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Development and transmission of saurian Plasmodium and Schellackia in bloodfeeding arthropods

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Title:
Development and transmission of saurian Plasmodium and Schellackia in bloodfeeding arthropods
Creator:
Klein, Terry Allen, 1946-
Publication Date:
Language:
English
Physical Description:
xvii, 216 leaves : ill. ; 28 cm.

Subjects

Subjects / Keywords:
Erythrocytes ( jstor )
Infections ( jstor )
Lizards ( jstor )
Malaria ( jstor )
Midgut ( jstor )
Oocysts ( jstor )
Parasitemia ( jstor )
Parasites ( jstor )
Salivary glands ( jstor )
Sporozoites ( jstor )
Blood -- Parasites ( lcsh )
Malaria -- Transmission ( lcsh )
Mosquitoes as carriers of disease ( lcsh )
Plasmodium ( lcsh )
Schellackia ( lcsh )
City of Gainesville ( local )
Genre:
bibliography ( marcgt )
theses ( marcgt )
non-fiction ( marcgt )

Notes

Thesis:
Thesis (Ph. D.)--University of Florida, 1985.
Bibliography:
Includes bibliographical references (leaves 204-214).
General Note:
Typescript.
General Note:
Vita.
Thesis:
Aleph has 1986 and should be 1985.
Statement of Responsibility:
by Terry A. Klein.

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University of Florida
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University of Florida
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Copyright [name of dissertation author]. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
Resource Identifier:
14388669 ( OCLC )
ocm14388669

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DEVELOPMENT AND TRANSMISSION OF SAURIAN PLASMODIUM AND
SCHELLACKIA IN BLOODFEEDING ARTHROPODS

















BY

TERRY A. KLEIN








A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA
IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY






UNIVERSITY OF FLORIDA 1985
































TO JACQUI, KEVIN, AARON, MICHELLE, AND ROBERT











ACKNOWLEDGEMENTS


This dissertation is the result of the cooperation, assistance, and collaboration of many people. While I accept the responsibility for the contents of this dissertation, I do not claim all the credit.

I am grateful to the members of my committee for their contributions and support throughout the study. They provided instructional guidance, enthusiasm, and continuous encouragement. In addition, each member gave freely of his time and offered valuable suggestions in his areas of expertise. They are Dr. David Young, chairman; Dr. Martin Young, cochairman; Dr. Jerry Butler, committee member; and Dr. Stephen Zam, committee member. Dr. Ellis Greiner, while not serving as a committee member, gave freely of his time and expertise in parasitiolgy and provided material support. I am also indebted to Dr. Sam Telford for the many discussions on lizard parasites and examination of histological preparations. Dr. Donald Forrester also provided assistance and laboratory space for the histology studies. I also appreciate the assistance and space for rearing mosquitoes that Dr. Larry Lacy and Dr. Al Undeen provided. Special gratitude is due to Ms. Debra Akin who provided her time, instructional guidance, expertise, and suggestions for the transmission electron studies and Dr. Robert Kimsey who kindly provided me with some infected








lizards for transmission studies. COL John Reinert served on the committee for the first year and, in concert with the other committee members, outlined a demanding schedule of courses.

Sincere thanks go to Ms. Dianna Simon, Mrs. Debra Boyd, Mr. Brooks Ferguson, and other members of the laboratory staff for their assistance in administrative and logistical details. I extend special thanks to Ms. Edna Mitchell for her assistance in maintenance of insect and lizard colonies. Mrs. Margo Duncan offered valuable suggestions in designing table and figures. I also thank Mr. Greg Piepel for his assistance with the statistical portions of this study.

I am grateful for the friendship of fellow graduate

students, Dr. Richard Johnson, Dr. Phillip Lawyer, Mr. Eric Milstrey, Mr. Bruce Alexander, Mr. Charles Beard, MAJ Richard Kramer, and Mr. Clay Smith. Their suggestions, humor, and encouragement were appreciated.

Last, I would like to express my deepest gratitude to my wife Jacqui who supported and encouraged me throughout the study. I wish to thank my children, Kevin, Aaron, Michelle, and Robert for their understanding and enjoyment that they gave me throughout my studies.












iv














TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS ................... ... ... ........ iii

LIST OF TABLES ....................................... viii
LIST OF FIGURES ................... . . . . . ix

ABSTRACT ............................... .. .............. xvi

CHAPTER

1 SAURIAN MALARIA AND Schellackia ................... 1
Saurian malaria ................................... 1
Schellackia ....................................... 12

2 EXPERIMENTAL TRANSMISSION OF Plasmodium mexicanum
BY BITES OF INFECTED Lutzomyia vexator (DIPTERA:
PSYCHODIDAE) ..................... *....0..... ....... 16

Introduction ...................................... 16
Materials and Methods ............................. 17
Lutzomyia vexator Colony Maintenance ......... 17
Collection and Laboratory Maintenance of
Lizards .................................... 18
Plasmodium mexicanum Transmission Studies .... 19 Course of Infection, Parasitemia ............. 20
Histology of Infected Lizards ................ 21
Results ........................................... 21
Plasmodium mexicanum Transmission Studies .... 21 Course of Infection, Parasitemia ............. 29
Histology of Infected Lizards ................ 49
Discussion ........................................ 49
Plasmodium mexicanum Transmission Studies .... 49 Course of Infection, Parasitemia ............. 57
Histology of Infected Lizards ................ 60

3 SPOROGONY, DEVELOPMENT, AND ULTRASTRUCTURE OF
EXTRINSIC STAGES OF Plasmodium mexicanum .......... 62
Introduction ...................................... 62
Materials and Methods ............................. 63
Sporogony of Plasmodium mexicanum ............ 63
Ultrastructure of Extrinsic Stages of
Plasmodium mexicanum ....................... 64


V









Results.............................. ... 64
Sporogony of Plasmodium mexicanum............ 64
Ultrastructure of Extrinsic Stages of
Plasmodium mexicanum ....................... 71
Discussion ........................................ 92
Sporogony of Plasmodium mexicanum............ 92
Ultrastructure of Extrinsic Stages of
Plasmodium mexicanum ....................... 94

4 EXPERIMENTAL TRANSMISSION OF Plasmodium floridense
BY BITE OF INFECTED Culex (Melanoconion) erraticus
(DIPTERA:CULICIDAE)............................... 105

Introduction.............................. ........ 105
Materials and Methods....... .... .. ......... .... .. 106
Field Studies and Collection of Mosquitoes... 106 Culex erraticus Colony Maintenance........... 110
CoTection and Laboratory Maintenance of
Lizards ......... ..... ..... ............ .... 112
Plasmodium floridense Transmission Studies... 112 Course of Infection, Parasitemia............. 115
Results........................................... 115
Field Studies ........ ......................... 115
Plasmodium floridense Transmission Studies... 118 Course of Infection, Parasitemia ............. 129
Discussion....... .......... ............... ........ 134
Field Studies ........................... ... 134
Plasmodium floridense Transmission Studies... 140 Course of Infection, Parasitemia............. 145

5 SPOROGONY, DEVELOPMENT, AND ULTRASTUCTURE OF
EXTRINSIC STAGES OF Plasmodium floridense IN Culex
erraticus......................................... 147

Introduction.......................... ............. 147
Materials and Methods............................. 148
Sporogony of Plasmodium floridense.......... 148
Ultrastructure of Extrinsic Stages of
Plasmodium floridense. .............. ...... 149
Results...................................... ..... 150
Sporogony of Plasmodium floridense........... 150
Ultrastructure of Extrinsic Stages of
Plasmodium floridense......... ........... 154
Discussion........................................ 166
Sporogony of Plasmodium floridense........... 166
Ultrastructure of Extrinsic Stages of
Plasmodium floridense............. ........ 168








vi








6 DEVELOPMENT AND EXPERIMENTAL TRANSMISSION OF
Schellackia golvani AND Schellackia occidentalis
BY INGESTION OF INFECTED BLOODFEEDING ARTHROPODS .... 171

Introduction ....... ........................ .. 171
Materials and Methods ............................ 172
Transmission Studies ......................... 172
Histological Studies .......................... 175
Results ........................................... 176
Schell lacki a goi vani .. . . .. . . 176
Schellackia occidentalis ................ 185
Discussion ..... .................................. 187
Transmission Studies ....................... .. 187
Histological Studies ......................... 199

7 SUMMARY AND RECOMMENDATIONS FOR FUTURE
INVESTIGATIONS .................................... 201
REFERENCES ................................. . . . 204

BIOGRAPHICAL SKETCH............ .......... 215
































vii













LIST OF TABLES


Table Page

1. Laboratory transmission data of Plasmodium
mexicanum to Sceloporus undulatus by bite of
infected Lutzomyia vexator females.............. 25

2. Linear regression analysis of acute Plasmodium
mexicanum infections with fewer than 500 parasites per 10,000 red blood cells (5%
parasitemia) .................................... 34

3. The effects of temperature on sporogony of
Plasmodium mexicanum in Lutzomyia vexator....... 66

4. Summary of Plasmodium floridense infections in
Anolis carolinensis and Sceloporus undulatus
coTllected from different localities in Florida
(1983-5) ................... .................... 113

5. Summary of CDC light and lizard bait trap
collections and bloodfeeding of feral mosquitoes in the field and laboratory from 31 April to 10
October, 1984 ................................... 117

6. Summary of laboratory transmission data of
Plasmodium floridense to Anolis carolinensis and
Sceloporus undulatus for a three year period.... 123

7. Laboratory transmission data of Plasmodium
floridense to Anolis carolinensis .............. 130

8. Summary of sporogony and transmission of saurian
malaria in bloodfeeding Diptera................ 131

9. Sporogonic development of Plasmodium floridense
on the midgut of Culex erraticus ................ 153

10. Experimental transmission data of Schellackia golvani and Schellackia occidentalis to wild
caught Anolis carolinensis and Sceloporus
undulatus by ingestion of arthropods............ 177




viii













LIST OF FIGURES


Figure Page

1. Bloodfed female Lutzomyia vexator resting on
Sceloporus undulatus upon which it had
previously fed on .............................. 23

2. Midgut of Lutzomyia vexator with oocysts of
Plasmodium mexicanum ........................... 23

3. Plasmodium mexicanum sporozoites from ruptured
oocysts ......................................... 23

4. Plasmodium mexicanum sporozoites in the
salivary gland of Lutzomyia vexator............. 23

5. Prepatent and patent period of Plasmodium
mexicanum infection and survival of individual Sceloporus undulatus infected with Plasmodium
mexicanum sporozoites ........................... 28

6. Percent of infected red blood cells during the course of Plasmodium mexicanum infection for six Sceloporus undulatus infected by bite of
Lutzomyia vexator............................... 31

7. Course of acute infection of Plasmodium mexicanum in nine Sceloporus undulatus infected
by bite of Lutzomyia vexator .................... 33

8. Course of Plasmodium mexicanum infection in Sceloporus undulatus infected with sporozoites
(S-51) .......................................... 38
9. Course of Plasmodium mexicanum infection in
Sceloporus undulatus infected with sporozoites
(S-42).......................................... 40
10. Course of Plasmodium mexicanum infection in Sceloporus undulatus infected with sporozoites
(S-8) ........................................... 42
11. Course of Plasmodium mexicanum infection in Sceloporus undulatus infected with sporozoites
(S-15) .......................................... 44
ix








12. Course of Plasmodium mexicanum infection in
Sceloporus undulatus infected with sporozoites
(S-25 ... ............................. .......... 46

13. Course of Plasmodium mexicanum infection in
Sceloporus undulatus infected with sporozoites
(S-14) .................... ..... ...... .... ...... 48

14. Bloodfilm of Sceloporus undulatus infected with
Plasmodium mexicanum parasites .................. 51

15. Spleen tissue impression of Sceloporus undulatus
infected with Plasmodium mexicanum parasites.... 51

16. Bone marrow smear of Sceloporus undulatus
infected with Plasmodium mexicanum parasites.... 51

17. Schizonts of Plasmodium mexicanum in
endothelial cells of capillaries in the brain... 51

18. Regression of Plasmodium mexicanum mean oocyst
size (and 95% confidence limits) in Lutzomyia
vexator and day post-feed for days 2 through 9.. 68

19. Midgut of Lutzomyia vexator with asynchronous
development of Plasmodium mexicanum oocysts..... 70

20. Sporoblast formation in 5 day old oocyst on the
midgut of Lutzomyia vexator..................... 70

21. Plasmodium mexicanum oocyst on the midgut of
Lutzomyia vexator...................... ......... 70

22. Sporozoites of Plasmodium mexicanum which
ruptured from oocysts........................... 70

23. Salivary gland of Lutzomyia vexator containing
numerous sporozoites of Plasmodium mexicanum.... 70

24. Cross section of a 4 day old Plasmodium
mexicanum oocyst on the midgut of Lutzomyia
vexator ......................................... 73
25. Cross section of Plasmodium mexicanum oocyst
undergoing differentiation...................... 76

26. Cross section of Plasmodium mexicanum oocyst
undergoing differentiation............ .......... 76

27. Cross section of Plasmodium mexicanum oocyst
undergoing internal vacuTolization of the
sporoblastoid................................... 76



x







28. Coelescence of the internal vaculization of
Plasmodium mexicanum oocysts produces
cytoplasmic clefts .............................. 76

29. Higher magnification of Plasmodium mexicanum
oocyst..................................... ... 76

30. Oocyst of Plasmodium mexicanum with developing
sporozoites.................................... 80

31. Higher magnification of developing sporozoites
of Plasmodium mexicanum......................... 80

32. Cross section of Plasmodium mexicanum oocyst
with developing sporozoites..................... 80

33. Plasmodium mexicanum sporozoites in the process
of penetrating the salivary gland of Lutzomyia
vexator................................. ........ 83

34. Another sequence of Plasmodium mexicanum
sporozoites in the process of penetrating the
salivary gland of Lutzomyia vexator............. 83

35. Plasmodium mexicanum sporozoites inside the
salivary gland cell of Lutzomyia vexator........ 86

36. Plasmodium mexicanum sporozoites in the lumen of
the salivary gland of Lutzomyia vexator......... 86

37. Magnification of sporozoites of Plasmodium
mexicanum in the salivary gland of Lutzomyia
vexator ....................................... .. 89

38. Sporozoites of Plasmodium mexicanum illustrating
polar rings...I............ ... .............. ... 89

39. Higher magnification of the cytostome of a
sporozoite of Plasmodium mexicanum.............. 91

40. Cross section of anterior of Plasmodium
mexicanum sporozoite in the lumen of the
salivary gland of Lutzomyia vexator............. 91

41. Magnification of the posterior end of a
Plasmodium mexicanum sporozoite with an elongate
U-shaped mitochondria with an associated
electron dense sphere.......................... 91

42. Photograph of lizard baited trap used to
attract and capture biting Diptera..........- .... 109

43. Culex erraticus bloodfeeding on a restrained
-o is carolinensis ............................. 109

xi







44. Number of Culex erraticus collected per light
trap during the period 1 May to 9 October,
1984 ............................................ 120

45. Percent Culex erraticus and Culex territans
collected per bait trap during the period 1
May to 9 October, 1984......................... 122

46. Midgut of Culex erraticus, (day 9 post-feed)
with asynchronous development of Plasmodium
floridense oocysts.............................. 126

47. Plasmodium floridense oocyst on the midgut of
Culex erraticus with many nearly mature
sporozoites (day 9 post-feed)................... 126

48. Culex erraticus salivary glands with sporozoites
of Plasmodium floridense.................. ....... 126

49. Magnification of living Plasmodium floridense
sporozoites from the salivary glands of Culex
erraticus............................ ........... 126

50. Plasmodium floridense oocyst on the midgut of
Culex erraticus on day 10 post-feed.
Melanization is beginning to occur along the
oocyst capsule and spread inward................ 128

51. Bloodfilm of Anolis carolinensis at the first
peak of a Plasmodium floridense infection....... 128

52. Course of acute infection of Plasmodium
floridense in two wild caught Anolis
carolinensis infected with sporozoites.......... 133

53. Course of acute infection of Plasmodium
floridense in Anolis carolinensis infected
with sporozoites (AA-59) ....................... 136

54. Course of acute infection of Plasmodium
floridense in Anolis carolinensis infected
with sporozoites (A-85) ....... ........... .... 138

55. Cross section of midgut of Culex erraticus with
asynchronous development of Plasmodium
floridense oocysts........ ...... .. ... ........... 152

56. Residual body of sporoblast with developing
sporozoites of Plasmodium floridense which
ruptured from an oocyst on the midgut of Culex
erraticus............... ....... ...... .......... 152
57. Sporozoites of Plasmodium floridense in the
salivary gland of Culex erraticus............... 152

xii








58. Sporozoites of Plasmodium floridense which
ruptured from the salivary glands of Culex
erraticus ....................................... 152

59. Midgut of Culex erraticus with melanized
oocysts and sporozoites (?) of Plasmodium
floridense ...................................... 152

60. Cross section of an oocyst of Plasmodium
floridense on the midgut of Culex erraticus..... 156

61. Higher magnification of the oocyst capsule of
Plasmodium floridense ......... .............. .. 156

62. Cross section of an oocyst of Plasmodium
floridense on the midgut of Culex erraticus..... 156

63. Virus particles in the midgut epithelium of
Culex erraticus .............................. .. ... ..... .. .. .... 156

64. Cross section of Plasmodium floridense oocyst
undergoing differentiation ...................... 159

65. Convex dense membranes, the precursors of the
developing sporozoites, form along the narrow
linear extensions............................... 159

66. Oocyst of Plasmodium floridense with developing
sporozoites..................................... 162

67. Longitudinal section of the posterior portion
of a developing sporozoite..................... 162

68. Higher magnification of developing sporozoites
of Plasmodium floridense ........................ 162

69. Higher magnification of developing sporozoites
of Plasmodium floridense........................ 162

70. Oocyst of Plasmodium floridense with developing
sporozoites ..................................... 165
71. Salivary gland of Culex erraticus infected with
sporozoites of Plasmod-ium floridense............ 165

72. Cross section of Plasmodium floridense
sporozoites in the salivary glands of Culex
erraticus ....................................... 165
73. Degenerating sporozoite of Plasmodium floridense
in an oocyst which is becoming melanized........ 165
74. Sporozoites of Schellackia golvani in white
blood cells of Anolis carolinensis .............. 181

xiii








75. Sporozoite of Schellackia golvani with two
chromatin bands in a white blood cell of Anolis
carolinensis .................................... 181
76. Sporozoite of Schellackia golvani teased from
the midgut of Culex erraticus ................... 181

77. Unstained sporozoite of Schellackia 9olvani
teased from the midgut of Culex erraticus....... 181

78. Cross section of the midgut of Culex erraticus
with a sporozoite of Schellackia golvani........ 181

79. Cross section of the midgut of Lutzomyia vexator
with a sporozoite of Schellackia golvani........ 181

80. Unstained sporozoites of Schellackia golvani in
a parasitophorus vacuole in the midgut
epithelium of Culex erraticus................... 181

81. Impression of small intestine of Anolis
carolinensis with macromerozoites of Schellackia
golvani ......................................... 184
82. Section of intestine of Anolis caorlinensis with
macroschizont and macromerozoites of Schellackia
golvani ......................................... 184
83. Section of intestine of Anolis carolinensis with
microschizont containing developing
micromerozoites of Schellackia golvani.......... 184

84. Section of intestine of Anolis carolinensis with
macro- and microgametocytes of Schellackia
golvani ......................................... 184
85. Sporozoites of Schellackia occidentalis teased
from the gut of a mite.......................... 184

86. Crescent-shaped sporozoite of Schellackia
occidentalis in the red blood cell of Sceloporus
undulatus ....................................... 189
87. Comma-shaped sporozoite of Schellackia
occidentalis in the red blood cell of Sceloporus
undulatus ....................................... 189
88. Sperical sporozoite of Schellackia occidentalis
in the red blood cell of Sceloporus undulatus... 189

89. Sporozoite of Schellackia occidentalis in a
white blood cell of Sceloporus undulatus........ 189



xiv









90. Numerous sporozoites of Schellackia occidentalis
in a white blood cell of Sceloporus undulatus ... 189 91. Section of intestine of Sceloporus undulatus
with a schizont of Schellackia O-Ccidentalii ..... 191 92. Section of intestine of Sceloporus undulatus
with an oocyst of STFeMaFMa occidentTais.. 191












































xv














Abstract of Dissertation Presented to the Graduate
School of the University of Florida in Partial
Fulfillment of the Requirements for the Degree of Doctor of Philosophy



DEVELOPMENT AND TRANSMISSION OF SAURIAN PLASMODIUM AND
SCHELLACKIA IN BLOODFEEDING ARTHROPODS


By

Terry A. Klein

August 1985

Chairman: Dr. David G. Young Co-chairman: Dr. Martin D. Young Major Department: Entomology and Nematology


A study was undertaken to (1)determine the vectors of two species of Plasmodium and two species of Schellackia parasites of lizards; (2) study the course of development of the parasite in the lizard host; and (3) study the growth and development of the extrinsic stages, including light and transmission electron microscopy (TEM). Lutzomyia vexator was shown to be an efficient vector of P. mexicanum, infecting 69.2% (9/13) of the Sceloporus undulatus lizards when fed on by infected flies. Culex erraticus was incriminated as a suspected vector of saurian malaria, P. floridense. Vector attraction and bloodfeeding propensity in the field was determined by using lizard baited traps. Both Cx. erraticus and Cx. territans were frequently


Xvi








collected in the trap. Culex territans is an early spring mosquito, while Cx. erraticus is abundant in the summer and fall months when transmission of P. floridense occurs. Transmission of P. floridense was accomplished by bite of Cx. erraticus and intraperitoneal injection of sporozoites. Sporogony of P. floridense was demonstrated in L. vexator, but sporozoites were rarely seen, and never in the salivary glands. The prepatent period for P. floridense and P. mexicanum is relatively long (>20 days) at 18-24oC. Plasmodium mexicanum is very pathogenic, invading the spleen and endothelial cells of the brain, in Sc. undulatus, while P. floridense rarely kills A. carolinensis.

Sporogony of both saurian malarias is similar to that described for other malarias. The gross morphology and ultrastructure of P. floridense sporozoites is similar to other mammalian and avian malarias. However, P. mexicanum sporozoites are short, stout, and have a subpellicular microtubular arrangement which is different from other malarias, but similar to P. agamae, another saurian malaria.

Transmission of Schellackia parasites was accomplished by ingestion of infected mosquitoes, sand flies, and mites that previously fed on conspecific hosts, i. e., S. golvani to A. carolinensis and S. occidentalis to Sc. undulatus. The prepatent period was reduced significantly by high temperatures. At room temperatures, the prepatent period for both Schellackia parasites was >20 days. However, at 320C, the prepatent period was decreased to as few as 7 days.
xvii



















CHAPTER 1
SAURIAN MALARIA AND SCHELLACKIA



Saurian Malaria




Species of Plasmodium, a hemosporidian genus of

protozoa that infect mammals, birds and reptiles, cause a disease generally referred to as "malaria." Malaria in man has been recorded in papyrus as early as 1500 BC in Egypt, but the causative agent was not known until 1880 when Laveran discovered Plasmodium parasites in human red blood cells. The persistent search for malaria parasites in mosquitoes by Ross (1897) eventually led to the discovery of oocysts on the midgut of a "dapple-winged" mosquito (Anopheles sp.) which fed on a human patient infected with malaria. The vector-host-parasite relationship was subsequently demonstrated for Plasmodium relictum Grassi and Felitti (avian malaria) (Ross, 1898) and for human malaria (Grassi et al., 1899). The cryptic exoerythrocytic stage was demonstrated much later by Rafaelle (1934) (Plasmodium


-1-





-2


elongatum Huff, an avian malaria) and by Shortt and Garnham (1948) (Plasmodium cynomolgi Mayer and Plasmodium vivax Grassi and Felitti, a monkey and human malaria, respectively). Literature on the mammalian and avian malarias is voluminous due to their medical and economic importance, but saurian malaria has received very little attention.

The first named saurian malaria parasites, P. mabuiae

Wenyon (1909) and Plasmodium (=Haemoproteus) agamae (Wenyon, 1909), were described from Mabuya quinquetaeniata (Lichenstein) (common skink) and Agama agama L., (rainbow lizard) respectively, from Africa (Wenyon, 1909). During that same year, Aragao and Neiva (1909) described Plasmodium diploglossi and P. tropiduri from Diploglossus fasciatus (Gray) and Tropidurus torquatus (Wied), respectively, from Brazil. Since that time, a total of 59 species of saurian malaria have been described from Africa (12), the Americas

(36), and Australasia (11) (Telford, 1982; Telford, 1983; Telford, 1984a; Telford 1984b; Garnham and Telford, 1984). A review of saurian malaria literature from 1909 to 1975 (Ayala, 1978) includes only 153 references, most of which emphasize taxonomy or distribution records. From 1975 to 1985, only 32 additional references have been added [including 4 previously overlooked by Ayala (1978)].

Garnham (1966) stated that, "A few puzzles in the life history of the malaria parasites remain, of which the most important are perhaps the exact nature of latency and vector





-3


of lizard malaria" (p. 13). In fact, no vectors, except for two species of phlebotomine sand flies (Ayala and Lee, 1970), have been incriminated as vectors for any of the species of Plasmodium infecting lizards and only limited attempts have been made to incriminate other vectors of saurian malaria. Previous suggestions that acarine ectoparasites transmit saurian malaria in Mexico (Pelaez et al., 1948) and in Africa (Pienaar, 1962) proved to be false (Russell et al., 1963; Ayala, 1977). Also, sporogonic development was not observed after triatomid bugs were fed on lizards infected with Plasmodium parasites (Ayala, 1977).

Although hematophagus Diptera are the most likely vectors of saurian malaria (Ayala, 1977), attempts to incriminate these insects have usually met with failure. The earliest recorded attempt to determine the vector of saurian malaria was by Huff (1941a) who fed both Culex pipiens L. and Aedes aegypti (L.) mosquitoes on Sceloporus undulatus Latreille (eastern fence lizard) infected with Plasmodium sp. The Plasmodium species, although not identified, was isolated from Sc. undulatus where P. floridense Thompson and Huff (1944b) is endemic. Culex pipiens females were negative; however, one of an unknown number of Ae. aegypti had one nearly mature oocyst on the midgut. Attempts to infect Ae. aegypti (61) by feeding them on Sc. olivaceaous Smith infected with numerous P. mexicanum Thompson and Huff (1944b) gametocytes were unsuccessful (Thompson and Huff, 1944b). Baker (1961) examined several





-4


species of sympatric biting Diptera (mosquitoes and sand flies) from areas of endemic saurian malaria in Africa after feeding them on lizards infected with P. agamae and P. giganteum Theiler. Gametes and ookinetes were described, but further development did not occur.

Jordan (1964) had little success demonstrating

sporogony in local mosquitoes collected in Georgia, USA, where Anolis carolinensis Voight and Sc. undulatus are naturally infected with P. floridense. Five species, Aedes atlanticus-tormentor Dyar and Knab (88), Aedes triseriatus (Say) (10), Psorophora confinis (Lynch Arribalxaga) and Psorophora ferrox (Von Humboldt) (20), and Coquillettidia (=Mansonia) perturbans (Walker) (25) were all negative. However, 1/80 Ae. aegypti had one oocyst, 4/70 Culex territans Walker had 1-23 oocysts, 2/150 Culex quinquefasciatus Say had 1-3 oocysts, and 1/3 unidentified Culex sp. had 70 oocysts. (Unfortunately the Culex sp. with 70 oocysts was not identified to species.) Anolis-baited traps, designed to attract and capture hematophagus insects, were ineffective (Jordon, 1964).

Landau et al. (1973) attempted to infect Ae. aegypti,

Anopheles stephensi Liston, Culex fatigans Weideman, and Cx. pipiens by feeding them on Tupinambus teguixin (L.) infected with Saurocytozoon (=Plasmodium) tupinambi Lainson and Shaw (1969), a plasmodiid. Only Cx. pipiens developed oocysts, but these were mostly abnormal, with sporozoites only observed in the oocysts, and never in the salivary glands.




-5


Attempts by Pessoa et al. (1974) to demonstrate snake malaria sporogony in Cx. fatigans were also unsuccessful.

Ayala (1970b) and Ayala and Lee (1970) were the first to report complete sporogonic development of a saurian malaria (P. mexicanum, from California, USA) in hematophagus insects [phlebotomine sand flies, Lutzomyia vexator (Coquillett) and Lutzomyia stewarti (Mangaberia and Galindo)]. In the laboratory, a large proportion of sand flies (61/72, 84.7%), dissected on day seven after feeding on an infected lizard, had developing stages of P. mexicanum. Some flies had more than 100 oocysts on the midgut. Sand flies dissected from days 11 to 14 after feeding had motile sporozoites in the hemocoel and salivary glands (Ayala and Lee, 1970). Transmission was demonstrated by inoculation of sporozoites (midgut and hemocoelic fluid) from wild caught females which had fed on infected Sceloporus occidentalis Baird and Girard (western fence lizard) in the laboratory. Although Ayala and Lee did not demonstrate transmission by insect bite, they showed that both sand fly species were susceptible to infection (complete sporogony), the sporozoites were infective to lizards when injected intraperitoneally, and that there is a close association of sand flies and western fence lizards in rodent burrows in California.

More recently, sporogony of a saurian malaria (P.

agamae) was observed in Culicoides nubeculosus (Meigen), an unnatural host (Petit et al., 1983). Although sporozoites





-6


were observed in the oocysts, the oocysts were intracellular and did not rupture. This suggests that this species of Plasmodium may be transmitted by ingestion. However, sporogony of malaria parasites in unnatural mosquito species has also been reported to be sometimes intracellular. Telford (1970b) stated that the current ". . finding(s) suggest that saurian malaria (parasites) may utilize a variety of hematophagus arthropods for transmission. Certainly, generalizations concerning vector relationships of this group of parasites are still premature" (p. 340).

Because the vectors of saurian Plasmodium are unknown, attempts to follow the life cycle of the parasite in lizards have been done only by experimentally inoculating susceptible lizards with blood from infected lizards (Thompson, 1944; Thompson and Huff, 1944a,b). These early reports demonstrated slow asynchronous parasite development in two species of saurian malarias (P. mexicanum and P. floridense) that is different from the synchronous, fast developing parasitemias of some avian and mammalian malarias (Thompson and Huff, 1944a,b; Thompson, 1944). Parasitemias of P. floridense (>1,000 parasites/10,000 erythrocytes in Sc. undulatus and A. carolinensis) peaked between 23-94 days, then gradually decreased. (The period of time between inoculation and peak parasitemia may be a function of parasite inoculation rate.) Blood-induced infections in Sc. olivaceous and A. carolinensis appeared to peak earlier, at approximately day 45. Thompson (1944) showed that there is





-7


much variability of parasite development within species and among different species of lizards infected with P. floridense and P. mexicanum. Later, Thompson and Winder (1947) established a relationship between parasite development rate and ambient temperature. Susceptible (?) A. carolinensis inoculated with a common pool of citrated blood, and maintained in an incubator at 20 C, had an average peak parasitemia at 55 days post inoculation, while those maintained in an incubator at 30 0C had an average peak parasitemia at 13 days. This indicated that environmental factors, host behavior and effect of the parasite on host behavior play an influential role in the progression of parasitemia in lizards under natural conditions.

Goodwin and Stapleton (1952) extended Thompson and Huff's observations by studying field collected Sc. undulatus infected with P. floridense. The initial bloodfilms of three Sc. undulatus were negative, thus allowing the authors to follow naturally acquired saurian malaria from the prepatent phase (assuming these were not relapses). In one case, parasites were not demonstrable until 27 days after capture. Goodwin (1951) also reported that wild caught lizards with initial negative bloodfilms and maintained in the laboratory developed parasitemia within two weeks. These reports were the first indication that a lengthy prepatent phase is present, at least for Sc. undulatus infected with P. floridense.





-8


Further advances in the host-parasite relationship were made by Ayala (1971) who infected lab reared Sc. occidentalis with P. mexicanum sporozoites. Blood parasites were not observed in the blood until 22 days post inoculation. This corresponds with the results of a 14 to 27 day prepatent period of P. floridense in Sc. undulatus (Goodwin, 1951; Goodwin and Stapleton, 1952). However, as previously indicated, the prepatent period may be influenced by ambient temperature (Thompson and Winder, 1947). Ayala (1970a,b) further noted that there is a spring relapse of gametocytes of P. mexicanum lasting from March to August. This corresponds to the period of activity of its suspected vector, L. vexator, indicating that most malaria transmission in California is limited to the spring and summer months.

Exoerythrocytic (EE) stages have been observed in a number of different saurian malaria species, usually in blood films (Garnham, 1950; Garnham and Duke, 1953; Bray, 1957, 1959; Lainson and Shaw, 1969; Telford 1970a; and Scorza 1971b). Huff (1969) provides an excellent review of the EE stages of avian and saurian malaria parasites and

points out the need for additional research in this area, especially the determination of the vectors of saurian malaria and subsequent observation of the life cycle of the

parasite in the saurian host.

The first and most detailed description of the EE

stages of saurian malaria parasites (P. mexicanum) was made




-9


by Thompson and Huff in the 1940's when similar forms of

avian plasmodia were also being studied. These EE studies were accomplished by examination of blood smears or other tissues of lizards having natural or experimental infections. Thompson and Huff (1944a) inoculated blood from Sc. erythrurus (=ferrariperzi) (Schinz) containing P. mexicanum parasites into five different species of lizards in three genera, Sceloporus, Phrynosoma, and Crotaphytus. Although there was much variation in the intensity of the infections among conspecific lizards, there were significant differences in the degree of parasitemia and the

distribution of the parasites in different types of cells of the five species of experimental hosts. The percentage of P. mexicanum parasites found in erythrocytes ranged from 95% (Sc. undulatus and Sc. olivaceous), <20% [Phrynosoma cornutum (Harlan)], <10% (Crotaphytus collaris Say) to <1% (Phrynosoma asio Cope). The majority of P. mexicanum parasites in Ph. cornutum and C. collaris were observed in the lymphocytes (45% and 35%, respectively) while for Ph. asio, the majority of parasites were observed in the thrombocytes (65%). Plasmodium mexicanum EE stages, unlike other species of saurian and avian malaria, are found in both the hemopoietic and reticuloendothelial tissues and therefore represent both P. elongatum and Plasmodium gallinaceum Brumpt types of EE cycles of avian malarias (Thompson and Huff, 1944a).




-10


The common occurrence of EE stages, the similarities in the types of non-erythrocytic cells invaded, and morphological similarities between avian and saurian Plasmodium suggest that malaria parasites in lizards and birds evolved from a common ancestor (Thompson and Huff, 1944a; Mattingly, 1965). So far, all avian and saurian malarias studied (except P. mexicanum) are restricted to the hemopoietic or reticuloendothelial tissues. Plasmodium mexicanum appears to be the most primitive of the Plasmodium species since its EE stages are not restricted to one type of tissue (Huff, 1945). According to Bray (1957, 1963), the genus Plasmodium is polyphyletic with the bird and reptilian plasmodia originating from a common ancestral stock and mammalian plasmodia originating from a different stock (Mattingly, 1965). This is based mostly on the types of EE tissues invaded by the different malarial parasites.

The elucidation of vectors of saurian malaria has been difficult. Sporogony, to the development of mature sporozoites, has been observed for only two species of saurian malaria (Ayala and Lee, 1970; Ayala, 1971; Petit et al., 1983). Giemsa stained sporozoites of both P. mexicanum and P. agamae appear to be similar and are very short, 5-7um (um = micron) and 4-6um, respectively, when compared to those of most other malarias. Only the fine structure of the development of oocysts and of mature sporozoites of P. agamae, in an unnatural host, Culicoides nubeculosus, has been described (Boulard et al., 1983). In general, oocyst





-11


development of P. agamae is similar to other Plasmodium. The sporozoites of P. agamae is also similar in structure to other Plasmodium species which have been examined. However, the organization and number of pellicular microtubules are different (Boulard et al., 1983).

The fine structure of blood forms of P. floridense

(Aikawa and Jordon, 1968), P. tropiduri (Scorza, 1971a), and P. mexicanum (Moore and Sinden, 1974) has been studied. In general, their morphology is similar to the avian malaria parasites examined.

Thompson (1946a) postulated that lizards might be used as a malaria model in chemotherapeutic research since the effects of anti-malarial drugs on malaria parasites maintained at different temperatures in poikilothermic lizards might provide valuable information. Thompson (1946a,b) studied the effects of atabrine on P. floridense and quinine on P. floridense and P. mexicanum. Both drugs were effective in reducing the parasitemias in lizards with P. floridense, but quinine did not appreciably lower the parasitemia of lizards with P. mexicanum. This was apparently due to the inability of the drug to destroy the EE stages. Reptilian malaria never became popular for study, largely because of the more demanding care required by the experimental animals, the difficulties in establishing cyclic transmission, and the many biological characteristics that separate them from mammalian species (Wernsdorfor, 1980). However, with advances in tissue




-12


culture techniques and relative ease with which reptilian cell lines can be established, the use of reptilian parasites for in vitro culture studies has increased (Wernsdorfor, 1980).

In summary, literature on various aspects of saurian malaria is limited. Morphology, electron microscopy, EE stages, chemotherapeutic drug studies, and parasite development in the vertebrate suggest a close phylogenetic relationship between the avian and saurian malarias. Aside from the possibility that the sand fly, L. vexator, may transmit P. mexicanum, no other arthropods have been incriminated as vectors of saurian malaria. Transmission by bite has not been demonstrated and no naturally infected arthropods have been discovered.



Schellackia



Schellackia parasites are transmitted mechanically by

ingestion of invertebrate hosts (mites, Diptera, or leeches) which previously fed on infected cold blooded vertebrates (reptiles or amphibians). The schizogonic stage occurs in the epithelial cells of the intestine while sporogony occurs usually in the lamina propria of the intestine. In some species, such as S. balli Lebail and Landau (1974), oocysts also form in the epithelial cells of the intestine (Lainson, et al., 1976). Levine (1980), however, extended the definition of Schellackia to include parasites with merogony





-13


in connective tissue and/or reticulendothelial systems. Sporozoites are released from the oocyst where they invade erythrocytes or lymphocytes or both. The host response may determine what type of cell is invaded by the sporozoite. For example, S. bolivari Reichenow (1919) invades the erythrocytes of one lizard, Acanthodactylus vulgaris (Schinz), while invading the lymphocytes of another, Psammodromus hispanicus Fitzinger (Manwell, 1977). (=Haemogregarina) weinbergi, parasites was made by Leger and Mouzels (1917) from South American lizards. Reichenow (1919) first described the genus Schellackia and named a new species, Schellackia bolivari from Spain. Reichenow's description included the complete life cycle of the parasite in the lizard and incriminated a mite, Lyponyssus saurarum, as the natural vector. Later, Bonnoris and Ball (1955) described Schellackia occidentalis from Sceloporus occidentalis in California, USA, and also incriminated a mite, Gekobiella texana (Banks), as the natural vector. There are four additional species of Schellackia in lizards in the western hemisphere: S. brygooi Landau (1973), S. landaue Lainson, Shaw, and Ward (1976), S. golvani Rogier and Landau (1975), and S. (=Lainsonia) iguanae (Landau, 1973), and one in toads, S. balli.

Experimental transmission of S. brygooi and S. landaue has been accomplished by ingestion of mosquitoes (Culex pipiens pipiens and Cx. D. fatigans) which bloodfed on




-14


infected hosts (Landau, 1973; Lainson et al., 1976). Jordan and Friend (1971) further incriminated mites (G. texana) as the natural vector of S. occidentalis by demonstrating that conspecific lizards, kept in glass gallon jars with infected lizards harboring mites, became infected. Lainson et al. (1976) further showed that mosquitoes remain infective for as long as 14 days following a bloodmeal from an infected lizard, and may remain infective for the life of the mosquito.

