DEVELOPMENT IN VITRO OF
MICROSPORIDIA FROM INSECT HOSTS
by
XIE WEI-DONG, M.Sc.
(Zhongshan University, Guangzhou)
Thesis submitted for the Degree of Doctor of Philosophy
University of London
and for the Diploma of Imperial College
Imperial College of Science & Technology
Department of Pure & Applied Biology
Silwood Park
Ascot
Berkshire November 1985 2
ABSTRACT
Five species of microsporidia derived from insect hosts, Nosema sp. from Spodoptera litura, N. heliothidis, N. locustae, Vairimorpha necatrix and Pleistophora opero- phterae, were successfully established in £5. frugiperda cell cultures by addition of haemolymph from larvae infected with the microsporidia. By using these systems, the growth and development of the 5 species of microsporidia and spread of infection by microsporidia from cell to cell were investi gated in vitro. Four of the species of microsporidia did grow at 30°C, but P. operophterae did not. The most favourable temperatures for growth of microsporidia were
20°C and 25°C. Extracellular vegetative stages of micro sporidia were capable of initiating infections in new cells, but their ability was almost completely lost within 30 min by the Nosema sp., 60 min by N. heliothidis and V. necatrix,
120 min by P. operophterae and 180 min by N. locustae.
Addition of ATP to the medium resulted in more cells becoming infected with Nosema sp. and N. heliothidis and further confirmed that extracellular infective stages are responsible for infection of new cells. Meronts were observed by light and electron microscopy attached to the surface of host cells. A dose dependent inhibition of uptake of both parasites and red blood cells into Spodoptera cells by Cytochalasin B, provided evidence for entry of parasites into cells by phagocytosis. Inhibition of 3
infection by lectins and carbohydrates suggested that glycoproteins and/or glycolipids on parasites and host cell
surfaces interact during parasites entry into cells.
Phagosome-lysosome fusion was prevented in cells
containing the vegetative stages of P. operophterae but not
in cells containing only spores.
Nosema locustae, a microsporidia infecting locusts was
shown to be dimorphic at low temperatures vivo producing
Nosema type free spores and membrane bound octospores. In
an ultrastructural study of N. locustae, it was revealed
that synaptonemal complexes occur in early stages of the
octosporoblastic sequence, implying that meiosis occurs at the beginning of this sporogony, so that 8 haploid spores are produced. This type of development is characteristic
of the genus Vairimorpha. Neither N. locustae nor V.
necatrix produced octospores jjn vitro and attempts to induce
dimorphism in vitro by applying juvenile hormone or p-ecdysone to the medium failed.
Spores hatched when subjected to a shift from alkaline
conditions (pH 11-13 for Nosema sp., and N. heliothidis or
pH 9.5 for Orthosoma operophterae, Glugea anomala and
Amblyospora sp., to acid on neutral conditions (pH 6-7) in
1M KC1 buffer solution. 4
ACKNOWLEDGEMENTS
I wish to express my gratitude to Professor E.U. Canning for her supervision and encouragement throughout the course of my study and for many helpful and stimulating discussions.
I am grateful to Dr. D.C. Kelly and his colleagues, of
Institute of Virology, NERC, Oxford, for supplying Spodoptera frugiperda cell line and Heliothis zea and Marnestra brassicae eggs and to Dr. R. Gordon for Heliothis virescens eggs,, originally from the Bioenvironmental Insect Laboratory,
Mississippi, U.S.A. I am indebted to the following people for the supply of samples of microsporidian spores: Professor E.U. Canning for
Pleistophora operophterae and Orthosoma operophterae spores;
Dr. J.V. Maddox of the Illinois Natural History Survey,
Urbana, Illinois U.S.A., for Vairimorpha necatrix spores; Dr.
J.E. Henry of the Rangeland Insect Laboratory, Montana State
University, Bozeman, Montana, U.S.A., for Nosema locustae spores; Dr. R. Ishihara, of the Faculty of Agriculture and
Veterinary Medicine, Nihon University, Kanagana, Japan and Mr.
Z. Wang, of the Department of Sericulture, South China
Agricultural University, China, for Nosema bombycis spores;
Dr. J. Weiser, of the Institute of Entomology, Czechoslovak
Academy of Sciences, Prague, Czechoslovakia, for Pleistophora schuberqi spores; Dr. W.M. Brooks of the North Carolina
University, Raleigh, North Carolina, U.S.A., for Nosema heliothidis spores. 5
I am grateful to Mr. J.P. Nicholas and Mr. R. Hartley for their valuable technical assistance. I would also like to thank Mr. Z. Liu, Institute of Entomology, Zhongshan
University, Guangzhou, China, for drawing the diagram of fine structure of microsporidian spore, and Mr. Ian Fosbrooke for generating three-dimensional graphs.
My thanks also go to all my friends and colleagues at
Imperial College, Silwood Park, for their help and advice.
I should like to thank Zhongshan University, Guangzhou,
China, for financial support during this period of my study in
U.K.
I would like to express my sincere thanks to Professor Pu
Zhelong, Institute of Entomology, Zhongshan University,
Guangzhou, China, whose careful and stimulating supervision developed my first interest in microsporidiology.
Finally, my love and thanks to my parents, my sisters, particularly my wife, for their constant encouragement and support over these years. 6
TABLE OF CONTENTS
Page ABSTRACT 2 ACKNOWLEDGEMENTS 4 TABLE OF CONTENTS 6 LIST OF FIGURES. 12 LIST OF TABLES 20
1 INTRODUCTION 21
2 LITERATURE REVIEW 22 2.1 Germination of spores jji vitro 22
2.2 Microsporidia in tissue cultures 26
2.2.1 Techniques for the establishment 26
of microsporidia in tissue
cultures
2.2.2 Factors affecting microsporidian 31 infectivity and multiplication in
vitro
2.3 Current taxonomic position of three 33 genera of microsporidia and their
developmental cycles
2.3.1 The genus Nosemaand its related 33
genera
2.3.2 The genus Vairimorpha and other 36
dimorphic genera 2.3.3 The genus Pleistophora and 39 related genera 7
Page 2.4 Interaction of parasitic protozoa with 42 their host cells
2.4.1 Invasion into the host cells 43
2.4.1.1. Microsporidia 43
2.4.1.2. Other protozoan parasites 51
2.4.2 Fate of parasitic protozoa in host 55
cells. 3. MATERIALS AND METHODS 59
3.1 Microsporidian species 59
3.2 Infection of insects 59 3.3. Collection and purification of spores 61
from infected insects.
3.4 Induction of spore hatching 62
3.5 Maintenance of insect cell line 64
3.6 Infection of cell cultures 65
3.6.1 Inoculation of cell cultures with 65
spores
3.6.2 Inoculation of cell cultures with 65
the haemolymph from infected
insects 3.7 Maintenance of raicrosporidia in cultures 66 of £5. frugiperda cells
3.8 The effect of temperature on development 67
and multiplication of microsporidia in
cultures 3.9 The effect of antibiotics and ATP on the 68 growth of Nosema sp. and Nosema
heliothidis 8
Page 3.10 Isolation of pre-spore stages (meronts 68 and sporonts). 3.11 Determination of viability of 70
extracellular parasites
3.12 Determination of the phagocytic ability 70 of 55. frugiperda cells
3.13 Determination of the effect of 71
antiphagocytic reagent on infection of
£5. frugiperda cells by N. heliothidis
3.14 Demonstration of possible receptor 72
involvement in microsporidian invasion of
cells
3.15 Preparation of electron microscopy 73
3.15.1 General method 73
3.15.2 Detection of acid phophastase by 73
electron microscopy in 55.
frugiperda cells infected with P.
operophterae and N. heliothidis
3.15.3 Detection of secondary lysosomes 74
in 55. frugiperda cells infected with P. operophterae and N.
heliothidis
3.16 Attempts to induce dimorphism ir\ vitro by 74
applying hormones 9
Page
4 RESULTS 77
4.1 Germination of spores jin vitro 77
4.1.1 pH optima for pretreatment and 77 treatment
4.1.2 Time for completion of spore 79
germination
4.2 Establishment of microsporidia in the 79
Spodoptera frugiperda cell line
4.2.1 Attempt to infect cell cultures 79
with spores
4.2.2 Infection of cell cultures with 81
infected haemolymph
4.3 The effect of temperature on multi- 81
plication of microsporidia in £>.
frugiperda cells 4.3.1 Nosema heliothidis 83 4.3.2 Nosema sp. 86 4.3.3 Nosema locustae 89 4.3.4 Vairimorpha necatrix 89
4.3.5 Pleistophora operophterae 94
4.4 The effect of antibiotics on infectivity 97
to J3. frugiperda of Nosema sp. and N.
heliothidis
4.4.1 The effect of antibiotics on 97
N . heliothidis
4.4.2 The effect of antibiotics on 97
Nosema sp.
4.5 Viability of extracellular stages of 99
microsporidia in TC-100 medium 10
Page
4.6 The effect of adenosine triphosphate 105 (ATP) on invasion of cells by Nosema sp.
and N. heliothidis
4.7 The effect of Cytochalasin B on entry of 107
N. heliothidis into £3. frugiperda cells
4.8 The effect of lectins and carbohydrates 110
on invasions of S. fruqiperda cells by different species of microsporidia
4.9 Attempts to induce dimorphism in vitro 114
4.10 The morphology and life cycle of micro- 114 sporidia studied by light and electron
microscopy
4.10.1 N. heliothidis in vitro 114 4.10.2 Nosema sp. from Spodoptera litura 118
in vitro
4.10.3 Pleistophora operophterae in vitro 119
4.10.4 Life cycle of N. locustae in vivo 121
4.10.5 Attempt to induce dimoirphism in 125
Nosema bombycis
4.10.6 Nosema locustae in vitro 125
4.10.7 Light microscope observation on 125
Vairimorpha necatrix in vitro 4.10.8 Phase contrast light microscope 126
and electron microscope
observations of infective stages of N . heliothidis
4.11 Fate of P. operophterae inside the 127
£>. frugiperda cells. 11
Page
4.11.1 Labelling of secondary lysosomes 127
in £3. fruqiperda cells infected
with P. operophterae 4.11.2 Detection of acid phosphatase in 128
£3. frugiperda cells infected with
P . operophterae 4.12 Fate of N. heliothidis after uptake into 128
£3. f ruqiperda 5 DISCUSSION 164
5.1. Microsporidia in cell cultures 164
5.1.1. Establishment of microsporidia in 164
cell cultures 5.1.2 The effect of temperature on 166
microsporidia
5.1.3 The effect of antibiotics on 168
microsporidia
5.2 Cell to cell transmission of micro- 169
sporidia 5.3 The ways of entry of infective stages of 174
microsporidia into cells
5.4 Fate of microsporidia within cultured 178
insect cell
5.5 Dimorphism and meiosis 182
5.6 Sporophorous vesicle 190
5.7 Factor controlling host and tissue 192
specifity of microsporidia
BIBLIOGRAPHY 196
APPENDICES 218 12
LIST OF FIGURES
Fig. Page 1 . Diagram of fine structure of microsporidian 44 spores. 2. The effect of pH on germination of spores 00 of Nosema sp. from S. litura. 3. The effect of pH on germination of 0. 78 operophterae spores. 4. The effect of pH on germination of N. 80 heliothidis spores. 5. Time for completion germination of spores 80 of Nosema sp. from S. litura VO 00 l The effect of temperature on growth of N. 85 heliothidis in S. fruqiperda cell cultures. 6. The percentage of cells infected with N. heliothidis when incubated at 20°C, 25°C and 30°C. 7. The number of parasites (vegetative stages and spores) per 200 cells when cultures were incubated at 20°C/ 25°C and 30°C. 8. The number of vegetative stages per 200 cells when cultures were incubated at 20°C, 25°C and 30°C.
9-11 The effect of temperature on the growth of 88 . Nosema sp. in S. frugiperda cell cultures. 9. The percentage of cells infected with Nosema sp. when incubated at 20°C, 25°C and 30°C. 10 . The number of parasites (vegetative stages and spores) per 200 cells when cultures were incubated at 20°C, 25° and 30°C. 11. The number of vegetative stages per 200 cells when cultures were incubated at 20°C, 25°C and 30°C. 13
Page
12 - 14 The effect of temperature on the growth of 91 N. locustae in S^. fruqiperda cell cultures. 12. The percentage of cells infected with N. locustae when incubated at 20°C, 25°C and 30°C • 13. The number of parasites (vegetative stages and spores) per 200 cells when cultures were incubated at 20°C, 25°C and 30°C. 14. The number of vegetative stages per 200 cells when cultures were incubated at 20°C, 25°C and 30°C.
15 - 17 The effect of temperature on the growth of 93 V. necatrix in £>. frugiperda cell cultures. 15. The percentage of cells infected with V. necatrix when incubated at 20°C, 25°C and 30°C. 16. The number of parasites (vegetative stages and spores per 200 cells when cultures were incubated at 20°Cf 25°C and 30°C. 17. The number of vegetative stages per 200 cells when cultures were incubated at 20°C, 25°C and 30°C.
18 - 20 The effect of temperature on the growth of 96 P. operophterae in £3. frugiperda cell cultures. 18. The percentage of cells infected with P. operophterae when incubated at 20°C, 25°C and 30°C. 19. The number of parasites (vegetative stages and spores) per 200 cells when cultures were incubated at 20°C and 25°C. 20. The number of vegetative stages per 200 cells when cultures were incubated at 20 C and 25°C. 14
Page 21. The effect of penicillin and streptomycin 98 on infectivity of N. heliothidis at 25°C. 22. The effect of kanamycin on infectivity of 98 N. heliothidis at 25°C. 23. The effect of penicillin & streptomycin on 100 infectivity of Nosema sp. 24. The effect of kanamycin on infectivity of 100 Nosema sp. 25. Viability of Nosema heliothidis in vitro. 102 26. Viability of Nosema sp. i_n vitro. 102 27 Viability of V. necatrix in vitro. 104 28. Viability of N. locustae in vitro. 104 29 Viability of P. operophterae in vitro. 104 30 The effect of ATP on infectivity of Nosema 106 sp. 31. The effect of ATP on infectivity of N. 106 heliothidis. 32. The effect of Cytochalasin B on entry of N. 109 heliothidis into cells. 33. The effect of Cytochalasin B on entry of 109 red blood cells into cells of £3. frugiperda. 34. The effect of Cytochalasin B^ on N. 109 heliothidis cell invasion. 35. The effect of sugars and lectins on entry 111 of N. heliothidis into cells of £3. fruqiperda. 36. The effect of lectins and sugars on entry 111 of N . heliothidis into cells of £5. frugiperda. 37. The effect of sugars and lectins on entry 113 of N_^ locustae into cells of £3. frugiperda. 38. The effect of sugars and lectins on entry 113 of V. necatrix into cells of £3. frugiperda. 39. The effect of sugars and lectins on entry 115 of P. operophterae into £3. frugiperda cells. 15
Page 40 - 48 Development of N. heliothidis in S. 131 fruqiperda cell cultures. 40-44. Meronts and sporonts. 45. Vegetative forms in vitro. 46. An infective form attached to the surface of cell. 47-48. Numerous spores and sporoblasts.
49 - 57 Electron micrographs of N. heliothidis. 133 49. Meront with a single nucleus. 50. Meronts with 2 nuclei in diplokaryon arrangement. 51. Meronts surrounded by tubular structures. 52. Early sporont. 53. Sporont with 2 nuclei in diplokaryon 135 arrangement. 54. Sporont with complete surface coat. 55. Binucleate sporoblast. 56-57. Immature spores.
58 - 64 Development of Nosema sp. in S. frugiperda 137 cells cultures. 58-59. Meronts with diplokaryon nuclei. 60-63. Sporonts in spindle-shaped with diplokaryon nuclei. 64. A heavily infected cell with numerous spores and sporoblasts.
65 - 70 Electron micrographs of Nosema sp. from S. 139 litura. 65-66. Elongate meronts with 2 nuclei in diplokaryon arrangement. 67. Meront surrounded by host ribosomes. 68. Sporoblast with 2 nuclei in diplokaryon arrangement. 69. Sporoblast with thick coat. 70. Mature spore with 2 nuclei. 16
Page
71 - 78 Development of P. operophterae in £3. 141 fruqiperae cell cultures. 71. Uninucleate meront. 72. Uninucleate meront and division stage. 73. Large binucleate meronts. 74. Sporonts dividing within a vesicle. 75. Numerous sporoblasts. 76. Clusters of spores and pre-spore stages. 77. Infective forms closely applied to the host cell surface. 78. Numerous mature spores.
79 - 86 Electron micrographs of P. operophterae. 143 79. Meront with a single large nucleus. 80 . Meront with a single nucleus lying within a vacuolar membrane.
81. Sporont with a single large nucleus and on the surface coat thickening. 82. Two meront, each lying in its own vacuole. 83. Sporoblasts within the sporophorous 145 vesicle. 84. Spores and sporpblasts with sporophorous vesicle. 85. A heavily infected cell with several sporo phorous vesicles, containing numerous spores. 86. . Mature spore stages.
87 - 99 Light micrographs of N. locustae 147
87. A meront v:ith 2 nuclei 88. Binucleate meront. 89. Two diplokaryotic meronts. 90 Fusiform binucleate sporont. 91 Fusiform tetranucleate sporont. 17
Page
92. Fusiform sporont with diplokarya at the 147 poles. 93. Cytoplasmic cleavage of a sporont. 94. Two sporoblasts. 95. Free elongate spores. 96. Binucleate sporont of the octosporoblastic sequence. 97. Tetranucleate sporont. 98. Octonucleate sporont. 99. Eight spores within sporophorous vesicle.
100-113 Electron microqraphs of N. locustae. 149 100 . Meront with 2 nuclei in diplokaryotic arrangement. 101 . Division of meront. 102. Elongate disporoblastic sporont. 103. Disporoblastic sporont with thickened wall. 104. Division of sporont. 105. Binucleate sporoblast. 106. Mature free spore. 107. Early stage of octosporoblastic sporont. 151 108. Early octosporblastic sporont with 2 nuclei. 109. Nucleus of octosporoblastic sporont with synaptonemal complexes. 110 . Octosporoblastic sporont with 3 nuclei. 111. Octosporoblastic sporont with lobed and isolated nuclei. 112. Sporoblast with early stages of spore organelles. 113. Mature spore of octosporoblastic sequence.
114-121 Development of N. locustae in S. fruqiperda cell cultures. 153 114. Diplokaryon meronts. 115-120. Meronts and sporonts with one or two diplokarya. 121.121. Numerous spores and pre-spore stages. 18
Page 122-129 Development of V. necatrix in £3. fruqiperda 155 cell cultures. 122. Early meronts 123-127. Meronts with diplokaryon arrangement. 128. A heavily infected cell with numerous sporonts or sporoblasts. 129. Stages released from cells into the medium.
130-133 Phase contrast microscopy of N. heliothidis 157 in S. frugiperda. 130. S • fruqiperda cell culture heavily infected with N. heliothidis. 131. An infective form closely attached to the S. fruqiperda cell.
132-133 Diplokaryotic infective form free in 157 medium. 134. Heavily infected cell, disrupted and libera ting different stages of N. heliothidis. 135. Infective form closely attached to the host cell membrane.
136-141 Electron micrographs of £3. frugiperda cell 159 infected with P. operophterae labelled with saccharated iron oxide. 136. Secondary lysosome and small vesicles. 137. Vesicle labelled with saccharated iron oxide, close to a vacuole containing a vegetative stage of P. operophterae. 138. Vesicle labelled with saccharated iron oxide, close to a vacuole containing a sporogonic stage of P. operophterae. 139. A vacuole containing mature spore labelled with saccharated iron oxide. 19
Page
140 A vacuole containing vegetative stages and 159 spores but no fusion with lysosomes. 141 A vacuole containing spores and pre-spore stages but no fusion with lysosomes.
142 147 Electron micrograph of S. fruqiperda cells 16! infected with P. operophterae stained for acid phosphatase. 142 A vesicle heavily stained for acid phosphatase. 143 147. Vacuole containing spores with acid phosphatase.
148 151 Electron micrographs of N. heliothidis. 163
148 A diplokaryon stages of N. heliothidis. enclosed in a vacuole. 149 A stage of N. heliothidis lying directly in host cell cytoplasm. 150 A vesicle containing the reaction product indicating acid phosphatase close to vegetative stage. 151 A spore within a lysosome.
152. Diagram of life cycle of Amblyospora sp. 187 20
LIST OF TABLES
Table Page
1. Summary of pH requirements for germination of 23 microsporidian spores.
2. Summary of microsporidia from invertebrate 30
hosts which have been established in tissue cultures.
3. Sources of spores. -60
4. Table of buffer solutions (ph 3 to 13) 63 5. Chemicals used to investigate receptor 76
involvement in microsporidian invasion. 6. Chemical concentrations for hormones. 76 7. Cultures of microsporidia in £3. frugiperda 82
cell line by addition of haemolymph from
infected H. zea. 8. A summary of dimorphic microsporidia 189 producing spores in infected tissue. 21
1. INTRODUCTION
Microsporidia are obligate intracellular parasites which have been recognized as important factors in natural regulation of insect pest populations and as agents of diseases of fish and mammals, including man. They are characterized by spores which have an unique extrusion apparatus, composed by a spirally arranged polar tube, a lamellar polaroplast and a posterior vacuole. In response to appropriate stimuli of a host, the infective agent or sporoplasm is extruded through the polar tube and injected into a host cell directly.
After initial infection by this mechanism, stages spread from cell to cell, until the burden of infection causes serious tissue damage and mortality. However, very little is known of the mechanism of spread of microsporidia from cell to cell. The aim of this project was to establish several species of microsporidia in cell cultures and use these systems to study cell to cell transfer and investigate the possible existence of short-lived extracellular invasion stages and their relationship to host cells. 22
2. LITERATURE REVIEW
2.1 Germination of spores jLri vitro
The extrusion of the polar tube has attracted the attention of microsporidiologists for many years.
Consequently, attempts have been made to induce the discharge of the polar tube in various microsporidia and also to explain the mechanism of spore hatching.
It is known that the polar tube can be forced from the spore, when pressure is applied to the coverslip on a water-mounted preparation (Weiser & Briggs, 1970).
Extrusion can also be achieved when a suspension of spores in water is dried and then rewetted (Kramer, 1960) and when spores are exposed to (°hshima, 1927; Lom & Vavra,
1963). Several factors can be used to effect polar tube extrusion iri vitro, which can be considered to correspond more closely to the conditions of spore hatching ir\ vivo. a) pH: most species of microsporidia require a change in external pH to initiate polar tube extrusion. A two-step treatment is required for most species with a normally high pH in buffer solution followed by a lower pH buffer solution or water. A few species require only a one-step treatment.
A summary of the pH requirements for germination of several microsporidia is given in Table 1. b) Ions: metal ions have been found to play an important role in both the stimulation and inhibition of the polar tube extrusion. In his studies of the germination of N. 23
Table Is Summary of pH requirements for germination of microsporidia.
