIERLEY, W. B. (1918). The microconidia oi Botrytis cinerea. Kew Bull. Misc. Inf., 129. BYRDE, R. J. W. (1954). Observations on the sporulation of Sclerotinia fructigena on mummified apples and plums in late spring and summer. Rep. agric. hort. Res. Stn Univ. Bristol, 163. DRAYTON, F. L. (1934). The sexual mechanism of Sclerotinia gladioli. Mycologia, 26, 46. DRAYTON, F. L. (1937). The perfect stage oi Botrytis convoluta. Mycologia, 29, 305. GREGORY, P. H. (1938). Sclerotinia polyblastis n. sp. on Narcissus, the perfect stage oi Botrytis polyblastis Dowson. Trans. Br. mycol. Soc, 22, 201. GREGORY, P. H. (1941). Studies on Sclerotinia and Botrytis 1. Trans. Br. mycol. Soc, 25, 26. GRIFFITHS, E. (1959). The cytology of Gloeotinia temulenta (blind seed disease of rye-grass). Trans. Br. mycol. Soc, 42, 132. GROVES, J. W. & DRAYTON, F. L. (1939)- The perfect stage oi Botrytis cinerea. Mycologia, 31, 485. HALL, R. (1963). Cytology of the asexual stages of the Australian Brown Rot Fungus Monilinia fructicola (Wint.) Honey. Cytologia, 28, 181. HEUBERGER, J. W. (1934)- Fruit-rotting Sclerotinias. IV. A cytological study of Sclerotinia fructicola (Wint.) Rehm. Bull. Midi, agric Exp. Stn, 371, 167. HONEY, E. E. (1936). North American species oi Monilinia. I. Occurrence, eroupins and life-histories. Am. J. Bot., 23, 100. t- 5, HUMPHREY, J. E. (1893). On Monilia fructigena. Bot. Gaz., 18, 85. JEHLE, R. A. (1914). Brown rot of orchard fruits. Ph.D. thesis, Cornell University MARCHANT R. & SMITH, D. G. (1967). Wall structure and bud formation in RJiodotorula glutinis. Arch. Mikrobiol., 58, 248. NiCKERSON, W J., FALCONE, G. and KESSLER, G. (1961). Polysaccharide-protein complexes of yeast cell wall. In: Macromolecular Complexes. (Ed. by M. V. Edds), 6th Symp. Soc gen. Physiol New York. REYNOLDS, h. b. (1963)- The use of lead citrate at high ph as an electron-opaque stain in electron microscopy. J. biophys. biochem. Cytol, 17, 208. TuBAKi, K. (1958). Studies of the Japanese Hyphomycetes. V. J. Hattori hot. Lab., 20, 142. ^

9 THE NEW PHYTOLOGIST, 68, i 2 W HW, HW, H, J. WILLETTS AND F. D, CALONGE SCLEROTINIA SPORE DEVELOPMENT (facing page 130)

10 THE NEW PHYTOLOGIST, 68, i H. J. WILLETTS AND F. D. CALONGE SCLEROTINIA SPORE DEVELOPMENT

11 Sclerotinia spore development 131 TuLASNE, L. E. (1851), Note sur I'appareil reproducteur des Lichens et des Champignons, Ann. ScL not nept, 3, 15. TURIAN G. (1966). The genesis of macroconidia of Neurospora. In: The Fungus Spore. (Ed by M F Madehn), p. 6i. Butterworths. a i- \ j.. WORONIN, M, (1900). Uber Sclerotinia cinerea und Sclerotinia fructigena. Mem. Acad. Sci. St Petersbs VIII e s6r, 10, Phys. Math., i.. '' EXPLANATION OF PLATES Key to symbols: HW, hyphal wall; HWj, electron-dense outer layer of wall; HW2, electrontransparent layer of wall; S, septum; P, septal pore; ER, endoplasmic reticulum or membranes; N, nucleus; M, mitochondrion; G, glycogen; L, lipid bodies; PL, plasmalemma- SW, spore wall; PH, phialide; PHW, wall of phialide; MUC, layer of mucilage; C, collar; NK, constricted neck; SP, microconidium; NM, nuclear membrane (surface view). PLATE I Development and structure of macroconidia, (Nos, 1-7, light micrographs; No, 12, Stereoscan electron micrograph, the rest transmission electron micrographs,) Nos, 1-6, 8-14, Sclerotinia fructigena] No. 7, S. fructicola. No, I, Conidiophore bearing a short chain of macroconidia. No. 2, Formation of a sub-apical branch from a macroconidium. No. 3, Branching of a conidiophore. No, 4, Branched chain of macroconidia. No, 5, Growth of a hypha from the most distal spore of a chain. No, 6, Origin of two branches from a triangular bud. No. 7, Budding off of new cells on one branch and liberation of macroconidia from the sister branch. No. 8, Early stage in the formation of a macroconidial bud. Note the absence of an electrondense outer layer at the tip, the poor contact between the plasmalemma and the hyphal wall, and the nucleus. No, 9, Aseptate chain with the most distal cell in process of being blown out; note nucleus moving into the young bud. No, 10, Part of a mature macroconidium. Note that at the end the eell wall is thicker and its outer layer is less electron-dense. No. II. Median section of a mature macroconidium. Note the septum with a simple pore. No. 12, Macroconidia about to separate. Note the connections between the spores in the position of the septal pores, and the thickened walls at the poles of the spores, Nos Successive stages, in the formation of a septum between macroconidial buds. No, 13, Slight thickening of wall at constricted region between buds, note nucleus with additional membranes; No. 14, centripetal growth of septum. PLATE 2 Development and structure of microconidia (lettering as in Plate i), (No, 15 Light micrograph; No, 16 Stereoscan electron micrograph, the rest transmission electron micrographs,) No, 15, A cluster of microconidia of Sclerotinia fructicola. No, 16, Small microconidial cluster of S. fructigena. No, 17. Longitudinal section through a phialide showing early stage in microconidium formation. No, 18. Late stage of development showing the formation of a septum between developing microconidium and phialide. The continuity of the contents of the microconidium and the mother cell is still maintained through a simple pore. Note the constriction of the neck of the phialide and the well-developed collar. No, 19, Longitudinal section to show septum between phialide and neighbouring cell; note the simple pore. No. 20. A mature microconidium loosely attached to a phialide, A thin wall extends over the tip of the phialide. No. 21. Longitudinal section of vegetative hypha showing hyphal wall, septum, nuclei, mitochondria, glycogen, lipid bodies and membranes.

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