The taxonomic characterization of Schellackia parasites which infect Anolis carolinensis and Sceloporus undulatus in Georgia and Florida is uncertain. Jordan and Friend (1971) and Telford (1978) have identified the Schellackia parasites of Sc. undulatus as S. occidentalis. Jordan and Friend report that it is doubtful that the Schellackia parasites in anoles is the same as that of S. occidentalis. Telford goes one step futher and indicates that Schellackia in the anoles resembles S. golvani described from Guadeloupe Anolis and common in other Caribbean anoles.

The fine structure of sporozoites of S. occidentalis has been reported by Moore and Sinden (1974). Sporozoites were contained in a parasitophorus vacuole as are the sporozoites of Eimeria and Lankesterella. Although there are similar structures in Schellackia and Lankesterella, there are also many similarities between Schellackia and Plasmodium sporozoites. Manwell (1977) remarked that while Schellackia have no known practical importance, they are of








interest to anyone who has speculated about the evolution of the malaria parasites. Their life history and ultrastructural organization may provide an insight as to how the coccidian ancestors may have adapted to an alternating existence in a vertebrate and arthropod host.













CHAPTER 2
EXPERIMENTAL TRANSMISSION OF Plasmodium mexicanum BY BITES
OF INFECTED Lutzomyia vexator (DIPTERA:PSYCHODIDAE)


Introduction


Saurian malaria research has received increasing

attention (primarily taxonomic) in the past few years, but the natural vectors remain unknown. Fifty-nine species of saurian Plasmodium have been described (36, Americas; 11 Australia, Asia and Oceania; 12, Africa), three of which occur north of Mexico (Ayala, 1978; Telford, 1982; Telford, 1983; Telford, 1984a; Telford, 1984b; Garnham and Telford, 1984). Ayala and Lee (1970), Ayala (1971) and Petit et al. (1983) are the only authors to describe the extrinsic cycle of lizard malaria (Plasmodium mexicanum and Plasmodium agamae, respectively) developing beyond the early oocyst stage. They demonstrated that P. mexicanum developed in phlebotomine sand flies while P. agamae developed in Culicoides nubeculosus, not in mosquitoes as previously suspected. Ayala (1971) further demonstrated sporogony and experimental transmission of P. mexicanum by intraperitoneal inoculation of sporozoites from wild caught Lutzomyia vexator females that had earlier fed on infected Sceloporus occidentalis lizards. He did not demonstrate transmission by bite of infected flies. Ayala (1971) further suggested

-16-




-17


that the natural route of infection is by bite because (1) sporozoites occur in the salivary glands (2) time for bloodfeeding was long and (3) infectivity of sporozoites occurred after intraperitoneal inoculation. Ayala did not rule out the possibility of transmission by ingestion of infected flies. Transmission of P. agamae is believed to be by bite but the sporozoites were retained in the oocyst and did not migrate to the salivary glands of C. nubeculosus (an unnatural vector) (Petit et al., 1983).

New developments in rearing phlebotomine sand flies (Endris et al., 1982) provided the opportunity for experimental transmission studies of P. mexicanum. This study describes the first successful experimental transmission of a saurian malaria by bite of a hematophagous insect other than mosquitoes. The incubation period and course of acute infection of P. mexicanum in Sceloporus undulatus, transmitted by bite of L. vexator, are also reported.



Materials and Methods


Lutzomyia vexator Colony Maintenance



The colony of Lutzomyia vexator, originating from wild caught females from Gulf Hammock, Levy Co., Florida, USA, in 1981, was maintained by methods similar to those described by Endris et al. (1982). Larvae, however, were provided




-18


finely ground horn fly larval medium (Greer and Butler, 1973) that decreased the larval development time from that observed in larvae fed on aged rabbit feces (Young et al., 1981). Larvae from individual females were transferred from 25 ml plastic oviposition vials (12-20 days post-eclosion) to 120 ml urine specimen containers. Approximately 200 larvae were placed in each large vial. Adults were released daily from the 120 ml containers into a modified glass aquarium (34x21x27 cm) and were provided slices of apple as a food source (Endris et al., 1982). All developmental stages, including bloodfed L. vexator females were maintained in a Hotpack incubator (temperature, 270 or 240 + 1C; relative humidity, 80 + 5%; and 16:8 LD photoperiod).



Collection and Laboratory Maintenance of Lizards



Sceloporus occidentalis (western fence lizard) were hand collected at Ramsey Canyon, three miles north of Ramsey, Yolo County, California, USA, and examined for the presence of Plasmodium mexicanum. Infected lizards were sent to the University of Florida for transmission studies. Sceloporus undulatus undulatus (eastern fence lizard),

collected from Austin Cary Forest, Alachua county, Florida, USA, were similarly examined for the presence of P. floridense parasites. Bloodfilms were made from a clipped

toe, air dried, fixed with absolute methyl alcohol, then stained with Giemsa. Subsequent bloodfilms were prepared by





-19


clipping the the tip of the tail. Sceloporus undulatus that did not show patent Plasmodium floridense infections within a 30 day period (from a minimum of three bloodfilms) were used in P. mexicanum transmission studies. Lizards were maintained in screened cages (50 x 25 x 25 cm) in the laboratory at room temperature and provided an external heat source from a 40 watt incandescent light bulb. Lizards were fed house flies (Musca domestica L.) and lepidoptera larvae (Galleria sp. and Spodoptera sp.). Water was provided by spraying the cages daily and by placing a water-filled petri dish in each cage.



Plasmodium mexicanum Transmission Studies



Lab reared L. vexator females were bloodfed on Sc.

occidentalis which demonstrated >1% of the red blood cells infected with P. mexicanum gametocytes. Bloodfed females were removed at 4 hr intervals, placed in 25 ml oviposition vials, provided a sugar source (1:1 mixture of Karo syrup and distilled water) and maintained in a temperaturehumidity controlled chamber as previously described. Midguts were dissected (Chaniotis and Anderson, 1968) at intervals from 2-7 days post-feed (period following initial bloodmeal on an infected lizard) and the number of oocysts counted. In addition, the salivary glands were examined subsequent to day five post-feed, and the sporozoite rate determined (+1, 1-10; +2, 11-100; +3, >100 sporozoites).




-20


One to six female L. vexator infected with P. mexicanum sporozoites were placed in a Plexiglas cage lined with plaster of Paris (Endris et al., 1982) and provided a second bloodmeal on a noninfected, wild-caught Sc. undulatus. Lizards fed on by one or more infected sand flies were placed in a screened cage and maintained as previously described or were placed in a temperature-humidity controlled chamber and maintained at 27 0C and 80% RH. Bloodfed females were dissected after the second bloodmeal and the sporozoite rate determined. To determine if transmission of P. mexicanum could also occur by the oral route, living L. vexator potentially infected with P. mexicanum sporozoites were force fed (placed in the back of the mouth with forceps) to Sc. undulatus.


Course of Infection, Parasitemia


Bloodfilms of Sc. undulatus previously fed on by

infected L. vexator or force fed infected sand flies were made at day 0 post-exposure (period of time from which non-infected lizards were exposed to bites of L. vexator with sporozoites of P. mexicanum) and at 2-4 day intervals subsequent to day 19 post-exposure. Parasites were counted and parasitemias expressed as the number of parasites per 10,000 red blood cells (RBC). Plasmodium mexicanum characteristically occupies all circulating blood cells (Jordan, 1970); therefore, the number of infected white




-21


blood cells per 10,000 red blood cells was also counted. A sufficient number of red blood cells was counted to keep the probable error within 10% according to the method formulated by Gingrich (1932).



Histology of Infected Lizards


Tissue impressions of various organs made subsequent to death for most of the lizards infected with P. mexicanum were fixed with methanol and stained with Giemsa. In addition, tissues from one lizard (S-51) were fixed in Carnoy's fluid, dehydrated, embedded in paraffin and sectioned at 5-6um (um = micron) on a rotary microtome. Thin sections were stained with hematoxylin-eosin or Giemsa-colophonium (Bray and Garnham, 1962).



Results



Plasmodium mexicanum Transmission Studies



Females of L. vexator readily feed on lizards in the

laboratory (Figure 1). Males are also attracted to lizards and mating frequently occurs during blood feeding. Exflagellation can be observed in a bloodmeal by removing the midgut contents within 30 minutes after a female sand fly has completed blood feeding. The length of time during which exflagellation occurs was not determined.


























Figure 1. Bloodfed female Lutzomyia vexator resting on
Sceloporus undulatus upon which it had previously
fed on.

Figure 2. Midgut of Lutzomyia vexator with oocysts (0) of
Plasmodium mexicanum. Day 5 post-feed).

Figure 3. Plasmodium mexicanum sporozoites (S) from
ruptured oocysts (0) on the midgut (Mg) of
Lutzomyia vexator. Day six post-feed on infected
Sceloporus occidentalis and maintained at 27 C.

Figure 4. Plasmodium mexicanum sporozoites (S) in the
salivary gland (Sg) of Lutzomyia vexator
(Nomarski interference contrast).






-23































































1"X




-24


The development time of P. mexicanum in the host, L. vexator, is relatively short at 270C. Oocysts were first seen on day two after feeding, often occurring in large numbers (Figure 2), and developing rapidly at 27 0C (Chapter 3). Sporoblasts with budding sporozoites are observed in some oocysts by day five post-feed. Sporozoites are free in the hemocoel by day six post-feed and are present in the salivary glands by day 6.5 post-feed (Figures 3 and 4). However, when sand flies were maintained at 240C, sporozoites were not observed in the salivary glands until days 8.5-9.0 after feeding. All sand flies used in the transmission study were maintained at 27 oC and provided second bloodmeals on noninfected Sc. undulatus subsequent to day 6.5 post-feed. Oocyst and sporozoite development is reported elsewhere.

The laboratory transmission data of P. mexicanum to non-infected Sc. undulatus by bite of infected L. vexator female(s) are shown in Table 1. A total of 13 Sc. undulatus were fed upon by 1-3 L. vexator females from days 7-10 following initial bloodmeals on Sc. occidentalis infected with P. mexicanum. All sand flies had sporozoites in the salivary glands 0-8 hrs subsequent to their second bloodmeal. Nine (69.2%) of the 13 lizards became infected with P. mexicanum. Two Sc. undulatus were each force fed more than five sand flies (10 days post-feed) potentially infected with P. mexicanum. Neither of these lizards became infected.







TABLE 1. Laboratory transmission data of Plasmodium mexicanum to Sceloporus
undulatus by bite of infected Lutzomyia vexator females.

No. Sporozolte Day Day post Duration No. para %RBC inf %WBC inf Lizard flies rate / patent feed of patent at death at death at death number fed (day post infect- lizard infection (killed) (killed) (killed)
feed) ion died (day)
(killed)

S-8 3 +3(7) 27 52 25 2095 20.3 25.0
+3(7) +3(8)
S-14 1 +3(9) 26 39 13 930 9.2 10.7
S-15 1 +3(8) 26 47 21 2130 20.2 44.4
s 25b 1 +3(9) 33 52 19 2780 25.9 34.3
S-42 1 +3(8) 33 61 28 8270 66.5 55.6
S-43 2 +2(7) 26 (66) (40) (7122) (53.0) (25.0)
+3(10)
S-47 3 +3(8) 23 (45) (22) (3070 (28.5) (37.5)
+3(8) +3(8)
S-50 3 +3(8) 23 (46) (23) (1370) (12.6) (N/D)
+2(9) +3(9)
S-51 I +1(9) 40 96 56 11,960 91.2 24.2
Averaged 28.6 50.2 27.0 4379 38.9 32.4

a +1, 1-10; +2, 11-100; +3, >100 sporozoites. b Yearling lizard.
c Lizard maintained in a temperature-controlled incubator at 27 0C.
Lizards which were killed are not included in the average.




-26


The F1 progeny of two species of mosquitoes, Culex

erraticus and Culex territans, collected in lizard baited traps were also provided bloodmeals on Sc. occidentalis infected with P. mexicanum during the same time as L. vexator. None of the mosquitoes developed oocysts while all of the L. vexator dissected had oocysts (range 9-54, mean 22.1).

Patent P. mexicanum infections were first observed in nine of the experimentally infected Sc. undulatus from days 23-40 post-exposure (mean 28.6 days) (Table 1 and Figure 5). Because lizards were bled only every third day, infections may have been patent as early as two days previous to the positive bloodfilm. The acute infection was allowed to run its course in each of six lizards. The remaining three lizards were killed when they became anorexic and lethargic and probably would have survived only a few days longer. The six lizards which were not killed died of fulminating infections by day 96 post-exposure and became lethargic and anorexic several days prior to death. Force feeding two of the lizards during this critical period did not appear to increase the survivability of the lizards. The period of survival for these lizards varied from 13-56 (mean 27.0) days following the detection of parasites in the bloodfilm and 39-96 (mean 57.8) days post-exposure. Two of the longest living lizards, S-42 and S-51, were adult females that deposited abnormal infertile eggs during the course of the infection.




















Figure 5. Prepatent and patent period of Plasmodium mexicanum infection
and survival of individual Sceloporus undulatus infected by bite
of Lutzomyia vexator infected with Plasmodium mexicanum
sporozoites.
a Killed during the course of the infection.
b Maintained in a temperature controlled chamber at 27 C.
c Yearling lizards.



























w a

U) S-47

I--a
S-43

0l S- 15
z

(0 6-14

0 6-42
0
-jC
w 8-26




0 10 20 30 40 50 60 70 80 90 100


DAY POSTEXPOSURE





-29


Course of Infection, Parasitemia



The number of parasites (expressed as the number of

parasites per 10,000 red blood cells), percent of infected red blood cells, and percent of infected white blood cells (per 10,000 red blood cells) for the day prior to death are shown in Table 1. Bloodfilms during the later course of the infections became increasingly difficult to obtain, apparently as a result of anemia. Excluding lizards S-42 and S-51, which had approximately 4x and 6x the number of parasites, respectively, as the other four lizards which died, the average number of parasites at time of death was 1,983 (19.8%). The percentage of infected red blood cells (number of parasites/10,000 RBC) approximated the percent parasitemia at levels below 25%. However, as the parasitemia increased, the number of multiple infected red blood cells also increased, as demonstrated by S-42 which had 82.7% parasitemia, but only 66.5% of the red blood cells infected (Table 1 and Figures 6 and 7). Although the parasitemia of S-51 did not increase greatly during the later part of the infection (11,420, day 89 post-exposure to 11,960, day 95 post-exposure), the percent of infected red blood cells continued to increase rapidly until nearly every RBC was parasitized (72.4 to 91.2%) (Figures 6, 7, and 14).

The transformed (Y=Log of number of parasites per 10,000 RBC) course of infection and linear regression analysis is shown in figure 7 and Table 2, respectively.




















Figure 6. Percent of infected red blood cells during the course of
Plasmodium mexicanum infection for six Sceloporus undulatus
infected by bite of Lutzomyia vexator.








100


go



080

70


60 *b81
0 -25 50 *8-42

*=8-51
S 40 S 30


20 10


1.0' 0.5


01
20 30 40 50 60 70 80 90 100


DAY POSTEXPOSUFIE






















Figure 7. Course of acute infection of Plasmodium mexicanum in nine
Sceloporus undulatus infected by bite of Lutzomyia vexator.









10000 5000






1000

a
o 500 a. 10


I



0
= S-4 10

a 0 = S50



5 S-51

=Mean






20 25 30 35 40 45 50 55 60 65 70 75 s0 85 90 9

DAY POSTEXPOSURE










TABLE 2. Linear regression analysis of acute Plasmodium mexicanum
infections with fewer than 500 parasites per 10,000 red
blood cells (5% parasitemia).

Lizard Days sur- Days sur- Linear regression equation number vived post- vived post- of acute infection wigh R Slope
exposure patent inf. <5% parasitemia/(SE)


S-8 52 25 Y= -11.35 + 0.42x .96 .417
(1.14) (0.03)
S-25 52 19 Y= -14.44 + 0.45x .96 .452
(1.94) (0.05)
S-42 61 28 Y= -10.05 + 0.36x .95 .361
(1.79) (0.05)
S-14 39 13 Y= -10.34 + 0.47x .96 .471
(1.93) (0.06)
S-15 47 21 Y= 9.74 + 0.42x .99 .420
(0.67) (0.02)
S-43a 66 40 Y= 4.57 + 0.31x .93 .309
(1.86) (0.06)
S-47 46 23 Y= -14.28 + 0.64x .97 .640
(2.11) (0.07)
S-50a 45 22 Y= 8.51 + 0.42x .86 .417
(2.86) (0.09)
S-51 96 56 Y= 5.71 + 0.23x .93 .230
(1.73) (0.04)
a Lizards were killed when lethargic and anorexic.
Y=Estimated number of P. mexicanum parasites per 10,000 RBC on day
post-exposure.





-35


The progression of the acute infection can be explained as a exponential linear relationship for parasitemia levels of fewer than 500 parasites/l0,000 RBC (5% parasitemia). In general, the slopes of the acute infections of the lizards with less than 5% parasitemia were similar. Except for lizard S-47 (patent infection on day 23 post-exposure), S-51 patent infection on day 40 post-exposure), and S-43, the slopes of parasitemia increase were not significantly different (p=.01). Lizards S-42, S-43 and S-51 that survived the longest and which had terminal parasitemias of >70% (71.2, 87.7, and 119.6%, respectively) had the lowest slopes (rate of increase in parasitemia) (Table 2). However, subsequent to parasitiemias of >500 parasites per 10,000 RBC, the parasitemia increases at a reduced rate and is better explained as a quadratic relationship. When considering all lizards that died or were killed (and probably would have died within a few days), there were significant differences (p=.Ol) in the curves of the quadratic equation. However, the curves are similar enough to average over all lizards (Figure 7). A regression of parasitemia over the course of the infection for all lizards was performed (R2=.88). The initial positive bloodfilm (patent infection) was adjusted to begin on the mean day of patent infection (28.6) since we were interested in the average course of infection. The predicted parasitemia over the course of the infection is shown by the dotted line (Figure 7).





-36


The number of trophozoites, schizonts, gametocytes and percent of white blood cells infected with P. mexicanum during the course of infection for each of the lizards which died is shown in figures 8 to 13. Both immature (single nucleated parasites larger than the nucleus of the host cell, but not displaceing the host cell nucleus) and "mature" gametocytes (as described by Garnham, 1966) are included together. The number of trophozoites increased logarithmically during the course of the infection. In general, the numbers of schizonts and gametocytes also increased logrithmically, but appeared to show more variation. This may be partially due to fewer numbers observed and greater chance of error. Schizogony appeared to develop synchronously in only one lizard (S-51), and then only after day 23 following the detection of parasites in the bloodfilm (Figure 8). From the limited numbers of bloodfilms, it appears that schizogony occurred at about 3-4 day intervals. In addition, thrombocytes and circulating white blood cells were also infected. The percentage of infected white blood cells generally increased as the number of red blood cells increased (Figures 8 to 13). However, the low and variable number and rupture of white blood cells during a bloodfilm preparation increased the potential for

error.




















Figure 8. Number of Plasmodium mexicanum trophozoites, schizonts,
gametocytes and percent of infected white blood cells per 10,000 red blood cells during the course of the infection
of Sceloporus undulatus (S-51) infected by bite of
Lutzomyia vexator.






-33









































cc

0
0.
x ui

0























0 0 x N W
CL
0 x 2
In






co cm

OSW 000'01/08M 03103ANI %







OSW 00WOt/S3iJSVkfVd :io tosnm 001



























Figure 9. Number of Plasmodium mexicanum trophozoites,
schizonts, gametocytes and percent of
infected white blood cells per 10,000 red
blood cells during the course of the
infection of Sceloporus undulatus (S-42)
infected by bite of Lutzomyia vexator.





-40







10000



5000 TROPHOZOITES

O= SCHIZONTS
=GAMETOCYTES
e= WBC


o 1000 co 50


Soo

co 60



CL 100 so
U
0
LU C.0
W 50
S40

00

-I '3010

10
w
5 U
z

10



1

25 30 35 40 45 50 55

DAY POSTEXPOSURE



























Figure 10. Number of Plasmodium mexicanum trophozoites,
schizonts, gametocytes and percent of
infected white blood cells per 10,000 red
blood cells during the course of the
infection of Sceloporus undulatus (S-8)
infected by bite of Lutzomyia vexator.




-42











10000



5000




*TROPHOZOITES
0= SCHIZONTS
O1000 *GAMETOCYTES

e= wec o 500



m0 0
0

w 5 I-0
an


0
40

10 0 zc
20


z 0





50 354w5 5 5 6
DA OT-PSR



























Figure 11. Number of Plasmodium mexicanum trophozoites,
schizonts, gametocytes and percent of
infected white blood cells per 10,000 red
blood cells during the course of the
infection of Sceloporus undulatus (S-15)
infected by bite of Lutzomyia vexator.





-44









1000



5000



TROPHOZOITES
0=SCHIZONTS

1000 GAMETOCYTES
e= wBC





w
I



0. 100 50.
U
0
wU 50
~40

00
0
10


10

5 W


10





25 30 35 40 45 50

DAY POSTEXPOSURE



























Figure 12. Number of Plasmodium mexicanum trophozoites,
schizonts, gametocytes and percent of
infected white blood cells per 10,000 red
blood cells during the course of the
infection of Sceloporus undulatus (S-25)
infected by bite of Lutzomyia vexator.














10000


5000





*=TROPHOZOITES
O= SCHIZONTS
a1000 =GAMETOCYTES
a=WBC S500




0
0 5 w 4



lo


02
5
40

U. 1 zz


1J L 3035445s556

DA OTEPSR



























Figure 13. Number of Plasmodium mexicanum trophozoites,
schizonts, gametocytes and percent of
infected white blood cells per 10,000 red
blood cells during the course of the
infection of Sceloporus undulatus (S-14)
infected by bite of Lutzomyia vexator.





-48






10000



5000 TROPHOZOITES
0= SCHIZONTS
= GAMETOCYTES
a= wBC


o 1000
0
0
0 50
CL

co



CL 100 U
0
WU 50


z

0 c

00
00


5


0 10
U




25 30 35 40 45

DAY POSTEXPOSURE





-49


Histology of Infected Lizards



Numerous parasites were observed in lymphocytes of

spleen tissue impressions of dead lizards (dead less than 2 hrs) (Figure 15). Immature erythrocytes and lymphocytes from bloodfilms of bone marrow extracts were also infected with P. mexicanum parasites (Figure 16). However, the number of parasites observed in the spleen tissue impressions was much greater and appears to be the primary site of attack. Schizogony was also observed in the endothelial cells of the brain in some lizards (Figure 17). Other tissues, i. e., lung, liver, kidney, intestine, pancreas, heart, and uterus were considered to be non-infected. Occasionally, cells of the above tissues appeared to be infected, but it could not be determined if the parasites were external from ruptured white blood cells or intracellular, since fixed or circulating lymphocytes within these tissues often had numerous parasites. Thin sections from one lizard, (S-5i) also failed to reveal parasites in these tissues.



Discussion


Plasmodium mexicanum Transmission Studies



Data reported herein demonstrate conclusively that P. mexicanum can be transmitted from Sc. occidentalis to S.























Figure 14. Red blood cells of Sceloporus undulatus (S-51)
infected with Plasmodium mexicanum parasites
during the later part of the infection
(Giemsa). Schizont (Sc), and trophozoite (T).

Figure 15. Spleen tissue impression of Sceloporus
undulatus infected with numerous Plasmodium
mexicanum parasites (Giemsa). (Merozoites
(M) and schizont (Sc).

Figure 16. Bone marrow smear of Sceloporus undulatus
infected with Plasmodium mexicanum parasites
(Giemsa). Schizont (Sc) in a white blood cell
and trophozoite (T) in a thrombocyte (Th).

Figure 17. Schizonts (Sc) of Plasmodium mexicanum in
endothelial cells (En) of capillaries in the
brain. Brain tissue impression stained with
Giemsa.










































S c













- loum 17





-52


undulatus by bite of L. vexator, a species that coexists with the parasite in California. Progeny of two species of mosquitoes, Cx. erraticus and Cx. territans, collected in lizard baited traps failed to transmit this parasite while as many as 50 oocysts developed on the midgut of L. vexator. feeding on the same lizard. This provides further evidence that sand flies are the natural vectors of P. mexicanum, especially since sporogony and transmission of P. floridense has been demonstrated for Cx. erraticus (Chapter 4).

Sporozoite infectivity of P. mexicanum transmitted by bite of L. vexator is relatively high and comparable with some other malarias (Coatney et al., 1945; Russell and Mohan, 1942). Transmission occurred in 62% (5/8) of the Sc. undulatus which were bloodfed on by one sand fly and 80% (4/5) of the Sc. undulatus which were bitten by two or more sand flies. Sporozoites were occasionally observed in the mouthparts of dissected sand flies, but only in small numbers (<5). One lizard (S-51), having the longest prepatent period, became infected when bitten by a sand fly with fewer than five sporozoites observed near the head and in the salivary glands following the second bloodmeal (Figures 5 and 8). Evidence indicating that only a few sporozoites are required to infect the host is provided by Fink (1968) for Plasmodium cathemerium and Vanderberg et al. (1968) for Plasmodium berghei. Vanderberg et al. and Fink, respectively, demonstrated that 26% of the mice in the study became infected when injected with approximately 10





-53


sporozoites of P. berghei, whereas 50% of the canaries became infected when inoculated intravenously with as few as five sporozoites. Also, intradermal injection of as few as 10 sporozoites of Plasmodium vivax is sufficient to infect man (Shute et al., 1976). A review of the kinetics of sporozoite injection by mosquitoes indicates that only a few sporozoites are injected during feeding (Vanderberg, 1977).

The influence of temperature on sporozoite development and maturation of malaria parasites is well documented (Vanderberg and Yoeli, 1966; Stratman-Thomas, 1940). For P. mexicanum, sporozoite development in lab-reared L. vexator maintained at 27 0C is rapid, with these forms being observed in the salivary glands by day 6.5 after feeding. However, sporozoites were not observed in the salivary glands until day 8.5-9.0 post feed for sand flies maintained at 240C. Observations by Ayala and Lee (1970) indicated that sporozoites from lab-infected wild caught sand flies were not observed in the hemocoel until day 11-14 post-feed when maintained at room temperature (24-26 oC). Maximum and minimum temperatures when sporozoite development ceases were not determined.

Vanderberg (1975) showed that sporozoites require a

period of maturation after their release from the oocyst and that P. berghei sporozoites in the salivary glands of a mosquito are 10,000 times more infective than sporozoites from the oocyst of the same mosquito. However, once released from the oocyst, the development of infectivity





-54


appears to be time-dependent rather than site-dependent, i. e., hemocoel sporozoites in some cases being equally infective as salivary gland sporozoites. Only one lizard, that did not become infected, was fed on by an infected sand fly on day 7.0-7.5 after its initial bloodmeal. The sand fly was dissected 0-8 hrs following the second bloodmeal, and had more than 100 sporozoites in the salivary glands. Nine of the 12 (75%) remaining lizards, which were bloodfed on by 1-3 infected sand flies 8-10 days after their initial bloodmeal on infected Sc. occidentalis became infected, indicating that sporozoites are infective within eight days of feeding (Table 1).

Both attempts to transmit P. mexicanum by ingestion of whole sand flies were unsuccessful. Transmission by oral ingestion of sporozoites has been reported to be occasionally successful under certain laboratory conditions for malarias transmitted by mosquitoes (Shortt and Menon, 1940; Young, 1941; Porter et al., 1952; Yoeli and Most, 1971) and may be the mode of transmission for P. agamae (Petit et al., 1983). However, it is believed that oral transmission will only occur if the sporozoites are released in the mouth and penetrate the tissues of the mouth and throat, since sporozoites are quickly killed in acid concentrations similar to that of the gut. Although hatchling lizards may eat sand flies, unrestrained yearlings were rarely observed feeding on sand flies and mature Sc. occidentalis and Sc. undulatus were never observed feeding





-55


on L. vexator in the laboratory. Since mastication of ingested flies does not normally occur, and both attempts to orally transmit P. mexicanum failed, transmission by the oral route is believed to have little epidemiological

significance.

Prepatent periods in experimentally transmitted P.

mexicanum by bite of infected L. vexator ranged from 23-40' (mean 28.6) days. In examining natural infections of another lizard malaria, P. floridense, Goodwin (1951) showed that parasites were not observed in bloodfilms until approximately two weeks after the lizards were collected. In another study, parasites were not observed in the bloodfilm of one lizard until 27 days after capture (Goodwin and Stapleton, 1952). Present studies on the transmission of P. floridense indicate that the prepatent period is more than 20 days for bite induced and IP induced infections at 18-240C (Chapter 4). These studies and those by Goodwin and Stapleton (1952), assuming that the naturally acquired infections were not relapses, and other studies on P. mexicanum laboratory transmission, support the hypothesis of a lengthy prepatent period for at least two of the saurian malarias.

Recent studies (Chapter 6) on other hemosporidians,

Schellackia golvani and Schellackia occidentalis, indicate that temperature may affect the length of the prepatent period. For lizards maintained at room temperature (18-240C), Schellackia sporozoites were not observed in




-56


bloodfilms until day 21 and 37 post-ingestion, respectively. However, when lizards were maintained at 320C (900F), sporozoites were seen in the bloodfilms as early as day 10 and 7 post-ingestion, respectively. Similarly, the prepatent period of P. floridense was decreased by as much as seven days when maintained at 320C (Chapter 4).

In general, lizards which were fed on by more than one infected sand fly developed earlier patent P. mexicanum infections. One lizard (S-51) which was bloodfed on by only one sand fly in which fewer than five sporozoites were observed in the salivary glands and around the head 0-8 hrs after the second bloodmeal did not develop a patent infection until 40 days after feeding. Ayala (1971) showed that Sc. occidentalis inoculated with sporozoites from five sand flies (some having more than 100 oocysts on the midgut) had a prepatent period of 21 days. Observations on human, rodent and avian malarias show that, within limits, the higher the inoculum of sporozoites, the shorter the prepatent period (Boyd, 1940; Greenberg et al., 1950; Vanderberg et al., 1968). Based on the present studies, the length of the prepatent period appears to be a response of the number of sporozoites. However, host-temperature maintenance in relation to parasite growth and development may also play an important role in the early course of the infection.

Natural infections of P. mexicanum occur in both Sc. occidentalis and Sc. undulatus (Ayala, 1971; Greiner and





-57


Daggett, 1973). Separate studies indicate that both species are highly susceptible to P. mexicanum and often die of fulminating infections during the acute phase (Ayala, 1971; Jordan, 1970; Thompson and Huff, 1944a; and Thompson, 1944). The age of the lizard (Sc. occidentalis) also appears to have a significant effect on the course of the infection. When hatchling lizards (3-5 months old) were blood inoculated with P. mexicanum, all lizards died of fulminating infections but only three of 10 wild caught naturally infected yearling Sc. occidentalis died (Ayala, 1971). However, the course of the infection for blood inoculation of some malarias is often more severe than sporozoite inoculation. In the present studies, two yearling and seven mature S. undulatus collected in Florida were experimentally infected by bite of L. vexator. Six of the lizards (including the two yearling lizards) died of fulminating infections within 96 days after exposure. The other three lizards were killed when they became lethargic and parasitemias reached approximately 20-75%.



Course of Infection, Parasitemia


Parasitemias of P. mexicanum ranged from 930 (yearling) to 11,960 (mature female)/1O,000 RBC at the time of death (Table 1, Figure 7). Maximum parasitemias attained by P. mexicanum from previous studies ranged from 4,100 to 8,100 for Sceloporus olivaceaous, 2,812 for Sc. undulatus





-58


undulatus, and 2,750 for Sc. undulatus consobrinus (Thompson and Huff, 1944a; Thompson, 1944). Maximum parasitemias for unnatural hosts, Phrynosoma cornutum and Crytophytus collaris only reached 392 and 238, respectively (Thompson and Huff, 1944a). Results in the present studies are similar to those of Thompson and Huff (1944a) and Thompson (1944), except that several of the Sc. undulatus which were blood inoculated in the previous studies did not develop fulminating infections. The higher parasitemias in some lizards in the present study may be a result of a larger sample size.

The course of the acute infection for Sc. undulatus infected with P. mexicanum is shown in Figure 7. The predicted mean parasitemia for an "average" Sc. undulatus infected with sporozoites, plotted on the mean day that parasites were first observed in the bloodfilm, is shown by the dotted line (R2=.88). The coefficient of determination

(R2), linear regression equation and slope for parasitemia levels of less than 5% for each lizard are shown in Table 2. In general, it appears that lizards which survive for a longer period of time (>25 days) subsequent to patent infection, develop parasitemias at a slower rate (slope <0.40). However, these infections are observed over a longer period of time and lizards develop higher parasitemias (>70%). Although the curves of the transformed course of infection appear to be similar, there are significant differences between some of the curves. These





-59


differences may be attributed to age and sex of the lizards (adult females surviving the longest), host immune response to the parasite, adaptation to a laboratory environment, temperature [orientation to the light source (behavior)], length of time surviving patent infection, number of sporozoites inoculated during feeding, and other factors which were not determined.

According to Thompson and Huff (1944a), variation in

the course of infection, gametocyte production, and cellular distribution of P. mexicanum parasites is due to host differences rather than alteration of parasites. They found that P. mexicanum, a parasite found naturally in Sc. occidentalis, Sceloporus torquatus (=ferrariperzi), and Sc. undulatus, lost its gametocytes when transferred by blood inoculation to another lizard, C. collaris. However, gametocyte production resumed upon experimental passage of the parasite to a third host species, Sc. olivaceous. "Mature" gametocytes were observed in all lizards except S-14 and S-15 during the infection. Lutzomyia vexator that were bloodfed on S-51 and S-42 on days 11 and 21, respectively, following the detection of parasites and when mature gametocytes were present in the blood, developed low numbers of oocysts. Sporozoites developed normally and appeared viable, but were not injected into another lizard to determine infectivity. It was not determined if gametocyte production would be increased during the chronic phase of the infection since all lizards died (or were killed) during the acute phase.





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Histology of Infected Lizards



These studies indicate that P. mexicanum parasites primarily invade erythrocytes and lymphocytes of Sc. undulatus. Thrombocytes are less frequently invaded. No attempt was made to distinguish between granulocytes and macrophages. Determining the percentage of lymphocytes infected was often difficult since occasionally many of the cells ruptured in bloodfilm preparation, especially during the latter part of the acute phase. These results agree with those of Thompson and Huff (1944a) who observed that 93% of the circulating cells infected with P. mexicanum are in erythrocytes, with a small percentage in lymphocytes and thrombocytes.

Tissue impressions of the spleen and bone marrow revealed numerous asexual forms of P. mexicanum in lymphocytes (Figure 15 and 16). Although exoerythrocytic

(EE) forms are often seen in lymphocytes (Jordan, 1970; Thompson and Huff, 1944a), they are rarely observed in other tissues in Sc. undulatus. However, in unnatural hosts such as Phrynosoma cornutum and C. collaris, Plasmodium mexicanum EE forms are frequently observed in fixed connective tissue while occurring less frequently in the circulating cells (Thompson and Huff, 1944a). When studying fixed tissues, Jordan (1970) recovered schizonts and segmenters from impressions of internal organs, especially endothelial cells of the brain capillaries. In the present study, impressions





-61


of the brain of some lizards also demonstrated schizogony in the endothelial cells (Figure 17). Endothelial cells were heavily infected in some lizards. However, attempts to identify parasites in liver, lung, intestine, pancreas, heart, and uterus from tissue impressions of four Sc. undulatus were unsuccessful.














CHAPTER 3
SPOROGONY, DEVELOPMENT, AND ULTRASTRUCTURE OF EXTRINSIC
STAGES OF Plasmodium mexicanum


Introduction


The natural vectors of saurian malaria have eluded researchers for years. Attempts to incriminate culicine mosquitoes largely resulted in failure, with sporogony being limited and erratic in the mosquitoes. While there are more than 55 species of saurian malaria, only two vectors, Lutzomyia vexator and Lutzomyia stewarti have been incriminated (Ayala, 1971). Recent laboratory transmission studies (Chapter 2) provides more conclusive evidence that L. vexator is a natural vector of Plasmodium mexicanum. Attempts to incriminate other hematophagous insects have largely failed. However, Petit et al. (1983) demonstrated that Plasmodium agamae developed in Culicoides nubeculosus, an unnatural host, and speculated that another species of Culicoides which is sympatric with P. agamae may be the natural vector.

The details of the subcellular morphology and differentiation of many of the avian and mammalian sporozoites and sporogony as revealed by the transmission electron microscope (TEM) are well documented. The only

-62-




-63


ultrastructure of sporogony of a saurian malaria was that done by Boulard et al. (1983) on P. agamae in C. nubeculosus.

The only known vectors of mammalian and avian malarias are mosquitoes. In view of histological and vector differences between P. mexicanum and other malarias, TEM studies were undertaken. Data from this study are compared with those of other Haemosporina.


Materials and Methods


Sporogony of Plasmodium mexicanum


Laboratory reared phlebotomine sand flies, Lutzomyia vexator, were bloodfed on wild caught Sceloporus occidentalis infected with P. mexicanum as previously described (Chapter 2). Bloodfed females were removed at 4 hr intervals, placed in 25 ml plastic oviposition vials, provided a sugar source (1:1 mixture of Karo syrup and distilled water), and maintained in a temperature-humidity controlled chamber at 27 oC, 240C, or 190C, and 80% RH. Midguts were dissected (Chaniotis and Anderson, 1968) at 12 hr intervals from days two through nine, placed on a clean slide with a drop of cold-blooded Ringer's solution, covered with an 18mm circular coverslip, and the number and measurements of 25 (or fewer) oocysts recorded. Salivary glands were examined after day five following a bloodmeal




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and the sporozoite rate determined (+1, 1-10; +2, 11-100; and +3 >100 sporozoites).


Ultrastructure of Extrinsic Stages of Plasmodium mexicanum


For ultrastructural examination, sand flies were fixed in 2% glutaraldehyde in 0.1M sodium cacodylate buffer at pH

7.2 for 1 hr at room temperature. Post-fixation was with 1% osmium tetraoxide in O.1M sodium cacodylate buffer, pH 7.2, for 1 hr at room temperature. Specimens were dehydrated through a graded ethanol series and acetone and embedded in Spurr's resin (Spurr, 1969). Sections were cut on an LKB ultratone III with a Diatome diamond knife, floated on water, picked up on formvar-coated grids, and post-stained with aqueous 1% uranyl acetate followed by Reynold's lead citrate. To enhance the polar rings, sections with sporozoites were post-stained with aqueous 1% uranyl acetate for 1.5 hr at 600C. Material was examined and micrographs taken on a Hitachi 11-E or Jeol 100CX electron microscope.


Results


Sporogony of Plasmodium mexicanum


Oocysts of P. mexicanum developing on the midgut of L. vexator were observed from days 2-9 post-feed. The effects of two different temperatures on the sporogony of P.






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mexicanum in L. vexator are summarized in Table 3. The mean oocyst size, regressed against the days post-feed for sand flies maintained at 24C (R2= .85) and 270C (R2=.91), is shown in figure 18. Sand flies maintained at 190C and dissected at days four and five after feeding did not show any oocysts. The maximum and minimum temperatures under which oocyst development ceases were not determined.