Species Pretreat Treatment Host Reference ment pH pH Nosema High 6.0 -6.5 Bombyx mori Ohshima,1964 bombycis
Nosema - 10.8 Choristoneura Ishihara, fumiferanae fumiferana 1967 Nosema 10 9.5 Anopheles Undeen, 1978 alqerae stephensi
Nosema 10.5 9.4 Pseudaletia Undeen, 1978 (=Vairimorpha) necatrix unipuncta Nosema sp. 10.5 9.4 Malacosoma Undeen, 1978 americanum
Nosema sp. 10.5 9.4 Spinx moth Undeen, 1978 Nosema 11.0 7 Heliothis zea Undeen, 1978 heliothidis
Vavraia - 6.0-6 • 5 Culiseta Undeen, 1983 culicis longiareolata
Vavraia - 7 - 9 Culiseta Undeen, 1983 culicis incidens
Gluqea 7.0 9.5 Osmerus Weidner,1982 hertwiqi eperlanus
Ameson 10 Medium 199 Callinectes Weidner,1972 michaelis (7.2) sapidus
Amblyospora 9.0 Water Culex Undeen & sp. salinarius Avery, 1984
Vairimorpha 11.5 8.5 Plodia Malone ,1984a plodiae interpunctella 24
algerae, Undeen (1978) found that K+ was the most effective in stimulating extrusion, followed by Na+ , Rb+ and Cs+ .
However, in the case of N. fumiferanae, the effectiveness of metal ions appears to decrease with decreasing atomic weight
(Ishihara, 1967). Since recognized metabolic inhibitors such as F or CN do not prevent polar tube extrusion
(Ishihara, 1967; Undeen, 1978), it is likely that polar tube extrusion is a system independent of enzymes and energy
(Dali, 1983).
Ohshima (1964) showed that Ca++ counteracted the stimulatory effect of metal ions on the hatching of N. bombycis. Ishihara (1967) and Undeen (1978, 1983, 1984) further demonstrated for N. algerae, V. culicis, Amblyospora sp. and N. fumiferanae, that not only Ca++, but Mg++, NH^ + and Li+ also inhibited spore hatching. Malone (1984) also demonstrated that Mg and Ca inhibited polar tube extrusion. Recently, Weidner & Byrd (1982) showed, by using a calcium affinity molecule, fcalcium ionophore A23187, combined with Ca-bound fluorescence, chlorotetracycline, on
G. hertwigi spores, that calcium is associated with the polaroplast membranes, and maintains the polaroplast in a contracted state, when the spore is in an inactivated condition. The displacement of calcium induces swelling of polaroplast membranes which cause polar tube extrusion.
The Ca++ on the polaroplast membranes can be removed by ionophore A23187 (Weidner & Byrd, 1982), or by the chelating agents EDTA, EGTA (Malone, 1984a), or by cations, 25
such as K+ or Na+ (Ishihara, 1967? Undeen, 1978, 1983), which thus enhance spore hatching. c) Ionic concentration: since the diffusion of metal ions is involved in spore hatching processes, it is not surprising that ionic concentration also has an effect on polar tube extrusion. Undeen (1978, 1983) found that if the ionic concentration was too low, spores of N. algerae and V. culicis became inactivated and that, after this, hatching would not occur, even if the spores were then placed in optimal salt concentrations. The KC1 concentration for inactivation was 0.012M for N. algerae and
0.025M for V. culicis. However, the spores once inactivated could be reactivated by incubation in distilled water, when it was thought that the ions which were responsible for the swollen state of the polaroplast diffused out of the spore (Undeen, 1978). The ion concentration for polar tube extrusion in most species is high. Thus, optimal KC1 concentration for N. algerae is
0.2M and at least this for V. culicis (Undeen, 1978, 1983).
Ishihara (1967) also showed that, if the K+ concentration was either too high or too low, polar tube extrusion would be inhibited. d) Temperature: increasing temperature induces greater spore hatching, but when the temperature approaches 40°C, spore hatching ceases (Ishihara, 1967). Usually, the optimal temperature for spore hatching is 30°C (Ishihara,
1967; Undeen, 1978, 1983; Malone, 1984a). 26
The intervals required for spore germination to take place after stimulation vary between species. In V. culicis, spore hatching reaches a maximum in 5 min, while in
N. fumiferanae it takes 40 min. (Ishihara, 1967? Undeen,
1983).
It has generally been accepted that the polaroplast and posterior vacuole are the organelles which contribute primarily to polar tube extrusion (Lorn & Vavra, 1963?
Weidner, 1972).
2.2 Microsporidia in tissue cultures
Microsporidia are obligate parasites and only develop and multiply in the living cells of the host. Because of their potential as biological control agents, knowledge of the parasite itself and its relationship to host cells is extremely important. However, there are gaps in our knowledge about microsporidia. The use of in vitro tissue cultures to study microsporidia, in conjunction with other techniques, could potentially contribute to research in microsporidian biology, physiology and biochemistry.
2.2.1 Techniques for the establishment of microsporidia in
tissue cultures
A number of microsporidia have been grown in tissue cultures and three basic methods have been employed.
1) Inoculation of cells already infected with microsporidia into a cell line. Trager (1937), pioneering the work on 27
tissue culture of microsporidia, was the first to achieve a culture by inoculating the haemolymph from larvae of Bombyx mori, infected with N. bombycis, into cultures of ovarian tube lining cells. However, he could not repeat these experiments and assumed that the infective forms in the haemolymph were at too low a concentration to initiate cultures. Surprisingly, since then this method has not been used for establishing microsporidia from invertebrates in tissue cultures, although the inoculation of tissues or cells from infected mammals into cell lines has been widely used in setting up cultures of Encephalitozoon cuniculi
(lino, 1960? Kaljakin & Akinshina, 1970? Vavra, Bedrnik &
Cinatl, 1972? Waller, 1975). Infected cells from established cultures can also be used as inocula to infect other cell lines, even from unrelated hosts. Tsang, Brooks
& Kurtti (1982) and Kurtti, Tsang & Brooks (1984) have used a haemocyte cell line from Malacosoma disstria persistently infected with Nosema disstriae, to infect cell lines from
Triatoma infestans and Blatella germanica, which had been adapted to the IPL-45 moth cell culture medium.
2) Primary cultures can be initiated from infected host cells or tissues. Several microsporidia have been established in primary cultures from infected organs of hosts in which the microsporidia normally complete their life cycles (Gupta, 1964? Grobov & Zuman, 1972? Vavra &
Maddox, 1976). Sohi & Wilson (1976) developed a cell line from haemocytes and ovarian tissue of Malacosoma disstria 28
larvae, which were naturally infected with N. disstriae.
The infection continued for more than 100 passages of the cells. Occasionally, microsporidia have been found as unexpected contaminants of cell cultures. Thus, Bayne,
Owczarzak & Noonan (1975) established a cell line from
Biomphalaria glabrata, which was contaminated with a
Pleistophora-like microsporidium, and a choroid plexus cell culture and kidney cell culture from a rabbit, and a primary placental cell culture from a baboon were found to be infected with E. cuniculi (Shadduck, 1969; Armstrong, Ke,
Breinig & Ople, 1973; Shadduck, Kelsoe & Helmke, 1979).
These contaminated cultures provided an opportunity for investigators to study the microsporidia.
3) Spores, harvested from infected hosts can be used to infect cell cultures and this technique has been successful in establishing several microsporidia in tissue cultures.
Ishihara & Sohi (1966) demonstrated that pretreatment of purified spores with 0.1M KOH and transfer of spores to the culture medium for hatching in the presence of cells could initiate infections: they infected an ovarian tissue culture of Bombyx mori with N. bombycis. Several other workers have applied this method to infect insect cell cultures with microsporidia from Lepidoptera (Kurtti & Brooks, 1977;
Kawarabata & Ishihara, 1984). Cell cultures, infected with
N. alqerae can be obtained by addition of sterile spores which have been previously stored in water (Undeen, 1975;
Streett, Ralph & Hink, 1980; Smith, Barker & Lai, 1982). 29
Similarly, infections can be obtained in cell cultures using spores of E. cuniculi previously stored in water or phosphate buffered saline (Vavra, Bedrnik & Cinatl, 1972).
The spores of N. eurytremae, when exposed to "conditioned medium", i.e. medium already used to culture the cells, hatched when transferred to insect cell cultures and fresh medium and established infections (Higby, Canning, Pilley &
Bush, 1979).
Microsporidian spores, isolated directly from the hosts are usually contaminated with micro-organisms, and the preparation of pure, sterile spores is a central problem in establishing cultures by inoculation of spores. Kurtti &
Brooks (1977) recovered N. disstriae spores from an organ culture of larval salivary glands taken from living M. disstria and obtained bacteria-free spores to infect insect cultures. Streett, Ralph & Hink (1980) applied a similar method to obtain sterile spores of N. algerae from
Trichoplusia ni pupae. Antibiotics and differential centrifugation were used to remove contaminating micro organisms from spores of N. eurytremae and N. algerae, and the resulting sterile spores successfully infected cell cultures (Higby, Canning, Pilley & Bush, 1979; Smith, Barker
& Lai, L982). Purification of spores of N. algerae by
Ludex density gradient centrifugation was achieved by Undeen
(1975). Kawarabata & Ishihara (1984) have obtained highly purified spores of N. bombycis by Urografin (diatrizoate) density gradient centrifugation and have then established this species in a continuous insect cell culture. 30
Table 2: Summary of a microsporidia from invertebrate hosts which have been established in tissue cultures
Species Organ culture or Inoculum Reference cell line N. bombycis Ovarian tube lining Haemolymph Trager, 1937 cells of Bombyx mori N. bombycis Ovary of Bombyx mori Spores Ishihara & Sohi, 1966. N. bombycis Primary cell culture Spores Ishihara, 1969 of mammalian and chicken embryos
N. bombycis Insect cell line of Spores Kawarabata & Antheraea eucalypti Ishihara, 1984 N. algerae Pig kidney cell line Spores Undeen, 1975
N. algerae XTC-6 and Chang Spores Smith, Barker liver cell lines & Lai, 1982
N. algerae Insect cell lines of Spores Streett, Ralph, Trichoplusia ni, & Hink, -1980 Heliothis zea, and Mamestra brassicae.
N. eurytremae Cell lines of Spores Higby, Canning, Xenopus laevis and Pilley & Bush, Aedes pseudoscutel- 1979 laris
N. disstriae H. zea cell line Spores Kurtti & Brooks, 1977 N. disstriae Malacosoma disstria * Naturally Sohi & Wilson, cell line infected 1976
N. disstriae Insect cell lines Infected Kurtti, Tsang M. disstria, H. zea, culture & Brooks, 1984 Triatoma infestans, Blatella germanica
N. mesnili Gut and fat body Infected Gupta, 1964 of P. brassicae insect
N. apis Mid-gut of honey Infected Grobov & Zuman, bee. insect 1972
V. necatrix Fat body of Infected Vavra & Maddox, P. unipuncta insect 1976
Pleistophora- Cell line of Naturally Bayne, Owczarzak like micro Biomphalaria infected & Noonan, 1975 sporidia glabrata snail 31
A summary of microsporidia from invertebrate hosts, which have been established in tissue cultures is given in Table 2.
2.2.2 Factors affecting microsporidian infectivity and
multiplication in vitro.
Among many factors, temperature is apparently of great significance to microsporidia from invertebrate hosts. None of the microsporidia from invertebrate hosts estab lished in tissue cultures so far, can survive at 37°C.
Only E. cuniculi, a parasite, which has been established in cultures of mammalian cells (Waller, 1975), has been known to survive at 37°C. N. algerae did invade cells and began development at 37°C but failed to complete its life cycle and the cultures died out within 96 h (Undeen, 1975). No cell was found infected with N. algerae or N. eurytremae at
38°C (Undeen, 1975; Smith, Barker & Lai, 1982). Ishihara
(1969) found that N. bombycis did not grow in primary cell cultures of mammals or chicken embryos at 37°C, but development took place in the cell cultures at 28°C. A number of microsporidia from insects can develop in tissue cultures at 34 - 35°C (Undeen, 1975; Smith, Barker & Lai,
1982), but Wilson & Sohi (1976) reported that there were adverse effects on the viability and growth of N. disstriae, after incubation for one week at 35°C and within 28 days the 32
parasites were eliminated from the cell cultures. These results confirm that temperature provides a natural barrier to infection of warm-blooded vertebrates with microsporidia from invertebrates.
There are only a few reports concerning the effect of temperature on growth and development of microsporidia in vitro. Wilson &.Sohi (1976) found that there were no differences in percentage infection and spore production at
20, 25, 28 and 30°C after incubating N. disstriae in vitro for 4 weeks.
The temperatures suitable for microsporidian growth in vitro accord with those investigated for growth i_n vivo.
The most favourable temperature range for development of microsporidia from invertebrates is 20-30°C. Wilson (1974) reported that increased temperature beyond 23°C appeared to have a detrimental effect on the survival of N. fumiferanae in vivo; higher spore production was obtained at 23°C compared with 28°C. During studies'on the effect of temper ature on Drosophila willistoni infected with N. kingi,
Armstrong (1976) found that 90% of the flies were infected with parasites at 22°C. It was considered that the optimal temperature for development of larvae is also the optimal temperature for growth and development of the parasites.
Antibiotics have been widely used to control micro organism contaminants of cell cultures. Some investigators have, therefore, been concerned with determining the effect of various antibiotics on microsporidian growth in cell 33
cultures. Gentamycin (50 pg/ml) or penicillin (100 u.i./ml) plus streptomycin (100 pg/ml) were found to have no obvious effects on N. disstriae or E. cuniculi (Shadduck &
Polley, 1978? Sohi & Wilson, 1979). Fumagillin blocked the merogonic development of N. disstriae but the organisms were still viable and sporoblasts were not arrested and still developed into spores. When the fumagillin was removed, the infection was re-established (Kurtti & Brooks, 1977).
Bayne, Owczarzak & Noonan (1975) found that neither Benomyl nor fumadil B eliminated the microsporidian infection from a
B. glabrata cell line. Sohi & Wilson (1979) found that treatment with Benomyl and fumagillin for 35 days eliminated
N. disstriae from cell cultures without adverse effect on the host cells. However, if treatment was discontinued after 7 days, the infection would recover to control level.
Chloroquine, an anti-malarial drug, was found to have a strong effect on E. cuniculi, but did not completely arrest the infection (Waller, 1979).
2.3 Current taxonomic position of three genera of micro-
sporidia and their developmental cycles.
2.3.1 The genus Nosema and its related genera
Nosema Naegeli, 1857, is the oldest generic name in the history of microsporidia and is the genus most rich in species. Many species, originally attributed to this genus, have subsequently been removed, as new techniques have 34
revealed differences, which differentiate them. Perez
(1905) made a clear distinction between Nosema and other microsporidia known at that time by considering that the
"sporozoite" (=sporont) of Nosema developed into a single spore and he introduced the term "monosporous" for this type of sporogony. In other genera this stage develops into two or more spores. This distinction was perpetuated by Kudo
(1924, 1966). For many years, this definition was almost universally accepted as characteristic of the genus Nosema.
Lorn & Weiser (1969) re-examined the type species of the genus Nosema, Nosema bombycis Naegeli, 1857, and found that a sporont produces two sporoblasts, each of which develop directly into a spore. Unfortunately, these authors synonymized Glugea with Nosema, on the basis that Glugea had already been defined as producing two sporoblasts from a sporont. Subsequently, using electron microscopy Cali
(1970) pointed out that in the type species of Nosema, nuclei are in diplokaryon arrangement, and Glugea was later shown to have isolated nuclei and xenoma formation (Sprague
& Vernick, 1968). Recently it has further been shown that in Glugea sporogony occurs within sporophorous vesicles and is polysporoblastic (Canning, Lorn & Nicholas, 1982). The genus Perezia, which was also thought to produce two sporoblasts from a sporont, was synonymized with Glugea by
Doflein (quoted by Lorn & Weiser, 1969) and was often confused with Nosema. Lorn & Weiser (1969) proposed that the genus Perezia should be re-established for species in 35
which the mature spores remain joined together along their long edges and lie paired within a membrane. In a recent study of the ultrastructure of Perezia lankesteriae Leger & Duboseq, 1909, the type species of the genus Perezia, it was shown that Perezia is a valid genus, which differs from both Nosema and Glugea. Nuclei are in diplokaryon arrange ment in merogony, but separate in sporonts, which divide to produce several (probably 8) uninucleate sporoblasts
(Ormi&res, Loub&s & Maurand, 1977).
Due to a simple developmental life cycle and numerous new isolates, the identification of species of Nosema has been extensively based on host specificity. Several decades ago, it was considered that a microsporidian species usually infected only a single host genus, sometimes a single species (Steinhaus, 1949), but current experiments have shown that almost all Nosema species, especially those from Lepidoptera, have a relatively wide host range and are not very host specific. It is clearly too difficult comprehensively to test all hosts, which may be susceptable to a particular Nosema species. Nordin & Maddox (1974) evaluated a number of Nosema species from Lepidoptera and failed to distinguish between at least 8 isolates, some of which had been named as separate species. Similarly a
Nosema sp. from Spodoptera litura was indistinguishable from several other Nosema species in Lepidoptera (Watanabe,
1976). 36
2.3.2 The genus Vairimorpha and other dimorphic genera Nosema necatrix Kramer, 1965 and Thelohania diazoma
Kramer, 1965 were reported as a mixed infection in adipose tissue of Pseudaletia unipuncta. It is now known that those parasites represent different phases of a single species.
There are two sporulation sequences, one of which is only expressed at low temperatures (Maddox, 1966). This was later confirmed by Fowler & Reeves (1974a). N. necatrix was re-described by Pilley (1976) and a new genus Vairimorpha was established for N. necatrix. The genus was defined as developing into free spores with diplokaryon nuclei resembling spores of Nosema at 25°C, and additionally producing uninucleate octospores resembling those of
Thelohania at 20°C. Two additional species, Nosema plodiae
Kellen & Lindegren, 1968 and Nosema heterosporum Kellen &
Lindegren, 1969 were transferred to the genus Vairimorpha by Weiser (1977) and recently more Nosema species were recognized as being dimorphic (Streett & Briggs, 1982). In an ultrastructural study of V. plodiae it was revealed that synaptonemal complexes occur in binucleate stages of the octosporoblastic sequence, implying that meiosis occurs at the beginning of this sporogony, so that 8 haploid spores are produced (Malone & Canning, 1982). As in the genus
Nosema, it is difficult to identify the species. Malone
(1984) found that V. plodiae and V. necatrix are remarkably similar in life cycle, spore size and tissue specificities.
She failed to distinguish them and suggested that further 37
critical biochemical tests are required to determine whether they are distinct species. Recently, sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) has shown that each of four species of Vairimorpha has a unique and reproducable electrophoretic profile, which was not affected by development in different host nor by development at different temperatures (Streett & Briggs, 1982). In contrast the electrophoretic profile of the hydrophobic fractions of spore proteins from Vairimorpha species had been found indistinguishable (Fowler & Reeves, 1974b).
A parasite which is similar to the genus Vairimorpha and also has temperature-dependent dimorphism, is Burenella dimorpha (Jouvenaz & Hazard, 1978). In this genus, binucleate free spores are produced from disporous sporonts in hypodermal tissue at temperatures between 20°C and 32°C, but uninucleate octospores develop from octonucleate sporonts in the fat body at 28°C. The octospores are produced only within a narrow range*of temperature: higher or lower than 28°C, the number of octospores is low, and if the temperature is above 30°C or below 20°C, octospore development is completely suppressed. The authors claimed tha the differences between Burenella and Vairimorpha lie in the different hosts and in tissue specifities, and also that the pansporoblastic membrane is persistent in Vairimorpha but not in Burenella.
Unlike the above two genera in which dimorphism is temperature-dependent, other genera exhibit dimorphism in 38
different ways. Species of Amblyospora and Parathelohania, both parasitizing mosquitoes, show sex-dependent dimorphism, which again involves two different sporogonic sequences? the binucleate free spores are found in adult females, whereas the haploid octospores with single nuclei occur in male larvae. It has been shown that the binucleate diploid spores function, in transovarial transmission and that the uninucleate haploid spores are not perorally infectious to the mosquito hosts. In an Amblyospora sp. from Culex annulirostria the octospores have been shown to infect a copepod intermediate host, Mesocyclops albicans, in which they develop into a third type of spore, infective to mosquito larvae (Sweeney, Hazard & Graham, 1985). This is the first demonstration of an alternation of hosts in the microsporidia but it is likely that many dimorphic species will have similar cycles of development.
Stempellia was probably the first genus to have been recognized as dimorphic (Ldger & Hesse, 1910) and this has been confirmed by electron microscopy (Desportes, 1976). In the type species Stempellia mutebilis, there are two sequences: 8 microspores are produced from a small sporont after 3 divisions and 4 macrospores are produced from a large sporont after 2 divisions.
Another dimorphic species, Culicosporella lunata was originally placed in the genus Stempellia (Hazard & Savage,
1970). More recent studies have revealed two sporogonic sequences. One sporogonic sequence begins with 39
diplokaryotic meronts, in which there are repeated nuclear division to give sporogonial plasmodia with nuclei in diplokaryon arrangement: these divide to give binucleate spores, which are perorally infectious to new hosts. The second sporogonic sequence also begins with diplokaryotic meronts but these undergo karyogamy, then meiosis and finally develop.into octonucleate sporonts which give rise to 8 haploid spores (Hazard, Fukuda & Becnel, 1984).
Hazardia milleri was also originally placed in the genus Stempellia, when described by Hazard & Fukuda (1974). It was subsequently placed in a new genus, Hazardia by
Weiser (1977). The parasite produces two types of spores in fat body tissue: thick-walled binucleate spores are produced from disporous diplokaryotic sporonts and thin-walled, uninucleate spores are produced from multinucleate sporonts with isolated nuclei. The authors assumed that in diplokaryotic schizonts the nuclei fuse to form uninucleate sporonts which become multinucleate'sporonts after nuclear division.
2.3.3 The genus Pleistophora and related genera
The genus Pleistophora was described by Gurley (1893) and was distinguished from other pansporoblastic micro- sporidia by its sporulation sequence, in which more than 8 spores are produced within a pansporoblastic membrane. Since then, most microsporidia with a characteristic pansporoblastic membrane enclosing a variable and large 40
number of spores have been included in the genus
Pleistophora. Recent ultrastructural studies have shown that there are fundamental differences in the nuclear arrangement, development cycle and mode of formation of the pansporoblastic membrane between species attributed to the genus Pleistophora. This indicated that Pleistophora is a heterogenous assemblage of species rather than a well- defined genus (Canning & Hazard, 1982). The type species of
Pleistophora, P. typicalis Gurley, 1893, was re-examined at light and electron microscopic levels (Canning & Nicholas,
1980) and a more precise definition of the genus was provided. The major finding was that the nuclei are iso lated in all development stages and that there is a thick amorphous coat secreted on to the plasmalemma which surrounds all stages of the parasite. For comparison with
P. typicalis, Canning & Hazard (1982) studied Vavraia culicis, which had already been transferred from the genus Pleistophora by Weiser (1977). Canning & Hazard (1982) found that Vavraia closely resembled Pleistophora typicalis in having isolated nuclei throughout its developmental cycle and a persistent amorphous coat during merogony. They upheld Weiser's (1977) separation of Vavraia but used the mode of division of the sporogonial plasmodium and the structure of the sporophorous vesicle wall (= pansporo blastic membrane) as diagnostic characters. In the same paper, Canning & Hazard found that Pleistophora simulii
(Lutz & Splendore, 1907) did not belong either to 41
Pleistophora or to Vavraia and they proposed a new genus Polydispyrenia for this species. In Polydispyrenia vegetative and sporogonial plasmodia contain diplokaryon nuclei and meiosis occurs at the beginning of sporogony, resulting in haploid spores, which are surrounded by a fine cyst wall (= pansporoblastic membrane). Yet another species formerly assigned to the genus Pleistophora, P. debaisieuxi had already been transferred to the genus Tuzetia by Loubes
& Maurand (1976). They found that, following the division of the sporogonial plasmodia, each uninucleate sporoblast was encased in an individual pansporoblastic membrane. More recently, based on light and electron microscopy, a new genus Janacekia was proposed for debaisieuxi, which differs from Tuzetia in that meronts and sporonts are diplokaryotic. In Tuzetia all stages have isolated nuclei
(Larsson, 1983).