Figures 19 to 23 illustrate oocyst and sporozoite

development from days 2-7 in sand flies maintained at 270 C. By day two post-feed, P. mexicanum oocysts on the midgut of sand flies maintained at 270C are spherical, relatively uniform in size [range 10.8 to 13.5um (um = micron) in diameter] and undifferentiated. Differentiation, i. e., sporoblastoid formation and the formation of sporozoites for sand flies maintained at 27 0C is not evident until day five after feeding (Figures 19 and 20). Many oocysts contain nearly developed sporozoites by day 6.0 (Figure 21). Shortly after day 6.0, some oocysts have released sporozoites into the hemocoel while others are easily ruptured by pressure from the coverslip (Figure 22). Sporozites are observed in the salivary glands by day 6.5 post-feed (Figure 23). Sporogonic development culminating in the invasion of the salivary glands of sand flies maintained at 24C is much slower, not being observed in the salivary glands until day 8.5-9.0 post-feed. The infectivity of sporozoites from sand flies maintained at 240C was not determined. However, sand flies maintained at




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TABLE 3. The effects of temperature on sporogony of
Plasmodium mexicanum in Lutzomyia vexator.


240 C 270 C
Day Mean Mean
postfeed oocysj Rangeb SE oocysj Rangeb SE
size (mean) size (mean)
(microns) (microns)


2.0 ...... 11.6 11.6 -2.5 ...... 15.7 15.1-16.2 0.31
3.0 ...... 16.2 15.3-18.4 0.56
3.5 .-- -- 18.8 17.9-19.7 0.51
4.0 15.2 14.3-16.0 0.29 23.0 20.2-26.4 0.65
4.5 .-- -- 24.6 23.8-25.4 0.33
5.0 18.8 17.8-20.0 0.28 30.4 29.0-31.6 0.78
5.5 .-- -- 31.4 30.9-31.8 0.46
6.0 24.5 22.4-26.5 0.61 31.8 30.6-34.7 0.60
6.5 .-- -- 33.5 33.5 -7.0 29.1 27.2-30.7 0.60 30.6 27.5-34.1 1.93
8.0 29.1 26.6-32.4 0.87 -- -8.5 34.2 32.4-36.0 1.8
9.0 31.8 25.7-39.2 1.1
a Overall' mean oocyst size by day.
Mean oocyst size for sand flies by day.






























Figure 18. Regression of Plasmodium mexicanum mean oocyst
size (and 95% confidence limits) in Lutzomyia
vexator and day post-feed for days 2 through 9.





-68


















40







C30
z


~25
wU
N

1.-20 1


0
0 15 x =27.O0C LU 0 =24.C
> 10 S Sporozoltes



5


0
2 3 4 5 6 7 8 9

DAY POSTFEED























Figure 19. Midgut (Mg) of Lutzomyia vexator (day five
post-feed) with asynchronous development of
Plasmodium mexicanum oocysts (0).

Figure 20. Sporoblast (Sb) formation in 5 day old oocyst
on the midgut (Mg) of Ltzomyia vexator. Sand
flies were reared at 27 C.

Figure 21. Plasmodium mexicanum oocyst (0) on the midgut
(Mg) of Lutzomyia vexator with many nearly
mature sporozites T(S).

Figure 22. Sporozoites (S) of Plasmodium mexicanum which
ruptured from the oocysts (0) on the midgut
(Mg) of Lutzomyia vexator.

Figure 23. Salivary gland (Sg) of Lutzomyia vexator
containing numerous sporozoites (S) of
Plasmodium mexicanum.





-70






4:u




Sb, Mg



20


0












21




S







23





-71


27 C were highly infective by days 8-10 post-feed with 9 of 13 Sceloporus undulatus becoming infected with P. mexicanum when bitten by 1-3 sand flies with sporozoites in the salivary glands (Chapter 2).


Ultrastructure of Extrinsic Stages of Plasmodium mexicanum


Oocyst. Electron micrographs of oocysts of P.

mexicanum on the midgut of L. vexator were taken from days 4-7 post-feed. The early oocyst (day 4) appears as an undifferentiated solid sphere in which the oocyst capsule [range, 10.8-27.0um; mean 23.0um (Table 3)] is extracellular, directly applied to the oocyst plasma membrane, and protruding into the hemocoel (Figure 24). The oocyst capsule is in direct contact with the basement membrane of the midgut epithelial cells and is composed of an amorphous granular material which is slightly more electron dense than the basement membrane. Intracellular development of oocysts in the midgut epithelium was not observed. Changes in the oocyst capsule are similar to those described by Duncan et al. (1960) for P. cathemerium and Vanderberg et al. (1967) for P. berghei. The external and internal margin of the oocyst capsule is smooth in early oocysts, where it is not in contact with the midgut. The margin of the oocyst capsule in contact with the basement membrane is often thickened and invaginates into the plasma membrane of the oocyst (Figure 24). As the oocyst matures,

























Figure 24. Cross section of a 4 day old Plasmodium
mexicanum oocyst on the midgut of Lutzomyia
vexator. The solid, non-vacuolated oocyst
protrudes into the hemocoel (H), is bounded by
a capsule (C) and is in contact with the basement membrane of the sand fly midgut epithelium (E). Large nuclei (N) with a
distinct nucleolus (Nu), mitochondria (M), and
endoplasmic reticulum (Er) are scattered
throughout the cytoplasm. Inclusion bodies of
very dense granules associated with a lesser dense area (Di) are also scattered througout
the cytoplasm.








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-74


the capsular material usually appears thickened with irregular projections toward he sporoblastoid plasma membrane along the entire margin of the oocyst. The internal capsular projections remain after the separation of the sporoblastoid from parts of the oocyst capsule, giving it the appearance of a scalloped margin (Figures 25 to 27).

The nuclei in early oocysts are large, scattered

throughout the cytoplasm and enclosed in a nuclear envelope which is marked by occasional nuclear pores (Figure 24). The nucleoplasm is about the same density as the cytoplasm. A large distinct dense nucleolus is observed in some nuclei (Figure 24). Mitochondria appear to be in scattered clumps throughout the cytoplasm in early oocysts (Figures 24 to 29). They vary from circular to tubular in shape and have tubular cristae.

Both granular and smooth endoplasmic reticulum (ER) are found in the cytoplasm (Figures 24 to 26). However, it appears that smooth or rough ER is more numerous in more developed oocysts (Figures 24 to 26). Numerous typical ribosomes are also scattered throughout the cytoplasm, giving the cytoplasm a granular appearance. Three types of membrane bound inclusions similar to those observed by Terzarkis (1971), i. e., circular granules of moderate density, very dense granules associated with a space of low density, and lamellar forms usually associated with a irregular space of low density, are seen in the cytoplasm (Figures 24 to 27). These inclusions appear to be more numerous in more developed oocysts.












Figure 25. Cross section of Plasmodium mexicanum oocyst
undergoing differentiation. Early
vacuolization (V) with convex dense membranes
(Dm). Nuclei (N) with an electron dense
nucleolus (Nu), endoplasmic reticulum (Er), and numerous mitochondria (M) which appear in groups
are scattered throuout the cytoplasm.

Figure 26. Cross section of Plasmodium mexicanum oocyst
undergoing differentiation. The sporoblastoid
(Sbs) is completely separated from the oocyst capsule (C). Convex dense membranes (Dm) are present along the margin of the sporoblastoid
body. Inclusion bodies (I) are seen in groups
or scattered througout the oocyst.

Figure 27. Cross section of Plasmodium mexicanum oocyst
undergoing internal vacuolization of the
sporoblasoid (Sbs). Lamellar inclusion bodies
(Li),very dense granules associated with a
space of low density (Di) and golgi apparatus
(Go) are present. Sporozoite buds (B) from the convex dense membranes form along the periphery
of the sporoblastoid and along the internal
membranes caused by vacuolization of the
sporoblastoid.

Figure 28. Coelescence of the internal vacuolization of
Plasmodium mexicanum oocysts produces cytoplasmic clefts (Cc) which are the
precursors for the sporoblasts. A moderately dense inclusion body is often associated with
the convex dense membranes (Dm) during this
stage.

Figure 29. Higher magnification of Plasmodium mexicanum
oocyst. The sporoblast has pulled away from
the oocyst capsule, showing a convex dense membrane (Dm) beneath the plasma membrane.
Moderately dense vacuoles (Mi), which are
scattered throughout the oocyst, migrate and
come to lie beneath these convex dense
membranes. Microtubules (arrows) associated
with the convex dense membranes are seen in
cross section.






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-77


Sporozoite formation. Figures 25 to 28 illustrate

differentiation of the sporoblastoid into sporoblasts and the initial development of sporozoites. Differentiation of the sporoblastoid and development of sporozoites is a dynamic process within the oocyst and occur concurrently. Differentiation of the oocyst is initiated by the formation of irregularly-shaped vacuoles in the sporoblastoid body and immediately beneath the oocyst capsule (Figure 25). A subcapsular space is formed along the periphery of the sporoblastoid as the sporoblastoid plasma membrane contracts away from the oocyst capsule. Linear to convex dense membranes are sometimes present along the sporoblastoid membrane during vacuolization, but are more numerous following complete separation of the sporoblastoid from the oocsyt capsule (Figures 26 and 29). Large vacuoles form within the sporoblastoid body (Figure 27). Coelescense of the internal cytoplasmic vacuoles and their extension to the sporoblastoid surface produces clefts which subdivide the oocyst cytoplasm into several sporoblasts (Figure 28).

Figure 29 illustrates the initial sporozoite formation which is characterized by dense inner thickened convex membranes (= linear dense areas) which develops beneath the sporoblast plasma membrane. Immediately inside and along the newly condensed inner membrane, a single row of microtubules is formed. The outer sporoblast plasma membrane continues to evaginate and forms the outer sporozoite membrane, while the denser inner membrane forms





-78


along the junction of the newly evaginating sporozoite, becoming the inner sporozoite membrane. A moderately dense inclusion body forms or migrates immediately adjacent to the newly formed inner membrane and is the first cytoplasmic structure to enter the developing sporozoite (Figure 29). As the sporozoites mature, they elongate into stout cylindrical forms, in longitudinal section, which are loosely packed within the oocyst capsule (Figure 30). In more developed sporozoites, several inclusion bodies may be seen anterior to the nucleus (Figure 31). The anterior inclusion bodies become less pronounced as the rhoptries elongate and micronemes become denser in nearly formed sporozoites. Moderately dense anterior inclusion bodies are usually absent in sporozoites which are in the salivary glands of the sand fly.

Endoplasmic reticulum (granular and smooth) become more abundant as vacuolization and sporozoite formation is initiated and are frequently observed in association with the budding sporozoites (Figures 25 to 27). The nuclei are smaller and in the early stages of sporozoite formation migrate to the periphery of the sporoblast. As the sporozoite elongates and following the inclusion body entering the sporozoite, the nearly spherical nucleus moves into the sporozoite, becoming more elongate in shape as it takes the shape of the sporozoite (Figure 31). Following the nucleus is a spherical mitochondrion (Figure 31). While circular and tubular mitochondria are observed in the















Figure 30. Oocyst of Plasmodium mexicanum with developing
sporozoites. Sporozoites of P. mexicanum are loosely packed within the oocyst capsule (C).
Sporozoites bud off the sporoblast leaving a
residual body (Rb). The nucleus and
moderately dense inclusion body (Mi) are
present in the anterior of the sporozoite.
Short bag-like rhoptries (Rh) are seen in a few
sporozoites.

Figure 31. Higher magnification of developing sporozoites
of Plasmodium mexicanum, illustrating the migration of the nucleus (N) and spherical
mitochondrion (M) into the sporozoite. The
moderately dense inclusion bodies (Mi) appear to
be degenerating as the rhoptries (Rh) are
forming.

Figure 32. Cross section of Plasmodium mexicanum oocyst
with developing sporozoites. A cytostome (Cy),
anterior to the nucleus (N), is present in
sporozoites of Plasmodium mexicanum, which are
approximately 1/2 their mature length. The
oocyst capsule appears to be wrinkled in the later stages of sporozoite development. The sporoblastoid (Sb) often contains many dense
inclusions (Di).





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sporoblastoid, only spherical forms are observed migrating into the developing sporozoites. The cytostome, anterior to the nucleus, is observed in developing sporozoites which are not completely formed (Figure 32). Formation of the sporozoite is complete when it is pinched off from the sporoblast.

Sporozoites. Sporozoite formation is to some degree synchronous within a given oocyst but not between oocysts (Figure 19). Several sporoblasts with protruding sporozoites are observed in oocysts which are prematurely ruptured (Ayala, 1971). Living and Giemsa-stained sporozoites of P. mexicanum are stout, cresent shaped, and measure 6.3-7.3um (um = micron) (mean 6.6um) in length and 1.3-1.7um (mean 1.5um) in diameter at their greatest width (Figure 22). The mature sporozoites migrate through the hemocoel to the anterior of the sand fly until they reach the salivary glands. Sporozoites in the hemocoel have distinct micronemes, elongated rhoptries, and inclusion bodies. The salivary glands of L. vexator are hollow fluid filled sacs which are 1 cell thick and roughly spherical in shape (Figure 23). The anterior end of the sporozoite comes into contact with the salivary gland cell and causes the cell membrane to become invaginated (Figures 33 and 34). The sporozoite pushes through the salivary gland cell becoming constricted at the membrane-parasite junction, and carrying with it a portion of the salivary gland cell plasma membrane which forms a parasitophorous-like vacuole inside





















Figure 33. Plasmodium mexicanum sporozoites (S) in the
process of penetrating the salivary gland (Sg) of Lutzomyia vexator. The anterior end of the
sporozoite comes into contact and causes an
invagination (Iv) of the salivary gland
membrane. The salivary gland cell plasma
membrane (Pm) continues to invaginate until
the sporozoite is contained in a parasitophorus
vacuole (Pv) inside the salivary gland cell
(Sg). No distinct thickening along the
sporozoite-cell membrane junction was noted
during penetration. However, a notable
constriction (Cs) of the sporozoite at the
sporozoite-cell membrane junction was apparent
in most longitudinal sections of sporozoites.
Micronemes (Mn) and elongate rhoptries (Rh) are present in sporozoites which are migrating into
the salivary glands.

Figure 34. Another sequence of Plasmodium mexicanum
sporozoites in the process of penetrating the
salivary gland (sg) of Lutzomyia vexator.
Nucleus (N), micronemes (Mn), cytostome (Cy),
mitochondria (M) with associated dense body
(Db), parasitophorus vacuole (Pv) and Hemocoel
are labeled.







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Full Text
-24-
The development time of P_. mexi canum in the host, J_.
vexator, is relatively short at 27C. Oocysts were first
seen on day two after feeding, often occurring in large
numbers (Figure 2), and developing rapidly at 27C (Chapter
3). Sporoblasts with budding sporozoites are observed in
some oocysts by day five post-feed. Sporozoites are free in
the hemocoel by day six post-feed and are present in the
salivary glands by day 6.5 post-feed (Figures 3 and 4).
However, when sand flies were maintained at 24C,
sporozoites were not observed in the salivary glands until
days 8.5-9.0 after feeding. All sand flies used in the
transmission study were maintained at 27C and provided
second bloodmeals on noninfected Sc_. undul atus subsequent to
day 6.5 post-feed. Oocyst and sporozoite development is
reported elsewhere.
The laboratory transmission data of P_. mexi canum to
non-infected Sc_. undul atus by bite of infected vexator
female(s) are shown in Table 1. A total of 13 S£. undulatus
were fed upon by 1-3 L^. vexator females from days 7-10
following initial bloodmeals on S£. occidentalis infected
with P_. mexi canum All sand flies had sporozoites in the
salivary glands 0-8 hrs subsequent to their second
bloodmeal. Nine (69.2%) of the 13 lizards became infected
with P_. mexi canum. Two Sc. undul atus were each force fed
more than five sand flies (10 days post-feed) potentially
infected with P. mexicanum. Neither of these lizards became
infected.


TO JACQIJI, KEVIN, AARON, MICHELLE, AND ROBERT


58.Sporozoites of Plasmodium floridense which
ruptured from the salivary glands of Culex
erraticus 152
59.Midgut of Cu 1 ex erraticus with melanized
oocysts and sporozoites (?) of Plasmodium
f 1 ori dense 152
60.Cross section of an oocyst of PIasmodiurn
f1oridense on the midgut of Culex erraticus 156
61.Higher magnification of the oocyst capsule of
PI asmodi urn flor i dense 156
62.Cross section of an oocyst of PIasmodiurn
f1oridense on the midgut of Culex erraticus 156
63.Virus particles in the midgut epithelium of
Culex erraticus 156
64. Cross section of PIasmodiurn f1oridense oocyst
undergoing differentiation 159
65. Convex dense membranes, the precursors of the
developing sporozoites, form along the narrow
linear extensions 159
66. Oocyst of PIasmodiurn floridense with developing
sporozoi tes 162
67. Longitudinal section of the posterior portion
of a developing sporozoite 162
68. Higher magnification of developing sporozoites
of Plasmodium floridense 162
69.Higher magnification of developing sporozoites
of Plasmodium floridense 162
70. Oocyst of PIasmodi urn f 1 ori dense with developing
sporozoites 165
71. Salivary gland of Culex erraticus infected with
sporozoites of PI asmodi urn floridense 165
72.Cross section of PIasmodiurn f1oridense
sporozoites in the salivary glands of Culex
erraticus 165
73. Degenerating sporozoite of PIasmodiurn floridense
in an oocyst which is becoming melanized 165
74. Sporozoites of Schel1ackia golvani in white
blood cells of Anolis carolinensis 181
xiii


-187-
X 4.7um), 7.7-10.9 X 4.8-7.7um (mean 10.1 X 5.6um), and
9.0-10.8 X 3.8-5.lum (mean 9.6 X 4.4um), respectively. The
chromatin of sporozoites in RBC's of S. occidentalis often
consisted of dense droplets which occasionally concentrated
along the periphery of the sporozoite (Figures 86 and 87).
The chromatin of sporozoites in the WBC's was usually more
diffuse and similar to S>. go 1 vani (Figure 88). Sporozites
of S_. occi dental i s in circulating WBC's are usually elongate
to oval, while those observed in the RBC's are spherical,
bean shaped or teardrop shaped (Figures 86 to 90). One WBC
was seen with numerous sporozoites of S_. occidentalis
(Figure 90) .
Impressions and thin sections of the intestine revealed
very few stages in the development of !S. occidentalis in Sc
undulatus. A few schizonts (Figure 91) and oocysts (?)
(Figure 92) were seen. Tissue impressions and thin sections
of other tissues (brain, spleen, liver, and lung) were
negative for parasites.
Discussion
Transmission Studies
The specific status and mode of transmission of two
sympatric Schel1ackia parasites in A. carolinensis and Sc .
undulatus in Florida was previously uncertain. Jordan and


-202-
3. While Coi. erraticus was susceptible to infection of
£_ f 1 ori dense and sporozoites were frequently observed in
the salivary glands, attempts to transmit it by bite and IP
inoculation of sporozoites were only marginally successful.
4. Ultrastructure of the sporogonic stages of P_.
f1oridense indicates that it is very similar to other
mammalian and avian malarias previously studied but differs
from the other two saurian malarias that have been examined.
5. Attempts to transmit P_. mexi canum with l. vexato r
were highly successful; nearly 70% of all lizards which were
fed on by 1-3 infected sand flies became infected.
6. Post-patent exoerythrocyti c stages of P_. mexi canum
were abundant in the spleen, bone marrow, and endothelial
cells of capillaries in the brain of Seeloporus undulatus
that died of fulminating infections.
7. U11 ra s t r uc t u re of P_. mexi canum sporogonic stages
indicates that while development is very similar to other
Plasmodium spp., the gross morphology and microtubular
arrangement differs from mammalian and avian malarias.
Mature sporozoites are also extracellular in the lumen of
the salivary gland rather than intracellular.
8. Schel1ackia gol vani and Schellackia occidentalis
can be experimentally transmitted by a wide variety of
arthropods and time of development is noticably affected by
different temperatures. Transmission was only successful
when parasites from conspecific hosts were ingested.


LOG NUMBER OF PARASITES/10.000 RBC
-46-
DAY POSTEXPOSURE



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Figure 44.
Number of Cu 1 ex errat i cus collected per light
trap during the period 1 May to 9 October,
1984.


DAY POSTEXPOSURE
LOG NUMBER OF PARASITES/10.000 RBC
cn o
o o
o o
o
-££T-
5000


-153-
Table 9. Sporogonic development of PIasmodium floridense
on the midgut of Culex erraticus maintained at
25C.
Day
post -
feed
Number
mosqui
toes
Range
oocyst
size
(microns)
Mean
oocyst
size3
(mi crons)
Mean
oocys£
range0
(microns)
5
3
10.4 23.4
16.3
13.7 20.4
6
3
14.3 26.0
21.7
20.6 22.8
7
2
20.8 39.0
31.7
31.3 32.1
8
3
18.2 41.6
29.3
28.0 30.8
9
2
18.2 49.4
35.6
34.0 37.2
10
3
23.4 54.6
41.8
37.4 46.6
11
3
28.6 59.8
42.7
40.2 45.6
a Overall mean oocyst size by day post-feed for all
, mosquitoes.
D Average oocyst size range for different mosquitoes by
day post-feed.


REFERENCES
Aikawa, M. and H. B. Jordan. 1968. Fine structure of a
reptilian malarial parasite. J. Paras it o 1 .
54:1023-1033.
Aikawa, M., L. H. Miller, J. Johnson, and J. Rabbege.
1978. Erythrocyte entry by malarial parasites: A
moving junction between erythrocyte and parasite. J.
Cell Biol. 77:72-82.
Aragao, H. de B., and A. Neiva. 1909. A contribution to
the study of the intraglobular parasites of the
lizards. Two new species of PIasmodium, PI .
di pi ogl ossi n. sp. and PJ_. t r o p i d u r i n. sp. Mem. Inst.
Oswaldo Cruz 1:44-50.
Ayala, S. C. 1970a. Lizard malaria in California;
Description of a strain of PIasmodiurn mexicanum and
biogeography of lizard malaria in western North
America. J. Parsito!. 56:417-425.
Ayala, S. C. 1970b. PIasmodiurn mexicanum in California:
Natural history and development in phlebotomine
sandflies (Diptera:Psychodidae). J. Parasitol. 56:13.
Ayala, S. C. 1971. Sporogony and experimental transmission
of PIasmodiurn mexicanum. J. Parasitol. 57:598-602.
Ayala, S. C. 1977. Plasmodia of reptiles. In: Kreier, J.
P. ed., Parasitic Protozoa 3:267-309. New York:
Academic Press.
Ayala, S. C. 1978. Checklist, host index, and annotated
bibliography of PIasmodiurn from reptiles. J.
Protozool. 25:87-100.
Ayala, S. C., and D. Lee. 1970. Saurian malaria:
Development of sporozoites in two species of
phlebotomine sandflies. Science 167:891-892.
Baker, J. R. 1961. Attempts to find the vector of the
Plasmodiidae of the lizard Agama agama agama in
Liberia. Ann. Rept. Research Activities Liberian
Inst, of the American Foundation for Trop. Med. pp.
28-40.
-204-


-192-
Friend (1971) and Telford (1978) treated the Schel1ackia
parasites of Sc_. undul atus as occidentalis, while, Jordan
indicated that Schellackia parasites observed in A.
carolinensis were the same as those in Sc_. undulatus
(Personal communication). Telford (1978) suggested that the
parasites found in the anoles represented S^. gol vani T o
add to the confusion, S^. bol i vari Reichenow (1919) from
Spain invades the erythrocytes of one lizard,
Ac anthodacty1 us vulgaris but infects lymphocytes of
another, Psammodromus hispanicus, also from Spain (Manwell,
1977 ) .
There are morphological differences in Schel1ackia
sporozoites invading RBC's in S£. undulatus and those
invading lymphocytes of the anoles. However, Schellackia
parasites occuring in the lymphocytes of S£. undulatus are
similar to those observed in the anoles (Figures 74, 75 and
89). The effect of host variation on the type of cell
infected and progression of the disease is also evident for
other hemosporidia Schizogony of PIasmodiurn mexicanum is
primarily erythrocytic following the prepatent period in Sc.
occidentalis and _Sc. undul atus, but is primarily
exoerythrocytic in Ph rynosoma cornutum and Crotaphytus
co 11 aris (Thompson and Huff, 1944b). Though gametocytes are
produced in both Seeloporus lizards, none are produced in C.
col 1 ari s and only a few in P_. cornutum. It was therefore
suspected that the morphological differences and host cell
types which are invaded are due to differences in the host
species .


-126-
\ *TV
20um
V1 Kfc? ?*> -*
\i*"NVv S&.vv AJ
y&x
Sic:- ..
10um
50um
47
49


-210-
Pelaez, D., R. Perez-Reyes, and A. Barrera. 1948. Estudios
sobre hematozoarios I. PIasmodium mexicanum Thompson
and Huff 1944 en sus huespedes naturales. Anal. Esc.
Nac. Ciee. Bi o 1 5:197-215.
Pessoa, S. B ., P. de Bias i, and L. Sacchetta. 1974.
[Evolucao do Hepatozoon sp parasita do Leptophis
a h a e t u1 la (Lineau) (Serpentes Colubridae) no Culex
fatigan~s~J. Mem. Inst. Butantan (Sao Paulo)
38:119-122. Taken from Bio. Abstr. 60:6687 (abstract
no. 62491).
Petit, G., I. Landau, Y. Boulard, A. Gomes, and L.
Touratier. 1983. Sporogonie de PIasmodium agamae chez
C u1 ic o id e s nubeculosus au laboratoire: I -
Experimentation et description du cycle.
Protistologica 19:537-541.
Pienaar, U. 1962. Haematology of some South African
reptiles. Johannesburg: Witwatersrand University
Press.
Porter, R. J., R. L. Laird, and E. M. Dusseau. 1952.
Studies on malarial sporozoites. I. Effect of various
environmental conditions. Exp. Parasitol. 1:229-244.
Rafaelle, G. 1934. Un ceppo italiano du PIasmodium
elongatum. Riv. di Malarial. 13:3-8.
Reichenow, E. 1919. Der entwicklungsgang des Hamococcidian
Karyolysus und Sc hel1acki a nov. gen. Sitzungsber. Ges.
Naturforsch. Freunde, Berlin. 440-447.
Rogier, E., and I. Landau. 1975. Description de
Schel1ackia golvani n. sp. (Lankesterel1idae), parasite
de lezards de guadeloupe. Bull. Mus. Nat. Hist. Nat.
Series 3, Zoologie. 194:91-97.
Ross, R. 1897. On some peculiar pigmented cells found in
two mosquitoes fed on malarial blood. Br. Med. J.
2:1786-1788.
Ross, R. 1898. Report on the cultivation of Proteosoma,
Labbe, in grey mosquitoes. Indian Med. Gaz.
33:401-448.
Russell, P. F., and B. N. Mohan. 1942. The immunization of
fowls against mosquito-borne PIasmodium gal 1inaceum by
injections of serum and of inactivated homologous
sporozoites. J. Exp. Med. 76:477-495.
Russell, P. F., L. S. West, R. Manwell, and G. MacDonald.
1963. Practical Malariology. 2nd ed. London: Oxford
Univ. Press.


Figure 25. Cross section of PIasmodium mexicanum oocyst
undergoing differentiation. Early
vacuolization (V) with convex dense membranes
(Dm). Nuclei (N) with an electron dense
nucleolus (Nu), endoplasmic reticulum (Er), and
numerous mitochondria (M) which appear in groups
are scattered throuout the cytoplasm.
Figure 26. Cross section of PIasmodium mexicanum oocyst
undergoing differentiation. The sporobl astoi d
(Sbs) is completely separated from the oocyst
capsule (C). Convex dense membranes (Dm) are
present along the margin of the sporoblastoid
body. Inclusion bodies (I) are seen in groups
or scattered througout the oocyst.
Figure 27. Cross section of PIasmodium mexicanum oocyst
undergoing internal vacuolization of the
sporoblasoid (Sbs). Lamellar inclusion bodies
(Li),very dense granules associated with a
space of low density (Di ) and golgi apparatus
(Go) are present. Sporozoite buds (B) from the
convex dense membranes form along the periphery
of the sporoblastoid and along the internal
membranes calised by vacuolization of the
sporoblastoid.
Figure 28. Coelescence of the internal vacuolization of
P1asmodium mexicanum oocysts produces
cytoplasmic clefts [Cc) which are the
precursors for the sporoblasts. A moderately
dense inclusion body is often associated with
the convex dense membranes (Dm) during this
stage.
Figure 29. Higher magnification of PIasmodium mexicanum
oocyst. The sporoblast has pulled away from
the oocyst capsule, showing a convex dense
membrane (Dm) beneath the plasma membrane.
Moderately dense vacuoles (Mi), which are
scattered throughout the oocyst, migrate and
come to lie beneath these convex dense
membranes. Microtubules (arrows) associated
with the convex dense membranes are seen in
cross section.


-15-
interest to anyone who has speculated about the evolution of
the malaria parasites. Their life history and
ultrastructural organization may provide an insight as to
how the coccidian ancestors may have adapted to an
alternating existence in a vertebrate and arthropod host.


-208-
Hunninen, A. V. 1953. Comparative development of
P1 asmodiurn re!ictum oocysts in Anopheles
quadrimaculatus A. albimanus, and Cu 1 ex pipiens. J.
Parasitol. 39:28-32.
Huff, C. G. 1941a. Saurian malaria. J. Parasitol.
(Suppl .) 27:29 (Abstr .).
Huff, C. G. 1941b. Comparative importance of various
factors upon the regulation of size of avian malarial
oocysts in mosquitoes. Am. J. Hyg. 34:18-21.
Huff, C. G. 1945. A consideration of the problem of the
evolution of malarial parasites. Rev. Inst. Salubr.
Ent. Trop. 6:253-258.
Huff, C. G. 1969. Parasitological reviews.
Exoerythrocytic stages of avian and reptilian malarial
parasites. Exp. Parasitol. 24:383-421.
Jordan, H. B. 1964. Lizard malaria in Georgia. J.
Protozool. 11:562-566.
Jordan, H. B. 1970. The occurrence and development of
PIasmodiurn mexicanum in the western fence lizard,
Seeloporus occidentalis. J. Protozool. 17:86-89.
Jordan, H. B., and M. B. Friend. 1971. The occurrence of
Schellackia and PIasmodiurn in two Georgia lizards. J.
Parasitol. 64:1126-1127.
Karnovsky, M. J. 1965. A formaldehyde-glutaraldehyde
fixative of high osmolarity for use in electron
microscopy. J. Cell Biol. 27:1 37A.
Klei, T. R. 1972. The fine structure of Haemoproteus
columbae sporozoites. J. Protozool. 19:281-286 .
Klein, T. A., B. A. Harrison, R. G. Andre, R. E. Whitmire,
and I. Inlao. 1982. Detrimental effects of PIasmodiurn
cynomol gi infections on the longevity of Anophel es
dirus. Mosq. News 42: 265-271.
Lainson, R., and J. J. Shaw. 1969. New host records for
PI asmodi urn diploglossi £. t r o p i d u r i Aragao and Neiva,
1909, and P. cnemidophori Carini, 1941. Parasitology
59:163-170.
Lainson, R., J. J. Shaw, and R. D. Ward. 1976. Schellackia
1andauae sp. nov. (Eimeriorina:Lankesterl1idae) in the
Brazilian lizard Po 1 ychrus marmoratus (Iguanidae):
Experimental transmission by Culex pipiens fatigans.
Parasitology 72:225-243.


-64-
and the sporozoite rate determined (+1, 1-10; + 2, 11-100;
and +3 >100 sporozoites).
Ultrastructure of Extrinsic Stages of Plasmodium mexicanum
For ultrastructural examination, sand flies were fixed
in 2% glutara1dehyde in 0.1M sodium cacodylate buffer at pH
7.2 for 1 hr at room temperature. Post-fixation was with 1%
osmium tetraoxide in 0.1M sodium cacodylate buffer, pH 7.2,
for 1 hr at room temperature. Specimens were dehydrated
through a graded ethanol series and acetone and embedded in
Spurr's resin (Spurr, 1969). Sections were cut on an LKB
ultratone III with a Diatome diamond knife, floated on
water, picked up on formvar-coated grids, and post-stained
with aqueous 1% uranyl acetate followed by Reynold's lead
citrate. To enhance the polar rings, sections with
sporozoites were post-stained with aqueous 1% uranyl acetate
for 1.5 hr at 60C. Material was examined and micrographs
taken on a Hitachi 11-E or Jeol 100CX electron microscope.
Results
Sporogony of Plasmodium mexicanum
Oocysts of P_. mexicanum developing on the midgut of l.
vexator were observed from days 2-9 post-feed. The effects
of two different temperatures on the sporogony of P_.


AVERAGE OOCYST SIZE (MICRONS)
-68-


Figure
Figure
Figu re
30. Oocyst of PI asmodium mexicanum with developing
sporozoites. Sporozoites of P_. mexi canum are
loosely packed within the oocyst capsule (C).
Sporozoites bud off the sporoblast leaving a
residual body (Rb). The nucleus and
moderately dense inclusion body (Mi) are
present in the anterior of the sporozoite.
Short bag-like rhoptries (Rh) are seen in a few
sporozoites.
31. Higher magnification of developing sporozoites
of PIasmodium mexicanum, illustrating the
migration of the nucleus (N) and spherical
mitochondrion (M) into the sporozoite. The
moderately dense inclusion bodies (Mi) appear to
be degenerating as the rhoptries (Rh) are
forming.
32. Cross section of PIasmodium mexicanum oocyst
with developing sporozoites. A cytostome (Cy),
anterior to the nucleus (N), is present in
sporozoites of PIasmodium mexicanum, which are
approximately 1/2 their mature length. The
oocyst capsule appears to be wrinkled in the
later stages of sporozoite development. The
sporoblastoid (Sb) often contains many dense
inclusions (Di).


-o-


-181-


-8-
Further advances in the host-parasite relationship were
made by Ayala (1971) who infected lab reared Sc.
occi dental i s with P_. mexi canum sporozoites. Blood parasites
were not observed in the blood until 22 days post
inoculation. This corresponds with the results of a 14 to
27 day prepatent period of P_. flor i dense in S£. undul atus
(Goodwin, 1951; Goodwin and Stapleton, 1952). However, as
previously indicated, the prepatent period may be influenced
by ambient temperature (Thompson and Winder, 1947). Ayala
(1970a,b) further noted that there is a spring relapse of
gametocytes of P_. mexi canum lasting from March to August.
This corresponds to the period of activity of its suspected
vector, j_. vexato r, indicating that most malaria
transmission in California is limited to the spring and
summer months .
Exoerythrocytic (EE) stages have been observed in a
number of different saurian malaria species, usually in
blood films (Garnham, 1950; Garnham and Duke, 1953; Bray,
1957, 1959; Lainson and Shaw, 1969; Telford 1970a; and
Scorza 1971b). Huff (1969) provides an excellent review of
the EE stages of avian and saurian malaria parasites and
points out the need for additional research in this area,
especially the determination of the vectors of saurian
malaria and subsequent observation of the life cycle of the
parasite in the saurian host.
The first and most detailed description of the EE
stages of saurian malaria parasites (P_. mexi canum) was made


-33-
v


-87-
approximately 1/2 of the circumference of the sporozoite
while the remaining five are arranged equidistantly and
farther apart around the circumference of the other half
(Figure 40). The sporozoite pellicle is thickened and more
electron dense near the apical region where the
subpel1icular microtubles originate. Polar rings, present
in sporozoites of PIasmodiurn sp., were observed only after
staining in hot (60C) 1% aqueous uranyl acetate solution.
Moderately dense inclusion bodies, which appear to
degenerate in developing sporozoites in the oocyst, are
rarely observed in sporozoites in the hemocoel or salivary
glands. The rhoptries in developing sporozoites appear
short and sac-like (Figure 32), but become elongate, wider
in diameter basally, and extending nearly to the nucleus in
mature sporozoites (Figure 37). The usual number of
rhoptries is two. However, three rhoptries are seen in a
few. Numerous dense spherical to elongate micronemes are
located in the anterior 1/3 of the sporozoite (Figures 33,
34, 37 and 38). Micronemes were not observed in the
posterior of the sporozoite nor in sporozoites which were in
the oocyst.
The nucleus is subcentrally located, tubular, and only
slightly more electron dense than the cytoplasm (Figure 37).
Nucleoli become indistinct prior to migrating into the
developing sporozoite and are not observed in the nucleus of
mature sporozoites. The single spherical mitochondrion
becomes irregularly shaped, elongate, tubular, and appears


-93-
expossed to high temperatures for only a few hours following
the bloodmeal. Huff (1941b), further demonstrated that for
P. cathemeriurn Hartman, the two primary factors which affect
oocyst size are age of the oocyst and environmental
temperatures and that humidity, activity of the vector, and
oocyst rate had no significant effect on oocyst size.
Hunninan ( 1953) also indicated that for £. relictum, there
was no correlation between infectivity and oocyst size, but
he did observe differences in sporogonic development between
different vectors .
As previously indicated, sporogonic development of P_.
mexi canum in l^. vexator maintained at 27C is very rapid,
with sporozoites present in the hemocoel and salivary glands
by days 6.0 and 6.5 post-feed, respectivly. If the
longevity of laboratory reared sand flies is any indication
of longevity of sand flies in the wild, rapid development of
the parasite would be essential since the mortality rate is
>60% by day 8 post-feed and nearly 100% by day 15 post-feed
when maintained at 27C. The mortality rate of sand flies
maintained at 24C was lower, but the sporogonic development
rate was also slower, requiring 2.0 2.5 additional days
until sporozoites were observed in the salivary glands. A
few sand flies were reared at 19C and dissected on days
four and five post-feed. The bloodmeal of these flies was
largely undigested and no oocysts were observed in them.
Oocysts were not observed in flies which were returned to an
environmental chamber maintained at 27C and dissected


CHAPTER 7
SUMMARY AND RECOMMENDATIONS FOR FUTURE INVESTIGATIONS
During this study, the following objectives were
achieved or determined.
1. Field studies were conducted in an area near
Newnans Lake, Alachua Co., Florida, where Plasmodium
f1oridense was found in Anolis carolinensis. Lizard-bait
traps and CDC light traps established that two species of
mosquitoes, Cu 1 ex erraticus and Cu 1 ex territans were
abundant, attracted to, and bloodfed on lizards in the
field. Peak seasonal abundance of (lx. errati cus
corresponded to the time of active transmission of P_.
f1oridense to juvenile anoles.
2. A laboratory colony of Culex erraticus was
established in 1984 for transmission studies of P_.
f1oridense. A laboratory colony of Lutzomyia vexator was
also maintained for transmission studies. Both Cx.
erraticus and L. vexator developed oocysts following
bloodmeals on anoles with moderate to high gametocyte
numbers. Low levels of gametocytes (<1%) gave varying
responses. Sporogony of P_. flor i dense in C_x. errati cus
resulted in mature sporozoites in the salivary glands.
Sporozoites were infrequently observed in the hemocoel of L_.
vexator; they were never observed in the salivary glands.
-201-


86


LOG NUMBER OF PARASITES/10,000 RBC
10000
5000
1000
500
100
50
10
5
10
20
30
40
50
X
* = TROPHOZOITES
= SCHIZONTS
*= GAMETOCYTES
60 70 80 90
i
100 110 120
DAY POSTEXPOSURE


-112-
Collection and Laboratory Maintenance of Lizards
Ano 1is carolinensis and S^. undulatus were hand
collected from the Hatchet Creek, Austin Cary Forest, and
other localities near Gainesville (Alachua county), and from
Gulf Hammock (Levy county), Florida, USA (Table 4). In
addition, A. carolinensis were collected from Manchac swamp,
Tangipaho Parish, Louisiana, where P^. flor i dense infections
have not been observed. Bloodfilms were made from a clipped
toe, air dried, fixed with absolute methyl alcohol, stained
with Giemsa, and examined for the presence of blood
parasites. Subsequent bloodfilms were made by either
clipping the toe or tail. Anolis carolinensis were
maintained in the laboratory as previouly described (Chapter
2) or in a temperature controlled environmental chamber at
32C.
Plasmodium floridense Transmission Studies
Laboratory reared Dc. errati cus females were placed in
a plastic cylinder (4.5 X 15 cm) with a screened end and
containing a lizard (A. carolinensis or S£. undulatus)
infected with P_. floridense. The lizard was restrained on a
tongue depressor with two thin pieces of tape, one over the
shoulder, and the other over the pelvic girdle to prevent
movement. Bloodfed mosquitoes were removed the following
morning, placed in 100 ml plastic urine specimen containers
with a small amount of water, provided a 10% sugar solution,


£8


-19-
clipping the the tip of the tail. Seeloporus undu1atus that
did not show patent PIasmodiurn floridense infections within
a 30 day period (from a minimum of three bloodfilms) were
used in mexicanum transmission studies. Lizards were
maintained in screened cages (50 x 25 x 25 cm) in the
laboratory at room temperature and provided an external heat
source from a 40 watt incandescent light bulb. Lizards were
fed house flies (Musca domestica L.) and lepidoptera larvae
(Galleria sp. and Spodoptera sp.). Water was provided by
spraying the cages daily and by placing a water-filled petri
dish in each cage.
Plasmodium mexicanum Transmission Studies
Lab reared L. vexator females were bloodfed on Sc.
occidentalis which demonstrated >1% of the red blood cells
infected with P_. mexi canum gametocytes. Bloodfed females
were removed at 4 hr intervals, placed in 25 ml oviposition
vials, provided a sugar source (1:1 mixture of Karo syrup
and distilled water) and maintained in a temperature-
humidity controlled chamber as previously described.
Midguts were dissected (Chaniotis and Anderson, 1968) at
intervals from 2-7 days post-feed (period following initial
bloodmeal on an infected lizard) and the number of oocysts
counted. In addition, the salivary glands were examined
subsequent to day five post-feed, and the sporozoite rate
determined (+1, 1-10; +2, 11-100; +3, >100 sporozoites).


-12-
culture techniques and relative ease with which reptilian
cell lines can be established, the use of reptilian
parasites for in vitro culture studies has increased
(Wernsdorfor, 1980) .
In summary, literature on various aspects of saurian
malaria is limited. Morphology, electron microscopy, EE
stages, chemotherapeutic drug studies, and parasite
development in the vertebrate suggest a close phylogenetic
relationship between the avian and saurian malarias. Aside
from the possibility that the sand fly, L_. vexator, may
transmit P_. mexi canum, no other arthropods have been
incriminated as vectors of saurian malaria. Transmission by
bite has not been demonstrated and no naturally infected
arthropods have been discovered.
Schel1ackia
Schel1ackia parasites are transmitted mechanically by
ingestion of invertebrate hosts (mites, Diptera, or leeches)
which previously fed on infected cold blooded vertebrates
(reptiles or amphibians). The schizogonic stage occurs in
the epithelial cells of the intestine while sporogony occurs
usually in the lamina propria of the intestine. In some
species, such as S^. ba 11 i Lebail and Landau ( 1974), oocysts
also form in the epithelial cells of the intestine (Lainson,
et al., 1976). Levine (1980), however, extended the
definition of Sc he 11ac kia to include parasites with merogony


-157-
separated by a narrow electron dense layer (Figure 61). The
interior margin of the oocyst capsule is generally smooth
with portions of the capsule sometimes protruding into the
sporoblastoid (Figure 61).
The internal structure of the undifferentiated oocyst
of P_. f 1 ori dense is similar to that of P_. mexi canum and
other malarias which have been examined (Chapter 3; Duncan
et al., 1960; Vanderberg et al., 1967; Terzarkis et al.,
1967). The nucleoplasm of the large nuclei is only slightly
denser than the cytoplasm. A large electron dense nucleolus
is observed in some nuclei (Figures 60 and 62).
Mitochondria are tubular in shape, have tubular cristae, and
are scattered throughout the sporoblastoid. Mitochondria
may be in scattered clumps, as in other malarias, since they
are absent in some of the sections examined while numerous
in others. Amorphous electron opaque spherical shaped
"lipid-like" droplets, ribosomes, endoplasmic reticulum,
golgi bodies, and three forms of inclusion bodies (similar
to those described by Terzarkis et al. ( 1967 ) for P_.
gal 1inaceum) are scattered throughout the sporoblastoid
(Figures 62 and 64). Virus particles were seen in midgut
epithelium, adjacent to developing oocysts in some Cx.
erraticus examined (Figure 63).
Sporozoite development. Differentiation of the oocyst
appears to begin with internal vacuolization of the
sporoblastoid (Figure 64). Linear extensions of the
internal vacuoles extend to the surface of the sporoblastoid


Figure 60. Cross section of an oocyst of PIasmodium
floridense on the midgut of Culex errati cus 8
days after feeding on an infected lizard. The
solid, non-vaculated oocyst protrudes slightly
into the hemocoel and is bounded by a thick
capsule (C). Large nuclei (N) with a distinct
nucleolus (N u ) and mitochondria (M) are
scattered throughout the cytoplasm. Lamellar
inclusion bodies (Li) are occasionally seen.
Figure 61. Higher magnification of the oocyst capsule (C)
of PIasmodium f1oridense The oocyst capsule
consists of three layers, two amorphous layers
of similar electron density separated by a thin
electron dense layer (arrow). Dense inclusion
bodies (Di), mitochondria (M), and endoplasmic
reticullum (Er) are scattered throughout the
spo roblastoid.
Figure 62. Cross section of an oocyst of PIasmodium
f1oridense on the midgut of Culex erraticus 12
days after feeding. Internal vacuolization of
the sporoblastoid is just beginning. Large
nuclei (N) with a distinct nucleolus (Nu),
golgi bodies (Go), "lipid-like" globules (Lp),
and inclusion bodies are scattered throughout
the cytoplasm.
Figure 63. Virus particles (Vi) in the midgut (Mg)
epithelium of Cu 1 ex erratius An oocyst capsul
(C) containing developing sporozoites (S) is
adjacent to the infected epithelial cell.


-176-
Tissue impressions and thin sections of liver, spleen,
lung, kidney, bone marrow, intestine, and brain were made
from _Sc. undul atus maintained at 32C on days 2, 4, 6, and
8, following the ingestion of infected Aedes aegypti
mosquitoes and also when sporozoites were in the blood.
Tissue impressions and thin sections of the same tissues
were made from A. carolinensis maintained at room
temperature (18-24C) subsequent to the appearance of
sporozoites in the blood. Tissue impressions were stained
with Giemsa. Histological sections were made from tissues
which were fixed in Carnoy's fluid or Bouin's fixitive,
dehydrated, embedded in paraffin, sectioned at 5-6um on a
rotary microtome, and stained with hemotoxylin-eosin or
Giemsa-collophonium.
Results
Schellackia golvani
Transmission studies. A summary of the experimental
transmissions of S^. golvani by ingestion of l. vexator and
Cx. erraticus that had previously bloodfed on infected
lizards is shown in Table 10. Noninfected /\. carolinensis
demonstrated _S. golvani sporozoites in their bloodfilms from
days 10-81 after being fed mosquitoes and sand flies which
had previously fed on infected green anoles. Several Cx.
erraticus were maintained in a temperature controlled


-175-
methanol, stained with Giemsa, and examined for sporozoites
of S^. occi dental i s .
Histological Studies
Cross sections of mosquitoes and sand flies which
bloodfed on A. carolinensis with >70% polymorphonuclear
leucocytes infected with S^. gol vani sporozoites were made to
determine if the sporozoites remained in the lumen of the
midgut or invaded other tissues. Mosquitoes were fixed in
Carnoy's fluid or Bouin's fixitive, dehydrated, embedded in
parafin and sectioned at 6um (urn = micron) on a rotary
microtone. Thin sections were stained with
hemotaxy1in-eosin or Giemsa-col1ophoniurn (Bray and Garnham,
1962). Sand flies were fixed in 2% glutaraldehyde in 0.2M
sodium cacodylate buffer at pH 7.2 for 1 hr at room
temperature. Sections were post-fixed with 1% osmium
tetraoxide in 0.2M sodium cacodylate buffer, pH 7.2, for 1
hr at room temperature. Specimens were then dehydrated
through a graded ethanol series and acetone and embedded in
Spurr's resin (Spurr, 1969). Thick sections (2-3um) were
cut on a LKB ultratome III with a glass knife, floated on
water, transferred to a drop of water on a glass slide, and
heated at 100C on a hot plate for 1 hr. Sections were
stained with modified solutions of fuschin and methylene
blue as described by Di Sant-Agnese et a 1 (1984). Sections
were coverslipped with Klermount and examined for
sporozoites .


NUMBER MOSQUITOES COLLECTED PER LIGHT TRAP
200
180
MAY JUNE JULY AUG SEPT OCT
DAY/MONTH (1984)
-120-


ACKNOWLEDGEMENTS
This dissertation is the result of the cooperation,
assistance, and collaboration of many people. While I
accept the responsibli 1ity for the contents of this
dissertation, I do not claim all the credit.
I am grateful to the members of my committee for their
contributions and support throughout the study. They
provided instructional guidance, enthusiasm, and continuous
encouragement. In addition, each member gave freely of his
time and offered valuable suggestions in his areas of
expertise. They are Dr. David Young, chairman; Dr. Martin
Young, cochairman; Dr. Jerry Butler, committee member; and
Dr. Stephen Zam, committee member. Dr. Ellis Greiner, whil
not serving as a committee member, gave freely of his time
and expertise in parasitiolgy and provided material support
I am also indebted to Dr. Sam Telford for the many
discussions on lizard parasites and examination of
histological preparations. Dr. Donald Forrester also
provided assistance and laboratory space for the histology
studies. I also appreciate the assistance and space for
rearing mosquitoes that Dr. Larry Lacy and Dr. A1 Undeen
provided. Special gratitude is due to Ms. Debra Akin who
provided her time, instructional guidance, expertise, and
suggestions for the transmission electron studies and Dr.
Robert Kimsey who kindly provided me with some infected
iii


-166-
70), but were clumped together in sporozoites in the gland
(Figure 72). This, however, may be an artifact of the
fixation process. The nucleus is subcentrally located and
very elongate in micrographs (Figure 68 and 73). However,
when sporozoites are fixed with menthanol and stained with
Giemsa, the nucleus appears condensed and generally
spherical in shape (Figure 57 and 58). There are either
several mitochondria in the posterior 1/3 of the sporozoite
or there is one mitochondrion which is irregular in shape
and when sectioned longitudinally, appears to be more than
one (Figures 68 and 69). All three types of inclusion
bodies seen in the undifferentiated oocyst were also seen in
sporozoites. However, the most frequently observed
inclusion body is the very electron dense granule associated
with an area of less density. Crystalloid inclusions, as
seen in some hemoproteids and some Plasmodium sp. were not
observed.
Discussion
Sporogony of Plasmodium floridense
The sporogonic development of P_. f 1 ori dense is very
similar to that of other malarias. But, while most P_.
floridense oocysts appear to develop "normally", i. e.,
protruded into the hemocoel some developed between
epithelial cells or protruded into the lumen of the midgut.


-£Z~


-80-




Figure 52. Course of acute infection of PIasmodium floridense in two wild
caught Anolis carolinensis infected by bite C-85) of Cu 1 ex
erraticus and intraperitoneal inoculation (AA 5 9) of
sporozoites from the salivary glands of Culex erraticus.


wnoi
-681-


-143-
parti al bloodmeals. Mosquitoes were not sectioned or
dissected to determine if the integrity of the midgut was
damaged by invading parasites. But since few mosquitoes
died when fed on lizards with moderate numbers of
gametocytes, it is suspected that parasite invasion of the
midgut resulted in the death of the mosquitoes. Therefore,
the upper limit of infectivity may be approximately 100
oocysts. Klein et al (1982) demonstrated that large
numbers of oocysts (>100) had a significant effect on the
longevity of Anopheles di rus Peyton and Harrison infected
with £. cynomolgi after 11 days following the release of
sporozoites. However, there was no significant difference
in the survival of highly infected and noninfected
mosquitoes at 0-3 days after a bloodmeal.
The low number of infected anoles by bite of Cx.
er rati cus infected with P.. f 1 ori dense can not be explained
since the transmission rate of another saurian malaria, P_.
mexicanum by l. vexator to S£. undulatus, was nearly 70%.
However, nutritional requirements for proper maturation of
the parasites are unknown. It is suspected that the
addition of multivitamins to the sugar solution fed to
mosquitoes during sporogonic development shortened the
period when sporozoites were first observed in the salivary
glands by 2-3 days following an infective bloodmeal. The
effects of parasite growth during the period of maturation
by multiple bloodmeals is unknown. But since the
development of P_. flor i dense is relatively long (11-14


Abstract of Dissertation Presented to the Graduate
School of the University of Florida in Partial
Fulfillment of the Requirements for the Degree of
Doctor of Philosophy
DEVELOPMENT AND TRANSMISSION OF SAURIAN PLASMODIUM AND
SCHELLACKIA IN BLOODFEEDING ARTHROPODS
By
Terry A. Klein
August 1985
Chairman: Dr. David G. Young
Co-chairman: Dr. Martin D. Young
Major Department: Entomology and Nematology
A study was undertaken to (l)determine the vectors of
two species of PIasmodiurn and two species of Schel1ackia
parasites of lizards; (2) study the course of development of
the parasite in the lizard host; and (3) study the growth
and development of the extrinsic stages, including light and
transmission electron microscopy (TEM). Lutzomyia vexator
v
was shown to be an efficient vector of P. mexicanum,

infecting 69.2% (9/13) of the Seeloporus undulatus lizards
when fed on by infected flies. Cu 1 ex erraticus was
incriminated as a suspected vector of saurian malaria, P_.
f1oridense Vector attraction and bloodfeeding propensity
in the field was determined by using lizard baited traps.
Both £x. er rat i cus and Cx. territans were frequently
xvi


TABLE 10. Experimental transmission data of Schel1acki a gol vani and
Schel1ackia occidentalis to wild caught Ano 1is caroTTnensis and
$celoporus undulatus by ingestion of infected Lutzomyia vexator ,
Cu 1 ex erraticus Aedes aegypti and Geckobiella texana .
Lizard cFT No. Arthropod Day ^ Day
species lizards positive vector Post- Temp. patent
feed3 C infect.
Schel1ackia go 1vani
A.
carol inensis
7
5
Cx. erraticus
2-29
18-24
21-25,
carolinensis
3
3
L. vexator
5-6
18-24
21-23
A.
carolinensis
5
3
Cx. erraticus
4-6
32
10-12
S.
undulatus
2
0
Cx. erraticus
1-5
18-24
S.
undu1atu s
2
0
L. vexator
5
18-24
S.
undulatus
5
0
Cx. erraticus
5-6
32
Schellackia occidentalis
S.
undulatus
2
2p
Cx. erraticus
1
18-24
37
S.
undulatus
10
7 e
Cx. erraticus
1-7
32
12
s.
undulatus
8
8
L. vexator
1-7
32
8-17
s.
undulatus
3
2
G. texana

32
10
s.
undulatus
3
3
Ae. aegypti
6 hrs
32
7-9
A.
carolinensis
7
0
Cx. erraticus
1
32
-177-


LIST OF FIGURES
Figure Page
1. Bloodied female Lutzomyia vexator resting on
Seeloporus undulatus upon which it had
previously fed on 23
2. Midgut of Lutzomyia vexator with oocysts of
Plasmodium mexicanum 23
3. PIasmodiurn mexicanum sporozoites from ruptured
oocysts 23
4. PIasmodiurn mexicanum sporozoites in the
salivary gland of Lutzomyi a vexator 23
5.Prepatent and patent period of PIasmodiurn
mexicanum infection and survival of individual
Seeloporus undulatus infected with PIasmodiurn
mexi canum sporozoites 28
6.Percent of infected red blood cells during the
course of PIasmodiurn mexicanum infection for
six Seeloporus undulatus infected by bite of
Lutzomyi a vexator 31
7.Course of acute infection of PIasmodiurn
mexicanum in nine Seeloporus undulatus infected
by bite of Lutzomyi a vexator 33
8. Course of PIasmodiurn mexicanum infection in
Sceloporus undulatus infected with sporozoites
(S-51) ... 38
9. Course of PIasmodiurn mexican urn infection in
Sceloporus undulatus infected with sporozoites
( S 4 2) 7777 40
10. Course of Plasmodium mexicanum infection in
Sceloporus undulatus infected with sporozoites
(S 8) 42
11. Course of PIasmodiurn mexicanum infection in
Sceloporus undulatus infected with sporozoites
( S-15 ) . 44
IX


75.Sporozoite of Schel1ackia gol vani with two
chromatin bands in a white blood cell of Ano 1is
carol i nensi s 181
76.Sporozoite of Schel1ackia gol vani teased from
the midgut of Cu 1 ex erraticus 181
77.Unstained sporozoite of Schel1acki a g o 1v a ni
teased from the midgut of Culex erraticus 181
78.Cross section of the midgut of Culex erraticus
with a sporozoite of Schel1acki a golvani 181
79.Cross section of the midgut of Lutzomyia vexator
with a sporozoite of Schel1ackia go!vani 181
80.Unstained sporozoites of Schel1acki a golvani in
a parasitophorus vacuole in the midgut
epithelium of Culex erraticus 181
81.Impression of small intestine of Ano!is
carolinensis with macromerozoites of Schellackia
gol vani 184
82.Section of intestine of Anolis caorlinensis with
macroschizont and macromerozoites of Schel1ackia
g o 1 v a n i 184
83.Section of intestine of Ano 1is carolinensis with
microschizont containing developing
mi cromerozoi tes of Schel 1 acki a go! vani 184
84.Section of intestine of Ano 1is carolinensis with
macro- and mi c rogametocy tes of Schellackia-
g o 1 v a n i 184
85. Sporozoites of Schellackia occidental is teased
from the gut of a mite 184
86. Crescent-shaped sporozoite of Schel1ackia
occidentals in the red blood cell of Seel oporus
undul atus 189
87.Comma-shaped sporozoite of Schellackia
occidentalis in the red blood cell of Seeloporus
undu 1 atus 189
88.Sperical sporozoite of Schellackia occidentalis
in the red blood cell of Sceloporus undulatus... 189
89.Sporozoite of Schel1ackia occidentalis in a
white blood cell of Sceloporus undulatus 189
xiv


-20-
One to six female l. vex ato r infected with _P. mexicanum
sporozoites were placed in a Plexiglas cage lined with
plaster of Paris (Endris et al., 1982) and provided a second
bloodmeal on a noninfected, wild-caught Sc_. undul atus.
Lizards fed on by one or more infected sand flies were
placed in a screened cage and maintained as previously
described or were placed in a temperature-humidity
controlled chamber and maintained at 27C and 80% RH.
Bloodfed females were dissected after the second bloodmeal
and the sporozoite rate determined. To determine if
transmission of JP. mexi canum could also occur by the oral
route, living J_. vexator potentially infected with P_.
mexicanum sporozoites were force fed (placed in the back of
the mouth with forceps) to S_c. undul atus.
Course of Infection, Parasitemia
Bloodfilms of S£. undulatus previously fed on by
infected l. vexator or force fed infected sand flies were
made at day 0 post-exposure (period of time from which
non-infected lizards were exposed to bites of l. vexator
with sporozoites of P_. mexicanum) and at 2-4 day intervals
subsequent to day 19 post-exposure. Parasites were counted
and parasitemias expressed as the number of parasites per
10,000 red blood cells (RBC). PIasmodiurn mexicanum
characteristically occupies all circulating blood cells
(Jordan, 1970); therefore, the number of infected white


-182-
size for the sporozoites measured from different A.
carolinensis ranged from 8.8-10.0 X 4.6-5.4um (mean 9.5 X
5.2um). The chromatin of _S. gol van i intracellular (WBC)
parasites is usually diffuse, forming one or more distinct
bands, while the chromatin of extracellular parasites in the
blood is usually condensed into a distinct spherical nucleus
(Figures 76 and 77).
Sporozoites of S. go!vani are ingested during the
bloodmeal by hematophagous arthropods. Subsequent to the
bloodmeal, the intracellular sporozoites emerge from the
WBC's and assume their slender elongate form. The chromatin
condenses into a spherical mass and the sporozoite moves by
slow bending motions of the anterior portion. At least in
two insects, £x. errati cus and J_. vexator, prior to
digestion of the bloodmeal some sporozoites penetrate the
epithelial cells of the midgut where they remain quiescent
in what appears to be a parasitophorous vacuole until
ingested (Figures 78 to 80). In the laboratory, S. go 1vani
remain infective in £x. erraticus for up to 29 days
(probably longer) following a bloodmeal on an infected anole
and infective for up to six days in J.. vexator (Table 10).
Sporozoites teased from the gut of both species of insects
move as previously described and are elongate, pointed
anteriorly, and retain their condensed spherical chromatin
(Figures 76 and 77). Impressions of the midgut of one anole
on day 56 post-ingestion revealed several groups of
macromerozoites (Figure 81). Macroschizonts with


LIST OF TABLES
Table Page
1.Laboratory transmission data of PIasmodiurn
mexicanum to Seeloporus undulatus by bite of
infected Lutzomy i a vexator female's 25
2. Linear regression analysis of acute PIasmodiurn
mexicanum infections with fewer than 500
parasites per 10,000 red blood cells (5%
parasi temi a) 34
3. The effects of temperature on sporogony of
PIasmodiurn mexicanum in Lutzomyia vexator 66
4. Summary of PI a smodiurn floridense infections in
Anolis carolinensis and Sceloporus undulatus
collected from different localities in Florida
( 1983-5 ) 113
5. Summary of CDC light and lizard bait trap
collections and bloodfeeding of feral mosquitoes
in the field and laboratory from 31 April to 10
October, 1984 117
6. Summary of laboratory transmission data of
PIasmodiurn f1oridense to Anolis carolinensis and
Sceloporus undulatus for a three year period.... 123
7.Laboratory transmission data of PIasmodiurn
floridense to Anolis carolinensis 130
8. Summary of sporogony and transmission of saurian
malaria in bloodfeeding Diptera 131
9. Sporogonic development of PIasmodiurn f1oridense
on the midgut of Cu 1 ex er rati cus 153
10.Experimental transmission data of Schel1ackia
go!vani and Schellackia occidenta1is to wild
caught Anolis carolinensis and Sceloporus
undul atus by ingestion of arthropods 177
viii


Figure 81. Impression of small intestine of Anolis
carolinensis with macromerozoites (Mm) of
Schel1acki a g o1v a ni (Giemsa).
Figure 82. Section of intestine of Anolis carolinensis
with macroschizont (Ms) and macromerozoites
(Mm) of Schel1ackia golvani
(Giemsa-collophonium).
Figure 83. Section of intestine of Anolis carolinensis
with microschizont (Me) with developing
micromerozoites of Schellackia gol vani
(Giemsa-collophoni um)'.
Figure 84. Section of intestine of Ano 1is carolinensis
with macro- and mi crogametocytel {IT] of
Schel1ackia gol van i (Giemsa-col1ophonium).
Epithelium (E~) and lamina propria (Lpr).
Figure 85. Sporozoites (S) of Schel1ackia occidentalis
teased from the gut of a mite, Gekobi el 1 a
texana, which was feeding on an infected
Seeloporus undulatus (Giemsa).


-57-
Da g g e 11, 1973). Separate studies indicate that both species
are highly susceptible to P_. mexi canum and often die of
fulminating infections during the acute phase (Ayala, 1971;
Jordan, 1970; Thompson and Huff, 1944a; and Thompson, 1944).
The age of the lizard (S_c. occidental is) also appears to
have a significant effect on the course of the infection.
When hatchling lizards (3-5 months old) were blood
inoculated with P_. mexi canum, all lizards died of
fulminating infections but only three of 10 wild caught
naturally infected yearling Sc_. occidental is died (Ayala,
1971). However, the course of the infection for blood
inoculation of some malarias is often more severe than
sporozoite inoculation. In the present studies, two
yearling and seven mature S^. undul atus collected in Florida
were experimentally infected by bite of L.. vexator. Six of
the lizards (including the two yearling lizards) died of
fulminating infections within 96 days after exposure. The
other three lizards were killed when they became lethargic
and parasitemias reached approximately 20-75%.
Course of Infection, Parasitemia
Parasitemias of _P. mexi canum ranged from 930 (yearling)
to 11,960 (mature female)/10,000 RBC at the time of death
(Table 1, Figure 7). Maximum parasitemias attained by P_.
mexicanum from previous studies ranged from 4,100 to 8,100
for Seel oporus olivaceaous, 2,812 for Sc^. undul atus


TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS iii
LIST OF TABLES viii
LIST OF FIGURES ix
ABSTRACT xvi
CHAPTER
1SAURIAN MALARIA AND Schellackia 1
Saurian malaria 1
Schel 1 acki a 12
2 EXPERIMENTAL TRANSMISSION OF Plasmodium mexicanum
BY BITES OF INFECTED Lutzomyia vexator (DIPTERA:
PSYCHODIDAE) 16
Introduction 16
Materials and Methods 17
Lutzomy i a vexator Colony Maintenance 17
Collection and Laboratory Maintenance of
Lizards 18
PIasmodiurn mexicanum Transmission Studies.... 19
Course of Infection, Parasitemia 20
Histology of Infected Lizards 21
Results 21
Plasmodium mexicanum Transmission Studies.... 21
Course of Infection, Parasitemia 29
Histology of Infected Lizards 49
Discussion 49
PIasmodiurn mexicanum Transmission Studies.... 49
Course of Infection, Parasitemia 57
Histology of Infected Lizards 60
3 SP0R0G0NY, DEVELOPMENT, AND ULTRASTRUCTURE OF
EXTRINSIC STAGES OF Plasmodium mexicanum 62
Introduction 62
Materials and Methods 63
Sporogony of Plasmodium mexi canum 63
Ultrastructure of Extrinsic Stages of
Plasmodium mexicanum 64
v


-71-
27C were highly infective by days 8-10 post-feed with 9 of
13 Seel oporus undul atus becoming infected with J?. mexi canum
when bitten by 1-3 sand flies with sporozoites in the
salivary glands (Chapter 2).
Ultrastructure of Extrinsic Stages of Plasmodium mexicanum
Oocyst. Electron micrographs of oocysts of
mexicanum on the midgut of l. vexator were taken from days
4-7 post-feed. The early oocyst (day 4) appears as an
undifferentiated solid sphere in which the oocyst capsule
[range, 10.8-27.0um; mean 23.0um (Table 3)] is
extracellular, directly applied to the oocyst plasma
membrane, and protruding into the hemocoel (Figure 24). The
oocyst capsule is in direct contact with the basement
membrane of the midgut epithelial cells and is composed of
an amorphous granular material which is slightly more
electron dense than the basement membrane. Intracellular
development of oocysts in the midgut epithelium was not
observed. Changes in the oocyst capsule are similar to
those described by Duncan et al ( 1960) for P_. cathemeri urn
and Vanderberg et al ( 1 967 ) for P_. berghei The external
and internal margin of the oocyst capsule is smooth in early
oocysts, where it is not in contact with the midgut. The
margin of the oocyst capsule in contact with the basement
membrane is often thickened and invaginates into the plasma
membrane of the oocyst (Figure 24). As the oocyst matures,


-17-
that the natural route of infection is by bite because (1)
sporozoites occur in the salivary glands (2) time for
bloodfeeding was long and (3) infectivity of sporozoites
occurred after intraperitoneal inoculation. Ayala did not
rule out the possibility of transmission by ingestion of
infected flies. Transmission of P_. agamae is believed to be
by bite but the sporozoites were retained in the oocyst and
did not migrate to the salivary glands of C_. nubecul osus (an
unnatural vector) (Petit et al., 1983).
New developments in rearing phlebotomine sand flies
(Endris et al ., 1982) provided the opportunity for
experimental transmission studies of P_. mexi canum. This
study describes the first successful experimental
transmission of a saurian malaria by bite of a hematophagous
insect other than mosquitoes. The incubation period and
course of acute infection of P_. mexi canum i n Seel oporus
undul atus, transmitted by bite of L. vexator, are also
reported.
Materials and Methods
i
Lutzomyia vexator Colony Maintenance
The colony of Lutzomyia vexator, originating from wild
caught females from Gulf Hammock, Levy Co., Florida, USA, in
1981, was maintained by methods similar to those described
by Endris et al ( 1982). Larvae, however, were provided


-148-
non-mosquito, Cu 1 i coi des nubeculosus But the significance
of this in not known because £. nubeculosus is an unnatural
host. Another saurian malaria, P. floridense, transmitted
by mosquitoes, will also develop in a phlebotomine sand fly
(£. vexato r), and occasionally produce sporozoites (Chapter
4).
The purpose of this study was to examine the fine
structure of P_. flor i dense which has sporozoites
morphologically similar to avian and mammalian malaria
sporozoites. A comparison of sporogonic development and
sporozoite morphology to other saurian, avian, and mammalian
malarias is made.
Materials and Methods
Sporogony of Plasmodium floridense
Laboratory reared Cu 1ex erraticus were bloodfed on
Ano 1is carolinensis as previously described (Chapter 4).
Bloofed females were removed at 12 hr intervals, placed in
100 ml plastic urine specimen containers with a screened
lid, provided a 10% sugar solution, and maintained in a
temperature-humidity controlled chamber at 25C. Midguts
were dissected at periodic intervals, placed on a clean
slide with a drop of cold-blooded Ringer's solution, and
covered with an 18 mm circular coverslip. The number and
measurements of 25 (or fewer) oocysts were recorded.


-205-
Ball, G. H., and J. Chao. 1964. Temperature stresses on
the mosquito phase of Plasmodium relictum. J.
Parasitol. 50:748-752.
Barnett, H. C. 1962. The incrimination of arthropods as
vectors of disease. XI. Internationaler Kongress Fur
Entomol. Wien 1:341-345.
Bonorris, 0. S., and G. H. Ball. 1955 Schel 1 acki a
occidental is n. sp., a blood-inhabiting coccidian
found in lizards in Southern California. J. Protozool.
2:31-34.
!
Boulard, Y., G. Petit, I. Landau, A. F. Gomes, and L.
Touratier. 1983. Sporogonie de PIasmodiurn agamae chez
C u1 ic o id e s nubeculosus : II Observations
ultrastructurales. Protistoologica 19:543-551.
Boyd, M. F. 1940. The influence of sporozoite dosage in
vivax malaria. Am. J. Trop. Med. 20:279-286.
Boyd, M. F. 1949. Malariology. Philadelphia: W. B.
Saunders Co.,
Bray, R. S. 1957. Studies on the exo-erythrocytic cycle in
the genus PIasmodiurn. London Sch. Hyg. Trop. Med.
Memoir 12. London: H. K. Lewis.
Bray, R. S. 1959. On the parasitic protozoa of Liberia.
II. The malaria parasites of agamid lizards. J.
Protozool. 6:13-18.
Bray, R. S. 1963. The exo-erythrocyti c phase of malaria
parasites. Int. Rev. Trop. Med. 2:41-74.
Bray, R. S., and P. C. C. Garnham. 1962. The Giemsa-
colophonium method for staining protozoa in tissue
sections. Ind. J. Malariol. 16:153-155.
Chaniotis, B., and J. R. Anderson. 1968. Age structure,
population dynamics and vector potential of Phiebotomus
in northern California. II. Field population dynamics
and natural flagellate infection in parous females. J.
Med. Entomol. 5:273-292.
Chao, J., and G. H. Ball. 1962. The effect of low
temperature on Plasmodium relictum in Culex tarsalis.
J. Parasitol. 48:232-254.
Coatney, G. R., W. C. Cooper, and V. I. Miles. 1945.
Studies on Plasmodium gallinaceum. I. The incidence
and course of the infection in young chicks resulting
from single mosquito bites. Am. J. Hyg. 41:109-118.


-96-
size to approximately 36um and eventually protrude through
the basement membrane, extending into the hemocoel (Sterling
and Degiusti, 1974). In contrast, the oocysts of agamae
are intracellular in the epithelial cells of the midgut of
C. nubeculosus (Boulard et al., 1983). However,
intracellular development of malaria oocysts may be a
response of developing in an unnatural vector. Some oocysts
of PI a smodiurn b e r g h ei Vinke and Lips have also been reported
to undergo intracellular development in Anopheles stephensi
and An_. quadrimaculatus Say. Both are unnatural vectors of
P^. berghei but are natural vectors of human malarias.
Organelles and inclusion bodies of the solid oocyst and
vacuolated oocyst are similar to those described for JP.
cynomolgi (Terzarkis, 1971) and other malaria parasites.
The nuclei, with a prominent nucleolus, are large and
scattered throughout the cytoplasm in the solid oocyst
(Figure 24). Subsequent to vacuolization and cleft
formation, the nuclei migrate to the periphery of the
sporoblast. The mitochondria appear to be irregular in
shape, are often seen in clusters, and may be branched as
described by Duncan et al ( 1960) (Figures 25 and 26).
Oocyst differentiation and sporozoite formation.
Differentiation of the oocyst of P_. mexicanum is similar to
other malaria species. A subcapsular space which separates
the sporoblastoid from the oocyst wall is formed by
vacuolization along the periphery of the oocyst capsule.
The physiological changes which cause the separation of the


-113-
Table 4. Summary of Plasmodium f1oridense infections in
Ano 1is carolinensis and Sceloporus undulatus
collected from different localities in Florida
(1983-5). All are located in Alachua County
except for Gulf Hammock (Levy Co.) and Ocala
National Forest (Marion Co.)
Locality Species Number Number(%)
collected infected
Hatchet Creek
Gainesvi11e
Cross Creek
Austin Cary Forest
Ocal a Natl. Forest
Gu1f Hammock
San felasco Park
Total
A. carolinensis
Sc undulatus
A. carolinensis
A. carolinensis
Sc. undulatus
A. carolinensis
Sc undulatus
A. carolinensis
A. carolinensis
A. carolinensis
A. carolinensis
Sc. undulatus
45
17(37.8)
1
0
47
6(12.8)
18
3(16.7)
62
0
33
13(39.4)
51
6(11.8)
2
0
3
2(66.7)
6
0
154
41(26.6)
114
6(5.3)


-154-
salivary glands after more than 20 days post-feed. Apparent
host response to oocyst development was evident in some
oocysts after 10 days post-feed. In some mosquitoes which
were dissected 22 days after feeding, many oocysts and
sporozoites (?) appeared to be melanized (Figure 59).
Mosquitoes dissected from the same lot (fed on the same
lizard during the same time interval) did not demonstrate
any signs of host-parasite response, 9 and 11 days after
feeding (Figure 55 ) .
Ultrastructure of Extrinsic Stages of Plasmodium floridense
Oocyst. Electron micrographs of oocysts of P_.
floridense were taken at 8 and 12 days after the infective
bloodmeal. Because of the asynchronous development of
oocysts, midguts sectioned at 12 days after feeding had
oocysts in all stages of development, i. e.,
undifferentiated oocysts and oocysts with fully formed
sporozoites (Figures 55, 62, 64, and 66). The eight day old
oocyst appeared as a solid undifferentiated sphere enclosed
by a thick extracellular oocyst 'capsule (Figure 60). The
position of the oocyst in relation to the midgut was
variable. Most oocysts appeared to protrude slightly into
the hemocoel while others were tightly packed between midgut
epithelial cells or protruded into the midgut lumen (Figure
55). The thick oocyst capsule appeared to be composed of
two amorphous layers of moderately electron dense material


-185-
developing macromerozites mi croschizonts, and gametocytes
were also present in thin sections of the midgut (Figures 82
to 84.
Schellackia occidentals
Transmission studies. The results of experimental
transmission of S^. occidentals to noninfected wild caught
Sc. undulatus by ingestion of bloodfeeding arthropods which
previously fed on infected lizards are shown in Table 10.
The prepatent period for two Sc^. undul atus maintained at
room temperature (18-24C) was relatively long (38 days).
The prepatent period for lizards maintained at 32C was much
shorter (7-17 days). Nearly all of the noninfected Sc.
undu1atus that were force fed arthropods and which had
earlier bloodfed on infected lizards, demonstrated
sporozoites in their bloodfilms.. Two of the three Sc.
undulatus which were fed mites from an infected lizard and
none (0/3) of the lizards which were force fed mosquitoes
(Cx. erraticus) seven days subsequent to a bloodmeal, became
patent. However, all 12 fence lizards fed mosquitoes by day
one following a bloodmeal, became patent. None of the A.
carolinensis which were fed mosquitoes and sand flies
infected with S^. occidentals demonstrated sporozoites in
their bloodfilms. Sporozoites were not observed in the
bloodfilms of noninfected Sc. undulatus which were kept in a
screened cage with G. texana-infested Sc. undulatus with


-59-
differences may be attributed to age and sex of the lizards
(adult females surviving the longest), host immune response
to the parasite, adaptation to a laboratory environment,
temperature [orientation to the light source (behavior)],
length of time surviving patent infection, number of
sporozoites inoculated during feeding, and other factors
which were not determined.
According to Thompson and Huff (1944a), variation in
the course of infection, gametocyte production, and cellular
distribution of £. mexicanum parasites is due to host
differences rather than alteration of parasites. They found
that P.. mexi canum, a parasite found naturally in 5c.
occidentals, Seel oporus torqu atu s (=ferrariperzi ) and 5c .
undulatus, lost its gametocytes when transferred by blood
inoculation to another lizard, C_. col 1 ari s However,
gametocyte production resumed upon experimental passage of
the parasite to a third host species, Sc_. ol i vaceous.
"Mature" gametocytes were observed in all lizards except
S-14 and S-15 during the infection. Lutzomyia vexator that
were bloodfed on S-51 and S-42 on days 11 and 21,
respectively, following the detection of parasites and when
mature gametocytes were present in the blood, developed low
numbers of oocysts. Sporozoites developed normally and
appeared viable, but were not injected into another lizard
to determine infectivity. It was not determined if
gametocyte production would be increased during the chronic
phase of the infection since all lizards died (or were
killed) during the acute phase.


o
CD
DC
O
O
O
O
to
£
w
<
cc
<
0.
cc
LU
m
2
3
Z
O
o
DAY POSTEXPOSURE
-38-


Figu re
14.
Red blood cells of Sceloporus undulatus (S-51)
infected with Plasmodium mexicanum parasites
during the later part of the infection
(Giemsa). Schizont (Sc), and trophozoite (T).
Figure
15.
Spleen tissue impression of Sceloporus
undulatus infected with numerous Plasmodium
mexicanum parasites (Giemsa). (Merozoites
(M) and schizont (Sc).
Figure
16.
Bone marrow smear of Sceloporus undulatus
infected with Plasmodium mexicanum parasites
(Giemsa). Schizont (Sc) in a white blood cell
and trophozoite (T) in a thrombocyte (Th).
F i g u r e
17.
Schizonts (Sc) of Plasmodium mexicanum in
endothelial cells (En) of capillaries in the
brain. Brain tissue impression stained with
Giemsa.


-104-
transmission of the parasite by sympatric potential vectors
is essential for vector incrimination.
Thompson and Huff (1944a) and Telford (personal
communication) among others indicated that _P. mexicanum is
the most primitive of the known PIasmodiurn sp. It is also
noted that the vectors of P_. mexi canum, i. e., the
phlebotomine sand flies, are considered among the oldest of
Diptera, showing many primitive features. In view of this,
P_. mexi canum may represent an early branch of the ancestral
stock of PIasmodiurn and should perhaps be placed in a
separate genus subgenus. The elucidation of the vectors of
more saurian malarias and the histological and ultrastruture
of oacysts and sporozoites will undoubtedly provide more
information on the phylogeny of saurian PIasmodiurn in
relation to avian and mammalian malaria.