Pleistophora sp., a parasite from a marine polycheate egg, Armandia brevis, was described by Szollosi (1971).
This species in which the nuclei are in diplokaryon arrangement throughout sporogony, was later transferred to a new genus Pseudopieistophora as P. szollosii by Sprague
(1977).
Recently, a parasite from larch sawfly larvae,
Pristiphora erichsonii, was tentatively identified as
Pleistophora sp. (Percy, Wilson & Burke, 1982). In this species, the nuclei are in diplokaryon arrangement in the meronts, but are isolated in sporonts and spores. All 42
stages were observed within parasitophorous vesicles which the authors thought had originated from the host cells. The authors realised that the characteristics of the species from P. erichsonii differed from those of the type species of the genus Pleistophora and that a new generic assignment should be made.
Although more than 10 species of Pleistophora have been recorded from Lepidoptera, only two have been observed at electron microscopic level; one is tissue specific and the other not. A subspecies of P. schubergi, a species which has a wide range of hosts and only infects midgut epithelium was examined by Purrini (1982): 2 morphologically different schizonies (merogonies) were recognized. One generation produced multinucleate plasmodia and the other gave rise to large "ribbon-like" stages. Sporogony was polysporous, but no details were given to distinguish this species within the
Pleistophora complex.
Lastly Pleistophora operophterae has been studied by
Canning, Barker, Nicholas & Page (1985), who established a new genus Cystosporogenes for it. In this genus all stages have isolated nuclei, but meronts and sporonts lie within a membrane of enigmatic origin. Sporogony is completed within this membrane to give rise to numerous uninucleate spores.
2.4 Interaction of parasitic protozoa with their host
cells
As a consequence of advances in cell biology and 43
biochemistry, knowledge of the interaction of parasitic protozoa and their host cells has increased rapidly. Here, only those aspects of the interaction, which are considered to be relevant to invasion of microsporidia into host cells, are discussed.
The main events which are involved in interaction of protozoan parasites with their host cells are attachment and invasion into the host cells and survival or otherwise once inside the host cells.
2.4.1 Invasion into the host cells
2.4.1.1 Microsporidia
Microsporidia form small resistant spores with a remarkable, highly specialized, internal structure, which is shown in Fig. 1. The spores are typically ovoid or ellipsoid, generally less than 10 pm long and 5 pm in diameter and possess a three layered wall. There is an outer proteinaceous exospore, a middle chitinous endospore and an internal plasma membrane. Within the wall there is a mass of undifferentiated cytoplasm containing one or two nuclei and ribosomes. There is also an extrusion apparatus, consisting of polar cap and polar sac, polaroplast, polar tube and posterior vacuole. These specialized organelles play a role in the infection process. Although the mechanism for extrusion is still not quite clear. Current studies in vivo or in vitro have uncovered some details of the extrusion process. Under appropriate stimuli, usually 44
Fig. Is Diagram of fine structure of microsporidian
spores
AD = anchoring disc
CM = cytoplasmic membrane
EN = endospore
MNB = manubroid
N = nuclei
PB = posterior body (or posterior vacuole)
PL = polaroplast
PA = polar aperture
PF = polar filament (polar tube) Ex = exospore 45
in the gut lumen of new hosts, the spores are activated and the polar cap, which is believed to act as a permeability
barrier, undergoes a change in structure (Weidner & Byrd,
1972). This is followed by swelling of polaroplast membranes, activated by displacement of internal calcium
(Weidner, 1982). Pressure builds up inside the spore,
causing discharge of the polar tube through the polar cap.
The polar tube is now thought to re-assemble from protein
units during extrusion (Weidner & Byrd, 1982; Weidner,
1982). The sporoplasm then passes through the polar tube
and is inoculated into host cell. The polaroplast membrane
becomes the plasma membrane of the sporoplasm, while the
spore plasma membrane remains in the empty spore (Weidner et
al. , 1984). Spores do not generally hatch within the host
in which they are produced, so this method of infection does
not account for cell to cell spread within hosts. Little
information on this process is available in the literature.
(a) In vivo studies of cell infection
Early authors thought that so-called planonts were
responsible for the transmission of infection of N. bombycis
(Stempell, 1909). Some recent studies have suggested that,
for some species of microsporidia, infections are spread within the host by the transport of meronts in haemocytes, which have phagocytic activity, or simply in the haemolymph. In their studies of N. bombycis in silkworms B. mori and V. necatrix in Barathra brassicae, Weiser (1976) and Abe (1978) 46
proposed that, when parasites are released from disrupted cells into haemolymph, they circulate in haemolypmh and are eventually phagocytosed by the target tissues. Laigo &
Paschke (1966) reported that meronts of Nosema sp. in
Trichoplusia ni are released from infected cells and are taken up by "free circulating" haemocytes, mainly plasma- tocytes; the haemocytes were thought to spread the parasitic infection inside the body. A similar observation was made on the coccinellid beetle, Coccinella septempunctata infected with Nosema tracheaphila (Cali & Briggs, 1967). The presence of infective stages in haemocytes or haemolymph was confirmed by Ohshima (1975) who infected healthy silkworm larvae by injection of haemolymph from infected larvae. The haemolymph was known to contain only meront stages. Hazard,
Fukuda & Becnel (1984) found that meronts of Culicosporella lunata undergo repeated divisions in haemocytes and this causes them to burst and release meronts into the haemo lymph. The parasites migrate to and invade fat body cells.
A similar process was found to occur in Culex pilosus infected with Hazardia milleri (Hazard & Fukuda, 1974).
When parasites are restricted to the midgut cells, spread of infection must occur in a different way. Abe &
Fujiwara (1979) showed that a Pleistophora sp. multiplied in the anterior part of the midgut of silkworm larvae. After division into binucleate forms, stages which were presumptive secondary infective forms, were extruded into 47
the gut lumen and these survived and migrated to invade other cells. A similar phenomenon was observed at the electron microscopic level in larvae of Choristoneura fumiferana, infected with N. fumiferanae (Nolan & Clovis,
1984).
Some authors have provided evidence that in some species of microsporidia, spores can hatch intracellularly.
The germination of mature spores of N. algerae was observed in the midgut of mosquito larvae, Anopheles albimanus, 72 h after infection (Avery & Anthony, 1983). These authors concluded that, either sporoplasms extruded from spores or sporonts from heavily infected cells, were responsible for secondary infection.
In some species of microsporidia there may be different modes of spread of infection according to the stages of development. Thus in Amblyospora sp., a microsporidian parasite of mosquitoes, there are different developmental cycles in male and female hosts. The earliest stages in male larvae develop in haemocytes. After multiplication of the parasites the haemocytes rupture and release numerous diplokaryotic meronts into the haemolymph. These meronts were believed to migrate and invade adipose tissue. In female hosts, the infection is restricted to oenocytes until sporulation is initiated, when the adult females take a blood meal. After sporulation, the spores extrude their sporoplasms into the ovaries and infective stages enter eggs to transmit the infection to the next generation (Andreadis & Hall, 1979; Andreadis, 1983; Hazard, 1984). 48
(b) In vitro studies of cell infection
Cultures of microsporidia jji vitro offer an easier means of studying the mechanism of transfer of the parasites from cell to cell. The results from several studies suggest that extracellular infective forms are responsible for the spread of infection from cell to cell. Trager (1937), who obtained an infection of cultured cells by inoculating haemolymph from silkworm larvae infected with N. bombycis, believed that initiation of infection was by means of extracellular forms. This suggestion was supported by
Ishihara (1969), who observed that binucleate parasites resembling sporoplasms leave the host cells, migrate to and invade other cells. He called these secondary infective forms. Based on Ishihara's findings and their own studies of N. disstriae in M. disstria cell line, Sohi & Wilson
(1976) proposed that there could be two possible mechanisms of spread of infection of Nosema in cell cultures:
(a) They suggested that since the Nosema spores require a medium of high pH for germination, only the vegetative stages could be infective forms. They observed the release of such stages from disintegrating, heavily infected cells together with other stages of N. disstriae.
The infective stages were thought to invade a new cell after random contact with the cell surface.
(b) Transmission of infection by division of infected cells. They showed that cells infected with N. disstriae could undergo mitosis. This has also been found by Grobov & 49
Zuman (1972) in honey bee tissue cultures infected with N. apis and by Kurtti & Brooks (1977) in H. zea cell line infected with N. disstriae. Kurtti & Brooks (1977) also observed that there were extracellular parasites of N. disstriae, similar to the secondary infective forms of N. bombycis. However, they failed to determine whether these were expelled by, or migrated from the host cells.
Recently, Kawarabata & Ishihara (1984) have shown that there were extracellular infective forms of N. bombycis in an Antheraea eucalypti cell line at 42 h postinfection.
During the stages of sporoblast formation, stages were released into the medium without disruption of the host cell. They suggested that spread of infection of N. bombycis between cells was neither by germination of mature spores nor by division of infected cells but by extra cellular infective forms. Kurtti, Tsang & Brooks (1984) also came to the same conclusion, that infective stages rather than spores are involved in the spread of infection by N. disstriae in insect cell lines. However, they found that vegetative stages and immature spores were enclosed in vesicles within host cell cytoplasm and they thought that these vesicles extruded from the infected cells by exo- cytosis. They did not determine whether the membranes of the vesicles, containing the infective forms, were able to fuse with the membranes of uninfected cells nor did they determine whether the spores could be taken up by phago cytosis, and hatch during fusion of phagosome and lysosome 50
within the cell. Tsang, Brooks & Kurtti (1982) provided strong evidence for spread of infection by extracellular infective forms. They demonstrated that N. disstriae was unable to infect Blatella germanica cells when these were grown in cockroach cell medium, but that B. germanica cells became infected with N. disstriae when the cell line was adapted to grow in moth cell medium. They pointed out that condition in the culture medium, such as osmotic pressure and ion concentration may be important to the survival of extracellular infective forms.
With E. cuniculi it was found the polar tubes extruded from extracellular spores and from intracellular spores which were thought to have been phagocytosed (Pakes,
Shadduck & Cali, 1975). These authors proposed that sporoplasms of JE. cuniculi could initiate infection either when spores are taken up by phagocytosis or by injection.
However, it has been demonstrated that the sporoplasms of
Arneson michaelis in vitro require extraneous energy in the form of ATP; and other metabolites in the medium to retain their integrity (Weidner & Trager, 1973). Sporoplasm of N. al^ erae, which were allowed to germinate in the culture medium before addition of the medium to cells in culture, were unable to infect the cells (Undeen, 1975). Streett,
Ralph & Hink (1980) observed neither hatching of spores of
N. algerae nor extracellular infective forms of N. algerae in culture medium. 51
2.4.1.2 Other protozoan parasites In the spread of many intracellular parasites in mammalian hosts, it is assumed that heavily infected cells break down and release their parasites, some of which then enter into other cells. Successful invasion of mammalian cells by parasitic protozoa is generally considered to involve at least two steps; a) attachment to the host cell membrane, via some kind of specific recognition-receptor and ligand binding and b) enclosure of the organism in a membrane derived from the host cell.
a) Attachment
It is known that interaction between host cells and parasitic protozoa involves receptor and ligand binding.
Alternation in the host structure can be demonstrated by pretreatment with some substances, after which the attachment of protozoa is either inhibited or increased.
The presence of immunological receptors, such as the Fc receptor can be demonstrated by blocking the receptor site with either polyclonal antiserum, raised against the cell type, or more specific monoclonal antibodies to the Fc receptor. This has been demonstrated in Trypanosoma cruzi attachment to mouse peritoneal macrophages, since opsoniz ation with hyperimmune mouse serum enhanced the uptake of blood trypomastigotes but the parasites proliferated inside the macrophages (Nogueira, Chaplan & Cohn, 1980). Alter natively immune sera obtained from rabbits infected with T 52
brucei promoted attachment and subsequent ingestion of the parasite by rabbit peritoneal cells (Cook, 1981). Ward and
Jack (1980) demonstrated the presence of C^b receptors, either on the erythrocyte membrane or on Babesia rodhaini merozoites. These receptors were thought to play a critical role in facilitating the entry of B. rodhaini into red cells, but this has not been shown for other Babesia spp. (Callow & Dalgliesh, 1982). Alternatively, the interaction of protozoan parasites and host cells may involve non-immunological ligands and specific receptors, which may be by means of lectin-like glycoprotein receptors binding to specific carbohydrate residues. Dwyer (1974) was the first to report the existence of specific terminal saccharide residues on the surface of a parasitic protozoa (promastigotes of
Leishmania donovani). Since then lectins binding specific — i sugars have been extensively employed in investigation of receptors on the surface of a variety of other parasitic protozoa. These can be shown either by competitive binding of the lectin-like ligand with the specific sugars, or by competitive binding of the receptor sugar moiety with the lectins.
Different protozoa display different lectin binding characteristics. Dwyer (1977) found that promastigQtes of
Leishmania donovani were specifically agglutinated by the plant lectin concanavalin A (Con A), soybean agglutinin
(SBA), wheat germ agglutinin (WGA), fucose binding protein 53
and phytohaemagglutinin-M. However, Dawidawicz, Hernandex
& Infente (1973) demonstrated that promastigotes of L. braziliensis were only agglutinated by Con A and Ricinus communis agglutinin (RCA) and not by SBA, WGA and phytohaemagglutinin-p.
The ability to bind different lectins may even be useful in characterization of parasite strains; this has been shown in strains of T. cruzi by Arauju, Handman &
Remington (1980).
The complex life cycles of parasitic protozoa require that parasites recognize different cell types. The carbohydrates on protozoan cell surfaces may be stage specific. Amastigotes of L. donovani were shown to bind wheat germ agglutinin (WGA) but not peanut agglutinin (PNA).
However, during the process of conversion from amastigotes to promastigotes, the WGA binding receptor was lost, and the
PNA binding residue was newly displayed (Wilson & Pearson,
1984). In some strains of T. cruziit has been found that avirulent epimastigotes obtained from cultures, but not highly virulent trypomastigotes from blood, agglutinate with
Con A. (Alves & Colli, 1974; Arauju, Handman & Remington,
1980).
The involvement of glycoproteins, such as glycophorin, in attachment of Plasmodium falciparum merozoites to red cells has been reported (Weiss, Oppenheim & Vanderberg 1981;
Pasvol & Jungery, 1983). It was found that there was significant inhibition of invasion of P. falciparum into red 54
cells by N-acetyl-D-glucosamine, N-acetyl-galactosamine and
N-acetyl-neuraminic acid. This also has been demonstrated in the binding of promastigotes of L. mexicana mexicana to macrophages: it has been found that attachment to mouse macrophages, in the absence of serum in vitro, is governed by a wheat germ agglutinin-like ligand on the surface of the promastigotes and that this is linked to an N-acetyl- glucosamine moiety, which is part of the receptor site on the macrophages. (Bray, 1983).
b) Entry of parasite into cells Once attached to their host cells the so-called zippering mechanism hypothesis postulated that parasitic protozoa became internalized, by the sequential and circumferential interaction between host cell receptors and parasite-ligands (Silverstein, 1977). According to this hypothesis, entry of parasites into phagocytes is an energy-dependent process, for which energy is provided by the hydrolysis of ATP in the binding of actin and myosin microfilaments in the phagocytes, without any assistance from the parasites. Cytochalasins, a class of fungal metabolites, affect a wide range of motile functions of eukaryotic cells. One of these is the reversible suppression of phagocytosis, by inhibition of the action of microfilaments. A number of workers have speculated that the inhibition of entry of parasites into phagocytes by cytochalasins implicates phagocytosis as the major means of 55
uptake of parasites by the phagocyte. (Alexander, 1975;
Nogueira & Cohn 1976; Ryning & Remington 1978; McCabe,
Remington & Arauju, 1984). Bray (1983) also demonstrated
that uptake was not affected by glutaraldehyde treatment of
promastigotes, but was proportionally reduced by treatment
of macrophage with increasing concentration of glutaralde
hyde.
Another model for invasion of erythrocytes by malarial
parasites, proposed by Aikawa, Miller, Johnson & Rabbage (1978), is the so-called attachment-detachment or modified
zipper model. These workers suggested that new junctions
are formed between host cell and parasite which move back along the parasite surface, as the previous junctions
separate. Russell (1983) proposed another model for entry
of parasites into cells, based on contractile elements in
the parasite surface, which can also be inhibited by
cytochalasins. Russell suggested that the "capping mechanism", which he proposed, might apply to all
Apicomplexa. The parasites, which possess a locomotory
system, are capable of capping the junction between parasite and host cell down their bodies. This is an active process of entry by the parasite not involving host cell energy.
(Turner & Gregson 1982; Russell, 1983).
2.4.2 Fate of parasitic protozoa in host cells
Once the parasites have been internalized, they take in with them the invaginated cell membrane which forms a 56
parasitophorous vacuole, of which the membrane is derived from the plasma membrane of the host cell. Parasitic protozoa have evolved a variety of mechanisms to enable them to survive within host cells.
Trypomastigotes of T. cruzi, which are able to survive in macrophages and other cell types, avoid destruction within host cells by escape from the parasitophorous vacuoles. Phagosome-lysosome fusion is thus prevented and the parasites multiply in the host cell cytoplasm (Kress et al. , 1975? Nogueira & Cohn, 1976? Grimaud & Andrade, 1984). It was also shown that zoites and meronts of Sarcocystis, prior to tissue cyst formation in the intermediate host, escape from phagocytic vacuole and lie free within host cell cytoplasm (Dubey, Speer & Douglas, 1980).
It has been suggested that the mechanism of parasite escape from the parasitophorous vacuole (PV) into the cytoplasm is possibly by lysis of the PV membrane by some factors released from the parasite. (Kress et. al., 1975?
Nogueria & Cohn, 1976).
Toxoplasma gondii has a different mechanism for survival in host cells. The parasite remained within the vacuole, but the membrane of the vacuole appears to be altered, so that phagosome-lysosome fusion cannot take place. Host mitochondria and endoplasmic reticulum surround the vacuole closely. However, if Toxoplasma zoites are coated with antibody prior to being engulfed by macrophages, the lysosomes are able to fuse with the phagosome and the 57
parasites are digested, after being killed by phagosome acidification and the "oxidative burst". It has further been demonstrated that Toxoplasma zoites, which are highly susceptible to acidic pH conditions, are capable of inhibiting phagosome acidification by blocking the fusion of the acid delivery-vesicles (Sibley, Weidner, & Krahebuhl,
1985).
Prevention of lysosomal fusion with parasitophorous vacuoles had also been shown in macrophages infected with the microsporidium, Encephalitozoon cuniculi. If the vacuole contains vegetative stages or spores which have developed from vegetative stages, which have entered the macrophage, lysosomal fusion is prevented. However, if hatching spores, are phagocytosed by macrophages they can be digested (Weidner, 1975).
Another way in which the parasites can survive in an unfavourable cell environment is demonstrated by Leishmania.
Amastigotes survive within phagolysosomes apparently without major damage (Alexander & Vickerman, 1975? Chang & Dwyer,
1978; Brazile, 1984). The mechanism of survival in phagolysosomes is still obscure. Recently, the very high proteinase activity in L. m. mexicana amastigotes was found and it is suggested that the enzymes may play a major role in survival and growth of the parasite within host macrophage (Coombs, 1982). Excretion of amines, such as ammonia and urea into the parasitophorous vacuole by
Leishmania promastigotes and amastigotes might also play a 58
part in protecting the parasite against the microbiocidal activity of the host. (Coombs & Sanderson, 1985).
Alternatively, the survival and growth of amastigotes of Leishmania within the acid environment of phagolysosomes may also account for their metabolic function activating at high
H+ concentration (pH 4.0-5.5) (Mukkada, Meade, Glaser & Bonventre, 1985). 59
3. MATERIALS AND METHODS
3.1 Microsporidian species
The species of microsporidia and the hosts from which they were derived, the geographical locations and the
investigators supplying them are given in Table 3. The microsporidia were made available as purified suspensions
of spores in water or were harvested from cadavers of
infected hosts.
3.2 Infection of insects
Three insect hosts, the cabbage moth, Mamestra
brassicae, the corn earworm, Heliothis zea and the tobacco budworm, H. virescens, were used in this study. Eggs of
M. brassicae and H. zea were kindly supplied by Dr D.C.
Kelly, NERC, Institute of Virology, Oxford, England; and additionally by Dr R. Gordon, Department of Pure and
Applied Biology, Imperial College at Silwood Park, Ascot,
Berks, England. The latter had been sent originally as pupae from the U.S.A. Eggs were received attached to pieces of muslin and were left to hatch on moistened paper
towels in sealed plastic boxes. After hatching, the
larvae were maintained individually in 30 ml plastic cups with 4-6 ml of semi-synthetic diet (Appendix 1), and kept
at a constant temperature of 25°C.
Spores were purified as given in Section 3.3. From a 6 7 spore suspension, containing 10-10 spores/ml, a 50 pi Table 3: Sources of spores
Species Host(s) Location Supplier
Nosema heliothidis Heliothis zea U.S.A. Dr.W.M.Brooks
Nosema locustae Melanoplus spp. U.S.A Dr.J.E.Henry
Nosema sp. Spodoptera litura China Collected by author
Vairimorpha necatrix unknown U.S.A. Dr.J.V.Maddox
Orthosoma operophterae Operophtera brumata U.K. Prof.E.U.Canning
Pleistophora operophterae * n U.K. Prof.E.U.Canniing
Pleistophora shuberqi unknown Czechoslovakia Dr.J.Weiser
Glugea anomala Gasterosteus aculeatus U.K. Collected by author
Amblyospora sp. Simulium ornatum U.K. Collected by author Nosema bombycis Bombyx mori Japan & China Dr.R.Ishihara and
Mr.Z.Wanq.
* Propagated in Mamestra brassicae and H. zea 61
volume was dispensed by an automatic pipette onto the
surface of a 3 mm 3 piece of semi-synthetic diet and allowed to dry* The 2nd or 3rd instar larvae were starved
for 12 h and introduced into 5 ml disposable plastic tubes which contained a piece of spore-contaminated diet. The
larvae were allowed to feed until the whole piece of diet had been consumed and they were then transferred to cups containing a 5 mm layer of fresh spore-free diet. Cups were incubated at 25°C or 20°C. At various intervals post-treatment, larvae were removed, their tissues
individually smeared on glass slides and 10% (v/v) Giemsa
stained, until 25 days had passed.