-134-
and IP inoculation of sporozoites) range from 24-25 days and
13-15 days for A. carolinensis maintained at room
temperature (18-24C) and 32C, respectively. The
parasitemia of both lizards rose rapidly, and peaked by 55
and 74 days after feeding. Peak parasitemias were 1,780 and
4,280/10,000 RBC's for each of the lizards (Table 7 and
Figure 51).
The number of trophozoites, schizonts, and gametocytes
seen during the course of P_. floridense infection for each
of the lizards is shown in figures 53 and 54. Both immature
(single nucleated parasites larger than the nucleus of the
host cell, but not displacing the host cell nucleus and with
pigment granules clumped) and "mature" gametocytes (as
described by Garnham, 1966) are included together. The
number of trophozoites and schizonts increased
1ogrithmically until the peak parasitemia was reached.
Thereafter, parasitemias became erratic (AA-59). The number
of gametocytes fluctuated over the course of the infection
and were not observed in several of the bloodfilms from
lizard number AA-59.
Discussion
Field Studies
The present study indicates that a large number of
species of biting Diptera are present at the Hatchet Creek
study site where anoles are commonly found infected with P_.


-51-


-162-


Figure 8. Number of PIasmodium mexicanum trophozoites, schizonts,
gametocytes and percent of infected white blood cells per
10,000 red blood cells during the course of the infection
of Seeloporus undulatus (S-51) infected by bite of
Lu t zomyia vexator .


CHAPTER 3
SPOROGONY, DEVELOPMENT, AND ULTRASTRUCTURE OF EXTRINSIC
STAGES OF Plasmodium mexicanum
Introducin' on
The natural vectors of saurian malaria have eluded
researchers for years. Attempts to incriminate culicine
mosquitoes largely resulted in failure, with sporogony being
limited and erratic in the mosquitoes. While there are more
than 55 species of saurian malaria, only two vectors,
Lutzomyia vexator and Lutzomyia stewarti have been
incriminated (Ayala, 1971). Recent laboratory transmission
studies (Chapter 2) provides more conclusive evidence that
L. vexator is a natural vector of PIasmodiurn mexicanum.
Attempts to incriminate other hematophagous insects have
largely failed. However, Petit et al. (1983) demonstrated
that PIasmodiurn agamae developed in Cul i coi des nubeculosus,
an unnatural host, and speculated that another species of
Cul i coi des which is sympatric with P^_ agamae may be the
natural vector.
The details of the subcellular morphology and
differentiation of many of the avian and mammalian
sporozoites and sporogony as revealed by the transmission
electron microscope (TEM) are well documented. The only
-62-


Figure 64. Cross section of PIasmodium floridense oocyst
undergoing differentiation. Early
vacuolization (V) with narrow linear
extensions extending to the surface of the
sporoblastoid. Nuclei (N) with a electron
dense nucleolus (Nu) are smaller than in
non-differentiating oocysts. "lipid-like"
globules (Lp) are scattered throughout the
cytoplasm.
Figure 65. Convex dense membranes (Dm), the precursors of
the developing sporozoites, form along the
narrow linear extensions. A moderately dense
(Mi) inclusion body, which is the first observed
stucture to enter the developing sporozoite,
is seen adjacent to the dense membranes.


-123-
Table 6.
Summary of
PIasmodiurn
Seeloporus
laboratory transmission data of
f1oridense to Anolis carolinensis and
undulatus for a three year period.
Li zard
Number
Mode of
Number/
Year
species
Lizards
t ran s -
(percent
mission
in fected
1983
A.
carolinensis
5
bloodmeal
1(20.0)
A.
carolinensis
2
IP
0
1984
A.
carolinensis
5
bloodmeal
0
A.
carolinensis
1
IP
1(100.0)
1985
A.
carolinensis
11
bloodmeal
2(18.2)
A.
carolinensis
10
IP
1(10.0)
Sc.
undulatus
1
bloodmeal
0
Sc.
undulatus
2
IP
0
Total
A.
carolinensis
34
_
5(14.7)
Sc .
undulatus
3
0


CHAPTER 6
DEVELOPMENT AND EXPERIMENTAL TRANSMISSION OF Schellackia
golvani AND Schellackia occidental's BY INGESTION OF
INFECTED BLOODFEEDING ARTHROPODS
Introduction
Examination of bloodfilms of Anolis carolinensis (green
anole) and Sceloporus undulatus (eastern fence lizard) from
Florida revealed natural infections of Schellackia golvani
Rogier and Landau ( 197 5 ) and S^. occidental is Bonorris and
Ball (1955), respectively. Schellackia parasites from the
western hemisphere have been described from bloodfilms from
lizards and amphibians but received little attention until
the mid 1970's. The natural vector of only one species of
Schellackia, S^. occi dental i s, in the western hemisphere is
known (Lainson et al., 1976). Yet, transmission by
ingestion of mosquitoes previously bloodfed on infected
lizards was demonstrated for S^. brygooi Landau ( 197 3 ) and S^.
1andaue Lainson, Shaw and Ward. In view of new evidence
given in the present report, a number of biting arthropods
may serve as passive vectors for Schel1ackia parasites.
Reichenow (1919) first described schizogony and
microgametogony of Schel1ackia in the epithelial cells of
the small intestine of lizards and then incriminated mites
as the natural vector. Bonorris and Ball (1955) were the
-171-


Figure 54. Number of Plasmodium floridense trophozoites, schizonts, and
gametocytes per 10,000 red blood cells during the course of
infection in Anolis carol inensis (A-85) infected by bite of
Culex erraticus.


-173-
the Insects Affecting Man and Animals Laboratory, USDA,
Gainesville, Florida. Mites (Gek o b i e 11 a texana) were
brought into the laboratory on wild caught Seeloporus
undul atus and maintained on lizards kept in the laboratory.
An o 1is carolinensis and S£. undulatus were hand collected in
Florida from Gainesville, Austin Cary Forest (10 km N
Gainesville), Hatchet Creek (15 km NE Gainesville) or near
Cross Creek and maintained in the laboratory as previously
described (Chapter 4). In addition, A. carolinensis were
collected from Manchac swamp, Tangipahoa, Parish,
Louisianna. Bloodfilms were made from a clipped toe, air
dried, fixed with absolute methyl alcohol, stained with
Giemsa, and examined for the presence of blood parasites.
Subsequent bloodfilms were made by either clipping the toe
or the tip of the tail.
Laboratory reared C^. errati cus, Ae. aegypti and j_.
vexator were allowed to feed on either A. carolinensis or
Sc undulatus which demonstrated Schel1ackia parasites in
the bloodfilms. Bloodfed mosquitoes were removed at 12 hr
intervals, placed in a screen topped pint carton and
provided with a 10% sugar solution. The bloodfed sand flies
were removed, placed in a screen-topped urine specimen
container partially filled with plaster of Paris, and
provided with a 1:1 solution of distilled water and Karo
syrup. All bloodfed females were maintained in a
temperature-humidity controlled environmental chamber at
27C and 80% RH. Midguts of IL. vexator and Cjc. errati cus


-167-
This "abnormal" development might indicate an unnatural
host. However, evidence indicates that this type of
"abnormal" development might be normal for some Plasmodium
sp. For example, P_. hermani (turkey malaria) oocysts often
develop between the midgut epithelium similar to that of
Leucocytozoon sp., and occasionally protrude into the lumen
of the gut in both C_x. s a 1 i n a r i u s and C_x. nigripalpus -
natural vectors of P_. hermani (M. Young, personal
communication; Nayar et al., 1981).
Sporogonic development of P_. floridense in Cx.
erraticus is much slower, and was not observed in the
salivary glands until at least 11 days, and usually not
until 13-14, days after feeding. Sporogonic development of
P_. floridense in L_. vexato r, a vector of P_. mexi canum, was
equally as long as in C_x. errati cus, but was never observed
in the salivary glands. The time of parasite development
may vary according to the parasite-vector relationship based
on the longevity of the vector. Lutzomyia vexator survival
in the laboratory is very short with most of the flies dying
15-20 days after eclosin, whereas Cx. erraticus adults
survived more than 90 days in the laboratory. The
asynchronous development of oocysts would allow for a
gradual release of sporozoites to the salivary glands. But,
in these studies, it appeared that the slower developing
oocysts often become melanized. However, this may be the
result of the laboratory conditions (temperature) at which
the mosquitoes were maintained.


-95-
Oocyst. The oocysts of malaria species and other
hemoproteids, i. e., Leucocytozoon and Haemoproteus, are
very similar. The solid oocyst of _P. mexicanum and other
mammalian and avian malaria species are extracellular in
location and extend into the hemocoel, at least in the
mature oocyst. There is some question as to the initial
location of the early malaria oocyst, 4. e., lying external
of the basement membrane of the midgut and protruding into
the hemocoel (Terzarkis, 1971; Terzarkis et al., 1967;
Vanderberg et al 1967 ) or between the basement membrane
and epithelial cells of the midgut (Melhorn et al., 1980).
Studies by Melhorn et al on P_. gal 1 i nace urn indicated that
early oocysts lie between the epithelial cell and basement
membrane, but by 100 hours post-infection, the oocysts had
ruptured the basement membrane and protruded into the
hemocoel with only a small region of the oocyst in contact
with the midgut epithelial cells. The oocysts of other
hemoproteids e. g., Leucocytozoon sim o n di Mathis and Leger,
Leucocytozoon dubreui 1i Mathis and Leger, Leucocytozoon
tawaki Fallis, Bissett and Allison, and Haemoproteus
metchnikovi (Simond), also develop between the basement
membrane and epithelial cell of the midgut (Desser and
Wright, 1968; Wong and Desser, 1976; Desser and Allison,
1979; and Sterling and DeGiusti, 1974). Oocysts of
Leucocytozoon only slightly increase in size to maturity and
remain deeply invaginated in the epithelial cell of the
midgut, while mature oocysts of ji. metchni kovi increase in


-196-
were neither observed feeding on parasitic mites in the
laboratory nor became infected when they were maintained in
screened cages with infected lizards harboring numerous
mites (£. texana).
Although mosquitoes and sand flies remain infective
following digestion of the the bloodmeal, there is
essentially no extrinsic incubation period for Schel1ackia.
While transmission of S_. occi dental i s and Si. g o 1 v a n i has
been accomplished by directly feeding blood from infected
lizards to conspecific non infected lizards (Jordan, personal
communication), Lainson ( 1 976 ) demonstrated that Cx^. pi pi ens
fatigans remained infective for at least 14 days subsequent
to a bloodmeal on Polychrus marmoratus infected with .
1andaue.
In the present study, l. vexator that fed on Sc.
undul atus infected with S_. occi dental i s remained infective
for seven days. Lutzomy i a vexator and C_x. errat i cus that
fed on A_. carolinensis infected with Si. go 1 vani remained
infective for six and 29 days, respectively, after feeding
on infected lizards. The reason that C_x. errat i cus was not
infective on day seven after a bloodmeal on Sc. undulatus
infected with S_. occidentalis can not be explained. The
sample size was small and only included three lizards, but
the question "do some or all mosquitoes remain infective
following digestion of the bloodmeal?" must be raised.
Whereas malaria transmission requires the survival of
the vector for a certain extrinsic incubation period (until


Figure 37. Magnification of sporozoites of PIasmodium
mexi canum in the salivary gland of Lutzomyia
vexator. The anterior moderately dense
inclusion bodies are absent. Micronemes (Mn)
and elongate rhoptries (Rh) are in the
anterior of the sporozoite. The nucleus (N)
is slightly subcentral with a mitochondrion
(M) and associated dense body (Db) in the
posterior of the sporozoite.
Figure 38. Sporozoite of PIasmodium mexicanum
illustrating polar rings (Pr ) MTcron ernes
(Mn), nucleus (N), mitochondrion (M), and
dense inclusion bodies (Di) are also observed.


-163-
and tubular as it takes the shape of the sporozoites.
Mitochondria observed migrating into the sporozoite were
tubular and very long (Figure 69). The mitochondrion is
U-shaped and in some sections it appears that there are
several mitochondria (Figure 67). Subpel1icular
microtubules were infrequently seen in developing
sporozoites. They were only in the anterior 1/3 of the
sporozoite and ranged from 10-11 in number (Figure 70)
Micronemes were also infrequently observed in sporozoites in
the oocysts.
Sporozoites. Sporozoites escape from the oocyst and
migrate to the salivary glands. Sporozoites are
intracellular and appear to be in groups (Figure 71).
Living and Giemsa stained sporozoites are narrow and
elongate, measuring 16-19 X l-2um (Figure 54). Micronemes
are large and scattered throughout the anterior 1/3 of the
sporozoites (Figure 72). Micronemes were only observed in
nearly mature sporozoites in the oocysts and sporozoites in
the salivary glands. Micronemes appeared to be abundant in
some sporozoites which were apparently trapped in abnormal
oocysts, i. e., were becoming melanized (Figure 73). The
rhoptries are long and sinuous, extending nearly to the
nucleus. A cytostome was not observed in either developing
or mature sporozoites. The subpellicular microtubules were
infrequently observed and are apparently only in the
anterior of the sporozoite. Subpellicular microtubules are
arranged asymetrically in developing sporozoites (Figure


-4-
species of sympatric biting Diptera (mosquitoes and sand
flies) from areas of endemic saurian malaria in Africa after
feeding them on lizards infected with IP. agamae and P_.
giganteum Theiler. Gametes and ookinetes were described,
but further development did not occur.
Jordan (1964) had little success demonstrating
sporogony in local mosquitoes collected in Georgia, USA,
where Anolis carolinensis Voight and S£. undulatus are
naturally infected with P_. f 1 ori dense. Five species, Aedes
atlanticus-tormentor Dyar and Knab (88), Aedes triseriatus
(Say) (10), Psorophora confinis (Lynch Arribalxaga) and
Psorophora ferrox (Von Humboldt) (20), and Coqui 11ettidia
(=Mansonia) perturbans (Walker) (25) were all negative.
However, 1/80 Aj. aegypti had one oocyst, 4/70 Cu 1 ex
territans Walker had 1-23 oocysts, 2/150 Cu 1 ex
quinquefasciatus Say had 1-3 oocysts, and 1/3 unidentified
Cu 1 ex sp. had 70 oocysts. (Unfortunately the Cu 1 ex sp. with
70 oocysts was not identified to species.) Anolis-baited
traps, designed to attract and capture hematophagus insects,
were ineffective (Jordon, 1964).
Landau et al ( 197 3) attempted to infect Ae^. aegypti ,
Anopheles stephensi Liston, Culex fatigans Weideman, and Cx.
pi piens by feeding them on Tupinambus teguixin (L.) infected
with Saurocytozoon (=P1asmodiurn) tupinambi Lainson and Shaw
( 1969), a plasmodiid. Only C_x. p i p i e n s developed oocysts,
but these were mostly abnormal, with sporozoites only
observed in the oocysts, and never in the salivary glands.


-197-
the sporozoites are in the salivary glands), Schel1ackia
parasites depend upon the lizard ingesting the saurian
bloodfeeding arthropods, either shortly after or sometime
after the bloodmeal. Sporozoites of S_. gol v a n i invade the
intestinal epithelium of both C_x. erraticus and L. vexator
(Figures 78 to 80) where they remain quiescent in a
parasitophorus vacuole similar to that shown by Lainson
( 1976 ) for S_. 1 andaue. It is also believed that S_.
occidentalis sporozoites invade the intestinal epithelium of
j_. vexator since they remain infective in l. vexator after
digestion of the bloodmeal. However, the survival of
Schel1ackia sporozoites in other potential arthropod vectors
is unknown. Furthermore, the effect of subsequent
bloodmeals on the sporozoites remains unstudied.
Temperature has a marked influence on the prepatent
period for both S.. occidentalis and S_. gol vani The
prepatent period for S_. occidentalis at room temperature
(18-24C) was slightly longer, 37 days post-ingestion (n=2),
than that for _S. go 1vani 21-25 days post-ingestion (n = 7).
At room temperature, one A. carolinensis did not demonstrate
sporozoites in the blood until day 81 post-ingestion. No
explanation for this extended prepatency is provided except
that it may be dose related as well as an individual host
response. However, at 32C (90F), the prepatent period was
as short as seven days post ingestion for S_. occidentalis and
10 days for S_. gol vani Sporozoites were not observed in
the blood of Sc. undulatus on day six, and only a few


TABLE 1. Laboratory transmission data of PIasmodium mexicanum to Seeloporus
undu1atus by bite of infected Lutzomyia vexator females.
No.
Lizard flies
number fed
Sporozoite Day
rate3/ patent
(day post infect-
feed) ion
Day post Duration
feed of patent
lizard infection
died (day)
(killed)
No. para %RBC inf %WBC inf
at death at death at death
(killed) (killed) (killed)
S-8
3
+ 3(7)
+ 3(7)
27
52
25
2095
20.3
25.0
+ 3(8)
S-14D
1
+ 3(9)
26
39
13
930
9.2
10.7
S-15
1
+ 3(8)
26
47
21
2130
20.2
44.4
S 2 5
1
+ 3(9)
33
52
19
2780
25.9
34.3
S -42
1
+ 3(8)
33
61
28
8270
66.5
55.6
S 4 3
2
+ 2(7)
+3(10)
26
(66)
(40)
(7122)
(53.0)
(25.0)
S 47
3
+ 3(8)
+ 3(8)
+ 3(8)
23
(45)
(22)
( 3070
(28.5)
(37.5)
S-50
3
+ 3(8)
+ 2(9)
+ 3(9)
23
(46)
(23)
(1370)
(12.6)
(N/D)
S 51
1
+ 1(9)
40
96
56
11,960
91.2
24.2
Average^
28.6
50.2
27.0
4379
38.9
32.4
^ +1, 1-10; + 2, 11-100; +3, >100 sporozoites.
c Yearling lizard.
d Lizard maintained in a temperature-controlled incubator at 27C.
Lizards which were killed are not included in the average.


-101-
mexi canum and P_. agamae. However, the microtubules of P.
agamae are arranged asymetrically with approximately 2/3
(14 of 23) of the microtubules in approximately 1/3 of the
circumference and the other 1/3 (9 of 23) of the
microtubules in the other 2/3 of the circumference (Boulard
et al., 1983). The microtubules of mexicanum are
arranged asymetrically with approximately 2/3 (9 of 14) of
the microtubles in 1/2 (0.42%) of the circumference of the
sporozoite and 1/3 (5 of 14) in the other half of the
circumference (Figure 3-22). Microtubules of non-malarian
hemoproteids are arranged symetrically around the
circumference and extend the full length of the sporozoite.
The significance of the differences in the subpellicular
microtubule arrangement of saurian and other types of
malaria is not understood.
Mature oocysts are either ruptured by the sporozoites
or are perforated by the escaping sporozoites as shown by
Sinden ( 1975) for P^. y o e 1 i i ni geri ensi s Sporozoites in the
hemocoel are similar to those observed in the salivary
glands except rhoptries are not as elongate and only a few
micronemes are present. Living and Giemsa-stained
sporozoites of P_. mexi canum and P^. agamae (Petit et al.,
1983) are short and stout (6.5xl.5um and 5.7xl.5um,
respectively) when compared to mammalian and avian malarias
which range from 9-15x1.5um. Sporozoites of P^. f 1 ori dense,
a saurian malaria of Ano lis carolinensis, are similar in
gross morphology to the mammalian malarias, i. e., >10um


Figure 7
Course of acute infection of PIasmodium mexicanum
Seeloporus unduiatus infected by bite of Lutzomyia
in nine
vexator.


-169-
closely parallel that of P_. berghei (Vanderberg et al.,
1967), but is also very similar to that of other malarias
which have been described (Chapter 3; Boulard et al., 1983;
Garnham et al., 1963; Duncan et al., 1960; Terzarkis 1971;
and Terzarkis et al 1967 ).
The position of the oocyst on the midgut, as already
discussed, is quite variable and may effect the development
and growth of the oocyst. The oocyst capsule of
flor i dense is usually very thick and similar to that of P_.
agamae. Internal vacuolization of the oocyst and separation
of the sporoblastoid from the oocyst capsule appears early
in P_. agamae and P_. mexi canum. However, while internal
vacuolization of P_. floridense occurs, the sporoblastoid
does not pull away from the oocyst capsule until some time
after sporogonic development is initiated. The
sporoblastoid does not initially form sporoblasts until late
in sporogonic development, similar to that of P_. berghei ,
whereas sporoblasts appear to be formed during the initial
development of sporozoites in P_. mexi canum. The
significance of these relationships is unknown, since most
malaria parasites studied form sporoblasts early in
vacuolization.
The arrangement of the subpel1icular microtubules of
developing sporozoites appear to be similar to other
non-saurian malarias which have been studied. However, the
subpellicular microtubules were clumped in sporozoites in
the salivary gland, apparently as a result of poor fixation.


-106-
of many mammalian and avian malaria parasites has been
described, only two species of phlebotomine sand flies,
Lutzomyia vexator and Lutzomyia stewarti have been
incriminated as vectors of a saurian malaria, PI a smodiurn
mexicanum (Ayala and Lee, 1970; Chapter 2). Attempts to
incriminate mosquitoes as a vector of this, or other,
s a u rian PI asmodium have failed or have been unsuccessful.
In some cases, a few mosquitoes developed small numbers of
oocysts, but none showed sporozoites the infective stage.
This study describes the first successful laboratory
transmission of a saurian malaria, P_. flor i dense, by bite of
a mosquito. Also, the incubation period and course of acute
infection of P_. f 1 ori dense i n An ol i s carolinensis are
reported for the first time following infected bites of Cx.
erraticus and intraperitoneal (IP) inoculation of
sporozoites from the salivary glands of erraticus.
Materials and Methods
Field Studies and Collection of Mosquitoes
The Hatchet Creek study site, located near highway 26,
approximately 15 km NE Gainesville, Florida, USA, was
selected because of the relativley high incidence of
PI asmodium floridense in Anolis carolinensis previously
collected from this area (D. Young, personal communication).
Hatchet Creek is a permanent stream, varying from 2-7 meters
across and emptying into Newnans Lake. At the study site,


-76-


-128-


BIOGRAPHICAL SKETCH
Terry A. Klein was born in Salem, Oregon, on 1 June
1946. He was raised in a rural farm environment in the
Willamette Valley, Oregon, attended Aumsville Elementary
School, Aumsville, Oregon, and graduated from Cascade High
School, Turner, Oregon, in 1964. He attended Oregon College
of Education, Monmouth, Oregon, where he received the
Bachelor of Science degree in 1968 with a major in secondary
education. He returned to Cascade Junior High School, where
he taught earth and life sciences and coached wrestling
until 1973. He accepted a research grant to attend Oregon
State University in 1973 where he received his Master of
Science degree in 1975.
Terry entered military active duty (US Army) in
November, 1976, and was assigned to Regional Division West,
Fitzsimmons Army Medical Center, Denver, Colorado, where he
coordinated with state and local persons and recommended
control measures for arthropod and other pests on military
installations. His next assignment took him to Bankok,
Thailand, where he assisted in coordinating a Dengue fever
epidemiological study, investigated the effects of monkey
malaria parasites on Anopheles diru s initiated the human
use protocol for malaria vector studies, conducted
-215-


Figure 66. Oocyst of Plasmodium f1oridense with
developing sporozoites. Sporozites of P_.
floridense appear to be loosely packed within
the oocyst capsule (C). Sporozoites bud off
the sporoblastoid, leaving a redi dual body (Rb)
Figure 67. Higher magnification of longitudinal section of
the posterior portion of a developing
sporozoite. In longitudinal section, several
mitochondria (M) appear to be present. The
nucleus (N) appears long and slender and takes
the shape of the sporozoite.
Figure 68. Developing sporozoites of PIasmodiurn floridense
The rhoptries (Rh) become elongate as the
moderately dense inclusion bodies (Mi)
degenerate. The previosly nearly spherical
nucleus (N) takes the shape of the sporozoite
and becomes tubular and elongate. Dense
inclusion bodies (Di) are frequently seen in
sporozoites.
Figure 69. Developing sporozoites of PIasmodiurn f1oridense
The mitochondrion becomes elongate as it takes
the shape of the developing sporozoite. Only
one mitochondrion (M) was seen migrating into
the sporozoites. Rhoptries (Rh) appear as
electron dense tubules.


22
20
ie
18
14
12
10
8
6
4
2
0
1 8 15 22 29 5 11 19 28 3 10 17 24 31 8 14 21 28 5 10 17 24 1 9
MAY JUNE JULY AUG SEPT OCT
M
N)
t-O
DAY/MONTH (1984}


-160-
where it begins to pull away from the oocyst capsule (Figure
64). There is no apparent subdivision of the sporoblastoid
into separate sporoblasts in the early differentiating
oocyst. However, in later oocysts where sporozoites are
nearly formed, several sporoblasts are often seen in
ruptured oocysts.
|
Sporozoite formation for P_. f 1 ori dense is similar to
that described for other malarias (Chapter 3; Terzarkis et
a 1 1967; Vanderberg et al., 1967). Linear to convex dense
membranes which initiate the formation of sporozoites are
observed along the length of the linear vacuole extensions
(Figure 65). The convex membranes (= linear dense areas)
are characterized by a dense inner thickened membrane which
develops immediately beneath the sporoblast membrane. The
outer membrane continues to evaginate while the inner
membrane forms along the junction of the evaginating
sporozoite. A moderately dense inclusion body was observed
associated with the early formed convex membranes (Figure
65), similar to that for £. mexicanum (Chapter 3). As
sporozoites elongate, a central residual body which
eventually separates to form several sporoblasts in late
sporogony is formed (Figure 66). The anterior inclusion
bodies begin to degenerate early in the developing
sporozoite as the rhoptries are formed (Figures 68 and 69).
Initially the rhoptries are short and sac-like, but later
elongate until they nearly reach the subcentral position of
the nucleus. The nearly spherical nucleus becomes elongate


-152-


-103-
mosquitoes and therefore the constriction observed in P..
mexicanum passing through the salivary gland ceil can not be
said to be typical. A junctional attachment site at the
point of contact of P_. mexicanum sporozoites and the
salivary gland cell as observed in merozoites of _P. know! es i
(Aikawa et al., 1978) entering a red blood cell was not
observed. Salivary glands of infected J_. vexator dissected
subsequent to a bloodmeal were often filled with many
sporozoites, indicating that only a few were probably
injected during feeding. Examination of the mouthparts of
females which had bloodfed and had >100 sporozoites in the
salivary glands revealed only a few sporozoites (five or
fewe r).
Based on a different vector (ph1ebotomine sand fly) and
histological and TEM examination of the sporozoite, P_.
mexicanum differs from other avian and mammalian malarias
and another saurian PI asmodi urn, P_. f 1 ori dense (Chapters 4
and 5), now believed to be transmitted by a culicne mosquito
(Cu 1 ex erraticus) The natural vector of P_. agamae, also
infecting lizards, is uncertain, although sporogony occurred
in one species of Cu1 icoides (Petit et al., 1983). We
observed sporogony of _P. flor i dense in Lutzomy i a vexator
with a few sporozoites being occasionally observed in the
hemocoel Attempts to transmit P_. flor i dense to lizards
with l. vexator failed, but transmission by a culicine
mosquito was successful. Speculation on the vector status
of a species can be made when sporogony is observed but


-115-
Cou rse of Infection ,Parasitemia
Bloodfilms of A. carolinensis previously fed on by
infected C_x. errati cus, or that were injected IP with
sporozoites, were made at day 0 and at 2-4 day intervals 10
days after exposure to infected bites or IP inoculation of
sporozoites. Parsitemias were expressed as the number of
parasites per 10,000 red blood cells (RBC). A sufficient
number of RBC's was counted to keep the probable error
within 10% according to the method of Gingrich (1932).
Results
Field Studies
A summary of P^. f 1 ori dense infections in wild caught A.
carolinensis and Sc_. undul atus from Hatchet Creek and other
localities near Gainesville, Florida is shown in Table 4.
During 1983-4, 17 of 45 (37.8%) of A. carolinensis collected
at Hatchet Creek were infected with £. f1oridense. The
average infection rate for all anoles collected was 26.6%
(41/154). Only 5.3% (6/114) of the bloodfilms of Sc.
undul atus demonstrated parasites of P_. f 1 ori dense Although
16.7% (3/18) A. carolinensis from Cross Creek demonstrated
parasites of P^. f 1 ori dense none (0/62) of the S£. u ndu 1 atu s
were infected.
A summary of mosquito species collected in CDC light
and lizard-baited traps at Hatchet Creek and bloodfeeding


-216-
hybridization studies on sibling mosquioto species, and
collaborated with local Thai nationals on cytogenetics of
potential malaria vectors. Following the assignment in
Thailand, Terry was assigned to the Advanced Medical Service
Corps Officer Training Course, Fort Sam Houston, Texas, and
Regional Division North, Fort Meade, Maryland.
Terry entered the Graduate School at the University of
Florida in August, 1982. He will be accompanied by his
wife, Jacqui three sons, Kevin, Aaron, and Robert, and
daughter, Michelle, to his next assignment in Brazilia,
Brazil, where he will continue to work with mosquitoes and
malaria.


Figure 9. Number of PIasmodiurn mexicanum trophozoites,
schizonts, gametocytes and percent of
infected white blood cells per 10,000 red
blood cells during the course of the
infection of Seeloporus undulatus (S-42)
infected by bite of Lutzomyia vexator .


-49-
Histology of Infected Lizards
Numerous parasites were observed in lymphocytes of
spleen tissue impressions of dead lizards (dead less than 2
hrs) (Figure 15). Immature erythrocytes and lymphocytes
from bloodfilms of bone marrow extracts were also infected
with P_. mexi canum parasites (Figure 16). However, the
number of parasites observed in the spleen tissue
impressions was much greater and appears to be the primary
site of attack. Schizogony was also observed in the
endothelial cells of the brain in some lizards (Figure 17).
Other tissues, i. e., lung, liver, kidney, intestine,
pancreas, heart, and uterus were considered to be
non-infected. Occasionally, cells of the above tissues
appeared to be infected, but it could not be determined if
the parasites were external from ruptured white blood cells
or intracellular, since fixed or circulating lymphocytes
within these tissues often had numerous parasites. Thin
sections from one lizard, (S-51) also failed to reveal
parasites in these tissues.
Discussion
Plasmodium mexicanum Transmission Studies
Data reported herein demonstrate conclusively that P_.
mexicanum can be transmitted from Sc. occidentals to S.


-58-
undul atus and 2,750 for S_c undul atus consobri nus (Thompson
and Huff, 1944a; Thompson, 1944). Maximum parasitemias for
unnatural hosts, Phrynosoma cornutum and Crytophytus
col 1aris only reached 392 and 238, respectively (Thompson
and Huff, 1944a). Results in the present studies are
similar to those of Thompson and Huff (1944a) and Thompson
(1944), except that several of the S£. undulatus which were
blood inoculated in the previous studies did not develop
fulminating infections. The higher parasitemias in some
lizards in the present study may be a result of a larger
sample size.
The course of the acute infection for S£. undulatus
infected with £. mexicanum is shown in Figure 7. The
predicted mean parasitemia for an "average" S£. undulatus
infected with sporozoites, plotted on the mean day that
parasites were first observed in the bloodfilm, is shown by
2
the dotted line (R =.88). The coefficient of determination
(R ), linear regression equation and slope for parasitemia
levels of less than 5% for each lizard are shown in Table 2.
In general, it appears that lizards which survive for a
longer period of time (>25 days) subsequent to patent
infection, develop parasitemias at a slower rate (slope
<0.40). However, these infections are observed over a
longer period of time and lizards develop higher
parasitemias (>70%). Although the curves of the transformed
course of infection appear to be similar, there are
significant differences between some of the curves. These


DAY POSTEXPOSURE
PERCENT PARASITEMIA
-12-
100


-18-
finely ground horn fly larval medium (Greer and Butler,
1973) that decreased the larval development time from that
observed in larvae fed on aged rabbit feces (Young et al ,
1981). Larvae from individual females were transferred from
25 ml plastic oviposition vials (12-20 days post-eclosion)
to 120 ml urine specimen containers. Approximately 200
larvae were placed in each large vial. Adults were released
daily from the 120 ml containers into a modified glass
aquarium (34x21x27 cm) and were provided slices of apple as
a food source (Endris et al., 1982). All developmental
stages, including bloodfed l. vexator females were
maintained in a Hotpack incubator (temperature, 27 or 24
+ 1C; relative humidity, 80 + 5%; and 16:8 LD photoperiod).
Collection and Laboratory Maintenance of Lizards
Seeloporus occidental is (western fence lizard) were
hand collected at Ramsey Canyon, three miles north of
Ramsey, Yolo County, California, USA, and examined for the
presence of PIasmodiurn mexicanum. Infected lizards were
sent to the University of Florida for transmission studies.
Seeloporus undulatus undulatus (eastern fence lizard),
collected from Austin Cary Forest, Alachua county, Florida,
USA, were similarly examined for the presence of £_.
f1oridense parasites. Bloodfilms were made from a clipped
toe, air dried, fixed with absolute methyl alcohol, then
stained with Giemsa. Subsequent bloodfilmswere prepared by


-Ill-
treated as above for laboratory colonization. The remaining
mosquitoes were killed, identified, counted, and the
proportion of bloodfed females determined.
Eggs from each feral female (£x. errati cus or Cx.
territans) were transferred to enamel larval rearing pans
(18 X 30 cm) with deionized water. A small amount of larval
food (2% solution of Tetra fish food) was added to the
water of the larval rearing pans one day after oviposition.
Larvae hatched 2-3 days after oviposition and were fed a
mixture of the food daily. Approximately 100-150 larvae
were reared in each pan. Pans were skimmed occasionally
with a paper towel when scum on the water surface appeared.
Pupae were removed from the larval rearing pans and
placed in 250 cc glass culture dishes (12 cm in dameter)
containing water. The and future generation adults were
provided a 10% sucrose solution soaked in cotton. Chicks
served as a blood source for ovarian development. Culture
dishes with circular sections of azalea leaves floating on
the water surface were used for oviposition. Eggs were
normally attached to the periphery of the azalea leaf.
However, beginning with the third generation, it was
determined that the adults would also oviposit on the water
surface or the side of the culture dish. Therefore, the use
of' leaves was discontinued.
The Cx. erraticus colony was maintained in an insectary
at 27C and 80% RH. The insectary is illuminated with a
combination of natural daylight and fluorescent lighting
with a 16:8 LD photoperiod.


Figu re
19.
Midgut (Mg) of Lutzomyia vexator (day five
post-feed) with asynchronous development of
Plasmodium mexicanum oocysts (0).
Figure
20.
Sporoblast (Sb) formation in 5 day old oocyst
on the midgut (Mg) of Lutzomyia vexator. Sand
flies were reared at 27C.
Figure
21.
Plasmodium mexicanum oocyst (0) on the midgut
(Mg) of Lutzomyia vexator with many nearly
mature sporozites (S).
Figure
22.
Sporozoites (S) of Plasmodium mexicanum which
ruptured from the oocysts (0) on the midgut
(Mg) of Lutzomyia vexator.
Figure
23.
Salivary gland (Sg) of Lutzomyia vexator
containing numerous sporozoites (S) of
Plasmodium mexicanum.


-186-
circulating sporozoites of S. occidentalis for more than 60
days. Also, the control group of Sc_. undul atus which were
not forcefed infected flies or mites, did not become
positive for occidental is sporozoites.
Histological studies. Intracellular sporozoites of S^.
occidentals emerge from the circulating blood cells
following ingestion by either Diptera or mites and move in a
manner similar to S^. go 1 vani Because of the low numbers of
sporozoites in the red blood cells, attempts were not made
to determine if the sporozoites penetrated the epithelial
cells of the midgut of the mosquitoes or sand flies. Cu 1 ex
erraticus which fed on infected S£. undulatus were not
infective on day seven post-ingestion, and it is unknown
whether the sporozoites were able to penetrate the
epithelial cells of the mosquito midgut. Elongate
sporozoites, similar to S_. gol vani were teased from the gut
of G. texana (Figure 85). The chromatin was very condensed
and looked similar to S^. go 1 vani sporozoites teased from the
gut of £x. erraticus.
Sporozoites of S^. occi dental is are observed in red
blood cells (RBC's) and WBC's of Sc^. undul atus. However,
the preponderance of parasites in the circulating blood
appears to be in the RBC's (81.5% RBC, 15.2% WBC, and 3.3%
free in circulating blood; n=26 bloodfilms from newly
infected Sc_. undul atus. The average size of intracellular
(RBC and WBC) and extracellular sporozoites measured for
different lizards ranged from 6.9-8.7 X 4.1-7.0um (mean 8.2


-36-
The number of trophozoites, schizonts, gametocytes and
percent of white blood cells infected with P_. mexi canum
during the course of infection for each of the lizards which
died is shown in figures 8 to 13. Both immature (single
nucleated parasites larger than the nucleus of the host
cell, but not displaceing the host cell nucleus) and
"mature" gametocytes (as described by Garnham, 1966) are
included together. The number of trophozoites increased
logarithmically during the course of the infection. In
general, the numbers of schizonts and gametocytes also
increased 1ogrithmical 1y but appeared to show more
variation. This may be partially due to fewer numbers
observed and greater chance of error. Schizogony appeared
to develop synchronously in only one lizard (S-51) and then
only after day 23 following the detection of parasites in
the bloodfilm (Figure 8). From the limited numbers of
bloodfilms, it appears that schizogony occurred at about 3-4
day intervals. In addition, thrombocytes and circulating
white blood cells were also infected. The percentage of
infected white blood cells generally increased as the number
of red blood cells increased (Figures 8 to 13). However,
the low and variable number and rupture of white blood cells
during a bloodfilm preparation increased the potential for
error.


-74-
the capsular material usually appears thickened with
irregular projections toward he sporoblastoid plasma
membrane along the entire margin of the oocyst. The
internal capsular projections remain after the separation of
the sporoblastoid from parts of the oocyst capsule, giving
it the appearance of a scalloped margin (Figures 25 to 27).
The nuclei in early oocysts are large, scattered
throughout the cytoplasm and enclosed in a nuclear envelope
which is marked by occasional nuclear pores (Figure 24).
The nucleoplasm is about the same density as the cytoplasm.
A large distinct dense nucleolus is observed in some nuclei
(Figure 24). Mitochondria appear to be in scattered clumps
throughout the cytoplasm in early oocysts (Figures 24 to
29). They vary from circular to tubular in shape and have
tubular cristae.
Both granular and smooth endoplasmic reticulum (ER) are
found in the cytoplasm (Figures 24 to 26). However, it
appears that smooth or rough ER is more numerous in more
developed oocysts (Figures 24 to 26). Numerous typical
ribosomes are also scattered throughout the cytoplasm,
giving the cytoplasm a granular appearance. Three types of
membrane bound inclusions similar to those observed by
Terzarkis (1971), i. e., circular granules of moderate
density, very dense granules associated with a space of low
density, and lamellar forms usually associated with a
irregular space of low density, are seen in the cytoplasm
(Figures 24 to 27). These inclusions appear to be more
numerous in more developed oocysts.


Figu re
86.
Cresent-shaped sporozoite (S) of Schellackia
occidentalis in the red blood cell of
Sceloporus undulatus (Giemsa).
Figure
87.
Comma-shaped sporozoite (S) of Schellackia
occidentalis in the red blood cell of
Sceloporus undulatus (Giemsa).
Figure
88.
Spherical sporozoite (S) of Schellackia
occidentalis in the red blood cell of Sceloporus
undul atus ("Gi emsa ) .
Figure
89.
Sporozoite (S) of Schellackia occidentalis in
a white blood cell of Sceloporus undulatus
(Giems a).
Figure
90.
Numerous sporozoites (S) of Schellackia
occidentalis in a white blood cell of
Sceloporus undulatus (Giemsa).


-98-
membranes (= linear dense areas) are present in hemoproteids
after vacuolization has commenced, Terzarkis (1971) reported
that thickened membranes are apparent in P_. gal 1 i naceum
preceeding the formation of the subcapsular space. At any
rate, sporozoite development is initiated just prior to or
soon after the initiation of vacuolization of the oocyst.
Microtubules are associated with the pair of convex
membranes of V_. mexi canum and have also been reported in
other species of PIasmodium and Leucocytozoon (Terzarkis,
1971; Boulard et al 1983 ; and Wong and Desser, 1976 )
(Figure 29). These microtubules are not observed in mature
sporozoites.
Three types of inclusion bodies observed in P_.
cy nomol gi (Terzarkis, 1971), are seen in the oocysts of P_.
mexicanum. Terzarkis reported that inclusion bodies
observed in the non-vacuolated oocyst were never observed in
the sporozoites and that the rhoptries (= paired organelle)
arise de nova. However, in our observations moderately
dense inclusion bodies are the first cytoplasmic structures
that are incorporated in P_. mexi canum sporozoites (Figure
29). Moderately dense inclusion bodies are also
incorporated in other PIasmodiurn sp. and Leucocytozoon sp.
(Terzarkis et al., 1967; Boulard et al., 1983; and Sterling
and Degiusti, 1974). Wong and Desser (1976) suggest that
the inclusion bodies are the precursors of the anterior
organelles (rhoptries) of L^. Dubreui 1 i and are also the
precursors of the anterior organelles of PIasmodiurn. The


-10-
The common occurrence of EE stages, the similarities in
the types of non-erythrocytic cells invaded, and
morphological similarities between avian and saurian
PIasmodiurn suggest that malaria parasites in lizards and
birds evolved from a common ancestor (Thompson and Huff,
1944a; Mattingly, 1965). So far, all avian and saurian
malarias studied (except P_. mexi canum) are restricted to the
hemopoietic or reticuloendothelial tissues. PIasmodiurn
mexicanum appears to be the most primitive of the PIasmodiurn
species since its EE stages are not restricted to one type
of tissue (Huff, 1945). According to Bray (1957, 1963), the
genus PIasmodiurn is polyphyletic with the bird and reptilian
plasmodia originating from a common ancestral stock and
mammalian plasmodia originating from a different stock
(Mattingly, 1965). This is based mostly on the types of EE
tissues invaded by the different malarial parasites.
The elucidation of vectors of saurian malaria has been
difficult. Sporogony, to the development of mature
sporozoites, has been observed for only two species of
saurian malaria (Ayala and Lee, 1970; Ayala, 1971; Petit et
al., 1983). Giemsa stained sporozoites of both IP. mexi canum
and P_. agamae appear to be similar and are very short, 5-7urn
(urn = micron) and 4-6um, respectively, when compared to
those of most other malarias. Only the fine structure of
the development of oocysts and of mature sporozoites of P_.
agamae in an unnatural host, C u1 ic oid e s nubeculosus has
been described (Boulard et al., 1983). In general, oocyst


Figu re
74.
Sporozoites (S) of Schellackia golvani in
white blood cells of Anolis carolinensis
(Giemsa).
Figure
75.
Sporozoite (S) of Schellackia govani with two
chromatin bands in a white blood cell of Anolis
carolinensis (Giemsa).
Figure
76.
Sporozoite (S) of Schellackia golvani teased
from the midgut of Culex erraticus and stained
with Giemsa.
Figure
77.
Unstained sporozoite (S) of Schellackia
golvani teased from the midgut of Culex
erraticus (Nomarski differential interference
contrast).
Figure
78.
Cross section of the midgut of Culex erraticus
with a sporozoite (S) of Schellackia golvani
in a parasitophorus vacuole (Pv) in the midgut
epithelium (Giemsa-collophonium).
Figure
79.
Cross section of the midgut of Lutzomyia
vexator with a sporozoite (S) of Schellackia
golvani in a parasitophorus vacuole (Pv) in the
midgut epithelium (modified methylene blue and
fu sc hin ) .
Figure
80.
Unstained sporozoites (S) of Schellackia
golvani in a parasitophorus vacuole (Pv) in the
midgut epithelium of Culex erraticus seven days
after feeding (Nomarski differential
interference contrast).


-165-


91

,2um


89


-149-
Salivary glands were examined after 10-13 days following the
initial bloodmeal and the sporozoite rate determined (+1,
1-10; + 2, 11-100; +3,101-1,000; +4, >1,000 sporozoites).
Ultrastructure of Extrinsic Stages of Plasmodium floridense.
For ultrastructural examination, midguts and salivary
glands of infected C_x. errati cus were fixed in 2%
glutaraldehyde in 0.1M sodium cacodylate buffer at pH 7.2
for 1 hr at room temperature. In addition, a portion of the
thorax and head and last three segments of the abdomen were
severed from living mosquitoes and fixed in Karnovsky's
fixative at pH 7.2 for 1 hr at room temperature (Karnovsky,
1965). Both were post-fixed with 1% osmium tetroxide in
0.1M sodium cacodylate buffer, pH 7.2, for 1 hr at room
temperature, dehydrated through a graded ethanol series and
acetone, and then embedded in Spurr's resin (Spurr, 1969).
Sections were cut on an LKB Ultratome III with a Diatome
diamond knife, floated on water, picked up on formvar-coated
grids, and post-stained with aqueous 1% uranyl acetate
followed by Reynold's lead citrate. Material was examined
and micrographs taken with a Jeol 10OCX electron microscope.


-211-
Scorza, J. V. 1971a. Electron microscope study of the
blood stages of PIasmodiurn tropiduri a lizard malaria
parasite. Parasitology 63:1-20.
Scorza, J. V. 1971b. Asexual and sexual stages of a
malaria parasite in the thrombocytes of Tropidurus
torquatus (Iguanidae) infected with Plasmodium
tropiduri J. Protozool 18:403-410.
Shortt, H. E. and K. P. Menon. 1940. Experimental
production of monkey and avian malaria by an unusual
route of infection. J. Malaria Inst. India 3:195-198.
Shortt, H. E., and P. C. C. Garnham. 1948. The
pre-erythrocytic development of Plasmodium cynomo!gi
and Plasmodium vivax. Trans. R. Soc. Trop. Med. Hyg.
41:785-795.
Shute, P. G., G. Lupascu, P. Branzei M. Maryon, P.
Constantinescu, L. J. Bruce-Chwatt, C. C. Draper, R.
Ki11ick-Kendrick and P. C. C. Garnham. 1976 A
strain of PIasmodiurn vivax characterized by prolonged
incubation: The effects of numbers of sporozoites on
the length of the prepatent period. Trans. R. Soc.
Trop. Med. Hyg. 70:474-481.
Sinden, R. E. 1975. The sporogonic cycle of PIasmodiurn
y o e1 i i nig e rie n sis : A scanning electron microscope
study. Protistologica 11:31-39.
Sinden, R. E., and P. C. C. Garnham. 1973. A comparative
study on the ultrastructure of PIasmodiurn sporozoites
within the oocyst and salivary glands, with particular
reference to the incidence of the micropore. Trans. R.
Soc. Trop. Med. Hyg. 67:631-637.
Sinden, R. E., and J. Moore. 1974. Fine structure of the
sporozoite of Schellackia occidentalis J. Parasitol.
60:666-673.
Spurr, A. R. 1969. A low-viscosity epoxy resin embedding
medium for electron microscopy. J. Ultrastruct. Res.
26:31-43.
Sterling, C. R., M. Aikawa, and J. P. Vanderberg. 1973.
The passage of PIasmodiurn berghei sporozoites through
the salivary glands of Anopheles Stephensi: An electron
microscope study. J. Parasitol 59:593-605.
Sterling, C. R., and D. L. DeGiusti. 1974. Fine structure
of differentiating oocysts and mature sporozoites of
Haemoproteus metchnikovi in its intermediate host
Chrysops cajlidus. J. Protozool. 21:276-283.


-81-
sporobl astoi d, only spherical forms are observed migrating
into the developing sporozoites. The cytostome, anterior to
the nucleus, is observed in developing sporozoites which are
not completely formed (Figure 32). Formation of the
sporozoite is complete when it is pinched off from the
sporoblast.
Sporozoites. Sporozoite formation is to some degree
synchronous within a given oocyst but not between oocysts
(Figure 19). Several sporoblasts with protruding
sporozoites are observed in oocysts which are prematurely
ruptured (Ayala, 1971). Living and Giemsa-stained
sporozoites of P_. mexi canum are stout, cresent shaped, and
measure 6.3-7.3um (urn = micron) (mean 6.6um) in length and
1.3-1.7um (mean 1.5um) in diameter at their greatest width
(Figure 22). The mature sporozoites migrate through the
hemocoel to the anterior of the sand fly until they reach
the salivary glands. Sporozoites in the hemocoel have
distinct micronemes, elongated rhoptries, and inclusion
bodies. The salivary glands of L^. vexator are hollow fluid
filled sacs which are 1 cell thick and roughly spherical in
shape (Figure 23). The anterior end of the sporozoite comes
into contact with the salivary gland cell and causes the
cell membrane to become invaginated (Figures 33 and 34).
The sporozoite pushes through the salivary gland cell
becoming constricted at the membrane-parasite junction, and
carrying with it a portion of the salivary gland cell plasma
membrane which forms a parasitophorous-1ike vacuole inside


TABLE 3
The effects of temperature on sporogony of
Plasmodium mexicanum in Lutzomyia vexator .
Day
post feed
Mean
oocyst
size5
(microns)
24C
Range,
(mean)D
SE
Mean
oocyst
si ze5
(microns)
27C
Range,
(mean)
SE
2.0
11.6
11.6
..
2.5


15.7
15.1-16.2
0.31
3.0


16.2
15.3-18.4
0.56
3.5
- -

18.8
17.9-19.7
0.51
4.0
15.2
14.3-16.0
0.29
23.0
20.2-26.4
0.65
4.5
- -

24.6
23.8-25.4
0.33
5.0
18.8
17.8-20.0
0.28
30.4
29.0-31.6
0.78
5.5


31.4
30.9-31.8
0.46
6.0
24.5
22.4-26.5
0.61
31.8
30.6-34.7
0.60
6.5
- -

33.5
33.5
7.0
29.1
27.2-30.7
0.60
30.6
27.5-34.1
1.93
8.0
29.1
26.6-32.4
0.87
8.5
34.2
32.4-36.0
1.8
9.0
31.8
25.7-39.2
1.1
P Overall mean oocyst size by day.
Mean oocyst size for sand flies by day.


Figure 6. Percent of infected red blood cells during the course of
PIasmodium mexicanum infection for six Seeloporus undulatus
infected by bite of Lutzomyia vexator.


-100-
Plasmodium sporozoites, but are found throughout sporozoites
of Leucocytozoon Haemoproteus and Schel1ackia (Moore and
Sinden, 1974; Wong and Desser, 1976; Klei, 1972).
The cytostome (= micropyle) is observed in developing
sporozoites of £. mexicanum in which the nucleus is
migrating into it from the sporoblastoid. The cytostome is
|
rarely observed in IP. berghei (Vanderberg et aj ., 1967 ) and
is absent in PIasmodiurn vinckei Rodhain (Sinden and Garnham,
1973) while being a common feature in other PIasmodiurn,
including P_. agamae. A cytostome is also present in
sporozoites of Haemoproteus col umbae (Klei, 1972) and jl.
metchnikovia (Sterling and DeGuisti, 1974), but has not been
seen in three species of Leucocytozoon. The exact nature
and fuction of the cytostome is unknown, but is believed to
have the same function as the cytostome seen in the
merozoites (Aikawa et al., 1978). However, host cellular
material in the salivary gland cell or in other tissue cells
was not observed being ingested by the sporozoite.
The number and length of the subpel1icular microtubules
of P_. mexi canum is similar to that of other mammalian and
avian malarias. However, the arrangement of pellicular
microtubules of P_. mexi canum is different from that observed
for avian, mammalian, and a saurian PI asmodi urn, P_. agamae.
Although the gross histological examination of P_. mexi canum
is similar to P^. agamae, the number of microtubules (usually
14) are small when compared to P_. agamae (26) (Figure 39).
The arrangement of the microtubules is similar in


Figure 10. Number of PIasmodium mexicanum trophozoites,
schizonts, gametocytes and percent of
infected white blood cells per 10,000 red
blood cells during the course of the
infection of Seeloporus undulatus (S-8)
infected by bite of Lutzomyia vexator.


Figure 18. Regression of PIasmodium mexicanum mean oocyst
size (and 95% confidence limits) in Lutzomyia
vexator and day post-feed for days 2 through 9.


-178-
Table 10--continued
a
b
c
d
e
Day force fed to noninfected lizard subsequent to bloodfeeding on
an infected lizard.
One group maintained at room temperature (18-24)
Since bloodfilms were not taken on a daily basis, patent
infections could have been as much as 1-3 days previous to
positive slide. However, bloodfilms from lizards maintained at
32C did not demonstrate any parasites prior to day eight,
post-ingestion .
Reason for apparent prolonged prepatent period unknown.
All seven of the S£. undul atus fed infected Cx.. er rati cus on day one
post-feed became patent by day 12 post-ingestion, while none of the
Sc. undulatus fed infected mosquitoes on day seven post feed became
patent.


-150-
Results
Sporogony of Plasmodium floridense
Oocysts of J?. f 1 ori dense were observed on the midgut of
Cx. erraticus 3->20 days after a bloodmeal. Early oocysts
were difficult to discern and were easily overlooked.
Development among oocysts was very asynchronous (Figure 55),
with some having fully developed sporozoites nine days after
feeding. Translucent lipid-like droplets were apparent in
undifferentiated oocysts (Figure 44), but were less
frequently seen in oocysts which had nearly mature
sporozoites. The range and mean oocyst size for different
mosquitoes maintained at 25C on 10% sugar + multivitamin is
shown in Table 9. While the mean oocyst size of mosquitoes
not supplemented with multivitamin was similar, sporozoite
development appeared to be delayed by 2-3 days.
Pressure from the coverslip often ruptured nearly
mature oocysts, releasing free sporozoites and sporozoites
still attached to the sporoblast (Figure 56). Sporozoites,
measuring 16-19 X l-2um (urn = micron), were present in the
salivary glands 11 days after feeding in two of three
mosquitoes which had >20 oocysts (Figures 57 and 58). The
time at which sporozoites were detected in the salivary
glands for mosquitoes with <10 oocysts was quite variable.
Some had sporozoites in the salivary glands as early as 13
days after feeding; others did not have sporozoites in the


-21-
blood cells per 10,000 red blood cells was also counted. A
sufficient number of red blood cells was counted to keep the
probable error within 10% according to the method formulated
by Gingrich ( 1932).
Histology of Infected Lizards
Tissue impressions of various organs made subsequent to
death for most of the lizards infected with £_. mexi canum
were fixed with methanol and stained with Giemsa. In
addition, tissues from one lizard (S-51) were fixed in
Carnoy's fluid, dehydrated, embedded in paraffin and
sectioned at 5-6um (urn = micron) on a rotary microtome.
Thin sections were stained with hematoxylin-eosin or
Giemsa-colophoniurn (Bray and Garnham, 1962).
Results
Plasmodium mexicanum Transmission Studies
Females of L. vexator readily feed on lizards in the
laboratory (Figure 1). Males are also attracted to lizards
and mating frequently occurs during blood feeding.
Exflagellation can be observed in a bloodmeal by removing
the midgut contents within 30 minutes after a female sand
fly has completed blood feeding. The length of time during
which exflagellation occurs was not determined.


-168-
The obvious differences between the saurian malarias in
which sporogony has been demonstrated are observed in the
gross morphology of the sporozoites. Those of P_. mexi canum
and P_. agamae are short and stout (4-7 X l-2um), in contrast
to _P. flor i dense sporozoites which are very long and sinuous
(16-19 X l-2um). Known vectors of IP. mexi canum are
phlebotomine sand flies and the vectors of P_. agamae are
also suspected to be a non-mosquito. Both have similar
subpel 1 i cul ar microtubular arrangements, which are different
from that of other non-saurian malaras examined. PIasmodiurn
floridense sporozoites resemble those of other
mosquito-transmitted malarias. The similarities of the
sporozoites of P_. agamae with P_. mexi canum and the
differences seen between P_. agamae and P_. f 1 ori dense provide
further evidence that P_. agamae may not be transmitted by a
mosquito. Sporozoites of P_. f 1 or i dense in the salivary
glands are intracellular, similar to other mosquito
transmitted malarias, but unlike P_. mexi canum which is
extracel1ualr in the lumen of the salivary gland. This
relationship may be more related to the morphological
structure of the salivary gland since the gland of L_.
vexator consists of a hollow fluid filled sac.
Ultrastructure of Extrinsic Stages of Plasmodium floridense
The u1trastructure of the oocyst, formation of the
sporoblast, and sporozoites of P_. f 1 ori dense appears to


Figure 91. Section of intestine of Seeloporus undulatus
with a schizont (Sc) of Schel1ackia
occidentalis in the intestinal epithelium (E)
(Giemsa-col1ophonium).
Figure 92. Section of intestine of Seeloporus undulatus
with an oocyst (0) (?) of Schel1ac kia
occidentals in the lamina propria (TTpr)
(Giemsa-collophonium).


-99-
degen e rat i on of the moderately dense anterior inclusion
bodies during the development of the rhoptries and
micronemes and the disappearance of the moderately dense
inclusion bodies in mature sporozoites (salivary glands)
support such a hypothesis. Although dense granular
inclusion bodies associated with a space of low density and
an occasional lamellar inclusion body are observed in
sporozoites, it is unclear whether they were incorporated
into the sporozoite from the sporoblast cytoplasm during
formation since none were observed migrating into the
developing sporozoites. Crystaloid material observed in
Haemoproteus and Leucocytozoon and "virus-like" crystaloid
material seen in some PIasmodiurn sp. (Terzarkis, 1969,1972;
Garnham et al., 1962) were not observed in the oocyst or
sporozoites of _P. mexicanum.
Micronemes and sac-like rhoptries are common features
of PIasmodiurn. Usually the rhoptries are relatively long,
tubular and have an expanded distal end (sac). The
rhoptries are present early in the developing sporozoites,
but become more elongated and more pronounced in the mature
sporozoite. The rhoptries are also present in other
hemoproteids, but in contrast, are rarely observed in some
(J_. tawaki and j_. dubreui 1 i ) until they enter the salivary
glands of the vector (Wong and Desser, 1976 and Desser and
Allison, 1979). A review of the micronemes of several
PIasmodiurn species is contained in Garnham et al., 1963.
Micronemes are only observed anterior of the nucleus in


-139-
f1 oridense. Although several species of mosqitoes will feed
on lizards in the laboratory, only a few were attracted to
and fed on lizards in lizard-baited traps in the field. The
design of the lizard-bait trap allowed mosquitoes to feed on
the lizard when captured in the trap and provided further
evidence that those mosquitoes collected in the trap
j
normally feed on lizards in nature. Only two species of
mosquitoes, £x. errati cus and Cjc. terri tans, were collected
in the bait trap with relative high frequency. While Cx.
erraticus has been reported to feed primarily on birds and
mammals (Edman, 1979), both species readily fed on lizards
in the traps and in the laboratory. Although Jordan (1964)
collected many of the same mosquito species at the
Fargo-Okefenokee Swamp where P_. f 1 o ri dense is relatively
abundant, Cj<. errati cus was not reported to be collected.
Culex erraticus has a wide geographic distribution (from
Michigan, USA to Bolivia) and probably occurs in the
Fargo-Okefenoke Swamp. Culex erraticus may have been
represented by one or more of the three unidentified Culex
sp. collected by Jordan, especially since one of them
developed 70 oocysts after feeding on an anole infected with
P_. flor i dense.
Early investigations by Huff (1941) and extensive
studies by Jordan (1964) indicated that P_. f 1 ori dense is
transmitted to lizards in the late summer early fall, with
new infections becoming more abundant in August, peaking in
November, and sharply declining in December. Although data


-11-
development of P^. agamae is similar to other PI asmodi um.
The sporozoites of P_. agamae is also similar in structure to
other PIasmodium species which have been examined. However,
the organization and number of pellicular microtubules are
different (Boulard et al., 1983).
The fine structure of blood forms of P_. f 1 ori dense
(Aikawa and Jordon, 1968), IP. t r o p i d u r i (Scorza, 1971a), and
P_. mexi canum (Moore and Sinden, 1974 ) has been studied. In
general, their morphology is similar to the avian malaria
parasites examined.
Thompson (1946a) postulated that lizards might be used
as a malaria model in chemotherapeutic research since the
effects of anti-malarial drugs on malaria parasites
maintained at different temperatures in poikilothermic
lizards might provide valuable information. Thompson
(1946a,b) studied the effects of atabrine on P_. floridense
and quinine on P_. f 1 ori dense and P_. mexi canum. Both drugs
were effective in reducing the parasitemias in lizards with
P_. f 1 ori dense, but quinine did not appreciably lower the
parasitemia of lizards with P_. mexi canum. This was
apparently due to the inability of the drug to destroy the
EE stages. Reptilian malaria never became popular for
study, largely because of the more demanding care required
by the experimental animals, the difficulties in
establishing cyclic transmission, and the many biological
characteristics that separate them from mammalian species
(Wernsdorfor, 1980). However, with advances in tissue


-116-
propensity of feral mosquitoes on lizards in the field and
laboratory is shown in Table 5.
Bait traps were operated from June through October
during 1983 and in May through October during 1984, because
Jordan (1964) indicated that P_. f1 oridense transmission
occurred in late summer early fall. During both trapping
periods, mosquitoes were the only bloodfeeding arthropods
collected in the lizard-baited traps. Excluding Cx.
territans, Cx. erraticus accounted for more than 80% of the
mosquitoes collected in the bait traps. Although Cx.
territans were not collected during 1983, they accounted for
more than 65% of the mosquitoes collected in 1984. More
than 30% of the C_x. terri tans and C_x. errati cus bloodfed on
lizards in the bait trap. The other species (Cx.
salinarius/nigripalpus and C£. perturbans ) were infrequently
collected but readily fed on the lizards in the
lizard-baited traps (Table 5).
Two species of Co rethrel 1 a C^. b rak 1 ey i (Coquillett)
and C_. wi rthi Stone, and more than 14 species (seven genera)
of mosquitoes were collected in the CDC light traps during
1984 (Table 5). Culex territans, the most frequently
collected mosquito in the lizard-baited trap (1984), was
rarely collected in the light trap. In the laboratory, more
than 70% and nearly 50% of the wild caught C_x. errati cus and
Cx. territans, respectively, fed on lizards (Table 5; Figure
43). Only 40% of the C><. salinarius/nigri palpus and
approximately 20% of C^_. perturbans and Aedes sp. fed on


-212-
Stratman-Thomas, W. K. 1940. the influence of temperature
on PIasmodiurn vivax. Am. J. Trop. Med. 20:7 03-1 5 .
Telford, S. R., Jr. 1970a. Exoerythrocytic gametocytes of
saurian malaria. Q. J. Florida Acad. Sci. 33:77-79.
Telford, S. R., Jr. 1970b. Comments on the vector
relationships of saurian malaria. J. Parasitol.
56:340.
Telford, S. R., Or. 1978. A hemoparasite survey of Florida
lizards. J. Parasitol. 64:1126-1127.
Telford, S. R., Jr. 1982. PIasmodiurn 1 ionatum sp. n., a
parasite of the flying gecko, Ptychozoon 1ionatum, in
Thailand. J. Parasitol. 68:1154-1157.
Telford, S. R., Jr. 1983. PIasmodiurn saurocaudatum sp. n.,
a parasite of Mabuya multifasciata in southeast asi a.
J. Parasitol. 69:1150-1155.
Telford, S. R., Jr. 1984a. Studies on african saurian
malarias: Three PIasmodiurn species from gekkonid
hosts. J. Paras it. 70:343-354.
Telford, S. R., Jr. 1984b. Haemoparasites of reptiles.
In: Huff, G. L., F. L. Frye, and E. R. Jacobson ed.,
Diseases of Amphibians and Reptiles. pp. 385-517. New
York: Plenum Publishing Corp.
Terzakis, J. A. 1969. A protozoan virus. Mil. Med.
(Suppl 10) 134:916-921 .
Terzakis, J. A. 1971. Transformation of the PIasmodiurn
cynomolgi oocyst. J. Protozool 18:62-7"3T
Terzakis, J. A. 1972. Virus-like particles and sporozoite
budding. Proc. Helm. Soc. Wash. (Special Issue:
Basic Research in Malaria) 39:129-137.
Terzakis, J. A., H. Sprinz, and R. A. Ward. 1967. The
transformation of the Plasmodium gal 1inaceum oocyst in
Aedes aegypti mosquitoes. J. Cell Biol. 57:311-326.
Thompson, P. E. 1944. Changes associated with acquired
immunity during initial infections in saurian malaria.
J. Inf. Dis. 74:138-150.
Thompson, P. E. 1946a. Effects of quinine on saurian
malarial parasites. J. Inf. Dis. 78:160-166.
Thompson, P. E. 1946b. The effects of atabrine on the
saurian malarial parasite, Plasmodium floridense. J.
Inf. Dis 79:282-288.


-184-
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-3-
of lizard malaria" (p. 13). In fact, no vectors, except for
two species of phiebotomine sand flies (Ayala and Lee,
1970), have been incriminated as vectors for any of the
species of PIasmodiurn infecting lizards and only limited
attempts have been made to incriminate other vectors of
saurian malaria. Previous suggestions that acarie
ectoparasites transmit saurian malaria in Mexico (Pelaez et
al., 1948) and in Africa (Pienaar, 1962) proved to be false
(Russell et al., 1963; Ayala, 1977). Also, sporogonic
development was not observed after triatomid bugs were fed
on lizards infected with Plasmodium parasites (Ayala, 1977).
Although hematophagus Diptera are the most likely
vectors of saurian malaria (Ayala, 1977), attempts to
incriminate these insects have usually met with failure.
The earliest recorded attempt to determine the vector of
saurian malaria was by Huff (1941a) who fed both Cu 1 ex
pipiens L. and Aedes aegypti (L.) mosquitoes on Seeloporus
undulatus Latreille (eastern fence lizard) infected with
PIasmodiurn sp. The PIasmodiurn species, although not
identified, was isolated from Sc_. undul atus where P_.
f1oridense Thompson and Huff (1944b) is endemic. Culex
pi piens females were negative; however, one of an unknown
number of Ae^. aegypti had one nearly mature oocyst on the
midgut. Attempts to infect Ae^. aegypti (61) by feeding them
on S£. ol i vaceaous Smith infected with numerous P_. mexi canum
Thompson and Huff (1944b) gametocytes were unsuccessful
(Thompson and Huff, 1944b). Baker (1961) examined several


DAY POSTEXPOSURE
LOG NUMBER OF PARASITES/10,000 RBC
in
in o
O o
in
O
o
o
ro
cn
% INFECTED WBC/10,000 RBC
to
o
,
CO
o
4
o
Ol
o
<*>+
II II II II
-p.
I
cn
o
10000


Figure 1
Figure 2
Figure 3
Figure 4
Bloodfed female Lutzomyi a vexator resting on
Seeloporus undulatus upon which it had previously
fed on.
Midgut of Lutzomyia vexator with oocysts (0) of
PIasmodium mexicanunu (Day 5 post-feed).
Plasmodium mexicanum sporozoites (S) from
ruptured oocysts (0) on the midgut (Mg) of
Lutzomyia vexator. Day six post-feed on infected
Seel opoTITs occi dental i s and maintained at 27C.
Plasmodium mexicanum sporozoites (S) in the
salivary gland (Sg) of Lutzomyia vexator
(Nomarski interference contrast) .


-140-
are limited, most new infections in the present study were
also observed in October and November. Based on this
evidence and the bait trap data, it was concluded that Cx.
erraticus was the most likely vector since its seasonal
abundance corresponded well with the period of suspected
transmission of P_. flor i dense. This mosquito is relatively
long-lived in the laboratory and is relatively abundant in
the field when natural transmission is suspected to occur.
Plasmodium floridense Transmission Studies
While field data suggest that C_x. erraticus may be a
natural vector of P_. f 1 ori dense, other arthropods can not be
ruled out. Based on new evidence by Petit et al ( 1984),
members of the Ceratopogonidae may be involved in the
transmission of a saurian malaria, P_. agamae. In addition,
members of the Psychodidae, L^. vexator and probably l.
stewarti are natural vectors of P_. mexicanum (Table 8).
Present studies and those of Young and Perkins (1984)
indicate that L. vexator readily develop oocysts of _P.
f1oridense. Approximately 50% of more than 600 sand flies
which fed on infected lizards developed oocysts. Sporogonic
development of P_. f 1 o ri dense is relatively long, 11-14 days
for mature sporozoites, and most of the lab-reared sand
flies (>90%) died within 14 days of feeding. Also, only a
few (<10) sporozoites (probably as a result of mechanical
damage to oocysts during dissection) were seen in the


CHAPTER 2
EXPERIMENTAL TRANSMISSION OF Plasmodium mexicanum BY BITES
OF INFECTED Lutzomyia vexator (DIPTERA:PSYCHOD I DAE )
Introduction
Saurian malaria research has received increasing
attention (primarily taxonomic) in the past few years, but
the natural vectors remain unknown. Fifty-nine species of
saurian PIasmodiurn have been described (36, Americas; 11
Australia, Asia and Oceania; 12, Africa), three of which
occur north of Mexico (Ayala, 1978; Telford, 1982; Telford,
1983; Telford, 1984a; Telford, 1984b; Garnham and Telford,
1984). Ayala and Lee ( 1970), Ayala (1971) and Petit et al .
(1983) are the only authors to describe the extrinsic cycle
of lizard malaria (PIasmodiurn mexicanum and PIasmodiurn
agamae, respectively) developing beyond the early oocyst
stage. They demonstrated that P_. mexi canum developed in
phlebotomine sand flies while _P. agamae developed in
C u1 ic o id e s nubecu1osus not in mosquitoes as previously
suspected. Ayala (1971) further demonstrated sporogony and
experimental transmission of P_. mexicanum by i nt raperi toneal
inoculation of sporozoites from wild caught Lutzomyia
vexator females that had earlier fed on infected Seeloporus
occidentals lizards. He did not demonstrate transmission
by bite of infected flies. Ayala (1971) further suggested
-16-


LOG NUMBER OF PARASITES/10,000 RBC
10000
5000
*= TROPHOZOITES
i
CT>
DAY POSTEXPOSURE


-60-
Histology of Infected Lizards
These studies indicate that P_. mexi canum parasites
primarily invade erythrocytes and lymphocytes of Sc.
undulatus. Thrombocytes are less frequently invaded. No
attempt was made to distinguish between granulocytes and
macrophages. Determining the percentage of lymphocytes
infected was often difficult since occasionally many of the
cells ruptured in bloodfilm preparation, especially during
the latter part of the acute phase. These results agree
with those of Thompson and Huff (1944a) who observed that
93% of the circulating cells infected with P_. mexi canum are
in erythrocytes, with a small percentage in lymphocytes and
thrombocytes .
Tissue impressions of the spleen and bone marrow
revealed numerous asexual forms of P_. mexi canum in
lymphocytes (Figure 15 and 16). Although exoerythrocytic
(EE) forms are often seen in lymphocytes (Jordan, 1970;
Thompson and Huff, 1944a), they are rarely observed in other
tissues in Sc_. undul atus. However, in unnatural hosts such
as Ph rynosoma cornutum and C_. col 1 ari s PI asmodi urn mexi canum
EE forms are frequently observed in fixed connective tissue
while occurring less frequently in the circulating cells
(Thompson and Huff, 1944a). When studying fixed tissues,
Jordan (1970) recovered schizonts and segmenters from
impressions of internal organs, especially endothelial cells
of the brain capillaries. In the present study, impressions


Figu re
Figu re
42. Photograph of lizard baited trap used to
attract and capture biting Diptera.
43. Cu 1 ex erraticus bloodfeeding on a restrained
Anolis carolinensis.


-146-
mexicanum (Chapter 2). However, the prepatency and course
of infection also appears to be affected by host behavior in
its temperature regulation.
The initial course of infection of P_. f 1 ori dense is
similar to that of P_. mexican urn (Figures 12 and 52-54).
However, unlike P_. mexi canum, experimental infections of P.
f1oridense appear to rarely kill the lizard. Experimentally
infected lizards, AA-59 and A-85, continue to appear healthy
and have survived in the laboratory for one and two years,
respectively, following experimental infections.
Parasitemias have fluctuated throughout the course of the
infections with some bloodfilms appearing negative or with
only a few parasites. Both lizards have been used in
experimental P_. f 1 ori dense transmission studies and have
been good donor lizards for the demonstration of sporogonic
development in Cx. erraticus.


Figure 50. P1asmodium flor i dense oocyst (0) on the midgut
(Mg ) of Cu 1 ex errati cus on day 10 post-feed.
Melanization (arrow) Ts beginning to occur
along the oocyst capsule and spread inward.
Figure 51. Bloodfilm of Anolis carolinensis at the first
peak of a PIasmodium floridense infection.
Trophozoites (T) and schizonts (Sc) are seen in
the red blood cells.


-9-
by Thompson and Huff in the 1940's when similar forms of
avian plasmodia were also being studied. These EE studies
were accomplished by examination of blood smears or other
tissues of lizards having natural or experimental
infections. Thompson and Huff (1944a) inoculated blood from
Sc. erythrurus (=ferrariperzi ) (Schinz) containing P_.
mexicanum parasites into five different species of lizards
in three genera, Seeloporus Phrynosoma and Crotaphytus .
Although there was much variation in the intensity of the
infections among conspecific lizards, there were significant
differences in the degree of parasitemia and the
distribution of the parasites in different types of cells of
the five species of experimental hosts. The percentage of
P_. mexi canum parasites found in erythrocytes ranged from 95%
(Sc undulatus and S£. olivaceous ), <20% [Ph rynosoma
cornutum (Harlan)], <10% (Crotaphytus co 11 aris Say) to <1%
(Ph rynosoma asi o Cope). The majority of P_. mexi canum
parasites in PJh. cornutum and £. col 1 ari s were observed in
the lymphocytes (45% and 35%, respectively) while for Ph.
as io, the majority of parasites were observed in the
thrombocytes (65%). PIasmodiurn mexicanum EE stages, unlike
other species of saurian and avian malaria, are found in
both the hemopoietic and reticuloendothelial tissues and
therefore represent both V_. el ongatum and Plasmodium
gal 1inaceum Brumpt types of EE cycles of avian malarias
(Thompson and Huff, 1944a).


-144-
days), the additional bloodmeals may provide nutritional
requirement necessary for maturation.
While sporogonic development appeared to be normal in
most £x. erraticus maintained at 25C, melanization of some
oocysts occurred in a few mosquitoes, usually beginning 10
or more days following an infective bloodmeal. The
influence of temperature on melanization of oocysts has not
been established, but when mosquitoes were maintained at
32C, nearly all oocysts were melanized or were becoming
melanized eight days after feeding. Melanization of oocysts
and sporozoites (?) also occurred in slowly developing
oocysts in heavily infected mosquitoes (>30 oocysts) which
were supplemented with multivitamins. However, up until day
11, all oocysts appeared normal, no melanization could be
detected, and at 22 days after feeding, all mosquitoes
dissected from this lot had sporozoites in the salivary
glands. It is uncertain whether the temperature at which
the mosquitoes were maintained or whether other factors are
involved in this defensive reaction to parasite growth.
Rearing £x. erraticus at temperatures below 25C may produce
more infective/vi a b1e sporozoites than in the present study.
More intensive studies on the temperature and nutritional
requirements on the development of P. f1oridense in Cx.
erraticus and an extensive search for other potential
vectors of P. f1 oridense are indicated. Certainly, it will
be important to find naturally infected mosquitoes.