3.3 Collection and purification of spores from infected
insects
Infected larvae, were collected 15-20 days after
inoculation. After rinsing with distilled water, the
cadavers were titrated in an equal volume of distilled water in a glass tissue grinder. The homogenate was
filtered through 4 layers of muslin several times until
all large pieces of debris were removed. The filtrate was centrifuged at 1000 g for 10 min to sediment the spores,
the supernatant was discarded, and the sediment was
resuspended in distilled water and centrifuged again.
This procedure was repeated at least twice.
The final spore suspension was stored at 4°C in antibiotic solution, containing 100 i.u/ml penicillin, 100 jjg/ml streptomycin and 50 pg/ml natamycin. 62
Although these methods gave adequate purification for general use, further purification was needed to provide a sterile spore suspension for infection of tissue cultures.
Discontinuous gradient centrifugation was used for this purpose. Stock Percoll (Pharmacia Fine Chemicals) was diluted with sterile distilled water to provide a series of different densities of 20%, 40%, 60%, 80% and 100%
Percoll. The Percoll solutions were carefully layered into a 20 ml centrifuge tube so that the 100% layer was at the bottom and the 20% layer was at top. The semi- purified spore preparation was carefully layered onto the top of the gradient which was then centrifuged at 1500 g for 30 min. The spores settled at 80%-100% boundary. The highly purified spores were removed and stored at 4°C in antibiotic solution as described above. The purity of the spores was ascertained by examination under a phase contrast microscope.
3.4 Induction of spore hatching
A series of buffer solutions of different pH values were made up as given in Table 4.
For pre-treatment at one pH value, 0.2 ml aliquots of 8 purified spore suspension, containing 3-5 x 10 spores per ml were added to 1 ml of each buffer solution in a 1.5 ml plastic tube. After shaking with a whirlimixer, the solutions were maintained at 30°C for 30 min. The pre-treated spore suspensions were centrifuged at 800 g \
Table 4: Table of buffer solutions (pH 3 to 13)
pH 3 4 5 6 7 8 9 9.5 10 10.5 11 12 13
Chemicals (ml)* Na2HP04 20.5 38.6 51.5 63.2 82.5 97.3
Citrate 79.5 61.4 49.5 36.8 17.8 2.7
Glycine 82.6 72.8 61.7 53.7 59 UJ
NaOH 12.4 27.2 38.3 46.2 50
NaOH ** * 6.0 36
NaCl______25 25
* all buffer solutions at 0.02M concentration solution to which was added 1M KC1
** diluted to 100 ml with H20 64
for 2 min. Two-thirds of the supernatant were removed, the remainder was agitated gently with the spore pellet, and 0.2 ml samples were added to buffer solutions of another pH value, or to TC-100 tissue culture medium. The solutions were incubated at 30°C for 60 min. Drops of the solution were examined by phase contrast microscopy.
Hatched spores were recognized by their phase dark appear ance and usually also by the presence of the extruded polar tube. The percentage of hatched spores was assessed by counting 200 spores from each sample, and there were 3 replicates for each sample.
3.5. Maintenance of the insect cell line. The insect cell line was kindly supplied by Dr D.C.
Kelly, NERC, Institute of Virology, Oxford, England. It was originally derived from pupal ovaries of the fall armyworm, Spodoptera frugiperda. The cells were grown . 2 either m 25 cm disposable plastic tissue culture flasks
(GIBCO Ltd) containing 5 ml TC-100 medium (GIBCO Ltd) or in 400 ml flat bottles containing 50 ml of medium. The
TC-100 medium contained 100 i.u./ml penicillin, 100 pg/ml streptomycin and 100 pg/ml kanamycin, and was supplemented with 10% heat-inactivated foetal calf serum (F.C.S.).
Cultures were incubated at 25°C and were subcultured every
5-7 days. To subculture the monolayer, cells were removed from the surface of the culture flask with a rubber scraper and the medium containing the free cells was 65
centrifuged in universal tubes at 750 g for 5 min. The supernatant was discarded and 5 ml of fresh medium was added to the tubes. The cells were stirred in the whirlimixer and transferred to fresh flasks with a sterile pipette.
3.6 Infection of cell cultures 3.6.1 Inoculation of cell cultures with spores
Infection of cultures by direct addition of highly purified spores was attempted using spores of Orthosoma operophterae, Nosema sp. from S3, litura and Glugea 7 anomala. 2.5 - 5.0 x 10 spores/ml were pre-treated in buffer solution at the required pH value for 30 min and one or two drops were added to a confluent monolayer of 53. frugiperda cells grown on a 7mm coverslip in a 5 ml flat bottom plastic tube containing 1 ml of TC-100 medium. The tubes were centrifuged at 500 g for 3 min, then incubated at 20°C or 25°C.
The spores of Nosema sp. from 53. litura were pre-treated with buffer solution at pH 12 and the spores of O. operophterae and G. anomala were pre-treated with buffer solution at pH 9.5.
3.6.2 Inoculation of cell cultures with the haemolymph
from infected insects.
Fifth instar larvae with 10-15 days infection of the microsporidia were surface-sterilized by immersing them in
0.5% Hyamine dissolved in 70% alcohol for 5 min. They 66
were then placed on sterile filter paper in sterile petri dishes to dry. Each larva was lifted out and carefully held on a sterile surface with the aid of the thumb and middle finger. One leg or proleg was cut with sterile scissors and the haemolymph was allowed to drip into a sterile 5 ml plastic tube containing a coverslip covered with a monolayer of £5. frugiperda cells. The tube contained 1 ml of TC-100 medium, plus 2 mg/ml phenylthio- urea which inhibit melanization. The tube was centrifuged at 750 g for 5 min to allow the haemocytes to settle and attach to the S. frugiperda cells on the coverslip. The cultures were incubated at 25°C for 24 h and then the coverslip was transferred to a 25 cm plastic flask containing 5 ml fresh tissue culture medium. The cultures were incubated at 25°C or 20°C. The medium was changed weekly and the cultures were examined weekly. Contaminat ed and uninfected cultures were discarded. Infection of cultures with Nosema sp. from £3. litura, Nosema helio- t M d i s , Nosema locustae, Vairimorpha necatrix and
Pleistophora operophterae were achieved.
3.7 Maintenance of microsporidia in cultures of £3. frugiperda cells.
Infected cells were maintained in 25 cm disposable plastic flasks containing 5 ml TC-100 medium at 20°C or
25°C as in Section 3.5. For subculture, one part of heavily infected cells was mixed with 3-5 parts of 67
uninfected cells. The medium was changed weekly when the cultures became heavily infected and only a few healthy cells remained. The uninfected cells were supplemented to . . 5 maintain a concentration of viable cells at 4 x 10 cells/ml.
3.8. The effect of temperature on development and
multiplication of microsporidia in cultures.
The effects of temperature on the growth character
istic^ in culture of Nosema sp., N. heliothidis, N. locustae, V. necatrix and P. operophterae were examined.
In these experiments, one part of infected cells was mixed with 5 parts of uninfected cells. The final levels of
infection ranged from 10 to 15% of cells infected in different experiments. 1.5 ml of the mixture containing 5 2-3 x 10 cells/ml was placed into 5 ml flat-bottomed plastic tubes each containing a 7 mm round coverslip.
Fifteen cultures were incubated at each of 4 temperatures, 20°, 25°, 30° or 37°C, respectively and the medium was renewed after 4 days.
The effects of temperature were assessed by counting
(a) the number of cells infected, and (b) the number of parasites in 200 cells. Every 2 days, 3 coverslips from each temperature group were removed, air dried, fixed in methanol and stained in 10% (v/v) Giemsa. 68
3.9 The effect of antibiotics and ATP on the growth of
Nosema sp. and Nosema heliothidis
Penicillin at 100 and 500 i.u./ml, streptomycin at
100 and 500 pg/ml and kanamycin at 100 and 500 pg/ml, in TC-100 medium were added to cultures. Adenosine tripho sphate (ATP) was added to provide a concentration of 2mM and 4mM in culture medium, without any added antibiotics.
No antibiotics or ATP were added to control cultures. The effects were assessed by counting the number of infected cells in 200 cells, and there were 3 replicates for each treatment.
3.10 Isolation of pre-spore stages (meronts and sporonts)
Several methods were used in attempts to prepare viable vegetative stages, free of their host cells, of
Nosema sp., N. heliothidis, N. locustae, V. necatrix and
P. operophterae.
(1) Immuno-lysis 2 ml of suspension of uninfected S. frugiperda cells, at 1 x 10 ^ cells/ml, was inoculated intravenously into a rabbit. This was followed by a similar inoculation 2 weeks later and 2 further inoculations of 2 ml containing 5 5 x 10 cells/ml at intervals of 2 weeks. The rabbit was bled 10 days after the final inoculation, the blood was allowed to clot, then centrifuged at 1100 g. The cells were discarded and the serum stored at -20°C. 69
S. frugiperda cells infected with N_^ heliothidis were centrifuged at 20°C for 5 min at 100 g then resuspended in
3 times their volume of fresh medium. 0.7ml of the anti-
S.fruqiperda serum, at several dilutions from 1/10 to 1/1000, was added to 2.5 ml of the infected cell suspen
sion in 5 ml plastic tubes. The mixtures were incubated
for 1.5 h at 20°C, then transferred to 10 ml centrifuge
tubes. The centrifuge was run at 50 g for 3 min at 4°C.
The sediment containing the cell debris was discarded.
The supernatant containing the parasites was further
centrifuged at 500 g for 15 min and used immediately to
inoculate the cultures.
(2) Mechanical lysis S. frugiperda cells infected with Nosema. sp., N. heliothidis, N. locustae, V. necatrix and P. operophterae
for 10-15 days were centrifuged in plastic universal tubes at 150 g for 15 min at 20°C, then resuspended in 3 times
their volume of ice-cold fresh medium for 5 min. The
infected cells were agitated for 30 seconds at room temperature with 2mm sterile glass beads using a whirli- mixer, then centrifuged at 50 g for 8 min at 4°C to remove the cell debris and most of the spores. The pellet was discarded. The suspension was centrifuged at 500 g for 10 min at 4°C and the harvested parasites were resuspended in
2 times their volume of fresh medium at 4°C. Examinations of purified parasite preparations and counts of the number of parasites per ml were carried out with a phase contrast microscope. 70
3.11 Determination of the viability of extracellular
parasites
Sterile 10 mm round coverslips were introduced into
12 wells of a 24-well plastic plate. 1.5 ml TC-100 medium 4 and 2 x 10 uninfected S3, frugiperda cells were added per well and the cells were incubated at 25°C for 48 h. 0.5 . 5 ml of the suspension of parasites containing 8-11 x 10 meronts and sporonts was added to the wells at time intervals of 0, 30, 60, 120 and 180 min after their isolation. There were replicated for each time group.
The percentage of cells infected was estimated after 48 h from the Giemsa stained coverslips.
3.12 Determination of the phagocytic ability of S3. frugiperda cells.
Uptake of calf red blood cells was used to assay the normal phagocytic activity Of S. frugiperda cells and their phagocytic activity, after addition of Cytochalasin B.
The cells with and without Cytochalasin B were 7 prepared, on coverslips before 1.4 x 10 calf red blood cells were added into each well. Cultures were incubated at 25°C and were removed after 2 h and washed twice in PBS for examination. The number of cells ingesting erythro cytes was estimated. Three replicates of each treatment were included 71
3.13 Determination of the effect of antiphagocytic
reagent on infection of S. frugiperda cells by N.
heliothidis. S. frugiperda cells were grown on sterile 10 mm round coverslips introduced into the wells of a 24-well tissue culture plate. To test the effect of the reagent on invasion by N. heliothidis, the cultures were divided into 2 groups. 4 In group I, 5 x 10 £3. frugiperda cells per well were incubated for 2 h, the medium was discarded and replaced with Cytochalasin B prepared at concentrations of 5, 7.5, and 10 jjg/ml. The cultures were incubated for 1 h at 25°C before 7 x 10^ isolated pre-spore stages (meronts or sporonts). (Section 3.10.) were added. The cultures were incubated at 25°C for a further 48 h when the coverslips were removed, stained with Giemsa and examined for percentage of cells infected.
In group II, one part of infected cells were mixed with 4 parts of uninfected cells. The initial levels of infection varied from 15 - 20% in different experiments.
The cells were added to coverslips in the wells of a 5 24-well plate at 7 x 10 cells/well. After incubation for 20 h to allow the cells to attach, the medium was discarded and replaced with medium containing Cytochalasin
B at concentrations as (b) above. Sample coverslips were removed at 96 h, stained with Giemsa and examined for percentage of cells infected. 72
In control cultures no Cytochalasin B was added, and to one group of control cultures 0.1% (v/v) DMSO was added in the concentration used to dissolve the Cytochalasin B.
The percentage of cells infected was estimated in the controls and compared with the test wells. Three replicates for each treatment were included.
3.14 Demonstration of possible receptor involvement in
microsporidian invasion of cells.
All chemicals were purchased from Sigma Ltd and dissolved at required concentrations in TC-100 medium. jS. frugiperda cells were grown on sterile 10 mm round coverslips introduced into the wells of a 24-well plate.
To test the effect of the chemicals on invasion by vegetative stages, the experiments were carried out in two ways: a) 4.5-5 x 10 £5. frugiperda cells were incubated for 2 h; the medium was discarded and replaced with the medium containing the chemicals at concentrations which are given in Table 5. After 7-8 xlO^ isolated meronts and sporonts were added, the cultures were incubated at 25°C for 72 h when the coverslips were removed., stained with Giemsa and examined for percentage of cells infected. b) One part of infected cells were mixed with 4 parts of uninfected cells. The initial level of infection was estimated by sampling the mixture. The cells were added to coverslips in the wells of a 24-well plate at 7.0 x 73
5 10 cells/well. After incubation for 2 h to allow the cells to attach, the medium was discarded and replaced with medium containing the chemicals. After incubation for 96 h the coverslips were removed, stained with Giemsa and examined for percentage of cells infected. Three
replicates for each treatment were included.
3.15 Preparation for electronic microscopy 3.15.1 General method
The infected cells were fixed iji situ in flasks, using Karnovsky's fixative for 5-10 min at room temper
ature and for a further 1 h at 4°C. After washing twice
in 0.12 M cacodylate buffer pH 7.4, the cells were removed with a rubber scraper, spun in embedding capsules (TAAB) and were then postfixed in 1% OsO^. The dehydration,
staining and embedding procedures are given in Appendix 2.
Sections were cut with a glass knife on a Reichert ultra microtome, then mounted on copper grids, stained in 2% uranyl acetate for 10-15 min and 2% lead citrate for 5 min. The sections were examined with a Phillips EM 300 electron, microscope at accelerating voltages of 60-100 KV.
3.15.2 Detection of acid phosphatase by electron micro
scopy in S . frugiperda cells infected with P.
operophterae and N. heliothidis.
The infected S. frugiperda cells were fixed in
Karnovsky's fixative iji situ for 1 h at 4°C, washed once
in 0.1 M cacodylate buffer, pH 7.4, then washed twice in 74
Tris/maleate buffer pH 5.0. Cells were incubated in an
incubation solution, which contained 40 mM Tris/maleate buffer, 2.4mM lead nitrate and 8mM ^-glycero-phosphate, at 25°C for 1 h. After washing twice in Tris/maleate buffer,
the cells were removed from the surface of the flask with
a rubber scraper. The suspension was transferred to
embedding capsules of Eppendorf tubes with a sterile
pipette and the cells were spun at 500 g in a centrifuge.
Cells were postfixed in 2% OsO^ for 1 h at 4°C, and
processed as before.
3.15.3 Detection of secondary lysosomes in £>. fruqiperda
cells infected with P. operophterae and N. heliothidis.
Secondary lysosomes in £5. fruqiperda cells were either labelled with 2 mg of saccharated iron oxide (TAAB)
per ml in TC-100 medium, added to the culture for 1 h, or
with 5 mg of Ferritin per ml added to the culture for 5 h.
After washing twice in TC-100 medium. The cells were incubated for a further 2 h, then scraped from surface of the flask, centrifuged to form a pellet and processed as
before.
3.16 Attempts to induce dimorphism Jjl vitro by applying hormone.
All chemicals were purchased from Sigma Ltd. and
dissolved at required concentrations in TC-100 medium, which are given in Table 6. 75
S. frugiperda cells, infected with either V. necatrix or N. locustae, were grown on sterile 10 mm round coverslips introduced into the wells of a 24-well tissue culture plate. The TC-100 medium contained B-ecdysone or juvenile hormone at different concentrations. Incubation was at 20°C. 76
TABLE 5
Chemicals used to investigate receptor involvement in microsporidian invasion
Chemicals Concentration Trypsin 100 pg/ml Concanavalin A . 50 pg/ml Wheat germ agglutinin 100 pg/ml Soybean agglutinin 50 pg/mi N-acetyl-D-galactosamine 10 mM N-acetyl-D-glucosamine 10 mM D-glucose 10 mM D-mannose 10 mM
TABLE 6 Chemical concentrations
J3 - ecdysone Juvenile hormone 10 ng 50 ng 50 ng 100 ng 100 ng 500 ng 500 ng 1000 ng 77
4. RESULTS
4.1 Germination of spores in vitro
The spores of a species of Nosema isolated from
Spodoptera litura (hereafter referred to as Nosema sp.),
Nosema heliothidis, Orthosoma operophterae, Glugea anomala and Amblyospora.sp. isolated from Simulium ornatum all germinated in a two step procedure, in which the spores were pre-treated with 1M KC1 buffer solution at one pH value and then treated in 1M KC1 buffer solution at another pH value.
The results are summarised in Figs. 2 to 4.
4.1.1 pH optima for pretreatment and treatment
The spores of the two species of Nosema, both derived from lepidopteran hosts, hatched when pretreated at a high pH value and treated at a lower pH value. The data for
Nosema sp., represented three dimensionally in Fig. 2, show that there are two germination peaks. One was induced by pretreatment at pH 11 - 13 and treatment at pH 5 - 7? the other was induced by pretreatment at pH 6 - 8 and treatment at pH 11 - 13. The most effective combination was pH 13 followed by pH 6. Fig. 4 shows that spores of N. heliothidis have pretreatment optima similar to those of Nosema sp. from S. litura and hatched when the spores were transferred to TC-100 medium at pH 6.8 or buffer at pH 6 or 7. 60% of 78
FIG. 2: The effect of pH in germination of spores of Nosema sp.
from J3. litura.
FIG.
in buffer at different pH. 79
spores of Nosema sp. (Fig. 2) and 40% of spores of N.
heliothidis (Fig. 4) hatched in TC-100 medium/ after pre
treatment at pH 13 or pH 12, respectively. The optimum pH
for the pre-treatment of spores of 0. operophterae, G.
anomala and Amblyospora sp. from S. ornatum was pH 9.5.
Hatching occurred after transfer to buffer at pH 6.5 - 7 or to TC-100 medium at pH 6.8. The maximum germinations were 30 - 40% for O. operophterae (Fig. 3) and 20% for G. anomala
and Amblyospora sp.
4.1.2 Time for completion of spore germination
The spores of Nosema sp. from S5. litura were sampled at
several time intervals, when they were placed in TC-100 medium at pH 6.8, after pretreatment in buffer solution at pH 13. More than 40% of spores hatched within 1 min. The number germinating rose to 60% at 20 min after which no further hatching took place (Fig. 5).
4.2 Establishment of microsporidia .in the Spodoptera
frugiperda cell line.
4.2.1 Attempts to infect cell cultures with spores Attempts were made to infect £5. frugiperda cells with
purified spores of the Nosema sp. from S. litura, 0.
operophterae and G. anomala. When spores had been exposed
to pH 13 (Nosema sp.) or pH 9.5 (O. operophterae, G.
anomala) then added to the cell cultures no intracellular parasites were seen within 48 h, although extruded polar 80
Fig. 4: The effect of pH on germination of
N. heliothidis spores
O Treatment with buffer * at pH 7.
Fig. 5: Time for completion germination of spores
of Nosema sp. from.S. litura 81
tubes and extracellular sporoplasms were present in the medium. After 48 h, the contaminated cultures were discarded.
4.2.2 Infection of cell cultures with infected haemolymph
Five microsporidia, belonging to the genera Nosema (3 species), Vairimorpha (1 species) and Pleistophora (1 species), all of which infect haemocytes of Heliothis zea larvae, were successfully transferred to S_. frugiperda cell cultures, by addition of infected haemolymph to the cultures. P. schuberqi, which only infects midgut tissue of host larvae, did not infect the cultures. The details are shown in Table 7. The infected cultures were incubated at 20°C or 25°C. The infected haemocytes from the donor larvae gradually died and finally disappeared from the cultures after 1 or 2 weeks. After 2 weeks, 21-64% of cultured cells were infected with the microsporidia. Cont inuous cultures were maintained by weekly addition of 3 - 5 parts of healthy cells into 1 part of heavily infected cells. Infected cultures were maintained for more than 1 year and were subcultured through 21-64 passages according to species. (Table 7).
4.3 The effect of temperature on multiplication of micro
sporidia in S. frugiperda cells
Infections of N. heliothidis, Nosema sp., N. locustae,
V. necatrix and P. operophterae were established in !3. frugiperda cells by mixing 5 parts of healthy cells with 1 Table 7: Cultures of microsporidia in £3. fruqiperda cell line by addition of haemolymph from infected H. zea
Cell Temperature No. of Duration of % Cells Species Cultures of cultures passages continuous infected * infected ° c cultures (months)
Nosema sp. + 25 43 18 14
N. heliothidis + 25 64 32 33
N. locustae + 25/20 23 6 38 V. necatrix + 25/20 21 6 26 P. operophterae + 20 32 12 34 P. schubergi** — 20 0 0 0
* Percentages of infected cells were estimated 2 weeks after inoculation with infected haemolymph
P. schubergi only infects larvae midgut tissue 83
part of infected cells. The initial levels of infection were estimated and the cultures were then incubated at 37°Cf 30°C, 25°C and 20°C. The number of cells infected and the number of parasites (estimated as total parasites or veget ative stages only) per 200 cells were estimated every 48 h for 8-14 days according to species. None of the micro- sporidia grew at 37°C. The data for the other temperature are presented in Figs. 6 to 20.
4.3.1 Nosema heliothidis
The growth curves over 10 days for 3 temperatures, in terms of number of cells infected arefshown in Fig. 6. The initial level of infection was 17%. No differences were observed for 2 days, but by 6 days there were significant differences in the percentage of cells infected (P<0.05).
At 30°C, the percentage rose to 41% after the 4th day and fluctuated slightly below this level for the remainder of the experimental period. At 20°C, the percentage of cells infected was 31% after the 4th day, but rose to 63% by the end of the experiment. At 25°C the infection increased steadily throughout the 14 days of incubation and by 10 days almost all of the cells were infected.