-65-
mexicanum in L. vexator are summarized in Table 3. The mean
oocyst size, regressed against the days post-feed for sand
flies maintained at 24C (R2=.85) and 27C (R2=.91), is
shown in figure 18. Sand flies maintained at 19C and
dissected at days four and five after feeding did not show
any oocysts. The maximum and minimum temperatures under
which oocyst development ceases were not determined.
Figures 19 to 23 illustrate oocyst and sporozoite
development from days 2-7 in sand flies maintained at 27C.
By day two post-feed, P_. mexi canum oocysts on the midgut of
sand flies maintained at 27C are spherical, relatively
uniform in size [range 10.8 to 13.5um (urn = micron) in
diameter] and undifferentiated. Differentiation, i. e.,
sporoblastoid formation and the formation of sporozoites for
sand flies maintained at 27C is not evident until day five
after feeding (Figures 19 and 20). Many oocysts contain
nearly developed sporozoites by day 6.0 (Figure 21).
Shortly after day 6.0, some oocysts have released
sporozoites into the hemocoel while others are easily
ruptured by pressure from the coverslip (Figure 22).
Sporozites are observed in the salivary glands by day 6.5
post-feed (Figure 23). Sporogonic development culminating
in the invasion of the salivary glands of sand flies
maintained at 24C is much slower, not being observed in the
salivary glands until day 8.5-9.0 post-feed. The
infectivity of sporozoites from sand flies maintained at
24C was not determined. However, sand flies maintained at


-13-
in connective tissue and/or reticulendothelial systems.
Sporozoites are released from the oocyst where they invade
erythrocytes or lymphocytes or both. The host response may
determine what type of cell is invaded by the sporozoite.
For example, S^. bol i vari Reichenow (1919) invades the
erythrocytes of one lizard, Acanthodactylus vulgaris
(Schinz), while invading the lymphocytes of another,
Ps ammod romu s h i s p a n i c u s Fitzinger (Manwell, 1977 ).
(=Haemogregarina) weinbergi, parasites was made by Leger and
Mouzels (1917) from South American lizards. Reichenow
(1919) first described the genus Schel1ackia and named a new
species, Schellackia bol i vari from Spain. Reichenow's
description included the complete life cycle of the parasite
in the lizard and incriminated a mite, Lyponyssus sau rarum,
as the natural vector. Later, Bonnoris and Ball (1955)
described Schellackia occidentals from Seel oporus
occidental is in California, USA, and also incriminated a
mite, Gekobiel 1 a texana (Banks), as the natural vector.
There are four additional species of Schel1ackia in lizards
in the western hemisphere: S_. brygooi Landau ( 1973), S_.
1 andaue Lainson, Shaw, and Ward ( 1976 ), S.. gol van i Rogier
and Landau ( 197 5 ), and S_. (=Lainsonia) i guanae (Landau,
1973), and one in toads, S_. bal 1 i .
Experimental transmission of S_. brygooi and S^. 1 andaue
has been accomplished by ingestion of mosquitoes (Cu 1 ex
p i p i e n s p i p i e n s and Cx.. £. f ati gans ) which bloodfed on


LOG NUMBER OF PARASITES/10.000 RBC
-40-
DAY POSTEXPOSURE


-102-
long (Chapter 4). The size differences of P_. mexi canum and
P_. agamae may be an adaptation to a non-mosquito vector
since, at least, the sporozoites are extracellular in P_.
mexi canum. The vector of P_. f 1 ori dense is a culicine
mosquito and the sporozites are intracellular in the
salivary glands (Chapter 5).
The salivary glands of l. vexator, unlike mosquitoes,
consist of a one cell thick fluid filled spherical sac. In
mosquitoes, sporozoites enter the secretory cells of the
salivary glands and remain intracellular until they enter
the narrow salivary duct (Sterling et al., 1973). The
narrow duct limits the number of sporzoites which are able
to pass during bloodfeeding. Sporozoites of P_. mexi canum
pass through the one cell thick salivary gland and into the
lumen of the salivary gland (Figures 33 to 36). While
passing through the salivary gland cells, a notable
constriction occurs in the sporozoite at the point of
entering the cell (Figures 33 and 34). A portion of the
gland cell membrane surrounds the sporozoite, forming a
parasitophorus vacuole which appears to be lost prior to
penetrating the lumen side of the gland cell. Sporozites
which enter the lumen of the salivary gland are again bound
in a parasitophorus vacuole, but the host membrane is
readily lost is only seen in a few sporozoites (presumably
those which just passed into the lumen) (Figure 36). In the
study done by Sterling et al. (1973), sporozoites were not
observed migrating through the salivary gland cell in


Results 64
Sporogony of PI asmodi um mexi canum 64
Ultrastructure of Extrinsic Stages of
PI asmodi um mexi canum 71
Di scussi on 92
Sporogony of PI asmodi um mexi can um 92
Ultrastructure of Extrinsic Stages of
Plasmodium mexicanum 94
4 EXPERIMENTAL TRANSMISSION OF PIasmodium floridense
BY BITE OF INFECTED Culex (Melanoconion) erraticus
(DIPTERA:CULICIDAE) 105
Introduction 105
Materials and Methods 106
Field Studies and Collection of Mosquitoes... 106
Culex errati cus Colony Maintenance 110
Collection and Laboratory Maintenance of
Lizards 112
PIasmodium f1oridense Transmission Studies... 112
Course of Infection, Parasitemia 115
Results 115
Field Studies 115
PIasmodium floridense Transmission Studies... 118
Course of Infection, Parasitemia 129
Di scussi on 134
Field Studies 134
PIasmodium floridense Transmission Studies... 140
Course of Infection, Parasitemia 145
5 SPOROGONY, DEVELOPMENT, AND ULTRASTUCTURE OF
EXTRINSIC STAGES OF PIasmodium floridense IN Culex
erraticus 147
Introduction 147
Materials and Methods 148
Sporogony of PI asmodi um floridense 148
Ultrastructure of Extrinsic Stages of
PI asmodi um f 1 ori dense 149
Results 150
Sporogony of PI asmodi um floridense 150
Ultrastructure of Extrinsic Stages of
PI asmodi um floridense 154
Di scussi on 166
Sporogony of PI asmodi um fl ori dense 166
Ultrastructure of Extrinsic Stages of
Plasmodium floridense 168
vi


-195-
Schel1ackia go 1van1 was also transmitted by mosquitoes
(Cx_. errati cus) and sand flies (L. vexator) (this chapter).
Further support for mosquito transmission of Schel1ackia was
demonstrated by Landau ( 1973) using S_. brygooi and Lainson
( 1976 ) studying S.. 1 andaue. These were transmitted by Cx.
pi pi ens p i p i e n s and Cx_. £. f at i gans respectively. In the
present paper, both A. carolinensis and S£. undulatus fed
readily on mosquitoes (especially after a bloodmeal) in the
laboratory. Also, A. carolinensis fed readily on sand flies
(L_. vexator) However, Sc_. undul atus yearlings were only
occasionally observed feeding on sand flies. Hatchling Sc.
undulatus may also feed on sandflies, but none was available
to test.
The age distribution of new infections of Schel1ackia
among lizards may be very skewed because (1) the potential
for many arthropods serving as vectors is great; and (2)
arthropods of different sizes would be ingested by lizards
of different age groups. Smaller arthropods, such as sand
flies would most likely be ingested by young lizards while
larger insects, such as mosquitoes, may present a more even
age distribution of newly acquired infections since they
would be eaten by both small and large lizards. However,
there are some indications that old lizards do not become
infected as readily as young lizards. It is our opinion
that bloodfeeding Diptera are the more likely natural
vectors of ,S. occidental i s and S_. go! vani and that mites are
less important as natural vectors since noninfected lizards


-179-
chamber at 27C and provided a 10% sugar solution following
their bloodmeals on an anole infected with S_. go 1 vani On
the 29th day, the one surviving mosquito was force fed to a
non infected anole (maintained at room temperature) which
developed a patent infection 25 days after ingestion. The
prepatent period varied from 10-12 days post-ingestion for
anoles maintained at 32C while those maintained at room
temperature varied from 21-25 days post-ingestion. One
anole, however, did not develop a patent infection until day
81 post-ingestion.
Differences in the transmission rate of A_. carol inensis
collected from central Florida and Louisiana were also
noted. Eight A. carolinensis from central Florida and seven
A. carolinensis from Louisianna were fed from 1-8 Cx.
erraticus or l. vexator which had previously bloodfed on A.
carolinensis showing >50% of the polymorphonuclear cells
infected with S^. gol van i sporozoites. All eight of the
anoles from central Florida demonstrated sporozoites of S^.
go!vani in their bloodfilms while only three of the seven
anoles from Louisiana became patent. In addition, none of
the Sc_. undul atus which were force fed mosquitoes or sand
flies infected with S_. gol vani became patent (Table 10).
Hi stol ogi cal studies. Sporozoites of S^. gol vani have
been observed only in circulating white blood cells (WBC),
primarily the polymophonuclear series, of A_. carolinensis
(Figure 74 and 75). Sporozoites in the leucocytes vary from
elongate (11.5 X 4.5um) to oval (9.0 X 7.7um). The average


-78-
along the junction of the newly evaginating sporozoite,
becoming the inner sporozoite membrane. A moderately dense
inclusion body forms or migrates immediately adjacent to the
newly formed inner membrane and is the first cytoplasmic
structure to enter the developing sporozoite (Figure 29).
As the sporozoites mature, they elongate into stout
cylindrical forms, in longitudinal section, which are
loosely packed within the oocyst capsule (Figure 30). In
more developed sporozoites, several inclusion bodies may be
seen anterior to the nucleus (Figure 31). The anterior
inclusion bodies become less pronounced as the rhoptries
elongate and micronemes become denser in nearly formed
sporozoites. Moderately dense anterior inclusion bodies are
usually absent in sporozoites which are in the salivary
glands of the sand fly.
Endoplasmic reticulum (granular and smooth) become more
abundant as vacuolization and sporozoite formation is
initiated and are frequently observed in association with
the budding sporozoites (Figures 25 to 27). The nuclei are
smaller and in the early stages of sporozoite formation
migrate to the periphery of the sporoblast. As the
sporozoite elongates and following the inclusion body
entering the sporozoite, the nearly spherical nucleus moves
into the sporozoite, becoming more elongate in shape as it
takes the shape of the sporozoite (Figure 31). Following
the nucleus is a spherical mitochondrion (Figure 31). While
circular and tubular mitochondria are observed in the


-14-
infected hosts (Landau, 1973; Lainson et al., 1976). Jordan
and Friend (1971) further incriminated mites ((3. texana) as
the natural vector of S_. occidentalis by demonstrating that
conspecific lizards, kept in glass gallon jars with infected
lizards harboring mites, became infected. Lainson et al.
(1976) further showed that mosquitoes remain infective for
as long as 14 days following a bloodmeal from an infected
lizard, and may remain infective for the life of the
mosquito.
The taxonomic characterization of Schel1ackia parasites
which infect A n o 1is carolinensis and Seeloporus undulatus i n
Georgia and Florida is uncertain. Jordan and Friend (1971)
and Telford (1978) have identified the Schel1ackia parasites
of S_c undul atus as S_. occi dental i s Jordan and Friend
report that it is doubtful that the Schellackia parasites in
anoles is the same as that of ,S. occi dental i s. Telford goes
one step futher and indicates that Schellackia in the anoles
resembles _S. g o 1 v a n i described from Guadeloupe A n o 1 i s and
common in other Caribbean anoles.
The fine structure of sporozoites of S_. occi dental i s
has been reported by Moore and Sinden (1974). Sporozoites
were contained in a parasitophorus vacuole as are the
sporozoites of Eimeria and Lankesterella. Although there
are similar structures in Schel1ackia and Lankesterella ,
there are also many similarities between Schellackia and
Plasmodium sporozoites. Manwell (1977) remarked that while
Schel1ackia have no known practical importance, they are of


Figure 35. Plasmodium mexicanum sporozoites in the hemocoei
(H) and inside the salivary gland cell (Sg) of
Lutzomyia vexator. While some sporozoites still
remain in the host cell membrane (Pv), others
have shed the host cell membrane. U-shaped
mitochonrion (M) with associated dense body (Dm)
are seen in the posterior of the sporozoites.
Figure 36. PIasmodiurn mexicanum sporozoites in the lumen of
the salivary gland ["Sg) of Lutzomy i a vexator.
Parasitophorus vacuole (Pv) formed by host cell
membrane during penetration of sporozoite into
the lumen of the salivary gland. Rhoptries
(Rh), micronemes (Mn), nucleus (N), mitochondria
(M) and inclusion bodies (I) are seen.


lizards for transmission studies. COL John Reinert served
on the committee for the first year and, in concert with the
other committee members, outlined a demanding schedule of
courses.
Sincere thanks go to Ms. Dianna Simon, Mrs. Debra Boyd,
Mr. Brooks Ferguson, and other members of the laboratory
staff for their assistance in administrative and logistical
details. I extend special thanks to Ms. Edna Mitchell for
her assistance in maintenance of insect and lizard colonies.
Mrs. Margo Duncan offered valuable suggestions in designing
table and figures. I also thank Mr. Greg Piepel for his
assistance with the statistical portions of this study.
I am grateful for the friendship of fellow graduate
students, Dr. Richard Johnson, Dr. Phillip Lawyer, Mr. Eric
Milstrey, Mr. Bruce Alexander, Mr. Charles Beard, MAJ
Richard Kramer, and Mr. Clay Smith. Their suggestions,
humor, and encouragement were appreciated.
Last, I would like to express my deepest gratitude to
my wife Jacqui who supported and encouraged me throughout
the study. I wish to thank my children, Kevin, Aaron,
Michelle, and Robert for their understanding and enjoyment
that they gave me throughout my studies.
xv


-54-
appears to be time-dependent rather than site-dependent, i.
e., hemocoel sporozoites in some cases being equally
infective as salivary gland sporozoites. Only one lizard,
that did not become infected, was fed on by an infected sand
fly on day 7.0-7.5 after its initial bloodmeal. The sand
fly was dissected 0-8 hrs following the second bloodmeal,
and had more than 100 sporozoites in the salivary glands.
Nine of the 12 (75%) remaining lizards, which were bloodfed
on by 1-3 infected sand flies 8-10 days after their initial
bloodmeal on infected S_c. occidentals became infected,
indicating that sporozoites are infective within eight days
of feeding (Table 1).
Both attempts to transmit P_. mexi canum by ingestion of
whole sand flies were unsuccessful. Transmission by oral
ingestion of sporozoites has been reported to be
occasionally successful under certain laboratory conditions
for malarias transmitted by mosquitoes (Shortt and Menon,
1940; Young, 1941; Porter et al ., 1952; Yoeli and Most,
1971) and may be the mode of transmission for P_. agamae
(Petit et al., 1983). However, it is believed that oral
transmission will only occur if the sporozoites are released
in the mouth and penetrate the tissues of the mouth and
throat, since sporozoites are quickly killed in acid
concentrations similar to that of the gut. Although
hatchling lizards may eat sand flies, unrestrained yearlings
were rarely observed feeding on sand flies and mature Sc.
occidentals and Sc. undulatus were never observed feeding


I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as
a dissertation for the degree of Doctor of Philosophy.
Dr. David G. Y'ungl Chafrman
Assistant Professor of Entomology and
Nemotology
I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as
a dissertation for the degree of Doctor of Philosophy.
V^r^.i^Marti r D Yoiuing',^ Co-ojiai rman
Research Profess'oV of Immunology and
Medical Microbiology
I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as
a dissertation for the xfe-f^e of Doctor of Philosophy.
r Jerry F Butler
rofessor of Entomology and
Nematology
I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as
a dissertation for the degree of Doctor of Philosophy.
D7T S
Assoc
and Cell
hen £. Zam J
te Professof-'O
of Microbiology
Science


-129-
development. However, many slower developing oocysts were
melanized in heavily infected midguts after 20 days '(Chapter
5).
Attempts to transmitt £. floridense were only partially
successful. The rate of transmission by bite and IP
inoculation of sporozoites from the salivary glands of Cx.
erraticus was very low. Five of 34 (14.7%) of the A.
carol i nensi s and none of the S_c. undul atus became infected
(Tables 6 and 7) .
Although sporogonic development of £. floridense in
Lutzomyia vexator occurred, only a few free sporozoites with
similar gross morphological characteristics as those seen in
Cx. erraticus, were twice observed from dissected midguts
(n=>50). These sporozoites may have been released by
mechanical rupture during dissection. Sporozoites were
never observed in the salivary glands of sand flies (n=>20)
which survived more than 14 days after feeding.
Course of Infection, Parasitemia
The number of parasites per 10,000 RBC's in lizards fed
upon by an infected mosquito (A-85) or inoculated with
sporozoites from the salivary glands of Cj(. errati cus
(AA-59) was recorded for 120 days (Table 8 and Figure 52).
Due to technical difficulties, bloodfilms from lizard number
A-85 were not made at 3-4 day intervals. The prepatent
period in experimentally transmitted £. floridense (by bite


-124-
development within an oocyst was synchronous (Figure 46).
Sporozoites were observed in some oocysts nine days after
the initial bloodmeal (Figure 47). These oocysts were often
easily ruptured by pressure from the coverslip, releasing
sporozoites and developing sporozoites still attached to the
sporoblast. Beginning nine days after the infecting feed,
free sporozoites were observed from dissected midguts from
mosquitoes maintained at 25C on 10% sugar solution +
multivitamin. However, oocysts may have ruptured during the
dissection since sporozoites were not observed in the
salivary glands until 11 days after feeding. Sporozoites
were not observed in the salivary glands of mosquitoes until
13 days after feeding when maintained at 25C on sugar water
without multivitamins. In general, sporozoites were present
in the salivary glands 11-14 days following a bloodmeal in
mosquitoes that showed 20 or more oocysts (Figures 48 and
49). However, sporozoites were not seen in the salivary
glands of some mosquitoes more than 20 days after feeding
when only a few oocysts were present.
Occasionally, melanization of some oocysts would occur,
but usually was not observed (by light microscopy) until the
later part of sporogonic development (Figure 50). In one
experiment, when the mosquitoes were maintained at 32C,
nearly all the oocysts were partially to completely
melanized by seven days post-feed. Fewer melanized oocysts
were observed following the addition of a multivitamin
solution to the sugar water during the first 14 days of


-200-
cells) than for Sc_. undulatus (percent parasites in RBC's)
and thus it is difficult to make meaningful comparisons.
One of the anoles, demonstrating an increasing parasitemia
of >70% of the polymorphonuclear cells by day 41
post-ingestion, had many gametocytes, but there were few
schizonts (Figure 83). The other three anoles had
relatively few gametocytes with no other stages being
observed. Development of the parasite, as outlined by
Lainson (1976) for S. 1andaue in P. marmoratus, in other
tissues was not observed. The lack of different
developmental stages in both S. occidental is and S^. go! vani
in their respective hosts is confusing since Telford
(personal communication) reported that numerous
developmental stages have been observed in Anolis cybotes
Cope infected with S^. gol vani .


Figure 24. Cross section of a 4 day old PIasmodium
mexicanum oocyst on the midgut of Lutzomyia
vexator The solid, non-vacuolated oocyst
protrudes into the hemocoel (H), is bounded by
a capsule (C) and is in contact with the
basement membrane of the sand fly midgut
epithelium (E). Large nuclei (N) with a
distinct nucleolus (Nu), mitochondria (M), and
endoplasmic reticulum (Er) are scattered
throughout the cytoplasm. Inclusion bodies of
very dense granules associated with a lesser
dense area (Di ) are also scattered througout
the cytoplasm.


-199-
Histological Studies
Parasitemia levels of S^. occi dental i s in Sc_. undul atus
never reached more than 0.4%, even in chronic infections.
In general, parasitemias peaked early at about 0.2 0.3%
and then dropped off to <0.1% when examined at 3 7 day
intervals for more than 60 days. Although parasitemias
remained low, arthropods which fed on these lizards were
highly infective (Table 10). Seeloporus undu1atus
maintained at 32C were killed at two day intervals
subsequent to ingestion of £x. erraticus which had
previously bloodfed on lizards having S_. occi dental is
sporozoites in the blood. Thin sections of the small
intestine revealed few parasites. Parasitemia of the
lizard, sacrificed eight days after ingestion of infected
mosquitoes was less than 0.2% (20/10,000 RBC). The reason
for the lack of forms in the intestine is not clearly
understood.
Ano 1is carolinensis were sacrificed when parasites were
demonstrated in the blood in experimentally transmitted
infections and chronic infections were high in wild caught
lizards. Whereas, most infections in anoles showed the same
general pattern as those in S£. undulatus (a slight peak in
parasitemia and then a chronic phase), two individuals had
marked increases in parasitemia. Due to the type of cell
invaded, the parasitemia levels were recorded differently
for A. carolinensis (percent parasites in polymophonuclear


Table 8. Summary of sporogony and transmission of saurian malaria in
bloodfeeding Diptera.
PIasmodiurn
species
Vector
family species
Oocysts
Sporoz-
i tes
Sporozo
ites in
glands
Trans
mission
mexicanum9
Psychodidae
L. vexator
X
X
X
X
K
Psychodidae
L. stewarti
X
X
X
-
agamae
Ceratpogon-
i dae
C. nubeculosus
X
X
-
-
f1oridense0
Psychodidae
L. vexator
X
X
-
-
Culicidae
Cx. erraticus ,
X
X
X
X
Culicidae
Cx. territans0
X
-
-
-
Culicidae
Cx. quinque-j
fasciatus j
X
Culicidae
Ae. aegypti0
X
-
-
-
Ayala, S. C. Sporogony and experimental transmission of PIasmodiurn
k mexicanum. J. Parasitol. 57:598-602.
Petit, G., I. Landau, Y. Boulard, A. Gomes, and L. Touratier. 1983
Sporogonie de PIasmodiurn agamae c h e z Culi coi des nubeculosus a u
laboratoire: I Experimentation et description du cycTe.
Protistologica 19:537-541.
Attempts to infect £x. sal inarius Cx. nigripal pus, Cx. pi piens ,
Psorphora sp., ke_. atlanticus-tormentor, Ae. triseriatus, and two species
of Chaoboridae, Corthrel la wi rthi and C. E~rak1eyi in the present study
and other studiei (Jordan, 1964; Huff, 1941; Savage, unpublished data)
^ were negative.
Jordan, H. B. 1964. Lizard malaria in Georgia. J. Protozool.
11:562-566.
Telford, S. R., Jr. 1970. Comments on the vector relationships of
f saurian malaria. J. Parasitol. 56:340.
Huff, C. G. 1941. Saurian malaria. J. Parasitol. 27 (suppl.):29
(abst r.).
-131-


Figure 53. Number of PIasmodiurn floridense trophozoites, schizonts, and
gametocytes per 10,000 red blood cells during the course of
infection in Anolis carol inensis (AA-59) infected by
intraperitoneal inoculation of sporozoites from the salivary
glands of Cu 1 ex erraticus .


Figure
46.
Midgut (Mg) of Culex erraticus, (day 9
post-feed) with asynchronous development of
Plasmodium floridense oocysts (0). Lipid-like
globules (Lp) and budding sporozoites (S) are
observed.
Figure
47.
Plasmodium floridense oocyst (0) on the midgut
(Mg) of Culex erraticus with many nearly mature
sporozoites (S) (day 9 post-feed). Lipid-like
globules (Lp) are observed in mature oocysts.
Figure
48.
Culex erraticus salivary glands (Sg)with
salivary ducts (Sd) and sporozoites (S) of
Plasmodium floridense, day 14 post-feed.
Figure
49.
Magnification of living Plasmodium floridense
sporozoites from the salivary glands of Culex
erraticus, day 14 post-feed.


-61-
of the brain of some lizards also demonstrated schizogony in
the endothelial cells (Figure 17). Endothelial cells were
heavily infected in some lizards. However, attempts to
identify parasites in liver, lung, intestine, pancreas,
heart, and uterus from tissue impressions of four Sc.
undulatus were unsuccessful.


Figure 55. Cross section of midgut (Mg) of Culex errati cus
(12 days post-feed) with asynchronous
development of PIasmodiurn f 1 ori dense oocysts.
Oocysts (0) protrude into the lumen of the
midgut, between midgut epithelial cells, and
into the hemocoel. Sporozoites (S) are in some
oocysts.
Figure 56. Residual body (Rb) of sporoblast with developing
sporozoites (S) of PIasmodiurn f1 oridense which
ruptured from an oocyst on the midgut of Cu 1 ex
erraticus. Stained with Giemsa.
Figure 57. Sporozoites (S) of Plasmodium f1oridense in the
salivary gland of Culex erraticus Stained
with Giemsa.
Figure 58. Sporozoites (S) of PIasmodiurn floridense which
ruptured from the salivary glands of Cu 1 ex
erraticus.
Figure 59. Midgut (Mg) of Cu 1 ex erraticus with melanized
oocysts (0) and sporozoi tes (T)(S) of
PIasmodiurn f1oridense 22 days after feeding.
Unstained.


Course of Infection Parasitemia
The number of parasites (expressed as the number of
parasites per 10,000 red blood cells), percent of infected
red blood cells, and percent of infected white blood cells
(per 10,000 red blood cells) for the day prior to death are
shown in Table 1. Bloodfilms during the later course of the
infections became increasingly difficult to obtain,
apparently as a result of anemia. Excluding lizards S-42
and S-51, which had approximately 4x and 6x the number of
parasites, respectively, as the other four lizards which
died, the average number of parasites at time of death was
I,983 (19.8%). The percentage of infected red blood cells
(number of parasites/10,000 RBC) approximated the percent
parasitemia at levels below 25%. However, as the
parasitemia increased, the number of multiple infected red
blood cells also increased, as demonstrated by S-42 which
had 82.7% parasitemia, but only 66.5% of the red blood cells
infected (Table 1 and Figures 6 and 7). Although the
parasitemia of S-51 did not increase greatly during the
later part of the infection (11,420, day 89 post-exposure to
II,960, day 95 post-exposure), the percent of infected red
blood cells continued to increase rapidly until nearly every
RBC was parasitized (72.4 to 91.2%) (Figures 6, 7, and 14).
The transformed (Y=Log of number of parasites per
10,000 RBC) course of infection and linear regression
analysis is shown in figure 7 and Table 2, respectively.


-7-
much variability of parasite development within species and
among different species of lizards infected with P_.
f 1 ori dense and P_. mexi canum. Later, Thompson and Winder
(1947) established a relationship between parasite
development rate and ambient temperature. Susceptible (?)
A. carolinensis inoculated with a common pool of citrated
blood, and maintained in an incubator at 20C, had an
average peak parasitemia at 55 days post inoculation, while
those maintained in an incubator at 30C had an average peak
parasitemia at 13 days. This indicated that environmental
factors, host behavior and effect of the parasite on host
behavior play an influential role in the progression of
parasitemia in lizards under natural conditions.
Goodwin and Stapleton (1952) extended Thompson and
Huff's observations by studying field collected Sc.
undul atus infected with P_. flor i dense. The initial
bloodfilms of three S£. undulatus were negative, thus
allowing the authors to follow naturally acquired saurian
malaria from the prepatent phase (assuming these were not
relapses). In one case, parasites were not demonstrable
until 27 days after capture. Goodwin (1951) also reported
that wild caught lizards with initial negative bloodfilms
and maintained in the laboratory developed parasitemia
within two weeks. These reports were the first indication
that a lengthy prepatent phase is present, at least for Sc_.
undulatus infected with P. floridense.


-214-
Yoeli, M., and H. Most. 1971. Sporozoite-induced
infections of PIasmodium berghei administered by the
oral route. Science 173:1031-1032.
Young, D. G., and P. V. Perkins. 1984. Phlebotomine sand
flies of North America (Diptera:Psychodidae) Mosq.
News 44:263-304.
Young, D. G., P. V. Perkins, and R. G. Endris. 1981. A
larval diet for rearing phlebotomine sandflies
(Diptera:Psychodidae) J. Med. Entomol. 18:446.
Young, M. D. 1941. The oral transmission of PIasmodium
relictum in the pigeon. Pub. Health Rep. U. S. Pub.
Health Serv. 56:1439-1440.


-42-
o
CD
DC
O
O
o
o'
CO
LU
t
CO
<
DC
<
0.
u.
O
DC
Hi
CD
2
3
Z
o
o
DAY POSTEXPOSURE


-141-
hemocoel in only two of the sand flies. Thus full
development including salivary gland infections, was not
documented. Since only a few sporozoites of _P. f 1 ori dense
were observed in vexator, no attempt was made to
determine if they were infective by inoculating them into a
lizard. Therfore, the infectivity of these sporozoites is
unknown. However, since sporozoites were never observed in
the salivary glands, natural transmission would have to
occur by some other mechanism other than b1oodfeeding .
Previous attempts to infect mosquitoes with P_.
floridense by feeding them on infected A. carolinensis and
Sc. undulatus have been mostly unsuccessful (Jordan, 1964 ;
Telford, 1970b) Unfortunately, one of three unidentified
Culex sp. which fed on an infected lizard and developed
large numbers of oocyts (70) was never identified to species
(Jordan, 1964). Attempts to infect other mosquito species
were negative or resulted in a low percentage (5.7% or less)
of infected mosquitoes with only a few oocysts (Jordan,
1964; Huff, 1941).
The relatively high frequency of Cx^. territans
collected in the lizard baited traps, its feeding preference
for cold blooded vertebrates, and the development of oocysts
in four of 70 mosquitoes (Jordan, 1964), suggested that they
may be involved with transmission of JP. flor i dense.
However, attempts to infect F1 £x. territans in this
laboratory were unsuccessful and the peak seasonal abundance
appeared to be in early spring (Figure 45), rather than late


-194-
natural transmission of _S. occi dental i s (Jordan and Friend,
1971). However, in the present study, transmission of S_.
occidentalis did not occur when more than 10 noninfected Sc.
undulatus were maintained for more than 60 days in screened
cages with infected lizards harboring mites (G. texana).
Furthermore, lizards were never observed feeding on mites in
the laboratory. A possible explanation for transmission of
S. occidental is to S£. undulatus by Jordan and Friend (1971)
is the ability of sporozoites of S^. occidentalis to survive
in a broad range of arthropods and that the lizards may have
ingested infected mites which became detached and that had
crawled on other larger arthropods that were introduced into
the cage for lizard food.
The epidemiology of Sc he!1ackia is very complex and
probably varies considerably from one locality to another.
As previously indicated, a broad range of vectors may be
involved in the natural transmission of the parasite because
transmission of S^. occidentalis has been demonstrated by the
ingestion of parasitic mites ((1. texana) mosquitoes (Cx.
erraticus and Ae. aegypti ) and sand flies (l. vexator)
(Table 10). Except for Ae^. aegypti these arthropods often
feed on lizards in nature and two of them, l. vexator and
Cx. erraticus are the natural vectors of saurian PIasmodiurn,
P. mexi canum and £_. f 1 ori dense, respectively (Chapter 2 and
4). The primary vector of Schel1ackia in one locality may
be specific or else several vectors may be involved in the
epidemiology and maintenance of the disease.


Table 7. Laboratory transmission data of PIasmodiurn f1oridense to Anolis
carolinensis by bite of infected Culex erraticus and by
intraperitoneal (IP) inoculation of sporozoites from the salivary
glands of Culex erraticus.
Lizard
Number of
Sporozoite
T empe r-
Day
First day
Number of
number
mosquitoes
rate (day ^
ture
patent
peak para
parasites
bloodfed/IP
post-feed)
C
infection
sitemia
at peak
A-85
1
+3(16)
18-24
25
74
4,280
AA-59
1/IP
+3(18)
18-24
24
55
1,780
AB-23
2 c
+ 2,+3(17 )
32
13
AB-271
i/n>c
+3(17)
32
15
AB-9
13
+ 2 + 3 + 2 ,
32
21
+3(19-20)
3 Lizards number A
-85, AA-59, and
AB-271
are wi1d caught from
Louisiana.
AB-
23 was wild caught near Cross Creek
. AB-9 was
col 1ected
near Rock
, Creek Gainesvilie.
+1, 1-10; + 2, 11-100; +3, 101-1000 sporozoites.
c One salivary gland from Culex erraticus which bloodfed on AB-23 was
. inoculated IP into AB-271.
Only 4 mosquitoes were positive for sporozoites in the salivary glands.
-0£I-


Figure 5. Prepatent and patent period of PIasmodiurn mexicanum infection
and survival of individual Sceloporus undulatus infected by bite
of Lutzomyia vexator infected with PIasmodium mexicanum
sporozoites.
? Killed during the course of the infection.
Maintained in a temperature controlled chamber at 27C.
Yearling 1izards .


12. Course of PIasmodium mexicanum infection in
Sceloporus undulatus infected with sporozoites
(S 2 5 ) .. 46
13. Course of PIasmodium mexican um infection in
Sceloporus undulatus infected with sporozoites
(S-14 ) . 48
14. Bloodfilm of Sceloporus undulatus infected with
PI asmodi um mexi can um parasites 51
15.Spleen tissue impression of Sceloporus undulatus
infected with Plasmodium mexicanum parasites.... 51
16.Bone marrow smear of Seel oporus undulatus
infected with PIasmodium mexicanum parasites.... 51
17. Schizonts of PI asmodi um mexicanum in
endothelial cells of capillaries in the brain... 51
18. Regression of PIasmodium mexicanum mean oocyst
size (and 95% confidence limits) in Lutzomyia
vexator and day post-feed for days 2 through 9.. 68
19.Midgut of Lutzomyia vexator with asynchronous
development of Plasmodium mexicanum oocysts 70
20.Sporoblast formation in 5 day old oocyst on the
midgut of Lutzomyi a vexator 70
21.PIasmodium mexicanum oocyst on the midgut of
Lutzomyi a vexator 7 0
22. Sporozoites of PIasmodium mexicanum which
ruptured from oocysts 70
23. Salivary gland of Lutzomyia vexator containing
numerous sporozoites of PI asmodium mexicanum.... 70
24.Cross section of a 4 day old PIasmodium
mexicanum oocyst on the midgut of Lutzomyia
vexator 73
25. Cross section of Plasmodium mexicanum oocyst
undergoing differentiation 76
26. Cross section of Plasmodium mexicanum oocyst
undergoing differentiation 76
27. Cross section of PIasmodium mexicanum oocyst
undergoing internal vacuolization of the
sporobl astoi d 76
x


DAY POSTEXPOSURE
SCELOPORUS UNDULATUS NUMBER
-8Z-


-5-
Attempts by Pessoa et al ( 1974) to demonstrate snake
malaria sporogony in C_x. f a t i g a n s were also unsuccessful.
Ayala (1970b) and Ayala and Lee (1970) were the first
to report complete sporogonic development of a saurian
malaria (P_. mexicanum, from California, USA) in hematophagus
insects [phiebotomine sand flies, Lutzomyia vexator
(Coquillett) and Lutzomyia Stewarti (Mangaberia and
Galindo)]. In the laboratory, a large proportion of sand
flies (61/72, 84.7%), dissected on day seven after feeding
on an infected lizard, had developing stages of P_.
mexicanum. Some flies had more than 100 oocysts on the
midgut. Sand flies dissected from days 11 to 14 after
feeding had motile sporozoites in the hemocoel and salivary
glands (Ayala and Lee, 1970). Transmission was demonstrated
by inoculation of sporozoites (midgut and hemocoelic fluid)
from wild caught females which had fed on infected
Seeloporus occidental is Baird and Girard (western fence
lizard) in the laboratory. Although Ayala and Lee did not
demonstrate transmission by insect bite, they showed that
both sand fly species were susceptible to infection
(complete sporogony), the sporozoites were infective to
lizards when injected intraperitoneally, and that there is a
close association of sand flies and western fence lizards in
rodent burrows in California.
More recently, sporogony of a saurian malaria (P^.
agamae ) was observed in C u1 ic o id e s nubeculosus (Meig e n), an
unnatural host (Petit et al., 1983). Although sporozoites


156


6ST-


-118-
lizards in the laboratory. Females of the remaining
mosquito species either did not feed, or only a few fed on
lizards in the laboratory. In addition, none of the
Corethrella sp. fed on lizards in the laboratory. Lutzomyia
v exat o r, the vector of _P. mexicanum, was not collected, nor
had it been previously collected from Hatchet Creek (D.
Young, personal communication).
A comparison of the frequency of C)c. erraticus in CDC
light trap collections and percent of C_x. erraticus and Cx.
territans collected in the lizard-baited traps is shown in
figures 44 and 45. Cu 1 ex territans was most frequently
collected in the late spring early summer, while Cx.
erraticus was most frequently collected in the late summer -
early fall.
Plasmodium floridense Transmission Studies
Since £x. errati cus Cx sal i nari us and Cjc. terri tans
readily fed on lizards (Figure 43), attempts were made to
infect them with P_. floridense (Table 6). Cu 1 ex errati cus ,
however was the only mosquito in which sporogonic
development of P_. floridense was observed. Efforts to
infect Cjx. erraticus with P_. mexi canum (saurian malaria) and
IP. hermani (avian malaria) (Nayar et al 1981) were
unsuccessful. Sporogonic development and oocyst frequency
of P. f1oridense in Cx. erraticus was highly variable. The
development among oocysts was asynchronous, whereas


Figure 33. PIasmodium mexicanum sporozoites (S) in the
process of penetrating the salivary gland (Sg)
of Lutzomyia vexator. The anterior end of the
sporozoite comes into contact and causes an
invagination (Iv) of the salivary gland
membrane. The salivary gland cell plasma
membrane (Pm) continues to invaginate until
the sporozoite is contained in a parasitophorus
vacuole (Pv) inside the salivary gland cell
(Sg). No distinct thickening along the
sporozoite-cell membrane junction was noted
during penetration. However, a notable
constriction (Cs) of the sporozoite at the
sporozoite-cell membrane junction was apparent
in most longitudinal sections of sporozoites.
Micronemes (Mn) and elongate rhoptries (Rh) are
present in sporozoites which are migrating into
the salivary glands.
Figure 34. Another sequence of PIasmodium mexicanum
sporozoites in the process of penetrating the
salivary gland (sg) of Lutzomyia vexator.
Nucleus (N), micronemes (Mn), cytostome [Cy),
mitochondria (M) with associated dense body
(Db), parasitophorus vacuole (Pv) and Hemocoel
are 1abeled .


28. Coelescence of the internal vaculization of
PIasmodium mexican urn oocysts produces
cytopl asmi c cl efts 76
29. Higher magnification of PIasmodium mexicanum
oocyst 76
30. Oocyst of PIasmodium mexican um with developing
sporozoites 80
31. Higher magnification of developing sporozoites
of Plasmodium mexicanum 80
32. Cross section of PIasmodium mexicanum oocyst
with developing sporozoites 80
33. PIasmodium mexicanum sporozoites in the process
of penetrating the salivary gland of Lutzomyia
vexator 83
34.Another sequence of Plasmodium mexicanum
sporozoites in the process of penetrating the
salivary gland of Lutzomyi a vexator 83
35.PIasmodium mexicanum sporozoites inside the
salivary gland cell of Lutzomyia vexator 86
36.PIasmodium mexicanum sporozoites in the lumen of
the salivary gland of Lutzomy i a vexator 86
37.Magnification of sporozoites of PIasmodium
mexicanum in the salivary gland of Lutzomyia
vexator 89
38. Sporozoites of PIasmodium mexicanum illustrating
polar rings 89
39. Higher magnification of the cytostome of a
sporozoite of PI asmodi um mexicanum 91
40.Cross section of anterior of Plasmodium
mexican um sporozoite in the lumen of the
salivary gland of Lutzomyi a vexator 91
41. Magnification of the posterior end of a
PI asmodi um mexicanum sporozoite with an elongate
U-shaped mitochondria with an associated
electron dense sphere 91
42. Photograph of lizard baited trap used to
attract and capture biting Diptera 109
43. Cul ex erraticus bloodfeeding on a restrained
An o 1 i s carolinensis 109
xi


-117-
Tab 1 e 5. Summary of CDC light and lizard-bait trap
collections and bloodfeeding of feral mosquitoes
in the field and laboratory from 31 April to 10
October, 1984.
Species
Trap
type
Number
female/
(male)
col 1ected
Number
bloodfed
in trap/
percent
Number
bloodfed
in lab/
percent3
Cx. erraticusb
bai t^
19
7(36.8)
11(91.7)
light0
750(9)
-
226(72.2)
Cx. territanse
bait
43
14(33.3)
12(48.0)
light
2
-
-
Cx. salinarius/ f
bait
2
2(100.0)
-
nigripalpus'
light
915(10)
-
74(40.0)
Cq. perturbans
bait
1 i ght
2
428(11)
2(100.0)
30(20.1)
Cs. melanura
light
995(14)
-
5(2.7)
Psorophora sp.9
1 i ght
16(12)
-
0
Ur. sapphirina
light
443(131
-
0
An. perplexens
light
2
-
0
An. crucians
light
1527(32)
-
3(<0.1)
Ae. fulvus pallens
1 i ght
39
-
0
Aedes sp.
light
1065(54)
-
29(21.2)
a Provided a bloodmeal on Ano!is carolinensis or Seeloporus
undulatus in a 15 X 15 X 22 cm screened cage for a 24 hr.
, period.
Cx. erraticus were only captured from 26 June to 30
August.
c Total number of bait traps set out during the trapping
. period = 924.
0 Total number of light traps set out during the trapping
period = 43.
e Cx. territans were captured throughout the period 1 May to
30 August. However, 71% of those captured were collected
f from 1 May to 7 June.
' Cx. salinarius and Cx. nigripalpus were not separated
since adults could not be identified to species with
certainty.
9 Includes Ps. col umbi ae (8), Ps_. c i 1 i a t a (1), and Ps^.
howa rdi (TO).


-53-
sporozoites of P_. berghei whereas 50% of the canaries
became infected when inoculated intravenously with as few as
five sporozoites. Also, intradermal injection of as few as
10 sporozoites of PIasmodiurn vivax is sufficient to infect
man (Shute et a 1 1976). A review of the kinetics of
sporozoite injection by mosquitoes indicates that only a few
sporozoites are injected during feeding (Vanderberg, 1977).
The influence of temperature on sporozoite development
and maturation of malaria parasites is well documented
(Vanderberg and Yoeli, 1966 ; Stratman-Thomas, 1940). For P_.
mexicanum, sporozoite development in lab-reared l. vexator
maintained at 27C is rapid, with these forms being observed
in the salivary glands by day 6.5 after feeding. However,
sporozoites were not observed in the salivary glands until
day 8.5-9.0 post feed for sand flies maintained at 24C.
Observations by Ayala and Lee (1970) indicated that
sporozoites from lab-infected wild caught sand flies were
not observed in the hemocoel until day 11-14 post-feed when
maintained at room temperature (24-26C). Maximum and
minimum temperatures when sporozoite development ceases were
not determined.
Vanderberg (1975) showed that sporozoites require a
period of maturation after their release from the oocyst and
that P_. berghei sporozoites in the salivary glands of a
mosquito are 10,000 times more infective than sporozoites
from the oocyst of the same mosquito. However, once
released from the oocyst, the development of infectivity


Figure 70. Oocyst of PIasmodium f1oridense with
developing sporozoites. Subpellicular
microtubules (arrows) are present in the
anterior 1/3 of the sporozoites.
Figure 71. Salivary gland (Sg) of Cu 1 ex errati cus infected
with sporozoites (S) of PIasmodium
f1oridense Sporozoites are intracellular and
appear to be in groups within cells.
Figure 72. Cross section of PIasmodium floridense
sporozoites in the salivary glands of Culex
erraticus. Micronemes (Mn) and rhopt ri es ("Rh)
are present. Subpel1icular microtubules (Mt)
are in groups, apparently as a result of
improper fixation.
Figure 73. Degenerating sporozoite (S) of PIasmodium
f1 oridense in an oocyst which is becoming
melanized in Cu 1 ex erraticus Oocyst capsule
(C) is very thick. Many micronemes (Mn) are
present in an abnormal sporozoite.