The total number of parasites and numbes of vegetative stages per 200 cells are given in Figs. 7 and 8 respect ively. At 20°C and 30°C the multiplication rates were similar, reaching about 1000 parasites or 500 vegetative stages per 200 cells. A much higher growth rate was found 84
Figs. 6 - 8: The effect of temperature on growth of
N. heliothidis in S. fruqiperda cell
cultures.
Fig. 6 The percentage of cells infected with N. heliothidis when incubated at 20°C, 25°C
and 30°C.
Fig.7 The number of parasites (vegetative stages and spores) per 200 cells when cultures were incubated at 20°C, 25°C and 30°C.
Fig. 8 The number of vegetative stages per 200 cells when cultures were incubated at 20°C,
25°C and 30°C. No. of vegetative stages in No. of parasites in % infected cells 200 cells 200 cells 0 2 Time in days in Time 85 25°C 30°C 20C
i. 6 Fig. Fig. 8 86
at 25°C, with final figures for total parasites and vegetative stages more than double those at 20°C or 30°C.
4.3.2 Nosema sp.
The initial infection level was 17% (Fig. 9). At 30°C the number of cells infected reached a maximum at 4 days.
Thereafter, there was a sharp reduction to 19% at 10 days.
When the cultures were incubated at 20°C, there was an initial fall in infection level then, an increase at 4 days to a level which was maintained almost constant for the remaining period of observation. At 25°C the slight initial drop in infection level was recovered at 4 days, then levelled off at 45% at 6 days. There were significant differences in infection levels at the 3 temperatures on days 8 and 10 (P<0.05). Growth curves, shown by numbers of parasites per 200 cells, were similar when the cultures were incubated at 20°C and 25°C (Fig. 10). The exponential rate was for the first
2 days at 25°C and for the first 4 days at 20°C. At 30°C the number of parasites increased to nearly 700 parasites per 200 cells at 6 days and remained at this level. Examination of vegetative stages only, showed the same patterns of growth as for total parasites with a steady increase in numbers at 20°C and 25°C but not at 30°C. There were significant differences at 10 days between the numbers of vegetative stages produced at 30°C and the numbers at
20°C or 25°C (P<0.05) (Fig. 11). 87
Figs . 9-11: The effect of temperature on the
growth of Nosema sp. in £5. fruqiperda
cell cultures.
Fig.9 The percentage of cells infected with
Nosema sp. when incubated at 20°C, 25°C and
30°C.
Fig.10 The number of parasites (vegetative stages and spores) per 200 cells when cultures were incubated at 20°C, 25° and 30°C.
Fig.11 The number of vegetative stages per 200
cells when cultures were incubated at 20°C,
25°C and 30°C. 00 00 (Ti tO tO ^ % % infected cells o o o o o H-
0
0 H H
0 u> u> to to O O O O O cn 200 200 cells U> U> Ln *0 O O <-n O No. No. of parasites in H- iQ H
0
0 200 200 cells
0 to U1 O O O <~n O O O No. No. of vegetative stages in
Time in days 89
4.3.3 Nosema locustae The percentage of cells infected increased with time at all temperatures but was lower at 20°C than at 25°C or 30°C
(P<0.05). The numbers of infected cells did not differ significantly between cultures held at 25°C and 30°C for 10 days. After incubation at 30°C for 10 days, all the £5. frugiperda cells were infected. (Fig. 12). Differences in the numbers of parasites in the cells were not significant until day 6 (Fig. 13), when the number of parasites at 30°C was higher (P<0.05). All three groups were different at 8 days (P<0.05). At 10 days there was a significant difference between the numbers of parasites in the cultures held at 20°C and in the cultures held at 25°C or 30°C (PC0.05) .
The vegetative stages followed similar growth patterns to total parasites (Fig. 14) with multiplication at 25°C being higher than at 30°C and 20°C.
4.3.4 Vairimorpha necatrix The progession of V. necatrix infection in cell cultures is shown in Fig. 15, from an initial level of infection of 13%. The percentage of cells infected increased steadily at 25°C to 74% at 10 days. Differences in levels of infection between cultures held at 25°C and those held at 20°C or 30°C became significant at 6 days (P<0.05), but differences between cultures at 20°C and 30°C were not significant at any time during the 10 days incubation. 90
Figs. 12 - 14: The effect of temperature on the
growth of N. locustae in S.
frugiperda cell cultures.
Fig.12 The percentage of cells infected with N.
locustae when incubated at 20°C, 25°C and
30°C.
Fig.13 The number of parasites (vegetative stages and spores) per 200 cells when cultures were incubated at 20°C, 25°C and 30°C.
Fig.14 The number of vegetative stages per 200
cells when cultures were incubated at 20°C,
25°C and 30°C. % cells % infected 200 200 cells No. No. parasites of in 200 200 cells . of vegetative stages . in No
Time in days 92
Figs. 15 - 17: The effect of temperature on the
growth of V. necatrix in S.
frugiperda cell cultures.
Fig.15 The percentage of cells infected with V.
necatrix when incubated at 20°C, 25°C and
30°C •
Fig. 16 The number of parasites (vegetative stages and spores per 200 cells when cultures were incubated at 20°C, 25°C and 30°C.
Fig.17 The number of vegetative stages per 200
cells when cultures were incubated at 20°C,
25°C and 30°C. No. of vegetative stages in No. of parasites in % cells infected 200 cells 200 cells
to £* cn oo O M to cn oo O O O O O C cn O <-n O Ln
KD tjj
u> tO to 15 Fig. O o in o o 0 H- o o O iQ H u> to to CT» O O <-n 0 0 0 no n 94
Figs. 16 and 17 show the numbers of parasites and vegetative stages per 200 cells. After 2 days the increase in numbers of total parasites and of vegetative stages alone in cultures held at 25°C differed significantly from the numbers in the cultures held at 20°C or 30°C (P<0.05)f but the last two did not differ from one another.
4.3.5 Pleistophora operophtera
The effect of temperature on development and multi plication of P. operophterae in cell cultures is shown in
Figs. 18, 19 and 20. There was no development in cultures incubated at 30°C, during 8 days of observation. Develop ment was established at 20°C and 25°C (Fig. 18). From an initial level of 35% there was a rapid increase in the percentage of cells infected at 20°C, so that the infections were significantly different between 20°C and 25°C at 2 days
(P<0.05). Thereafter the two cultures were similar, and there was no significant difference in levels of infection during the last 6 days incubation. The total number of parasites and numbers of vegetative stages per 200 cells are given in Figs. 19 and 20. The total number of parasites increased at 20°C throughout 8 days of incubation. Although the parasites developed more slowly at 25°C than at 20°C before day 4 (P<0.05), there was a spurt of growth during the next 4 days of incubation. The final level of infection was 3 times greater than the initial at 25°C and 4.5 times greater at 20°C, over 8 days 95
Figs. 18 - 20: The effect of temperature on the
growth of P. operophterae in S.
frugiperda cell cultures.
Fig.18 The percentage of cells infected with P.
operophterae when incubated at 20°C, 25°C
and 30°C.
Fig.19 The number of parasites (vegetative stages and spores) per 200 cells when cultures were incubated at 20°C and 25°C.
Fig.20 The number of vegetative stages per 200
cells when cultures were incubated at 20°C
and 25°C. No. of vegetative stages in No of parasites in % cells infected 200 cells 200 cells l-1 to U> H I—1 to U> 00 cr» to o 10 on cry oo vo
to H* 3 CD H- 13 a cr> cn vo at
oo
*i H- H* H- iQ uQ • • M3 to H O VO oo 97
incubation. A higher number of vegetative stages was also obtained at 20°C, compared with 25°C (P<0.05).
4.4 The effect of antibiotics on the infectivity to S.
fruqiperda cells of Nosema sp. and N. heliothidis The effect of certain antibiotics on viability of
Nosema sp. and N. heliothidis was investigated by cultivating infected cells either in the absence of anti biotics or in the presence of penicillin plus streptomycin, or in the presence of kanamycin. These antibiotics were added routinely to TC-100 medium during cell culture, as a precaution against contamination. Thus, it was necessary to determine their effect on the microsporidia. The cells were incubated at 25°C.
4.4.1 The effect of antibiotics on N. heliothidis
The significant differences were observed due to antibiotic treatment throughout 10.days cultivation (Figs.
21, 22). The percentage of infected cells rose in both treated and control groups, from an initial level of 22% to more than 90% during 10 days incubation.
4.4.2 The effect of antibiotics on Nosema sp. A significant difference in the growth of Nosema sp. in
S. fruqiperda cells was found betweem control cultures and those treated with 500 i.u. penicillin and 500 pg strepto mycin per ml, from the 6th day of culture (P<0.05)(Fig. 23). cells infected % cells infected Fig. 22: The effect of kanamycin on infectivity of infectivity on ofkanamycin effect The 22:Fig. Fig. 21: The effect of penicillin and strepto and penicillin of effect The 21:Fig. N. heliothidis at 25°C at heliothidis N. mycin on infectivity of heliothidis N. infectivity on mycin t 25°C at 8 • 98
/ug/ml /ug/ml 99
The percentage of cells infected in the control group rose from an initial 18% to 55% after 10 days. This was 21% higher than in cultures treated with penicillin and streptomycin. The difference between the control cultures and those treated with 100 i.u. penicillin and 100 pg streptomycin were not significant. An effect of kanamycin at 500 pg per ml on growth of Nosema sp. was detected throughout 10 days of incubation (P<0.05) (Fig. 24). The percentage of cells in the cultures treated with 500 pg per ml kanamycin was 16% lower than in the controls. The difference in infectivity of Nosema sp. between control culture and those treated with 100 pg per ml kanamycin was not significant.
4.5 Viability of extracellular stages of microsporidia in
TC-100 medium
The ability of vegetative (non-spore) stages to enter host cells was investigated using five species of micro sporidia, N. heliothidis, Nosema sp., N. locustae, V. necatrix, P. operophterae. The vegetative stages, which had been separated from spores and from host cells, were added to S3. frugiperda cells at different time intervals after isolation. Two techniques for separation of the N. heliothidis vegetative stages in TC-100 medium at 4°C were investigated. These were immuno-lysis and mechanical disruption of host cells (to release parasites) followed by differential centrifugation (to separate the dense spores 100
Fig. 23: The effect of penicillin and streptomycin
on infectivity of Nosema sp.
/ug/ml /ug/ml
Fig. 24: The effect of kanamycin on infectivity of
a) p -o from the vegetative stages). Details of the methods are given in section 3.10. Viable extracellular parasites, which could infect cells, were only obtained after mechanical disruption. This technique was, therefore, employed for the remaining species. The isolated vegetative stages were either added immediately to £3. frugiperda cells or after incubation at 20°C in TC-100 medium for intervals of 30, 60, 120 or 180 min and the percentage of cells infected after 48 h was estimated. After addition of the isolated parasites to the cultures, the cultures were incubated at 20°C. A temperature of 20°C was selected for incubation of the extracellular stages, to reproduce the conditions under which the parasites were known to survive in cultures. Fig. 25 shows that the percentages of cells which became infected were directly dependent on the time intervals between isolation of the vegetative stages and their addition to the cultures. The* proportion of cells infected in the cultures at 48 h declined from nearly 5% when the vegetative stages had been added immediately after their isolation, to zero when the vegetative stages had been incubated in TC-100 medium for 180 min before their addition to the cultures. The extracellular survival time for the vegetative stages of N. heliothidis in TC-100 medium, at 20°C, is little more than 60 min. For Nosema sp., there was a similar steep decline in infectivity, with time from isolation, of the vegetative 102 Fig. 25: Viability of Nosema heliothidis in vitro Fig. 26: Viability of.Nosema sp. in vitro 103 stages (Fig. 26). 4.5% of cells showed infection at 48 h, when the vegetative stages were added immediately and all infectivity was lost by the vegetative stages 60 min after their isolation. The figure for V. necatrix was 3.2% of cells infected at 48 h, when vegetative stages were added immediately, 1% when the vegetative stages remained extracellular at 20°C for 60 min before addition to cultures, and 0 % when they had remained extracellular for 120 min (Fig. 27). With P. operophterae an initial rapid loss of infecti vity, between 0 and 30 min after isolation, was followed by a slower rate of decline in infectivity with some viability of extracellular stages remaining at 120 min, giving rise to 1% infection of cells at 48 h (Fig. 28). The infection level had declined further to 0.5% after 180 min. N. locustae was able to tolerate extracellular con ditions at 4°C and 20°C for longer than the other parasites. After an initial rapid decline in infectivity, shown by a fall in percentage of cells infected at 48 h, from 7%, when the isolated parasites were added immediately, to less than 3% after an interval of 60 min, some infectivity was retained even after incubation in TC-100 medium at 20°C for 180 min (Fig. 29). The results of these experiments provide evidence that extracellular vegetative stages of microsporidia are capable of initiating infection in new host cells but that the ability is almost completely lost within 30 min by the 104- Fig. 27: Viability of V. necatrix in vitro 105 Nosema sp., 60 min by N. heliothidis and V. necatrix, 120 min by P. operophterae and 180 min by N. locustae. 4.6 The effect of adenosine triphosphate (ATP) on invasion of cells by Nosema sp. and N. heliothidis As microsporidia lack mitochondria and thus lack an endogenous energy source, an attempt was made to extend the viability of extracellular stages in cultures by adding ATP to the medium supporting the cells. Initial levels of infection of Nosema sp. and N. heliothidis were established in £3. fruqiperda cells, as described in section 3.9. The cells were maintained at 25°C in TC-100 medium, to which was added 2 mM or 4 mM ATP. Control cultures were set up without ATP. No antibiotics were added to any of the cultures, in order to avoid the possibility of interference with the growth of the parasites. For the Nosema sp., there was a greater increase compared with control cultures in the number of infected cells, in the presence of 4 mM ATP during 10 days incubation (P<0.05) (Fig. 30). At the end of the experiment more cells were infected in the presence of 4 mM ATP. In the presence of 2 mM ATP there was no significant difference in the number of infected cells compared to the control cultures up to the 6th day of cultures, but the difference was significant on the 8th and 10th days (P<0.05). Thus, when ATP is made available, the extracellular infective stages of Nosema sp. have a greater ability to invade cells. cells infected % cells infected Fig. 31: The.effect of ATP on infectivity of N.of infectivity on ofATP The.effect 31:Fig. Fig. 30: The effect of ATP on infectivity of Nosema sp Nosema of infectivity on of ATP effect The 30:Fig. heliothidis Time in days in Time 106 107 Equally, when ATP was added to cultures of N. helio- thidis, there was a significant difference between the percentage of cells infected in control cultures and in those to which 2 mM or 4 mM ATP had been added (Fig- 31). The differences were significant (P<0.05) from day 4 and in cultures to which 4 mM ATP had been added, there was near significant difference (P<0.07) on day 2. There was an almost 10 % difference in the number of cells infected with N. heliothidis between cultures treated with 4 mM ATP and control cultures throughout the experiment. These results provided evidence that an exogenous supply of energy is required by extracellular stages of microsporidia to sustain them long enough to enter host cells and initiate a new developmental cycle. 4.7 The effect of Cytochalasin B on entry of N. heliothidis into S. frugiperda Cytochalasin B, a reagent which affects microfilament function, was used to determine whether entry of vegetative stages of microsporidia into host cells is a function of host cell or parasite activity. In one experiment, the £5. frugiperda cells were treated for at least 1 h in Cytochalasin B, before the extracellular microsporidia or bovine red blood cells were added. In a second experiment, the S. frugiperda cells were infected and the level of infection was determined, after which they were incubated in the presence or absence of the microfilament inhibitor for 108 specified times. The details of the methods are given in section 3.12 As Cytochalasin B was made up with 0.1% DMSO, the effect of DMSO was ascertained by adding the same concentration to cultures as an extra control. A dose-dependent inhibition of uptake of erythrocytes by £5. fruqiperda cells was observed (Fig. 33). After 24 h incubation, more than 2% of cells had taken up the erythrocytes in the control; groups with and without DMSO. There was a significant decrease in the uptake of r.b.c. with increasing concentrations of Cytochalasin B, with complete inhibition of uptake of erythrocytes at 10 pg per ml Cytochalasin B. A dose-dependent inhibition by Cytochalasin B of entry of parasites into S. frugiperda cells, was observed when the infective stages were added immediately after their isolation to a cell monolayer (Fig. 32). A concentration of 7.5 pg/ml of Cytochalasin B, significantly decreased the number of cells infected with N. heliothidis (P<0.01) compared with the control group or with the cells exposed to 0.1% DMSO alone. No parasites were observed inside the cells which were treated with 10 pg/ml of Cytochalasin B. A slight, but statistically significant, reduction in number of cells infected with N. heliothidis was seen (P<0.lf) in cultures treated with 5 pg/ml of Cytochalasin B. When the cells already infected with N. heliothidis, were incubated with Cytochalasin B, spread of N. heliothidis to other cells was inhibited (Fig. 34) (P<0.001). After 109 Fig. 32: The effect of Cytochalasin B on entry of N. heliothidis into S . frugiperda cells control DMSO 5 pg 7.5 pg 10 pg Fig 33: The effect of Cytochalasin B on entry of red blood cells into cells of S. frugiperda o' • c = control x • u D = DMSO tp p II E Ln •H "P td +> F = 7.5 pg in II G o P: Fig. 34: The effect of Cytochalasin B on N. heliothidis cell invasion initial infection control Q)p -.u DMSO 9.6 h incubation, the increase in the number of infected cells from the initial level was less than 14% in the cultures treated with 10 }ig/ml of Cytochalasin B, compared with more than 40% increase in the control cultures, with and without DMSO. 4.8 The effect.of lectins and carbohydrates on invasion of S. frugiperda cells by different species of micro- sporidia All experiments were conducted by addition of vegetative stages or infected cells into £5. frugiperda cell cultures in the presence of the treatments of lectins or carbohydrates. N . heliothidis The number of infected cells was reduced by almost 20% compared with untreated control cultures when 10 mM D-mannose or 50 jig/ml ConA were present in the medium (P<0.05). A near-significant inhibition of infection was found when 10 mM N-acetyl-D-galactosamine was present (P<0.07); this caused a 14% decrease in the number of infected cells compared with the controls (Fig. 35). A similar result was obtained when £3. frugiperda cells were treated first with the sugars or lectins for 1 h, then the newly isolated stages were added (Fig. 36). There was a significant decrease in the percentage of cells infected after treatment of cells with 10 mM D-mannose, 10 mM % cells infected % cells infected P- H1 to N-acetyl-D-galactosamine, or 50 pg/ml ConA (P<0.01). When the ConA plus N-acetyl-D-galactosamine or 50 jig/ml soybean agglutinin plus the D-mannose were present in medium together, the percentage of cells infected was decreased by a further 1 or 2%. Complete inhibition of invasion of N. heliothidis was not achieved. N . locustae The vegetative stages were isolated and added into S. frugiperda cells which had been pretreated with the lectins or sugars for 1 h. Parasite entry was reduced by 4% in the presence of 10 mM N-acetyl-D-galactosamine,4,6% in the presence of 10 mM D-mannose and $% in the presence of 50 jig/ml ConA (Fig. 37) (P<0.05). The other sugars and lectins had no effect on the entry. V. necatrix Isolated vegetative stages of V. necatrix were added to the cell cultures in the presence of the sugars or lectins. Only 10 mM N-acetyl-D-galactosamine and 50 pg/ml soy-bean agglutinin inhibited the entry of V. necatrix into cells. The percentage of cells infected was decreased by 5 - 6% compared with the controls (P<0.01). The other sugars and lectins, or trypsin had no effect on the parasite entry (Fig. 38). Fig. 38: The effect of sugars and lectins on entry of entry on lectins and sugars of effect The 38:Fig. H* ip• % cells infected % cells infected (jJ «• O to £s» • oo of intocells necatrix V. IZ• 3*► 3 ro H 0 ro 0 Hi c Hi W 0 rt ort (DP H- 0Hi 3rt to 0 3 i p 0 P tt> H M cn t—* H U) OT P 3 0Hi Cb S. |cnH a> frugiperda • o Hi rtH- H 3 3 cn lQ H* 0 P 3 (D H (D Pi 3 P ft H oHi 114 P . operophberae Fig. 39 shows the effect on entry into cells by P. operophterae when sugars or lectins or trypsin were present in the culture medium. In the presence of 100 jig/ml trypsin, entry into cells was reduced by 23% compared with the controls (P<0.01). In the presence of 10 mM N-acetyl- D-galactosamine ■, 10 mM D-mannose or 50 pg/ml soybean agglutinin, entry was reduced by 16% and in presence of 50 pg/ml ConA entry was reduced by 26% (P<0.05). 4.9 Attempts to induce dimorphism iri vitro S. frugiperda cell cultures infected with either N. locustae or V. necatrix were incubated in the presence or absence of 4 concentrations of p-ecdysone, ranging from 10-500 ng/ml or juvenile hormone from 50-1000 ng/ml (section 3.16). Cultures were incubated at 20°C for more than 20 days. Neither N. locustae nor V. necatrix produced octospores and only the disporoblastic sequence was observed. Attempts to induce dimorphism .in vitro failed. 4.10 The morphology and life cycles of the microsporidia studied by light and electron microscopy 4.10.1 N. heliothidis in vitro Light microscope observations The life cycle of N. heliothidis in the S. frugiperda cell line was determined by inoculation of infected blood cells from Heliothis zea into uninfected cell cultures. The H- 0 3 % cells infected • U) tO £* cr> VO o o o O M 0 H3 H i 3 * CD • CD H i 1 0 H i 0 0 0 H r t 0 0 3 * H i r t 0 tn H 3 PJ 0 p> H H* H- [0 H-* 3 L n r t P) O 3 CL |c n • I - 1 0 H i 0 H r t 3 H* iQ 3 H- cn t l 0 0 H 3 CL PJ 0 3 O r t (0 H H W 116 development was monitored over 10 days. As the morphology and development of N. heliothidis did not change with temperature the infected cultures were normally incubated at 25°C. Stages with pale cytoplasm and two nuclei and sometimes stages with apparently only one nucleus, were thought to be meronts (Fig. 40). They measured 2.0 + 0.11 pm in diameter. The majority of stages both merogonic and sporogonic were diplokaryotic. It was difficult to distinguish between meronts and sporonts but ovoid stages with pale cytoplasm and rounded division products were considered to be meronts, while spindle shaped stages with dense cytoplasm were thought to be sporonts. (Figs. 40 - 44). Division of both was by binary fission. After successive merogonic divisions, abundant stages were seen in the cell cytoplasm (Figs. 42 - 44). When parasites were released by breakdown of heavily infected cells (Fig. 45). Some of the free stages became closely associated with the surface of new host cells (Fig. 46). Sporogony began after 4 - 5 replications of meronts. The sporqnts, which are spindle shaped with dense poles and diplokarya, measuring 8.4 + 1.2 x 2.9 + 0.4 pm, gave rise to two sporoblasts which gradually developed into free, ovoid spores, measuring 5.6 _+ 0.54 x 2.8 + 0.35 pm (Figs. 47, 48). Usually, the division and morphogenesis were synchronized in a cell so that a single cell, contained either meronts and sporonts or spores. 117 Electron microscope observations The earliest stages of N. heliothidis were seen lying close to the host cell nucleus. Isolated parasites lay in contact with unaltered host cell cytoplasm (Fig. 49), but after some divisions, spaces adjacent to the parasite, over part or even all of its surface, were occupied by tubular structures (Figs. 50 - 52). The tubules measured 160 nm diameter and appeared to connect the surface of the parasite with the regions of unaltered host cytoplasm. During sporogony the tubules tended to disappear (Fig. 55) and at the stage of spore maturation the surrounding host cell cytoplasm was almost totally disorganized (Figs. 56, 57). Meronts were oval or elongate in form. Usually there were two nuclei in diplokaryon arrangement and where a single nucleus was observed, it could not be determined with certainty, whether or not this was one of a diplokaryotic pair. The cytoplasm contained abundant ribosomes and a few cisternae of endoplasmic reticulum. (Tig. 52). At the initiation of sporogony a thick electron dense layer was secreted onto the plasma membrane. (Fig. 55). Many ribosomes were membrane-associated as rough endoplasmic reticulum (Figs. 53, 54). Signs of nuclear division were provided by electron dense centriolar plaques (Fig. 53). Spindle microtubules and electron dense masses were interpreted as chromosomes (Fig. 54). Spore morphogenesis was not observed because of poor fixation of sporoblasts and spores. However, transverse and 118 longitudinal sections of the polar tube and diplokarya and a membrane bound structure consisting of tubules were observed in sporoblasts (Fig. 55) and spores (Figs. 56, 57). The electron lucent endospore was interpolated between the plasma membrane and the exospore coat (Fig. 57). 4.10.2 Nosema sp. from Spodoptera litura in vitro Light microscope observations Cultures of Nosema sp. were established by inoculating the heavily infected haemocyte cells of H. zea into uninfected cultures, which were then incubated at 25°C. The earliest stages in the cytoplasm of S. fruqiperda cells, were rounded and measured 2.6 + 0.3 pm diameter, their cytoplasm and nuclei were deeply stained but the diplokaryon arrangement of nuclei was recognized (Figs. 58, 59). Division occurred by migration of the pairs of nuclei towards the poles and constriction of cytoplasm in the centre (Fig. 61). Sporonts were spindle shaped with the diplokarya occupying the wider central region, divided^nto sporoblasts was by binary fission (Figs. 60, 61, 63). Spores measured 6.9 + 0.5 x 2.7 + 0.35 pm. (Fig. 64). Electron microscope observations Stages were poorly fixed but some details were observed. Meronts were surrounded by a simple plasma membrane in direct contact with host cell cytoplasm (Figs. 65-67). An electron dense surface coat was added to form 119 the sporonts. In the sporoblasts there were two nuclei (Fig. 69) and several concentric layers of endoplasmic reticulum around the nuclei (Fig. 68). Traces of the electron lucent endospore were laid down (Fig. 68). Spore structures were better preserved (Fig. 70)„. The nuclei were situated in the posterior half of the spore within 12 - 13 coils of the polar tube. Several concentric layers could be seen in cross sections of the polar tube. The polaroplast occupied most of the anterior half of the spore and was composed of closely packed membrane (Fig. 70) 4.10.3 Pleistophora operophterae in vitro Light microscope observations The earliest stages of the parasite observed were meronts with a single, compact nucleus, measuring 2-3 urn. (Fig. 11). Division of meronts was by binary fission and there was considerable increase in size. Meronts lay in close contact with the host cell cytoplasm (Figs. 72 - 73). Multinucleate sporogonial plasmodia underwent segmentation into uninucleate sporoblasts, during which it could be seen that these stages lay in distinct vesicles, the sporophorous vesicles (Figs. 70 - 75). The sporoblasts further developed into spores which once again packed the sporophorous vesicles, so that the vesicle boundary was obscured (Figs. 76, 78). Small rounded stages were often seen closely attached to the host cell surface; these were thought to be infective stages responsible for secondary invasion (Fig. 77). Spores measured 3.1 + 0.6 x 1.12 + 0.14 jam. 120 Electron microscope observations Uninucleate meronts were limited by a simple plasma membrane and lay within a vesicle, the membrane of which was closely fitted around the parasite membrane and followed the contours of the meront. Host cell mitochondria were aggregated close to these vesicles sometimes following the contours (Fig. 79). The vesicle divided with the meronts, as they underwent binary fission so that daughter meronts were isolated within their own vesicle membrane (Fig. 82). Occasionally, small vesicles with thickened membranes were present, close to the vesicle membrane surrounding a meront (Fig. 80). The membrane of the small vesicles and that of the vesicle round the meront were of similar thickness and density but the origin of the small vesicles could not be determined. At the onset of sporogony on electron dense coat, of uniform thickness was gradually deposited on the plasma membrane of the parasite (Fig. 81). Withdrawal of the parasite\ surface away from the vesicle membrane was followed by several divisions, so that numerous uninucleate sporoblasts came to lie within the vacuole, now a recog nizable sporophorous vesicle as defined by Canning & Hazard (1982) (Fig. 83). Often different sporogonic stages were seen within the same sporophorous vesicle, with those stages completely detached from the vesicle membrane and lying free in the vesicles being the first to undergo morphogenesis into spores (Fig. 84). Finally the sporophorous vesicles contained spores only (Fig. 85). In heavily infected cells, 121 a large part of the host cytoplasm was replaced by several sporophorous vesicles (Fig. 86). Mature spores were poorly preserved but some internal structures could be recognized. The single nucleus was situated near the middle of the spore, while the polaroplast, consisting of anterior close-packed and posterior loosely-packed membranes surrounded by the polar sac, occupied the anterior one-third of the spore. About 7 sections of the polar tube (PT) were seen at the posterior end (Fig. 87). 4.10.4 Life cycle of N. locustae in vivo N. locustae from grasshoppers was successfully g transferred to H. zea by per os inoculation of 10 spores into each 2nd instar larva. When the larvae were reared at 20°C two sporogonic sequences were observed. Light microscope observations in smears of fat tissue The youngest stages observed were small round, diplokaryotic meronts, measuring 3.9 + 0.4 pm, (Figs. 87 - 89). Division was by binary fission. Isolated nuclei as in Fig. 88 were thought to arise as an artifact of the smearing technique. In the disporoblastic sporogonic sequence, the sporonts were spindle-shaped either bi- or tetra-nucleate measuring 13 + 1.3 x 4.4 + 0.7 pm (Figs. 90 - 92). After migration of the diplokarya to the poles, division into sporoblasts was by binary fission (Figs. 93 - 94). Sporo- blasts measured 8.15 + 0.2 x 3.9 + 0.1 pm. They developed 122 into elongate spores (Fig. 95) which, when fixed and stained, measured 5.3 + 0.13 x 2.33 + 0.06 pm. The octosporoblastic sporogonic sequence appeared after about 18 days. A round binucleate sporont (Fig. 96) measuring 6.9 + 1.3 pm became tetranucleate (Fig. 97) then octonucleate (Fig. 98). The mature sporont divided into 8 uninucleate sporoblasts and ultimately into 8 spores which were packed together and encased in a sporophorous vesicle (Fig. 99). The fixed and stained spores measured 2.12 + 0.13 x 1.04 + 0.04 pm. Electron microscope observations At the time of examination of the H. zea larvae, the fat tissue was heavily parasitized. The host cells were severely disrupted and the parasites isolated in disorgan ized host cytoplasm in which few of the cell organelles were recognizable. Merogony Meronts were rounded cells each with a diplokaryon (Fig. 100). The plasma membrane was somewhat thickened by electron dense additions, but there were clear differences in the thickness of the surface coats between meronts and sporonts. Division was by binary fission of stages with two diplokarya (Fig. 101). The meront cytoplasm contained cisternae of rough endoplasmic reticulum and abundant ribosomes 123 Disporoblastic sporoqony Diplokaryotic sporonts were elongate cells with a 100-180 nm thick electron-dense surface coat, which was laid down first in patches on the plasma membrane, then as a complete coat (Figs. 102 - 104). After division of nuclei to form two diplokarya,cytoplasmic fission gave rise to two sporoblasts (Fig. 104). During their development (Fig. 105), the sporoblasts became elongate, the two nuclei, in tandem, occupied a position in the anterior half surrounded by cisternae of rough endoplasmic reticulum. There was a prominent reticulum, representing the Golgi apparatus at the posterior end. Several coils of the polar tube were seen during their synthesis from the Golgi network and a polar cap was seen at the anterior top. Spores were poorly preserved but it was possible to confirm the diplokaryotic condition and seen that there were 17 coils of the polar tube in one layer (Fig. 106). Octosporoblastic sporogony Sporonts of the octosporoblastic sequence were spherical. While still at the binucleate stage a second membrane began its separation from the plasma membrane to form a sporophorous vesicle (pansporoblast membrane). There was progressive incorporation of electron dense tubular structures between the plasma membrane and the sporophorous vesicle as the sporont passed through its maturation division (Figs. 107 - 111). Some sporonts appeared to be 124 uninucleate when the section passed through only one of the nuclei of the diplokarya but the early sporonts were known to be diplokaryotic from light microscopic observation. The plasma membrane of the sporont was progressively thickened by addition of electron dense material during its maturation (Figs. 110, 111). Paired structures resembling synaptonemal complexes (Fig..109), seen in some nulei and condensed chromatin in others (Fig. 108), were typical of meiotic configurations and indicated a reduction of chromosome number in the first division of each nucleus of the diplo- karyon. Constriction of the sporont gave rise to uninuc- leated sporoblasts (Figs. 110 - 112). The sporophorous vesicle membrane (Fig. Ill) was extremely fine and was not readily visible but persisted to retain the sporoblasts. The tubular structures within the sporophorous vesicle gradually decreased in amount (Fig. 112). During morpho genesis the sporoblasts were seen to possess a single nucleus with peripheral chromatin, a Golgi apparatus synthesising the polar tube, a posterior vacuole and polar cap and cisternae of rough endoplasmic reticulum (Fig. 112). Little could be seen in the poorly preserved mature spores, except that there were at least 18 coils arranged in bundles rather than a single layer as in the disporoblastic spores. (Fig. 113). 125 4.10.5 Attempt to induce dimorphism in Nosema bombycis at low temperature. N. bombycis from silkworm, Bombyx mori was transferred to H. zea larvae by per os, and the larvae were reared at 20° and 16°C. However, only diplokaryon sequence was observed as previously reported and an attempt to induce octosporogony sequence at low temperatures failed. 4.10.6 Light microscope observations on Nosema locustae in vitro S. fruqiperda cell cultures were infected by addition of haemolymph from infected H. zea, cultures were kept at 20°C and 25°C. Stages of merogony and of disporoblastic sporogony were seen in cell cultures but not the octosporoblastic sequence. The-earliest stages of the parasite were spherical with 2 nuclei in diplokaryon arrangement, (Fig. 114) measuring 3.7 + 0.6 pm. Subsequent stages were spindle shaped with one or two diplokarya and it was not possible to distinguish meronts from sporonts (Figs. 115 - 120). All stages divided by binary fission and ultimately sporoblasts developed into spores which measured 5.1 + 0.43 x 2.2 + 0.08 pm (Fig. 121). 4.10.7 Light microscope observations on Vairimorpha necatrix in vitro The cultures of £5. fruqiperda were infected by addition of haemolymph of H. zea containing infected blood cells. 126 Cultures were incubated at 20°C and 25°C. No octosporo- blastic sporogony was observed in the cell cultures at either temperature. All stages of the parasite were seen to have nuclei in diplokaryon arrangement. The youngest stages were rounded, measuring 2.1 + 0.2 pm (Fig. 122). Subsequent stages were elongate or spindle shaped and it was not possible to distinguish meronts from sporonts (Figs. 123 - 127) although ovoid stages with clear cytoplasm were thought to be meronts (Fig. 126) and spindle shaped stages were thought to be sporonts (Figs. 125, 127, 128). Spores were elongate ovoid and measured 5.7 + 0.35 x 2.7 + 4.7 pm. Stages were seen which had been released from heavily infected host cells into the medium. Some of these were seen closely attached to host cell surfaces (Fig. 129 arrow). 4.10.8 Phase contrast light microscope and electron micro scope observations of infective stages of N. heliothidis Numerous parasites were released from heavily infected cells into the medium (Figs. 130, 134). Among the abundant free spores, there were spherical diplokaryotic stages which were interpreted as infective stages (Figs. 132, 133). They were often seen closely attached to host cell surface (Fig. 131). A similar stage was seen by electron microscopy (Fig. 135), the parasite was closely attached to the host cell membrane. No ultrastructural differences were detected 127 between the free stages and intracellular meronts; both were cells with a simple plasma membrane and little internal cytoplasmic organization. No specific modifications were seen which could be associated with invasion and they were presumed to be taken into the host cell by phagocytosis. 4.11 Fate of P. operophterae inside the £3. frugiperda cells• 4.11.1 Labelling of secondary lysosomes in £3. frugiperda cells infected with P. operophterae After incubation of £3. frugiperda cells with sacchar- ated iron oxide or with ferritin for 1 h f secondary lyso somes were readily recognized by electron microscopy. Saccharated iron oxide gave superior results in labelling secondary lysosomes compared with ferritin. The cells retained a normal structure and labelling was restricted to the secondary lysosomes and pinocytotic vesicles (Fig. 136). When cells were infected with P. operophterae, the vesicles surrounding the vegetative stages remained-devoid of labelling with the marker (Figs. 137, 138), although the electron dense iron oxide particles were widely distributed in secondary lysosomes and pinocytotic vesicles. There was no fusion between secondary lysosomes and the vesicle membrane around the parasites. However, electron dense particles were seen in vesicles containing the spores (Fig. 139). The iron oxide particles were heavily deposited on the surface of spores. No marker was observed inside 128 vesicles as long as these contained vegetative stages as well as spores (Figs. 140, 141). 4.11.2 Detection of acid phosphatase in S • frugiperda cells infected with P. operophterae Detection of acid phosphatase was achieved by a modification o f .the method of Berka and Anderson (1962). Acid phosphatase was observed in S • frugiperda cells in small smooth vesicles, which were presumed to be primary lysosomes and in large vacuoles, which were presumed to be secondary lysosomes (Figs. 142, 143). The electron dense particulate reaction products of lead phosphate were easily seen in secondary lysosomes, lying in the vicinity of the vesicles containing sporonts and sporoblasts of P. operophterae, but no acid phosphatase marker was detected in the vesicles containing the pre-spore stages of the parasites. Acid phosphatase marker was only seen in the vesicles containing spores (Figs. 143 - 147). The electron dense particles were deposited over the spore surface. 4.12 Fate of N. heliothidis after uptake into £5. frugiperda cells Meronts of N. heliothidis lay directly in contact with host cell cytoplasm without an intervening host membrane. As multiplication progressed in the host cell, abundant tubular structures, which appeared to originate from the 129 parasite surface, extended into regions where the host cell cytoplasm was disorganized (section 4.10.1). Fig. 148 shows a stage enclosed by a second membrane, the host cytoplasm around the parasite has retained its integrity and organelles such as presumptive lysosomes, nucleus and mitochondria are intact. The parasite inside the vacuole appeared to have fewer ribosomes than normal meronts but its membrane structure appears normal. There were no tubular structures surrounding the parasite. When S. frugiperda cells were treated with ferritin to label secondary lysosomes, no labelled lysosomes were seen to have fused with N. heliothidis (Fig. 149). This is to be expected since the parasites are not surrounded by a host vacuolar membrane. Similar results were obtained with acid phosphatase labelled lysosomes (Fig. 150). Occasionally, a secondary lysosome was observed to have fused with a vacuole containing a nearly mature spore (Fig 151). The spore appeared to have been engulfed by a membrane, probably host endoplasmic reticulum, which enabled the lysosome to fuse with the enveloping membrane. 130 Figs. 40 to 48. Development of N. heliothidis in £3. frugiperda cell cultures at 25°C. The monolayers air dried methanol fixed and Giemsa stained. Scale bar = 10 pm. Figs. 40 to 44. Meronts (m) and sporonts(s) undergoing division by binary fission. Fig 45. Numerous vegetative forms released into the medium after breakdown of a host cell: some of these are thought to be infective stages (arrows). Fig 46. An infective form with diplokaryon nucleus closely attached to the surface of a cell (arrow head) . Figs. 47, 48. Numerous spores among which are sporonts and sporoblasts (arrow heads). 131 132 Figs. 49 to 57. Electron micrographs of stages of N. heliothidis. Fig 49. Early meront with apparently a single large nucleus: there is a simple plasma membrane abutting directly on the host cell cytoplasm. Scale bar = 0.5 pm. Fig. 50. £3. frugiperda cell, containing several well developed meronts, each with 2 nuclei in diplokaryon arrangement. The tubular structures (T) associated with the parasite membrane have begun to form. Scale bar = 2 pm. Fig. 51. Several meronts, now almost totally surrounded by tubular structures (T). Scale bar = 1 pm. Fig. 52. An early sporont showing deposition of the electron-dense coat on the plasma membrane at the poles, centriolar plaque (cp) at opposite poles of the two nuclei; dense structures in the nucleoplasm are possibly nucleoli (nu). Scale bar = 1 pm. 133 134 Figs. 53 to 55, Scale bars = 0.5 pm; Figs. 56, 57, Scale bars = 1 pm. Fig. 53. A sporont with 2 nuclei in diplokaryon arrangement, a large area of surface is covered by the electron dense coat; rough endoplasmic reticulum (rer); centriolar plaque (cp). Fig. 54. A sporont with complete surface coat. The rough endoplasmic reticulum is regularly arranged and there are a few ribosomes. Fig. 55. Binucleate sporoblast; polar tube (PT) partly within a membrane-bound structure containing tubules. Figs.56, 57 Immature spores with polar tube (PT) and membrane-bound tubular structures. The endospore (EN) is beginning to form in Fig•57. 135 136 Figs. 58 to 64 Development of Nosema sp. from Spodoptera litura in £3. fruqiperda cells cultures at 25°C. (Air dried, methanol fixed and Giemsa stained). Scale bar = 10 Jim. Figs. 58, 59. Meronts (m) s rounded or ovoid with diplokaryon nuclei. Figs. 60 - 63. Sporonts (S): spindle-shaped with central diplokarya; dividing sporont arrowed in Fig. 61. Fig. 64. A heavily infected cell with numerous spores and sporoblasts (sp). 137 138 Figs. 65 to 70. Electron micrographs of stages of Nosema \ sp. from S. lutura in S. frugiperda cell cultures. Scale bars = 1 pm. Figs 65, 66. Elongate meronts with 2 nuclei (N) in diplokaryon arrangement. Fig. 67. A meront surrounded by regularly arranged- - host ribosomes. Fig. 68. A sporoblast, with 2 nuclei (N) in diplokaryon arrangement, has a thick plasmalemma and the cytoplasm contains endoplasmic reticulum in regular arrangement• Fig. 69. Sporoblast: one of the 2 nuclei (N) is visible? there is an electron dense surface coat and concentrically arranged rough endoplasmic reticulum. Fig. 70. A mature spore with a diplokaryon nucleus, closely-packed membranes of the polaroplast and coiled polar tube. 139 140 Figs. 71 to 78. Development of P. operophterae in S. fruqiperda cell cultures at 20°C (Air dried methanol-fixed and Giemsa-stained). Scale bar = 5 jam Fig. 71. Uninucleate meront. Fig. 72. Uninucleate meronts and division stage (arrow heads)• Fig. 73. Large binucleate meronts. Fig. 74. The sporonts dividing within a prominent vesicle (arrow heads). Fig. 75. Numerous sporoblasts each with a single nucleus lying within a vesicle (arrow head)• Fig. 76. Clusters of spores and numerous separate meronts, each with a single nucleus. Fig. 77. Presumptive infective forms closely applied to the host cell surface (arrows head). Fig. 78. Numerous mature spores replacing large areas of host cytoplasm. 141 142 Figs. 79 to 86. Electron micrographs of P. operophterae in S. frugiperda cell cultures. Figs. 79-83, Scale bars = 0.5 pm? Fig. 84, Scale bar = 1 pm? Fig. 85, Scale bar = 2 pm? Fig. 86, Scale bar = 0.25 pm. Fig. 79. Meront with a single large nucleus. There is a simple plasma membrane (arrow head) and a closely- fitting thickened vacuolar membrane surrounded by several host mitochondria. Fig. 80. Meront with a single nucleus lying within a vacuolar membrane (arrow head). Numerous small vesicles (sv), which may be protrusions from the main vacuole, lie close to the parasite. Fig. 81. Sporont with a single large nucleus in the plane of section contracting in the vesicle. An electron dense surface coat is added in parts to parasite's plasma membrane (arrow heads). Fig. 82. Two meronts each lying in its own vacuole, possibly just after separation. Note the place where the vacuolar membranes may leave just complete separation (arrow), n = nucleus m = mitochondria v = vesicle er = endoplamic reticulum sv = small vesicle 143 144 Fig. 83 Formation of sporoblasts, some free stages attached to the sporophorous vesicle membrane. Fig. 84. Sporoblasts and spores within sporophorous vesicle. Note that the free stages have become spores while the attached phases may still be dividing. Fig. 85. A heavily infected cell with its cytoplasm largely replaced by several sporophorous vesicles, containing numerous spores. Fig. 86. A mature spore with a single nucleus showing polar tube and polaroplast. m = mitochondria n = nucleus pt = polar tube ps = polar sac ap = anterior polaroplast - close-packed membranes, pp = posterior polaroplast - loosely packed membranes. lf35 146 Fig, 87 to 99. Development of N. locustae at 25°C and 20°C from H. zea larvae. (Air dried methanol- fixed and Giemsa stained). Scale bar = 10 pm. Fig. 87. A meront with 2 nuclei. Fig. 88. Binucleate meronts: the isolated nuclei in one of them is probably an artifact of the smearing technique. Fig. 89. Two diplokaryotic meronts possibly just after their division. Fig. 90. Fusiform binucleate sporont. Fig. 91. Fusiform tetranucleate sporont. Fig. 92. Fusiform sporont with diplokarya at the poles. Fig. 93. Cytoplasmic cleavage of a sporont. Fig. 94. Two sporoblasts. Fig. 95. Free elongate spores. Fig. 96. Rounded binucleate sporont of the octosporoblastic sequence. Fig. 97. Rounded tetranucleate sporont. Fig. 98. Octonucleate sporont within a sporophorous vesicle. Fig. 99. Eight spores within a sporophorous vesicle. 147 98 99 148 Figs. 100 to 113. Electron micrographs of N. locustae from H. zea larvae. Figs. 100-104/ Scale bars : 1 urn? Figs 105/ 106/ Scale bars = 0.5 pm* Fig. 100 Meront with 2 nuclei in diplokaryotic arrangement. The cytoplasm is surrounded by a plasmalemma with some electron dense surface addition. Fig. 101. Division of meront into diplokaryotic products: cytoplasmic cleavage is almost complete. Fig. 102. Elongate disporoblastic sporont/ with 2 nuclei in diplokaryotic arrangement; the surface coat is thickening. Fig. 103. Transverse section of a disporoblastic sporont: the plasma membrane is thickened with electron-dense material in patches. Fig..104 . Sporont with thickened surface membrane undergoing fission into sporoblast: a spindle plaque (sp), seen in one nucleus (arrow head), is probably persisting from the last nuclear division. Fig. 105. Binucleate sporoblast: PC = polar cap; pt polar tube; G = Golgi network; rer = rough endoplasmic reticulum. Fig. 106. Mature diplokaryotic spore, with about 16 coils of the polar tube (PT) in a single layer. 6 17! 150 Figs. 107-112, Scale bars = 1 um; Fig. 113, Scale bars = 0.5 }im. Figs. 107. Early stage of octosporoblastic sporont. Note the appearance at the surface of electron dense tubular structures an additional membrane (arrow heads) which forms the sporophorous vesicle. Fig. 108. Early octosporoblastic sporont with masses of condensed chromatin in the nuclei. The sporophorous vesicle is clear and the dense tubular.structures are abundant (arrow head)• Fig. 109. Octosporoblastic sporont with extensive development of the dense tubular structures: paired structures resembling synaptonemal complexes (sc) are seen in the nucleus (N). Fig. 110. Constriction of a sporont of the octo sporoblastic sequence: 3 nuclei are visible and the surface membrane is thickened. Fig. 111. Almost complete separation of the sporo- blasts from a sporont of the octosporo blastic sequence. In four of the lobes, the single nuclei (N) of the future sporoblasts are seen. Fig. 112. Separate sporoblasts held together in the sporophorous vesicle. In one of them the primordia of the spore organelles can be seen. PC = polar cap PV = posterior vacuole N = nucleus Fig. 113. Mature spore of the octosporoblastic sequence; the polar tube (pt) coils are clumped. 151 152 Figs. 114 to 121. Development of N. locustae in £3. frugi- perda cell cultures at 20°C (Air dried, methanol-fixed. Giemsa-stained). Figs. 114 to 120, Scale bars = 10 pm; Fig. 121, Scale bar = 5 pm. Fig. 114. Spherical diplokaryotic meronts. Figs. 115-120. Elongate, spindle-shaped stages with one or two diplokarya, which may be meronts or sporonts. Smaller rounded stages (arrow head) may be recently invaded stages. Fig. 121. Numerous spores and pre-spore stages. 15 3 154 Figs 122 to 129. Development of V. necatrix in S3. fruqi- perda cell cultures at 20°C. (Air-dried, methanol-fixed and Giemsa-stained). Scale bar = 10 pm. Fig. 122. Early stage on entry into cell, probably dioplokaryotic. Figs 123-127. Ovoid and spindle-shaped diplokaryotic stages which may be meronts (e.g. Fig 126) or sporonts. Division is by binary fission. Fig. 128. A heavily infected cell with spindle shaped stages, probably sporonts or sporoblasts (arrow heads) and spores. Fig. 129. Stages released from cells into the medium. Some of them are closely attached to a new cell surface (arrow heads). 155 156 Figs. 130 to 133. Phase contrast microscopy of N. helio- thidis in S. frugiperda cells. Figs. 134 to 135. Electron micrograph of N. heliothidis in S. frugiperda cell cultures. Figs. 130. S. frugiperda cell culture heavily infected with N. heliothidis: numerous spores and other development stages have been released from infected cells into the medium. The spherical shaped parasites were thought to be infective forms (arrow heads). Scale bar = 10 pm. Fig. 131. A presumptive infective form, seen closely attached to the S. frugiperda cell surface. Scale bar = 5 um. Figs. 132, 133. Diplokaryotic presumptive infective forms free in the medium. Fig 132, Scale bar = 2.5 um; Fig. 133, Scale bar = 5 um. Fig. 134. Electron micrograph showing a host cell, disrupted and liberating spores and pre-spore stages. Scale bar = 1 um. Fig. 135. A presumptive infection form, closely attached to the host cell membrane. Scale bar = 1 urn. 157 158 Figs. 136 to 141. Electron micrographs of £3. fruqiperda cells infected with P. operophterae labelled with saccharated iron oxide. Fig. 136. Secondary lysosomes and small vesicles containing granules of saccharated iron oxide. Scale bar = 1 pm. Fig. 137. Vesicles presumptive secondary lysosomes labelled with saccharated iron oxide, close to but not fused with a vacuole containing a vegetative stage of P. operophterae (arrow head). Scale bar = 0.5 pm. Fig. 138. Vesicles, presumptive secondary lysosomes, labelled with saccharated iron oxide, close to a vacuole containing a sporogonic stage of P. operophterae, but there is no fusion. Scale bar = 0.5 pm. Fig. 139. A vacuole containing mature spores of P. operophterae : the vacuolar membrane is labelled with saccharated iron oxide. Scale bar = 1 pm. Fig. 140. A vacuole containing vegetative stages as well as spores of P. operophterae: there is no fusion of the saccharated iron oxide labelled secondary lysosome with the vacuole (SLV). Scale bar = 1 jam. Fig. 141. A vacuole containing different stages of P. operophterae: a small vesicle (arrow head) a presumptive seocndary lysosome labelled with saccharated iron oxide, lies close to but has not fused with the parasite vacuole. Scale bar =0.5 pm. 159 160 Figs. 142 to 147. Electron micrographs of S. frugiperda cells infected with P. operophterae stained for acid phosphatase. Figs. 142 to 146, Scale bars = 1 pm; Fig. 147, Scale bar = 0 • 5 pm. Fig. 142. A vesicle heavily stained for acid phosphatase close to a vacuole containing sporogonic stages of P. operophterae? no fusion has occurred. Fig. 143. A vacuole containing spores on the surface of which is deposited the reaction products, indicating acid phosphatase. Fig. 144. A spore on the surface of which is deposited the particles of reaction product, indicating acid phosphatase. Fig. 145. A large vacuole labelled with acid phosphatase reaction particles; it contains several spores. Small vesicles with phosphatase staining reaction particles lie near the vacuole (arrow heads). Fig. 146. Spores of P. operophterae within a large vacuole which also contains the reaction products indicating acid phosphatase. Fig. 147. High magnification of spores of P. operophterae in a vacuole: the spores are heavily encrusted with the reaction particles, indicating acid phosphatase. 161 v *♦ • -n r * , •/*' .. Si 'V* > •. . & * ■+' - ‘ A*- * .* •', • I------< • * ■ h- *■ "r*” f .a** ate*** 147 162 Fig. 148 to 151. Electron micrographs of N. heliothidis. Fig. 148, Scale bar = 0.5 pm; Figs. 149 to 151, Scale bars = 1 pm. Fig. 148. A diplokaryotic stage of N. heliothidis enclosed in a closely fitting vacuolar membranethe parasite has fewer ribosomes than normal in the cytoplasm. A large vesicle, presumptive lysosome lies near the parasite. Fig. 149. A stage of N. heliothidis lying directly in the host cell cytoplasm. A number of vesicles slightly stained for ferritin lie near the parasite (arrow head). Fig. 150. A vesicle containing the reaction product indicating acid phosphatase close to a vegetative stage of N. heliothidis : no fusion has occurred. Fig. 151. A spore apparently within a lysosome the parasite appears to have been engulfed. 1 (j 3 164 5. DISCUSSION 5.1 Microsporidia in cell cultures 5.1.1 Establishment of microsporidia in cell cultures. A number of microsporidian species have been established in cell cultures. In most cases, spores have been used as the inoculum to infect the cells (Higby et al., 1919, Kurtti & Brooks, 1976? Streett et al•, 1980; Smith et al., 1982; Undeen, 1975; Ishihara, 1966; Kawarabata & Ishihara, 1984). However, the prerequisites are: (a) preparation of highly purified and sterile spores, and (b) activation of spores for germination in the vicinity of cells, so that the sporoplasm is inoculated into the cytoplasm through the polar tube. In many cases, the absence of reliable methods to induce spores to hatch and difficulties in making suitably purified preparations of spores, which do-not contaminate the cultures, have handicapped the establishment of microsporidia in cell cultures. Some workers have established cell lines persistently infected with microsporidia, from host tissues which were naturally infected with the parasites. (Sohi & Wilson, 1976; Bayne et aJL. , 1975; Shadduck, 1969). However, the establishment of a cell line from a primary culture of host tissues is a lengthy process often requiring 6 to 9 months of effort. The parasites might even overgrow the cultures and destroy them before the cell line has been firmly established. 165 Trager (1937) successfully infected primary ovarian tissue cultures of silkworms, B. mori by inoculating the haemolymph of silkworm larvae infected with N. bombycis. Although he could not repeat the experiment, apparently because the number of parasites in the haemolymph was too low to establish further infections, this was the first demonstration that the haemolymph from hosts infected with microsporidia could be used as inocula to introduce micro- sporidia into cell cultures. This method has not been fully exploited for the establishment of microsporidia in inverte brate cell cultures. In the present experiments, 5 species of microsporidia, derived from insect hosts, have been successfully introduced into uninfected S. frugiperda cell cultures, by addition of the haemolymph from previously infected lepidopteran larvae. This has proved to be a reliable and convenient method which obviates the two difficulties associated with setting up the cultures. It is likely that this method could be applied to many other microsporidia derived from invertebrates, if enough haemolymph can be obtained. However, it is essential that the haemolymph contains either extracellular infective stages or infected haemocytes. If the microsporidia do not infect the haemocyte and the fat body, and do not release the stages into the haemolymph but are specific to other tissues, infection cannot be established by this method. This is the case with P. schubergi which only infects the gut of its hosts. Curiously, cell cultures did not become 166 infected with N. eurytremae when heavily infected locust fat body cells were implanted into cell lines. The authors speculated that fat body cells contained only spores, which failed to hatch in the cell medium (Higby et al.., 1979). 5.1.2 The effect of temperatures on microsporidia. Apart from microsporidia parasitizing homoeothermic hosts, such as E. cuniculi in mammals and N. helminthorum, a parasite of tapeworms in sheep, which can survive at 37°C, (Waller, 1975? Canning & Gunn, 1984), microsporidia apparently can only multiply below 37°C. Ishihara (1969) first reported that no multiplication of vegetative stages of N. bombycis occurred in cells, when the cultures were incubated at 37°C. Undeen (1975) provided further evidence to support this view, when he showed that N. algerae, a parasite of mosquitoes, cannot complete its life cycle in cell cultures at 37°C. Similarly, Smith et al. (1982) found that N. alqerae failed to develop in cultures, incubated at 38°C. In the present study the 5 species of microsporidia, derived from insect hosts, all failed to grow in S. frugiperda cell cultures incubated at 37°C. These observations indicate that microsporidia, derived from poikilothermic hosts, are unlikely to infect homoeothermic vertebrates since the temperature provides a natural barrier. The mechanism which prevents the microsporidia of poikilothermic hosts from surviving at 37°C is not understood but may be due to the absence of heat shock 167 genes. Heat shock genes have been demonstrated in some parasitic protozoa e.g. Trypanosoma cruzi and Leishmania major (Van der Ploeg, Giannini & Cantor, 1985); the heat shock genes are triggered when subjected to a temperature stress, as when the parasites are transferred from the non-temperature regulated insect vector to the temperature-regulated mammalian host. The result is an adaptation to life in mammalian host tissue. However, the tolerance to temperature below 37°C varies according to species of microsporidia. N. algerae was observed growing in cultures at 34°C or 35°C (Smith et al., 1982, Undeen, 1975), but Wilson & Sohi (1977) found that N. disstriae was completely eliminated from the cell cultures when exposed to 35°C for 28 days. As expected, in the present experiments N. locustae was well adapted to growth at a high temperature (30°C) in £5. frugiperda cells since their original hosts, locusts, survive best at 30° - 35°C. . In contrast, P. operophterae has evolved to live at lower temperatures (20°C).since their hosts, the winter moth, Operophtera brumata, are adapted to living between 15°C and 25°C. With an increase in temperature beyond 30°C, there is complete suppression of the growth of P. operophterae in £5. frugiperda cells. Armstrong (1976) observed that the optimal temperature for survival of insect hosts is also the optimal temperature for growth and development of its parasites. Present results support this suggestion. This may be of signific 168 ance in limiting microsporidia in their host range and geographic distribution. 5.1.3. The effect of antibiotics on microsporidia Present studies have shown that antibiotics routinely added to culture medium, at concentrations designed to minimize contamination (100 i.u. or 100 jig per ml), have no adverse effects on the growth and development of micro sporidia. This has been shown previously in N. disstriae (Sohi & Wilson, 1979) and E. cuniculi (Shadduck & Polley, 1978). However, when concentration of antibiotics was increased to 500 i.u. per ml or 500 pg per ml, some of the species of microsporidia tested were affected. These concentrations had a slight effect on the infectivity of Nosema sp. so that there was reduction in the percentage of cells infected but N. heliothidis was not affected. No attempt has been made to determine.the mode of action of antibiotics on microsporidia. But it has been found that fumagillin arrested the DNA replication of Nosema apis by autoradiography (Hartwig & Przelecka, 1971). Although fumagillin and benomyl did not eliminate N. disstriae and a Pleistophora-like microsporidium from cell cultures, when applied for more than 12 days (Kurtti & Brooks, 1977; Bayne et al. , 1975). Sohi & Wilson (1979) found that N. disstriae was completely eliminated from cell cultures, when treatment with these drugs was extended to 35 days. 169 5.2 Cell to cell transmission of microsporidia. Since the early days of microsporidian studies, authors have speculated on the mechanism of cell to cell transfer. The development of jji vitro techniques for microsporidia have provided an easier system to study this process, than in vivo. Evidence is accumulating that stages can survive extracellularly.long enough to be responsible for transmission from cell to cell. Ishihara (1969) studied the development of N. bombycis in cultures of ovarian cells of silkworms and observed that forms, similar to sporoplasms but larger, leave the host cells, migrate and invade new cells. He named these "secondary infective forms". Kurtti & Brooks (1977) also reported "secondary infective forms', but neither Ishihara nor Kurtti and Brooks were able to determine how the "secondary infective forms" could leave their host cells and invade others. Sohi & Wilson (1976) studied cell cultures infected with N. disstriae and - confirmed the presence of infective* forms. They showed that heavily infected cells disintegrated and developmental stages including infective forms burst out into the medium. They suggested that the infective forms invaded host cells by random contact with them. However, Kawarabata & Ishihara (1984) claimed that infective forms of N. bombycis were able to emerge into the medium without destroying the host cells and were immediately able to invade other host cells. They did not, however, explain exactly how the infective forms emerged nor how they invaded other cells. 170 Recently, Kurtti et al. (1984) suggested that infective stages were extruded in vesicles from infective cells by exocytosis. They found numerous developmental stages of N. disstriae encased within fragments of disintegrating cell cytoplasm, rich in mitochondria and membranes, but they did not determine whether these parasite-containing vesicles could fuse with uninfected cells and cause the infection. In the present studies, several pieces of evidence have been provided, which support the view that extracellular- infective stages are responsible for spread of micro- sporidian infection from cell to cell. Five species of microsporidia were established in continuous cell cultures by inoculation of infective haemolymph or infective cells in haemolymph. As the parasites had to pass from the original insect cells to the tissue culture cells. Free stages must have been involved in transmission of infection from one cell type to another... During development of the micro sporidia in £5. fruqiperda cell cultures, the initial level of infection of about 10% increased to 60-100% within 8 to 12 days, according to species. This increase is unlikely to have been due to division of infected cells because uninfected cells would have been dividing at the same rate and the proportions of infected and uninfected cells would have remained approximately the same. Futher the increase in level of infection cannot have been due to spores hatching in the presence of cells, because spores have never been seen hatching in culture medium. Results obtained by 171 Smith et al. (1982) with N. algerae and N. eurytremae also suggest that spores are not responsible for spread of infection; although 70-90% of spores hatched in the culture medium after pretreatment/ only 0.1-0.9% cells became infected. Since the contact of spores with monolayers was random, this lead the majority of sporoplasms to fail to penetrate into cells through the polar tubes. Weidner & Trager (1973) with N. mithaelis and Scarborough-Bull & Weidner (1985) with G. hertwigi showed that sporoplasms can survive at least 40 min in the presence of ATP in the medium. In the present studies, addition of ATP to the culture medium prolonged the survival of infective stages and resulted in a greater number of cultured cells becoming infected with Nosema sp. and N. heliothidis. Furthermore, when vegetative stages of the 5 species of microsporidia were separated from spores by differential.centrifugation and were added immediately or after various intervals to uninfected cultures, infections were obtained but'the infectivity declined sharply with time and was almost zero at 30 min to 180 min after isolation. This dempnstrated that infective stages other than spores (which are long-lived) were responsible for the infections. Further evidence was provided by light and phase contrast microscopic observations of the microsporidia, in which parasites were seen closely attached to the host cell surface. An identical result was obtained by electron microscopy, when the infective stages were observed closely attached to the host cell membrane. 172 Many authors, who have studied microsporidia iji vivo, have held the view that microsporidian cell to cell transfer is by non-spore free infective stages. Ohshima (1975), studying silkworms infected with N. bombycis, found that small round parasites, larger than sporoplasms were present in haemolymph. He separated these stages, injected them into uninfected.larvae and obtained infections proving that they were infective. He suggested that they were released from broken infected host cells. Andreadis (1983) and Hazard (1984) using Amblyospora sp. from mosquitoes and Hazard et al. (1984), using Culicosporella lunata from Culex pilosus, found that meront stages developed and multiplied in haemocytes and caused the cell to become greatly hypertrophied and to rupture releasing many diplokaryotic stages into the haemolymph. Those immediately migrated and invaded the fat body. Cali & Briggs (1967) and Abe (1978) provided evidence to show that Nosema spp. in Coccinella septempunctata and in silkworms B. iftori developed in haemocytes producing stages, which were released into the blood and spread the infection to other tissues. Abe & Fujiwara (1979) reported similar observations for a Pleistophora sp. in Bombyx mori larvae. They found that round binucleate "secondary infective forms" transferred the infection from cell to cell. In contrast, Avery & Anthony (1983) reported that some mature spores of N. algerae extruded their polar tubes in situ, inside cells of the host in which they were produced. 173 Although they commented that the hatching of these spores in situ may have been caused by fixation, they considered that spread of infection by N. algerae in vivo was effected both by hatching of spores and by sporonts release from heavily infected cells. There is evidence that diplokaryotic spores of Amblyospora sp.from mosquitoes, as opposed to the uninucleate octospores, hatch within the host in which they are produced as a normal procedure (Andreadis, 1983). Diplokaryotic meronts infect oenocytes of the mosquito larvae. In male larvae, they develop rapidly and spread the infection to cells of the fat body, where eventually 8 haploid spores are formed from diplokaryotic sporonts. In female larvae the infection remains restricted to oenocytes and no sporogonic development takes place until the female hosts become adults and take a blood meal. At this stage sporulation is initiated and results in the formation of diplokaryotic spores in the oenocytfes. These spores regularly extrude their sporoplasms through the polar tubes and these sporoplasm infect host oocytes and hence the infection is passed to the next generation. In the present studies no evidence was found that spores hatch within host cells. Tsang et al. (1982) reported that they were unable to transfer N. disstriae from and H. zea cell line to a B. qerminica cell line grown in UMN-B1 (cockroach) medium. They thought that the medium was unfavourable for survival 174 of extracellular infective stages. When the cockroach cells were adapted to a moth cell culture medium, cross infection was achieved and the B. germanica cell line became infected. Spores, which could survive extracellularly, even in the presence of UMN-Bl medium, were thus discounted as the source of infection to new cells and only the stages, which could be affected by the medium were thought to be involved in spread of infection. In summary the bulk of evidence supports the view that infective stages, which are released from disrupted heavily infected cells, are normally responsible for cell to cell transfer. However, some spores may be induced to hatch in situ by certain mechanisms, possibly some nutritional feed-back or hormonal trigger. Spore hatching is likely to be of regular occurrence in certain life cycles, such as the diplokaryotic spores which are responsible for transovarial transmission of Amblyospora. In adult female mosquitoes ... . after the blood meal, there is an increase in ecdysone level (Lord & Hall, 1983) which suggests that the host's hormones may control the hatching process. 5.3 The ways of entry of infective stages of microsporidia into cells. It is generally accepted that successful entry of most parasitic protozoa into their host cells is dependent on the presence of appropriate surface receptors on the host cells and on ligands on the surface of parasitic protozoan which 175 bind to those receptors. The receptor-ligand systems of host-parasite interactions can be divided into immunological ligands and non-imraunological ligands. However, little work has been done on the receptors for invertebrate parasites and their host cells. It is known that invertebrate 3 phagocytes do not have immunological receptor such as the C and Fc receptors known in vertebrate immunity systems (Anderson, 1976). This is obvious because invertebrates do not produce immunoglobulins. Several studies have shown that attachment of parasitic protozoa derived from vertebrates to host cells can be inhibited by a range of lectins or by the specific monosaccharides that constitute cell surface carbohydrates. This indicates that surface carbohydrates are involved in the invasion process (Bray, 1983; Dawidowicz, et aj^. , 1975; De Araujo Jorge, & De Souza 1984). The present results show that infective stages of microsporidia require an attachment to.the host cell, by.way of specific receptor-ligand binding/ before they can enter into host cells. Thus, there is a need for ligands which bind to glycoprotein or glycolipid receptors via terminal saccharides. Those interactions may vary with different kinds of host cells. Not surprisingly, the receptor-ligands were found to be distinct from species to species, suggesting that they may serve as specific markers for diversity of microsporidia. Previous studies have shown that the receptor-ligands for other parasites also vary from species to species (Dwyer, 1977), strain to strain (Arauju 176 et al., 1980) and even differ between various development stages, in order to adapt to different tissues or hosts (Arauju et al., 1980). This demonstrates a high degree of evolutionary adaption of the parasite to the host. The results presented here may also partly explain the phenomenon of tissue specificity in microsporidian infections. Differences in receptors or ligands between parasites and host cell types may result in different abilities to enter host tissues. Although the present study has not demonstrated the saccharides on the surface of the parasites by electron microscopy, a recent study using peroxidase-labelled ConA has shown that D-mannopyranose and D-glucopyranose are present on the surface of sporoplasms of G. hertwigi (Scarborough-Bull & Weidner, 1985). Host cell entry by a parasite involves a distinct and essential cellular event, that of internalization of the parasite within^a:parasitophorous vacuole. Three possible mechanisms for this process have been proposed. These have been summarized by Blackwell & Alexander (1983), namely "zipper process", "modified zipper process" and "capping process". Obviously, different parasites employ different mechanisms for entering host cells. In studies of micro- sporidia dji vivo, phagocytosis as a major means of spread of microsporidian infections has been suggested by Weiser (1976), Cali & Briggs (1967) and Laigo & Paschke (1966). They observed that the infective stages of microsporidia were circulated in haemolymph and were phagocytosed by 177 haemocytes or cells of the target tissues. The results presented here demonstrated that Cytochalasin B inhibited the entry of infected stages of N. heliothidis into S3. frugiperda cells, and that the concentrations of Cytochalasin B, that produced increasing inhibition of phagocytosis of red blood cells, was similar to that which resulted in increasing inhibition of entry of infective stages of N. heliothidis into those cells. This implicates phagocytosis as the major means of uptake of infective stages of N. heliothidis by S3, frugiperda cells. In studies of Leishmania mexicana by Alexander (1975), Trypanosoma cruzi by Nogueira & Cohn (1976) and by McCabe ^t al. (1984), using Cytochalasin B, the authors came to similar conclusions that endocytosis is a primary mechanism by which parasites enter into host cells. At the electron microscopic level, no special organelle which may be responsible .for-penetration . of host cells can be seen in the infective stages of N. heliothidis. There is nothing corresponding to-the apical complex of organelles which is used to initiate invagination of host cells, in such protozoa as Babesia, malaria parasites and Toxoplasma. This suggests that the infective stages of microsporidia enter into host cells by the so-called "zipper process" (Silverstein, 1977), characterized by interaction of receptors on the phagocyte with ligands on the parasite. An electron micrograph of an infective stage of N. heliothidis located inside a well-fitted parasitophorous vacuole further 178 confirmed this suggestion. Undoubtedly the process was without active participation of the parasite, and is dependent on host cell energy. 5.4. Fate of microsporidia within cultured host cells. It is known that the mechanisms by which the phagocytes of mammals eliminate microorganisms involved the triggering of an "oxidative burst". This leads to the generation of toxic metabolites of oxygen and fusion of lysosomes with the parasitophorous vacuoles (Mauel, 1984). However, very little is known about invertebrate phagocyte killing mechanisms and the survival mechanism of intracellular parasites. The results of the present experiments suggest that microsporidia may employ two mechanisms by which they avoid destruction by lysosomes and survive in host cells after invasion. Observation of S. frugiperda cells infected with P. operophterae, containing labelled primary and secondary lysosomes, very clearly shows that lysosomes readily fuse with parasitophorous vacuoles only when they contain spores. Lysosomes did Jiot fuse with vacuoles containing vegetative stages, indicating that vegetative stages of P. operophterae can inhibit phagosome-lysosome fusion. Prevention of lysosome fusion with the parasitophorous vacuole has also been observed in T. gondii (Jones & Hirsch, 1972) and E. cuniculi (Weidner, 1975). The mechanism of inhibition is 179 unclear. Ultrastructural evidence suggests that P. operophterae alters the structure of the parasitophorous vacuoles, allowing endoplasmic reticulum and mitochrondria to surround the vacuolar membrane. This, perhaps, may be the event which prevents the lysosomes having access to the parasitophorous vacuole. It seems that the nature of the surface of the parasites may also affect the fusion of parasitophorous vacuoles with competent lysosomes. It has been demonstrated that when T. gondii was coated with specific (heat- inactivated) antibody, lysosomal fusion with the parasito phorous vacuole containing the T. gondii readily took place phagosome/lysosome (Jones & Hirsch, 1972). On the other hand, inhibition of . A fusion in l . mexicana could be obtained by incubating macrophages with poly-D-glutamic acid (Alexander, 1981). It is generally accepted that the phagosome’s endogenous acidification characteristically begins shortly after their formation and before their fusion with lysosomes. More recently, it has been shown that the acidification can be blocked by live T. gondii which are higly susceptible to acidic pH within the vacuole. (Sibley, Weidner & Krahebuhle, 1985). This suggests that the parasites within the vacuole interfere in some way with phagosome acidifica tion. Weidner & Sibley (1985) found that in macrophages, fusion of lysosomes with phagosomes containing Glugea spores was blocked, and that the Glugea phagosomes also failed to 180 acidify. Fusion did occur when cationized ferritin was used to mask the spore surface which suggested that the anionic molecules on the spore surface of Glugea may partly be responsible for initiating the inhibition. The inhibition of lysosomal fusion with a parasito- phorous vacuole containing vegetative stages of P. operophterae might employ a similar mechanism. However, as indicated above, inhibition of lysosomal fusion disappeared when the vegetative stages of P. operophterae within the vacuole developed into spores. This implies that some factors, perhaps similar to the anionic component on the surface of Glugea spores are either on the surface of vegetative stages of P. operophterae or secrected into the vacuole. It might be important for the vegetative stages of P. operophterae to be in close proximity with the vacuolar membrane, if the properties on the parasite surface are significant in controlling lysosomal fusion. The occurrence of lysosome-phagosome fusion when parasitophorous vacuoles containing immune serum-treated E. cuniculi provides supportive evidence (Niederkorn & Shadduck, 1980), that the factor preventing lysosomal fusion may be blocked by antibodies or complements, thereby allowing lysosomes to fuse with the parasitophorous vacuoles. Electron micrographs of N. heliothidis show early infective stages within a closely fitting vacuolar membrane which is believed to have originated from the host plasma membrane by invagination. The Nosema parasites must escape 181 from the phagocytic vauole rapidly as the parasites are almost invariably found in direct contact with host cell cytoplasm and multiply in the cytoplasmic matrix. This is undoubtedly of significance to parasites in their evasion of destruction within phago-lysosome. It has been found that T. cruzi and Sarcocystis cruzi are capable of escaping from their parasitophorous vacuoles into the host cell cytoplasm (Kress et al., 1975? Nogueira & Cohn, 1976? Dubey et al. , 1980), but the mechanism by which the parasites cause the vacuolar membranes to disintegrate is obscure. It has been suggested, in the case of T. cruzi, that the host cell vacuolar membrane is disrupted by a parasite-released factor (Nogueira & Cohn, 1976). The process of escaping from the vacuole into host cytoplasm represents a strategy of survival within the potentially hostile host cell environment. In the present studies, it is interesting to note that some of the nearly mature spores of ‘N. heliothidis were fused with a presumptive secondary lysosome,. and a similar phenomenon was observed in the vacuoles containing spores of P. operophterae. It is possible that parasitophorous vacuoles, containing mature spores, become targets for lysosomal fusion and subsequent digestion of the spores, because spores have lost the ability to secrete intermediate metabolic products into the cytoplasm or the vacuole. The fusion of parasitophorous vacuoles containing nearly mature spores with lysosomes may be beneficial and aid the 182 formation of the spore wall. It has been observed that immature spores of N. heliothidis become chitinized after they were phagocytosed by amoebae (unpublished observation). Possibly some enzymes within the amoebic lysosomes promoted the synthesis of chitin, which is a major chemical component of the spore wall. 5.5 Dimorphism and meiosis The occurrence of dimorphism in microsporidia of lepidoptera was first suggested by Maddox (1967)/ who suspected that Thelchania diazoma and Nosema necatrix were dimorphic forms of the same species in Pseudaletia unipuncta. This was subsequently confirmed by Fowler & Reeves (1974a) and a new genus Vairimorpha was described, to which the species was transferred as Vairimorpha necatrix (Pilley, 1976). This genus exhibits two sporogonic sequences at low temperatures.Since then more Nosema .. species from lepidoptera have been.transferred to the genus Vairimorpha. (Maddox & Sprenkel, 1978; Malone, 1984b? ' Streett & Briggs, 1982). A parasite, Burenella dimorpha, which is similar to the genus Vairimorpha, was described from a tropical fire ant, Solenopsis geminata (Jouvenaz & Hazard, 1978). The authors believed that Burenella differed from Vairimorpha because of tissue specificity. The disporoblastic sequence developed in hypodermal tissue and the octosporoblastic sequence developed in the fat body. In Vairimorpha, both types of spores develop in the fat body. 183 Additionally, pansporoblast membranes of Burenella are not persistent, while those of Vairimorpha are highly persistent. Nosema locustae was first described by Canning (1953) from the African migratory locusts Locusta migratoria migratorioides, which were reared at temperatures varying between 21°C and 35°C daily. Subsequently, N. locustae has been shown to infect 58 species of Orthoptera (Canning, 1962; Henry, 1969), but dimorphism was not observed. Henry et al. (1979) failed to infect H. zea larvae by inoculating 4 the larvae with a low dose of N. locustae (10 spores per larva). In the present study, H. zea larvae were success fully infected with N. locustae, using a higher dose of Q spores (10 spores per larva), and it was shown that N. locustae exhibited the dimorphic sporogonic sequences characteristic of the genus Vairimorpha. Both sporogonic sequences occurred in;the fat body of H. zea at: 20°CV . It is, thus, necessary to transfer N . .Iocustae to the genus Vairimorpha. The species should be known as Vairimorpha locustae (Canning, 1953). This finding suggests that dimorphism of the Vairimorpha type may be commoner than previously suspected and that many more species of Nosema will ultimately have to be transferred to the genus Vairimorpha once the conditions which initiate the octo- sporoblastic sequence are known. So far dimorphism and meiosis have not been demonstrated in N. bombycis the type species of the genus 184 Nosema, nor in many other Nosema species, which have been studied by electron microscopy. Although there are several reports that uninucleate stages occur in meronts of Nosema (Ohshima, 1973; Weiser, 1961; Canning et al., 1983), this does not constitute evidence of nuclear fusion. It has been » suggested that karyogamy and meiosis in Nosema occur in unobserved parts of the life cycle, or are absent (Loubes, 1979). In the present studies, H. zea were infected with N. bombycis and reared under the same temperature conditions as H. zea infected with N. locustae (20°C). The N. bombycis did not exhibit the octosporoblastic sporogonic sequence. It is possible that in N. bombycis dimorphism may occur in alternative hosts or under conditions which are not yet known but at present there is no reason to believe that N. bombycis is dimorphic and thus Vairimorpha remains a valid genus. Synaptonemal; complexes have.been reported in a number of microsporidian species belonging to different genera, including the dimorphic microsporidia (Loubes et al., 1976; Loub&s, 1979; Vivares & Sprague, 1979; Hazard et al., 1979 Canning & Hazard, 1982, Malone & Canning, 1982). The authors found that synaptonemal complex-like structures occur in young sporonts of a variety .of microsporidia. In the present study, it was found that synaptonemal complexes occur in the early octosporoblastic sporonts of N. locustae, indicating that meiosis occurs during this sporogonic sequence. This also implies that uninucleate sporophorous 185 vesicle-bound spores are haploid. It it not known whether the haploid spores of N. locustae are infective to hosts, but it has been reported that haploid spores of V. plodiae are directly infective (Malone & Canning, 1982). Since synaptonemal complexes occur in both nuclei of the diplokarya of early sporonts, these nuclei must be diploid. Therefore, if haploid spores are infective to new hosts, two events must occur i.e. nuclear fusion, to restore the diploid condition and replication, to form the diplokarya, not necessarily in that order. Garaetogenesis and nuclear fusion have not been found in Vairimorpha but recent studies of dimorphic species belonging to several genera in mosquitoes have shed much light on developmental cycles of dimorphic microsporidia (Hazard et al., 1985). Karyogamy has been demonstrated in Amblyospora sp. in mosquitoes by DNA measurement of all stages of the life cycle in mosquitoes by Hazard & Brookbank (1984). They found that Karyogamy occurs soon after the diplokaryotic meronts enter the fat body (Fig. 152), Hazard & Broodbank, (1984) and Hazard et al. (1984) also demonstrated by chromosome studies that karyogamy and meiosis occur during sporogony in Amblyospora sp. and in Culicosporella lunata? Hazard et al. (1985) showed that gametogenesis and fusion of gametes occurs in early larval infections of Hazardia milleri, Culicosporella maqna and a Microsporidium sp. Although nuclear fusion and reduction have been reported in the mosquito host, the haploid spores which 186 Key to Fig. 152 1. transovarial transmission to progeny 2. invasion of oenocytes 3. merogony 4. invasion of oocyst 5. spore extrusion 6. ovarian infection 7. invasion of fat body 8. diplokaryotic meront 9. nuclear fusion 10. diplokaryotic Spororvt 11. sporont 12. mingling of pachytene chromosome 13. divison of sporont 14. quatrinucleate sporont 15. octonucleate sporont 16. haploid spores 17. copepod 18. uninucleate meront 19. binucleate sporont 20. tetranucleate spront 21. 4 sporonts 22. those spores infected with mosquito larvae 187 Fig. 152: Life cycle of Amblyospora sp. in mosquito and intermediate hosts, (Copepods). 14 188 result from the octosporoblastic sequence are not infective to mosquitoes. Sweeney et al. (1985) have shown that the haploid spores are infective to copepods? in the copepods there were uninucleate meronts and binucleate and tetranucleate sporonts, which give rise to uninucleate sporoblasts and spores. These spores were infective to mosquitoes. Karyogamy and meiosis were not reported in this part of the life cycle. It seems that in most dimorphic species, meiosis occurs in the octosporoblastic sporogonic sequence to form haploid spores in host adipose tissue (Table 8). In the present studies, the octosporoblastic sporogonic sequence did not occur in S. frugiperda cell cultures at 20°C, infected with either N. locustae or V. necatrix, with or without addition of juvenile hormone or ecdysone. This implies that meiosis in microsporidia occurs in specific tissues, such as fat body, triggered by certain unknown factors. These triggers, are not likely to be juvenile hormone or ecdysone as postulated by Pilley (1976), but may be built into the genetic complement of the microsporidia themselves. The genes could be switched on under certain condition such as low temperature or host sexuality and specific tissue. Further studies could elucidate those problems for example, by culturing infected fat body rn vitro. 189 Table 8: A summary of dimorphic microsporidia producing spores in infected tissues Dimorphic Tissue in which Tissue in which genus haploid spores diplokaryotic Reference * are produced spores are produced Burenella fat body hyperdermal Jouvenaz & tissue Hazard (1978) Vairimorpha fat body fat body Pilley (1976) Amblyospora fat body oenocyte Andreadis (male larvae) (female adults) (1979) Parathelohania fat body oenocyte Hazard & (male larvae) (female adults) Weiser (1968) Stempellia fat body fat body Desportes (1976) Hazardia fat body fat body Hazard & Faxuda (1974) Culicosporella fat body fat body Hazard et al. (1984) 190 5.6 Sporophorous vesicle Most genera of microsporidia do not lie in direct contact with host cell cytoplasm but are separated by a membrane, which encloses them in a vesicle either throughout their development or only at the sporogonic stage. The vesicle, termed a sporophorous vesicle, sporocyst or pansporoblast membrane provides another characteristic by which genera can be distinguished depending on the presence or absence of the vesicle during merogony: a) All stages bounded by a membrane separating them from the host cell cytoplasm. Usually, this is a simple vesicle membrane, with unit structure, interpolated between the parasites and the host cell cytoplasm. In the present study, this is true in P. operophterae. A similar finding in the genus Encephalitozoon ,was, reported Jjy Pakes et aJL. (1975) and in the genus Loma by Loubes et al. (1984). However, the origin of this membrane is enigmatic and may differ according to genus. The vesicle enclosing stages of Encephalitozoon originates from the host cell (Weidner, 1975) and is correctly termed a parasitophorous vacuole not a sporophorous vesicle. In present study, the infective stages of P. operophterae were closely associated with the host cell surface and infection has been initiated _in vitro by addition of the infective stages to uninfected cultures. It is likely that the vesicle membrane originates from the 191 host cell, as an invagination of the surface membrane which persists around the dividing parasites. In this case it would also be a parasitophorous vacuole. Furthermore, this vesicle divides with the parasite, so that there is never more than one meront present in a vesicle. This may be benefical to the meronts, as they can utilise the maximum surface area to obtain nutrients through the vesicle membrane. Another example of division of parasitophorous vacuoles is provided by Encephalitozoon lacertae (Canning, 1981). In this species the vacuoles, containing separate parasites, ultimately fuse to form a large parasitophorous vacuole, containing merogonic and sporogonic stages. Other examples of species, in which the sporophorous vesicles have precursors around the meronts are P. typicalis and Vavaria culicis (Canning & Hazard, 1982). Their meronts are surrounded by a thick amorphous layer, which is believed to be derived from the parasite. After addition of electron dense secretions from the parasite this amorphous layer becomes the sporophorous vesicle. b ) Sporogonic stages only bounded by a membrane separating them from the host cell cytoplasm. In these cases, the meronts lie in direct contact with the host cell cytoplasm, but a sporophorous vesicle membrane is generated around the sporonts and sporogonic divisions take place within it. Tubular structures are commonly found within the sporophorous vesicle membrane, suggesting that 192 they play an important role in transporting nutrients to the sporogonic stages within the vesicle. Although the sporophorous vesicle membrane is thought to be of parasite origin, the mechanism by which it arises is obscure. Malone & Canning (1982) suggested that the plasma membrane of V. plodiae was expanded into bubbles towards the cell cytoplasm and that some of these separated to form a layer of vesicles around the surface of the sporonts? when these vesicles fused they formed a double envelope, but eventually the inner of the two membranes became degenerate, leaving the outer-layer to form the sporophorous vesicle. In this situation the sporophorous vesicle membrane must be derived from a unit membrane. A similar situation was found in Polydispyrenia simulii (Canning & Hazard, 1982). In the present study of V. locustae (= N. locustae), it has been clearly shown that the membrane of the sporophorous vesicle is a thin electron-dense amorphous layer, and not a unit membrane. The envelope occurs on the surface of the early sporont, and appears to arise as a secretion from the sporont cytoplasm. 5.7 The factors controlling host and tissue specificity of microsporidia. In the past the host range has been used as one of the factors in the diagnosis of a microsporidian species. The factors which determine the host and tissue specificity of microsporidia are unknown. Recent work and the present study provide some insight into this problem. 193 The first step in normal infection is that spores are ingested by a potential host. The chemical and physical conditions within the gut determine whether a species of microsporidia can extrude their sporoplasms into a host cell through the polar tube. The pH is apparently one of the important factors. In the present study, with the exception of Orthosoma operophterae which requires exposure to pH 9.5 for hatching, it has been shown that spores of microsporidia infecting lepidopteran hosts require exposure to high alkaline pH (pH 11-12) before they will hatch. Malone (1984) also found that the spores of V. plodiae, a parasite of lepidoptera, require exposure to high pH (pH 11.5). Spores of species infecting aquatic hosts require pH 9.5. Aquatic host, such as mosquito larvae have a slightly alkaline pH of the midgut (pH 9.0 - 7.0) (Undeen, 1976). Lepidopteran larvae are herbivorous and normally have a strongly alkaline midgut (pH 9.0-11) (Ishihara, 1967? Malone, 1984a). It is not surprising that different host species from a variety of habitats adapt to their diet and that this has a marked influence on the conditions in digestive tract. This implies that microsporidia could be transmitted between host within a niche where the animals share the same food. Burges et al. (1971) have provided an example as they found that Nosema oryzaephili infects five species of beetles and three species of moths involving five insect families of stored produce insects. 194 Cations are other important factors which determine spore hatching. Most studies have shown that a high level of K+ is found in the gut of insects (Malone, 1984a? Gupta et al. , 1976) whereas a high level of Na+ is found in the gut of vertebrates (Prosser, 1973). Both cations can promote spore germination. Further studies are needed to compare ca , which act as inhibitors of spore hatching (Weidner & Byrd, 1982? Malone, 1984a? Undeen, 1978) in a variety of hosts. Previous studies have shown that microsporidia exhibits little specificity iri vitro and will develop in a variety of cell lines. Thus microsporidia derived from insect hosts, can grow in mammalian cell cultures (Undeen, 1975? Ishihara, 1969). However, rn vivo studies have shown that host tissues differ in their ability to support growth of microsporidia. Malone (1984a) found that, although the species of V. plodiae were able to hatch and extrude their sporoplasms into the midgut epithelial cells of P. brassicae, the parasite failed to develop further, but when spores were injected directly into the haemocoele of P. brassicae, the parasite developed in different tissues such as the fat body. This suggests that tissue specificity of micro sporidia is also controlled by biochemical factors in the host cells. However, some microsporidia, such as P. schubergi only develop in midgut epithelial cells and not in other tissues. This implies that these parasites cannot 195 1 survive in haemolymph or in other tissues. Tsang et al. (1982) found that high osmotic pressure and low K+ concentration of UMN-Bl (cockroach) medium made the cell to cell cross infection of N. disstriae into a B. germanica cell line impossible. 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Infect. Immun., 46, 128-134. 218 APPENDIX I Semi-synthetic diet for rearing lepidoptera (Modification of Hoffman's Tobacco Hornworm diet; Smith, 1966). 1. Mix together in a blender: Agar 15 g Casein 26.4 g Wheat germ 57.6 g Wesson's salts 7.5 g Yeast powder 11.4 g Sucrose 23.4 g Cholesterol 0.75 g Methyl para-hydroxy-benzoate 0.75 g Sorbic acid 1.2 g 2. Add and mix thoroughly: Water 690 ml 4 M KOH 3.75 ml Linseed oil 1.5 ml 3. Autoclave mixture in covered vessel for 20 min at 20 lbs per sq.in. Cool to 60-70°C 4. Add and mix thoroughly: Vitamin/antibiotic mix 4.5 g Choline chloride 0.75 g 5. Dispense into no.10 Cristaseal polypots to a depth of about 1 cm and store in refrigerator (3—4 weeks) or freezer (indefinitely). 219 Appendix I (contd) Vitamin/antibiotic mix: Nicotinic acid 5 g Calcium pantothenate 5 g Riboflavin 2.5 g Aneurine hydrochloride 1.25 g Pyridoxine hydrochloride 1.25 g Folic acid 1.25 g Biotin 0.1 g Cyanocobalamine 0.01 g For every gram of this mixture, add: Streptomycini sulphate 2 g Aureomyc in(veterinary soluble powder. 25g/lb) 18 g Ascorbic acid, 40 g Mix well and store in refrigerator 220 APPENDIX II Fixation, dehydration and embedding of specimens for transmission electron microscopy. Modified Karnovsky's fluid Mix: Paraformaldehyde 2 g CaCl2 25 mg Distilled water 20 ml Heat to 65°C? add 2 drops 1 M NaOH? Stir until clear; cool to room temperature; Add gluteraldehyde (25%), 10 ml; and Make up to 50 ml 0.2 M sodium cacodylate buffer, pH 7.4. 1. Modified Karnovsky's fluid, 10 min, room temperature 2. Modified Karnovsky's fluid, 1 hour, 4°C 3. 0.12 M sodium cacodylate, 15 min, 4°C 4. 0.12 M sodium cacodylate, 15 min, 4°C 5. 2.5% (w/v) OsO^ in 0.1 M sodium cacodylate, 1 hour, 4°C, on rotator 6 • 0.1 M sodium acetate, 10 min, 4°C, on rotator 7. 0.1 M sodium acetate, 10 min, 4°C, on rotator 8. 0.25% (w/v) uranyl acetate in water, 1 hour, 9. 0.1 M sodium acetate, 10 min, 4°C, on rotator 10. 0.1 M sodium acetate, 10 min, 4°*C, on rotator 11. 35% acetone, 5 min, 4°C 12. 50% acetone, 5 min, 4°C 13. 70% acetone, with 1% uranyl acetate and 1% phosphotungstic acid, overnight, 4°C 14. 90% acetone, 10 min, room temperature 15. 100% acetone, 15 min, room temperature 16. 100% acetone, 15 min, room temperature 17. 100% acetone, 15 min, room temperature 18. 50/50 acetone/Spurr's resin, overnight, room temperature, on rotator 19. Spurr's resin, all day, room temperature, on rotator 20. Embed in fresh Spurr's resin and polymerise in 60°C oven overnight