-213-
Thompson, P. E., and C. G. Huff. 1944a. A saurian malarial
parasite, PIasmodiurn mexicanum, n. sp., with both
elongatum-and gal 1inaceum- types of exoerythrocytic
stages. J. Inf. Dis. 74:48-67.
Thompson, P. E. and C. G. Huff. 1944b. Saurian malarial
parasites of the United States and Mexico. J. Inf.
Dis. 74:68-79.
Thompson, P. E., and C. V. Winder. 1947. Analysis of
saurian infections as influenced by temperature. J.
Inf. Dis. 81:84-95.
Vanderberg, J. P. 1975. Development of infectivity by the
Plasmodium berghei sporozoite. J. Parasitol.
61:43-50.
Vanderberg, J. P. 1977. PIasmodiurn berghei : quantitation
of sporozoites injected by mosquitoes feeding on rodent
host. Exp. Parasitol. 42:169-181.
Vanderberg, J. P., R. S. Nussenzweig, and H. Most. 1968.
Further studies on the PIasmodiurn berghei-Anopheles
stephensi-rodent system of experimental malaria. J.
Parasitol 54:1009-1016.
Vanderberg, J., J. Rdodin, and M. Yoeli. 1967. Electron
microscopic and histochemical studies of sporozoite
formation in Plasmodium berghei. J. Protozool.
14:82-103.
Vanderberg, J. P., and M. Yoeli. 1966. Effects of
temperature on the sporogonic development of PIasmodiurn
berghei J. Parasitol. 52: 559-564.
Ward, R. A. 1963. Genetic aspects of the susceptibility of
mosquitoes to malarial infection. Exp. Parasitol.
13:328-341.
Wenyon, C. M. 1909. (dated 1908). Report of traveling
pathologist and protozoologist. Balfour, A. ed. Third
Report, Wellcome Research Laboratory at the Gordon
Memorial College, Khartoum. London: Bailliere, Tindall
and Cox.
Wernsdorfer, W. H. 1980. The importance of malaria in the
world. Kreier, J. P. ed. Malaria (Vol 1). New York:
Acad. Press.
Wong, T. C., and S. S. Desser. 1976. Fine structure of
oocyst transformation and the sporozoites of
Leucocytozoon d u b r e ui 1i J. Protozool. 23:115-126 .


Figure 45. Percent Cu 1 ex erraticus and Cu1ex territans
collected per bait trap during the period T
May to 9 October, 1984.


-110-
trap, counted, and identified. Living mosquitoes were
treated differently, as discussed below. Adult Culex
sa 1 inarius Coquillett and Cu 1 ex nigripalpus Theobald were
often badly rubbed and could not be separated with
certainty, and are treated together for purposes of this
discussion. Many Aedes sp. were also often badly rubbed and
were not identified to species.
Culex erraticus Colony Maintenance
The Cjc. errati cus colony originated from wild caught
females collected in both lizard-baited and CDC light traps
during June August, 1984. Engorged Cx^ erraticus and Cx.
territans collected in the lizard-baited traps were returned
to the laboratory, placed in screen topped pint cardboard
cartons (0.5 1) and provided a 10% sucrose solution for 2-3
days. Subsequently, mosquitoes were removed from the
cartons and placed in 15 ml oviposition vials half full of
water and plugged at the top with cotton. A water plant,
duckweed (Lemna sp.), was initially placed on the water
surface to enhance ovipostion of C_x. erraticus, but in later
generations was replaced with 14 mm circular sections of
azalea leaves cut with a size nine cork borer. Unengorged
mosquitoes from the lizard-baited and CDC light traps were
placed in a screened cage (18 X 18 X 21 cm) with a lizard
(A. carol inensis or Sc_. undul atus ) Bloodfed errati cus
and Cx. territans were removed after a 24 hr period and


-55-
on L. vexator in the laboratory. Since mastication of
ingested flies does not normally occur, and both attempts to
orally transmit P_. mexi canum failed, transmission by the
oral route is believed to have little epidemiological
significance.
Prepatent periods in experimentally transmitted P_.
mexicanum by bite of infected l. vexator ranged from 23-40'
(mean 28.6) days. In examining natural infections of
another lizard malaria, P_. f 1 ori dense, Goodwin (1951) showed
that parasites were not observed in bloodfilms until
approximately two weeks after the lizards were collected.
In another study, parasites were not observed in the
bloodfilm of one lizard until 27 days after capture (Goodwin
and Stapleton, 1952). Present studies on the transmission
of P_. f 1 ori dense indicate that the prepatent period is more
than 20 days for bite induced and IP induced infections at
18-24C (Chapter 4). These studies and those by Goodwin and
Stapleton (1952), assuming that the naturally acquired
infections were not relapses, and other studies on P_.
mexicanum laboratory transmission, support the hypothesis of
a lengthy prepatent period for at least two of the saurian
malarias.
Recent studies (Chapter 6) on other hemosporidians,
Schel1ackia golvani and Schel1ackia occidental is, indicate
that temperature may affect the length of the prepatent
period. For lizards maintained at room temperature
(18-24C), Schellackia sporozoites were not observed in


-207-
Garnham, P. C. C., R. G. Bird, and J. R. Baker. 1962.
Electron microscope studies of motile stages of malaria
parasites. III. The ookinites of Haemamoeba and
Plasmodium. Trans. R. Soc. Trop. Med. Hy q.
56:116-120.
Garnham, P. C. C., R. G. Bird, and J. R. Baker. 1963.
Electron microscope studies of motile stages of malaria
parasites. Trans. R. Soc. Trop. Med. Hyg. 57:27-31.
Garnham, P. C. C., R. G. Bird, J. R. Baker, and R. S. Bray.
1961. Electron microscope studies of motile stages of
malaria parasites. II. The fine structure of the
sporozoite of Laverania (=P1asmodiurn) falcipara.
Trans. R. Soc. Trop. Med. Hyg. 55:98-102.
Garnham, P. C. C., and S. R. Telford, Jr. 1984. A mew
malaria parasite PIasmodiurn (Sau ramoeba ) h eis c hi in
skinks (Mabuya striata) from Nairobi, with a brief
discussion of the distribution of malaria parasites in
the family Scincidae. J. Protozool 31:518-521.
Gingrich, W. D, 1932. Immunity to superinfection and
cross immunity in malarial infections of birds. J.
Prev. Med. 6:197-246.
Goodwin, M. H. 1951. Observations on the natural
occurrence of PIasmodiurn floridense, a saurian malaria
parasite, in Seeloporus undulatus undulatus J. Nat.
Mai. Soc. 10:57-67.
Goodwin, M. H., and T. K. Stapleton. 1952. The course of
natural and induced infections of PIasmodiurn floridense
Thompson and Huff in Seeloporus undulatus undulatus
(Latreille). Am. J. Trop. Med. Hyg. 1:77 3-783.
Grassi, B. A. bignami, and G. Bastianelli. 1899. Ulteriri
ricerche sul ciclo parasitti malarici unamy nel corpo
del zanzarone. Atti Reale Accad. dei Lincei.
8:21-28.
Greenberg, J., H. L. Trembley, and G. R. Coatney. 1950.
Effects of drugs on PIasmodiurn gal 1inaceum infections
produced by decreasing concentrations of a sporozoite
inoculum. Am. J. Hyg. 51:194-199.
Greer, N. I., and J. F. Butler. 1973. Comparisons of horn
fly development in manure of five animal species. Fla.
Entorno 1 56:197-199.
Greiner, E. C., and P. M. Daggett. 1973. A saurian
PIasmodiurn in a Wyoming population of Seeloporus
undulatus. J. Herpetol. 7:303-304.


-170-
Therefore, conclusions about their similarities and
differences between other malarias can not be made.
While the basic organization of the sporozites of _P.
flor i dense is similar to other malarias, further examination
of material that is properly fixed is required to make
detailed comparisons.


CHAPTER 5
SPOROGONY, DEVELOPMENT, AND ULTRASTRUCTURE OF EXTRINSIC
STAGES OF Plasmodium floridense IN Culex erraticus
Introduction
Plasmodium floridense was first described in the
lizard, Seeloporus undulatus, by Thompson and Huff (1944b)
and has been extensively studied by Goodwin (1951), Jordan
(1964), and others. The fine structure of the erythrocytic
stage of £. floridense, PIasmodiurn mexicanum, and PI asmodi urn
tropiduri has been examined (Aikawa and Jordan, 1968; Moore
and Sinden, 1974; Scorza, 1971a). However, the histological
examination and the study of the fine structure of
sporogonic development and sporozoites has not been possible
because attempts to find the natural vectors of saurian
malarias have eluded researchers until recently (Ayala and
Lee, 1971; Chapters 2 and 4).
The study of the fine structure of P_. mexi canum in its
natural vector, Lutzomyi a vexator and P_. agamae in an
unnatural host, C u 1 i c o i d e s nubeculosus indicate that some
characteristics of avian and mammalian malarias are not
shared by either P_. mexi canum or P_. agamae. In addition,
the vector of P_. mexicanum is a phlebotomine sand fly, not a
mosquito. PIasmodiurn agamae has been shown to develop in a
-147-


-193-
We conducted experimental transmission studies to
determine if S^. go 1 vani (host = A. carolinensis) and S^.
occidentalis (host = S£. undulatu s ) in Florida were distinct
species. Data indicate that blood stages (sporozoites) of
Schellackia were observed only in previously non infected
wild caught lizards when they were force fed arthropods that
had bloodfed only on conspecific lizards, i. e., go!vani
was only transmitted to A. carolinensis and S^. occidental is
was only transmitted to Sc_. undulatus (Table 10).
Wild-caught lizards that had initial negative bloodfilms
from the same locality as the test lizards and that were not
force fed infected arthropods did not develop patent
infections. It was not determined whether schizogony of S^.
gol vani and occi dental i s occurred in the intestine of Sc.
undulatus and A. carolinensis, respectively. However, even
if schizogony occurs in the intestine, the lack of
sporozoites in the circulating lymphocytes and RBC's
indicates an unsuitable host.
Studies of Reichenow (1919) and Bonorris and Ball
(1955) strongly suggest that parasitic mites of lizards are
the natural vectors of Schellackia parasites. Successful
experimental transmission of S.. occi dental is (2 of 3) by
force feeding infected £. texana to S£. undulatus (this
chapter) and transmission of S^. occi dental i s (3 of 4) to
non infected lizards which were in glass-lined cages with
infected lizards harboring parasitic mites (G^. texana)
provide further evidence that mites are involved in the


-92-
to be U-shaped in the posterior of the sporozoite (Figure
41). An electron dense spherical body is associated with
the mitochondrion (Figure 41). While all three types of
inclusion bodies are seen scattered throughout the mature
sporozoite, the one most frequently seen in mature
sporozoites is the very electon dense granule associated
with a space of low density. Crystalloid inclusions as
seen in some hemoproteids and some P1asmodiurn sp. were not
observed.
Discussion
Sporogony of Plasmodium mexicanum
The effects of temperature on the sporogonic
development of several avian and mammalian malaria parasites
have been well studied and much of the literature is
summarized by Boyd (1949), Chao and Ball (1962), and Ball
and Chao (1964). General conclusions from studies on human
malaria indicate that sporogonic development occurs between
environmental temperatures of between 16 and 30C. Similar
conclusions were made for P_. rel i ctum, an avian malaria, by
Chao and Ball (1962). At low temperatures, sporogonic
development is often arrested in human malarias, with
oocysts developing normally when they are returned to
favorable temperatures (Young, personal communication).
However, parasites appear to be readily damaged when


73
i v


6 DEVELOPMENT AND EXPERIMENTAL TRANSMISSION OF
Schellackia golvani AND Schellackia occidentalis
BY INGESTION OF INFECTED BLOODFEEDING ARTHROPODS.... 171
Introduction 171
Materials and Methods 172
Transmission Studies 172
Histological Studies 175
Results 176
Schellackia golvani 176
Schel 1 acki a occidentalis 185
Discussion 187
Transmission Studies 187
Histological Studies 199
7 SUMMARY AND RECOMMENDATIONS FOR FUTURE
INVESTIGATIONS 201
REFERENCES 204
BIOGRAPHICAL SKETCH 215
vi 1


-145-
Course of Infection, Parasitemia
The effect of different temperatures on blood induced
infections of P_. f 1 ori dense indicated that infections became
patent earlier at 30C (mean = 3.2) than at 20C (mean =
10.0) and that the infections also increased at a much
faster rate at 30C (Thompson and Winder, 1947). While the
prepatent period for sporozoite induced infections was
generally longer than blood induced infections, the
prepatent period for lizards maintained at 32C was much
shorter than in lizards maintained at room temperature.
Thompson and Winder (1947) did not determine if there were
significant differences in the numbers of gametocytes
produced at the two different temperatures. While the
present study is incomplete, it appears that gametocytes
appear earlier in lizards which are maintained at 32C.
This corresponds to the results observed when wild caught
infected lizards were placed in the incubator at 32C. In
general, all wild caught infected lizards produced more
gametocytes when maintained at 32C. However, those with
light infections generally continued to be light and
produced few gametocytes. Those with moderate or heavy
infections, although fluctuating, continued to be moderate
to heavy and produced many gametocytes. The lengthy
prepatent period of P_. flor i dense at room temperature
(18-24C) corresponds to other studies on P_. f 1 ori dense in
wild caught lizards (Goodwin and Stapleton, 1952) and P_.


LOG NUMBER OF PARASITES/10.000 RBC
-43-
DAY POSTEXPOSURE


44. Number of Cu 1 ex errati cus collected per light
trap during the period 1 May to 9 October,
1984 120
45. Percent Cu 1 ex erraticus and Cu 1 ex territans
collected per bait trap during the period 1
May to 9 October, 1984 122
46. Midgut of Cu 1 ex erraticus (day 9 post-feed)
with asynchronous development of PIasmodiurn
f 1 ori dense oocysts 126
47. PIasmodiurn f1oridense oocyst on the midgut of
Cu 1 ex erraticus with many nearly mature
sporozoites (day 9 post-feed) 126
48. Culex erraticus salivary glands with sporozoites
of Plasmodium floridense 126
49.Magnification of living Plasmodium f1oridense
sporozoites from the salivary glands of Cu 1 ex
erraticus 126
50. PIasmodiurn f1oridense oocyst on the midgut of
Cu 1 ex erraticus on day 10 post-feed.
Melanization is beginning to occur along the
oocyst capsule and spread inward 128
51. Bloodfilm of Ano 1is carolinensis at the first
peak of a PIasmodiurn floridense infection 128
52.Course of acute infection of PIasmodiurn
floridense in two wild caught Ano 1is
carol inensis infected with sporozoites 133
53. Course of acute infection of PIasmodiurn
f1oridense in Ano 1is carolinensis infected
with sporozoites (AA-59) 136
54. Course of acute infection of PIasmodiurn
floridense in Ano 1i is carolinensis infected
with sporozoite! (A 8 5 ) 138
55. Cross section of midgut of Cu 1 ex erraticus with
asynchronous development of PIasmodiurn
f 1 ori dense oocysts 152
56.Residual body of sporoblast with developing
sporozoites of Plasmodium floridense which
ruptured from an oocyst on the midgut of Cu 1 ex
errati cus 152
57.Sporozoites of Plasmodium floridense in the
salivary gland of Culex~erraticus 152
Xll


-203-
Recommentdations for future studies include.
1. Establish the vector status of Cx^ erraticus and j^.
vexator by dissecting wild collected females, examining for
sporogonic stages, and injecting sporozoites (IP) into
noninfected anoles to confirm species of PIasmodiurn.
2. Continue P_. f 1 or i dense transmission studies and
determine potential vectors of _P. f 1 ori dense in other
localities by use of lizard-baited traps and CDC light
traps.
3. Determine nutritional and temperature conditions
for optimum P_. f 1 ori dense development.
4. Examine development of sporogonic stages of £.
f 1 ori dense in L_. vexator and compare with development stages
in C>c. er rati cus .
5. Determine the prepatent exoerythrocytic stages of
both P_. mexi canum and P^. f 1 or i dense .
6. Determine if S^. go! vani can be transmitted to other
closely related anoles. Examine the ultrastructure of S_.
golvani and compare it to other hemoproteiids.


-84-
the gland cell (Figure 35). Soon after entering the gland
cell, the host membrane is lost (Figure 35). The sporozoite
continues to migrate until it reaches the hollow cavity of
the salivary gland, again becoming constricted at the
membrane-parasite junction adjacent to the lumen of the
salivary gland. Remnants of the parasitophorous vacuole
which are present in sporozoites which have recently
migrated into the lumen of the salivary gland disappear
shortly after penetration (Figure 36).
The sporozoite pellicle is composed of an outer and
inner plasma membrane as previously discussed (Figure 37 and
38). The cytostome occurs as a midcentral invagination
(1.0-1.5 X 1.2-1.7um) and is located near the distal portion
of the nucleus (Figures 32, 34 and 39). The pellicular
membranes are continuous at the junction of the cytostome.
However, the inner membrane is thickened and more electron
dense along the sides of the cavity. Subpellicular
microtubules are arranged asymetrically around the
circumference of the sporozoite and extend anteriorly from
the nucleus to the apex (Figure 40). The number of
subpellicular microtubules usually observed in the anterior
1/3 1/2 of the sporozoite is 14. Fewer than 14
microtubles are sometimes observed near the middle of the
sporozoite. The subpellicular microtubles are arranged
asymetically around the circumference and with a
dorsal-ventral aspect. Nine of the 14 microtubules are
arranged equidistantly and relatively close together around


-142-
summer to early fall when transmission of P. f1oridense is
suspected to occur (Jordan, 1964).
The results of the following study demonstrate
conclusively that C_x. erraticus is capable of transmitting
P_. f 1 ori dense to A. carol inensis. However, there are many
unanswered questions regarding its role in the natural
transmission of saurian malaria. Sporogonic development and
oocyst frequency in £x. erraticus were highly variable, for
reasons not well understood [e. g., genetic, temperature, or
other factor(s)]. However, Ward (1963) demonstrated genetic
selection for susceptibility to malaria parasites and
derived a nonsuseptible colony of /\e. aegypti to P_.
gal 1inaceum from a susceptible colony. It has not been
determined whether the colony of C_x. erraticus consists of
two genetically different populations, one susceptible and
the other resistent to infection, or if other factors are
i nvol ved.
Large numbers of oocysts (>100), that are sometimes
seen in mosquitoes infected with mammalian and avian
malarias, have not been observed in £x. erraticus infected
with P_. flor i dense. This, however, may be related to the
smaller size (bloodmeal) of Cjx. erraticus and lower
gametocyte levels in the saurian hosts. Furthermore, large
numbers of gametocytes may have a detrimental effect on the
mosquito since 63% (12/19) of the Cx. errati cus which fed on
a lizard with numerous gametocytes died by the third day
after feeding. Those that did survive may have only taken


-6-
were observed in the oocysts, the oocysts were intracellular
and did not rupture. This suggests that this species of
PIasmodium may be transmitted by ingestion. However,
sporogony of malaria parasites in unnatural mosquito species
has also been reported to be sometimes intracellular.
Telford (1970b) stated that the current ". . finding(s)
suggest that saurian malaria (parasites) may utilize a
variety of hematophagus arthropods for transmission.
Certainly, generalizations concerning vector relationships
of this group of parasites are still premature" (p. 340).
Because the vectors of saurian PIasmodium are unknown,
attempts to follow the life cycle of the parasite in lizards
have been done only by experimentally inoculating
susceptible lizards with blood from infected lizards
(Thompson, 1944; Thompson and Huff, 1944a,b). These early
reports demonstrated slow asynchronous parasite development
in two species of saurian malarias (P. mexi canum and P_.
f1oridense) that is different from the synchronous, fast
developing parasitemias of some avian and mammalian malarias
(Thompson and Huff, 1944a,b; Thompson, 1944). Parasitemias
of P_. f 1 ori dense (>1,000 pa r as i t es/ 10,000 erythrocytes in
Sc. undu1atus and A. carolinensis ) peaked between 23-94
days, then gradually decreased. (The period of time between
inoculation and peak parasitemia may be a function of
parasite inoculation rate.) Blood-induced infections in Sc.
olivaceous and A. carolinensis appeared to peak earlier, at
approximately day 45. Thompson (1944) showed that there is


-56-
bloodfilms until day 21 and 37 post-ingestion, respectively.
However, when lizards were maintained at 32C (90F),
sporozoites were seen in the bloodfilms as early as day 10
and 7 post-ingestion, respectively. Similarly, the
prepatent period of P_. f 1 ori dense was decreased by as much
as seven days when maintained at 32C (Chapter 4).
In general, lizards which were fed on by more than one
infected sand fly developed earlier patent P_. mexi canum
infections. One lizard (S-51) which was bloodfed on by only
one sand fly in which fewer than five sporozoites were
observed in the salivary glands and around the head 0-8 hrs
after the second bloodmeal did not develop a patent
infection until 40 days after feeding. Ayala (1971) showed
that Sc_. occidental is inoculated with sporozoites from five
sand flies (some having more than 100 oocysts on the midgut)
had a prepatent period of 21 days. Observations on human,
rodent and avian malarias show that, within limits, the
higher the inoculum of sporozoites, the shorter the
prepatent period (Boyd, 1940; Greenberg et al., 1950;
Vanderberg et al 1968). Based on the present studies, the
length of the prepatent period appears to be a response of
the number of sporozoites. However, host-temperature
maintenance in relation to parasite growth and development
may also play an important role in the early course of the
infection.
Natural infections of P.. m e x i c a n u m occur in both Sc .
occidentals and Sc. undulatus (Ayala, 1971; Greiner and


-63-
ultrastructure of sporogony of a saurian malaria was that
done by Boulard et al ( 1983) on P_. agamae in C.
nubeculosus.
The only known vectors of mammalian and avian malarias
are mosquitoes. In view of histological and vector
differences between P_. mexicanum and other malarias, TEM
studies were undertaken. Data from this study are compared
with those of other Haemosporina.
Materials and Methods
Sporogony of Plasmodium mexicanum
Laboratory reared phlebotomine sand flies, Lutzomyia
vexator, were bloodfed on wild caught Seeloporus
occidentalis infected with £. mexicanum as previously
described (Chapter 2). Bloodfed females were removed at 4
hr intervals, placed in 25 ml plastic oviposition vials,
provided a sugar source (1:1 mixture of Karo syrup and
distilled water), and maintained in a temperature-humidity
controlled chamber at 27C, 24C, or 19C, and 80% RH.
Midguts were dissected (Chaniotis and Anderson, 1968) at 12
hr intervals from days two through nine, placed on a clean
slide with a drop of cold-blooded Ringer's solution, covered
with an 18mm circular coverslip, and the number and
measurements of 25 (or fewer) oocysts recorded. Salivary
glands were examined after day five following a bloodmeal


-52-
undul atus by bite of L_. vexator, a species that coexists
with the parasite in California. Progeny of two species of
mosquitoes, C_x. erraticus and Cx. territans, collected in
lizard baited traps failed to transmit this parasite while
as many as 50 oocysts developed on the midgut of l. vexator
feeding on the same lizard. This provides further evidence
that sand flies are the natural vectors of P_. mexi canum,
especially since sporogony and transmission of P_. f 1 ori dense
has been demonstrated for £x. errati cus (Chapter 4).
Sporozoite infectivity of P_. mexi canum transmitted by
bite of J_. vexator is relatively high and comparable with
some other malarias (Coatney et al., 1945; Russell and
Mohan, 1942). Transmission occurred in 62% (5/8) of the Sc.
undu1atus which were bloodfed on by one sand fly and 80%
(4/5) of the Sc. undulatus which were bitten by two or more
sand flies. Sporozoites were occasionally observed in the
mouthparts of dissected sand flies, but only in small
numbers ( < 5) One lizard (S- 51) having the longest
prepatent period, became infected when bitten by a sand fly
with fewer than five sporozoites observed near the head and
in the salivary glands following the second bloodmeal
(Figures 5 and 8). Evidence indicating that only a few
sporozoites are required to infect the host is provided by
Fink (1968) for PIasmodiurn cathemerium and Vanderberg et al.
( 1968) for PIasmodiurn berghei Vanderberg et al and Fink,
respectively, demonstrated that 26% of the mice in the study
became infected when injected with approximately 10


-209-
Landau, I. 1973. Diversite des mecanismes assurant la
perennite de 1'infection chez les sporozoaires
coccidiomorphes. Mem. Mus. Nat. Hist. Nat., Serie A,
Zoologie 77:1-62.
Landau, I., R. Lainson, Y. Boulard, Y. Michel, and J. J.
Shaw. 197 3. Developpement chez Cu 1 ex pipiens de
Saurocytozoon tupinambi (Sporozoaire Leucocytozoidae) ,
parasite de lezards bresiliens. C. R. Acad. Sci.
(Paris) 276:2449-2452.
Laveran, A. 1880. Note sur un nouveau parasite trouve dans
le sang de plusiers malades atteints de fievre
palustre. Bull. Acad. Med. (Paris) 9:1235-1236.
Lebail, 0., and I. Landau. 1974. Description et cycle
biologique experimental de Schel1ackia bal 1i n s p .
(Lankesterel1idae) parasite de Crapauds de Guyane.
Bull. Mus. Nat. Hist. Nat., Serie 3, Zoologie
194:91-97.
Leger, L., and P. Mouzels. 1917. Hemogregarine
intraleucocytaire d'un saurien, Tupinambis
nigropunctatus. Bull. Soc. Pathol. Exot. 10:283-284.
Levine, N. D. 1980. Some corrections of coccidian
(Api comp!exa:Protozoa) nomenclature. J. Parasitol.
66:830-834.
Manwell, R. D. 1977. Gregarines and haemogregarines. In:
Kreier, J. P. ed., Parasitic Protozoa. 3:1-32.
New York: Academic Press.
Mattingly, P. F. 1965. The evolution of parasite-arthropod
vector systems. I n: Taylor, Angela E. R. ed.,
Evolution of Parasites. 3rd Symposium of Brit. Soc.
for Parasitol. Oxford: Blackwell Sci. Publ.
Mehlhorn, H., W. Peters, and A. Haberkorn. 1980. The
formation of kinetes and oocyst in PIasmodiurn
gal 1inaceum (Haemosporidia) and considerations on
phylogenetic relationships between Haemosporidia,
Piroplasmida and other Coccidia. Protistologica
16:135-154.
Moore, J., and R. E. Sinden. 1974. Fine structure of
PIasmodiurn mexicanum J. Parasitol. 60:825-834.
Nayar, J. K., M. D.Young, and D. J. Forester. 1981. Cu 1 ex
restuans: An experimental vector for wild turkey
malaria, Plasmodium hermani. Mosq. News 41:748-750.


DEVELOPMENT AND TRANSMISSION OF SAURIAN PLASMODIUM AND
SCHELLACKIA IN BLOODFEEDING ARTHROPODS
BY
TERRY A. KLEIN
A DISSERTAT
IN PARTIAL F
D
ION PRESENTED TO THE GRADUATE
THE NIVERSITY OF FLORIDA
ULFILLMENT OF THE REQUIREMENTS
EGREE OF DOCTOR OF PHILOSOPHY
SC
F
HOOL OF
OR THE
UNIVERSITY OF FLORIDA
1985


TABLE 2. Linear regression analysis of acute PIasmodium mexicanum
infections with fewer than 500 parasites per 10,000 red
blood cells (5% parasitemia).
Lizard
number
Days sur
vived post-
exposure
Days sur
vived post
patent inf.
Linear regression equation
of acute infection with
<5% pa ras itemia/(SE )D
R2
Slope
S-8
52
25
Y =
-11.35 + 0.42x
(1.14) (0.03)
.96
.417
S 2 5
52
19
Y =
-14.44 + 0.45x
(1.94) (0.05)
.96
.452
S-42
61
28
Y =
-10.05 + 0.36x
(1.79) (0.05)
.95
.361
S-14
39
13
Y =
-10.34 + 0.47 x
(1.93) (0.06)
.96
.471
S -15
47
21
Y =
- 9.74 + 0.42x
(0.67) (0.02)
.99
.420
S-43a
66
40
Y =
- 4.57 + 0.31x
(1.86) (0.06)
.93
.309
S 47 3
46
23
Y =
-14.28 + 0.64x
(2.11) (0.07)
.97
.640
S-503
45
22
Y =
- 8.51 + 0.42x
(2.86) (0.09)
.86
.417
S 51
96
56
Y =
- 5.71 + 0.23x
(1.73) (0.04)
.93
.230
k Lizards were killed when lethargic and anorexic.
Y=Estimated number of P_. mexi canum parasites per 10,000 RBC on day
post-exposure.


-2-
elongatum Huff, an avian malaria) and by Shortt and Garnham
(1948) (PIasmodiurn cynomolgi Mayer and PIasmodium vivax
Grassi and Felitti, a monkey and human malaria,
respectively). Literature on the mammalian and avian
malarias is voluminous due to their medical and economic
importance, but saurian malaria has received very little
attention.
The first named saurian malaria parasites, P_. m a b u i a e
Wenyon (1909) and PIasmodium (=Haemoproteus) agamae (Wenyon,
1909), were described from Mabuya quinquetaeniata
(Lichenstein) (common skink) and Agama agama L., (rainbow
lizard) respectively, from Africa (Wenyon, 1909). During
that same year, Aragao and Neiva (1909) described PIasmodium
d i p 1 o g 1 o s s i and P_. tropi duri from Diploglossus f asci atu s
(Gray) and Tropidurus torquatus (Wied), respectively, from
Brazil. Since that time, a total of 59 species of saurian
malaria have been described from Africa (12), the Americas
(36), and Australasia (11) (Telford, 1982; Telford, 1983;
Telford, 1984a; Telford 1984b; Garnham and Telford, 1984).
A review of saurian malaria literature from 1909 to 1975
(Ayala, 1978) includes only 153 references, most of which
emphasize taxonomy or distribution records. From 1975 to
1985, only 32 additional references have been added
[including 4 previously overlooked by Ayala (1978)].
Garnham (1966) stated that, "A few puzzles in the life
history of the malaria parasites remain, of which the most
important are perhaps the exact nature of latency and vector


This dissertation was submitted to the Graduate Faculty of
the College of Agriculture and to the Graduate School, and
was accepted as partial fulfillment of the requirements for
the degree of Doctor of Philosophy.
August, 1985
Dean, Graduate School


Figure 39. Higher magnification of the cytostome (Cy)of a
sporozoite of PIasmodium mexicanum .
Figure 40. Cross section of anterior of PIasmodium
mexicanum sporozoite in the lumen of the
salivary gland of Lutzomyia vexator. Note
the asymetrical arrangement of the
subpellicular microtubules (arrows) and the
dense rhoptries (RH).
Figure 41. Magnification of the posterior end of a
PIasmodium mexicanum sporozoite with an
elongate U-shaped mitochondria (M) with an
associated electron dense sphere (Db).


CHAPTER 4
EXPERIMENTAL TRANSMISSION OF Plasmodium f1oridense BY BITE
OF INFECTED Culex (Melanoconion ) erraticus
(DIPTERA:CULICIDT}~
Introduction
The first saurian malaria parasite was discovered in
1909 by Wenyon. Since then, 58 more species of saurian
PIasmodiurn have been described. Mosquitoes, phlebotomine
sand flies, and biting midges (Ceratopogonidae) have, at one
time or another, been suggested as possible vectors of
saurian malaria but only mosquitoes have been incriminated
as vectors of mammalian and avian malarias. A number of
biting Diptera may be involved in saurian malaria
transmission because previous studies (Chapter 2) have
demonstrated that (1) PIasmodiurn mexicanum is efficiently
transmitted by bite of Lutzomyia vexato r, (2) sporogony of
PIasmodiurn agamae occurs in Cu 1 i coi des nubeculosus (B o u1 a rd
et al., 1983), and (3) Culex erraticus transmits Plasmodium
f1oridense (this chapter). Differences observed in the
ultrastructure of some saurian malaria sporozoites (Boulard
et al., 1983; Chapter 3) and the ability of saurian malaria
parasites to develop in several families of biting flies may
indicate primitive characters (Mattingly, 1965).
The criteria for vector incrimination were outlined by
Barnett (1960). While the mosquito vector-host relationship
-105-


-97-
sporoblastoid from the oocyst are not completely understood.
However, Terzarkis et al ( 1967 ) proposed that a change in
the ionic concentration in the subcapsular space could lead
to a net accumulation of fluid. In some Plasmodium sp.,
cytoplasmic subdivisions of the sporblastoid occurs when
vacuoles develop within the sporolastoid body which
eventually coalesce and form several sporoblasts.
Vanderberg et al ( 1967 ) reported that for P_. berghei the
cytoplasmic divisions were incomplete and formed clefts
which were joined to a central sporoblastoid. It is unclear
whether the sporoblastoid undergoes cytoplasmic division in
_P. agamae. While several sporoblasts or sporoblasts
attached to a central body are formed in most PIasmodium
species, only one sporoblastoid has been reported in oocysts
of Leucocytozoon and Haemoproteus (Sterling and Deguisti,
1974) .
Sporozoite development in P_. mexi canum is initiated by
the formation of a convex dense inner membrane along the
periphery of the sporoblastoid during or after vacuolization
of the oocyst has commenced and is similar to that observed
for other Plasmodium, Leucocytozoon, and Haemoproteus
parasites (Figures 25 to 29). Usually the pair of convex
membranes are only in oocysts where the subcapsular space is
nearly complete. However, in a few cases, the pair of
convex membranes are observed during the initiation of
vacuolization and where the oocyst membrane had separated
from the sporoblastoid (Figure 25). Although the convex


-172-
first to describe Schel 1 ack i a (. occi dental i s ) in the
western hemisphere, and like Reichenow, incriminated a mite
(Geckobiella texana) as the natural vector. In the western
hemisphere, three additional species of Schellackia in
lizards {S_. brygooi S^. 1 andaue and g o 1 v a n i ) and one
species of Schel1ackia in toads [S^ bal 1i LeBail and Landau
(1974)], have been described to date. Experimental
transmission of _S. brygooi and S^. 1 andaue has been
accomplished by ingestion of mosquitoes which bloodfed on
infected hosts (Landau, 1973; Lainson et al., 1976).
Transmission of S. bal 1 i and _S. go 1 vani to their respective
amphibian and saurian hosts has not been accomplished.
The present studies describe transmission of _S. go! van i
and S^. occi dental i s to A_. carol i nensi s and S£. undul atus ,
respectively, by ingestion of bloodfeeding arthropods which
had previously fed on conspecific infected lizards. The
prepatent period, development of the parasite, and
histological studies of the sporozoite in the vector and
lizard are reported.
Materials and Methods
Transmission Studies
Colonies of Lutzomyia vexator and Culex erraticus were
maintained as previously described (Chapter 2). Aedes
aegypti females were obtained from a colony maintained by


-198-
sporozoites were seen in the blood of several lizards at day
seven post-ingestion. Lizards were not maintained at higher
temperatures to determine if the prepatent period could be
shortened further. Temperature also affects the development
of other hemosporidia in poiki 1othermic animals. Thompson
and Winder (1947) demonstrated that there was a marked delay
in the peak parasitemia levels of Plasmodium floridense in
A. carolinensis and S£. undu 1 atus held at lower
temperatures. The behavior of the poikilothermic lizard in
maintaining body temperature in relation to ambient
temperature would greatly affect the prepatent period and
increase or decrease the potential for transmission.
The intensity of transmission of Schellackia largely
depends on the saurian host-vector contact, the period of
vector activity, the home range of the host, and flight
range of the vector. Some potentially suitable lizards, i.
e., Seeloporus woodi in Florida, may not serve as reservoir
hosts of the parasite because they bury themselves in sand
in the evening and are, thus, unavailable as bloodmeal
sources when most mosquitoes are biting. No naturally
infected Sc^. woodi (n = 2 01) (Schellackia o r PI asmodi urn) have
so far been found (Telford, 1978). Thompson and Huff
(1944b), however, reported the presence of an unidentified
sporozoan in S£. woodi. However, vectors which acquire
Schellackia from nearby hosts, i. e., Sc^. undul atus, may be
ingested by Sc_. woodi and thereby become infected (assuming
of course that Sc. woodi is susceptible).


CHAPTER 1
SAURIAN MALARIA AND SCHELLACKIA
Saurian Malaria
Species of PIasmodium, a hemosporidian genus of
protozoa that infect mammals, birds and reptiles, cause a
disease generally referred to as "malaria." Malaria in man
has been recorded in papyrus as early as 1500 BC in Egypt,
but the causative agent was not known until 1880 when
Laveran discovered PIasmodium parasites in human red blood
cells. The persistent search for malaria parasites in
mosquitoes by Ross (1897) eventually led to the discovery of
oocysts on the midgut of a "dapple-winged" mosquito
(Anopheles sp.) which fed on a human patient infected with
malaria. The vector-host-parasite relationship was
subsequently demonstrated for PIasmodium relictum Grassi and
Felitti (avian malaria) (Ross, 1898) and for human malaria
(Grassi et al., 1899). The cryptic exoerythrocytic stage
was demonstrated much later by Rafaelle (1934) (PIasmodium
-1-


Figure 12. Number of PIasmodium mexicanum trophozoites,
schizonts, gametocytes and percent of
infected white blood cells per 10,000 red
blood cells during the course of the
infection of Seeloporus undulatus (S- 2 5 )
infected by bite of Lutzomyia vexator.


-114-
and maintained in a temperature-hurn idity controlled
incubator at 25C and 80% RH. In the later part of the
investigation, the sugar solution contained 0.1% Poly-vi-sol
multivitamin syrup. Midguts were dissected at daily
intervals, beginning five days after feeding, and examined
for oocysts. Ten days after feeding, the salivary glands
were also examined and the sporozoite rate determined (+1,
1-10; +2, 11-100; +3, 101-1000; +4, >1000 sporozoites.
In addition, F ^ progeny of wild caught Cx^. territans
(collected in 1 izard-baited traps) and Lutzomyia vexator
(Chapter 2) were also provided bloodmeals on infected A.
carolinensis. Midguts and salivary glands were similarly
examined as for £x. erraticus.
Cu 1 ex erraticus that were potentially infective, i. e.,
those from lots of mosquitoes which developed oocysts and/or
sporozoites, were placed in the screened feeding cylinders
and provided a second bloodmeal on wild caught, noninfected
A. carolinensis. These lizards were determined to be
noninfected if they did not demonstrate any blood forms of
P_. flor i dense for more than 30-90 days. The noninfected
anoles were restrained similarly to that described for
infected bloodmeals. Following the second bloodmeal,
mosquitoes were removed, dissected, and the salivary glands
examined for sporozoites. Salivary glands with sporozoites
from mosquitoes that had taken second bloodmeals on
noninfected lizards were injected IP into other noninfected
lizards.


Figure 11. Number of PIasmodium mexicanum trophozoites,
schizonts, gametocytes and percent of
infected white blood cells per 10,000 red
blood cells during the course of the
infection of Seeloporus undulatus (S-15 )
infected by bite of Lutzomyia vexato r .


90. Numerous sporozoites of Schel1ackia occidental is
in a white blood cell of Sceloporus undulatus... 189
91. Section of intestine of Seeloporus undulatus
with a schizont of Schellackia occidentals 191
92. Section of intestine of Sceloporus undulatus
with an oocyst (?) of Schel 1 acki a occTdentlais.. 191
xv


-107-
Hatchet Creek separates into several channels which form
pools during dry periods. Portions of Hatchet Creek consist
of grassy margins and pools that intermittently flood during
heavy rains. A combination of deciduous, deciduous-pine,
and pine forests occur along the margins and length of
Hatchet Creek. The study site along the margin of the creek
was heavily deforested in 1982, and during 1983-5, consisted
primarily of young willows ($ a 1ix sp.). Adjacent to the
study site, several residences have permanent ponds of water
covered, or nearly covered, with floating plants (Lemna
sp.). A high incidence of P_. flor i dense was also noted in
lizards collected at these residences.
Vector attraction and bloodfeeding propensity for feral
biting arthropods was determined by the use of lizard-baited
traps in which the host lizard was exposed to the arthropods
(Figure 42). Traps were operated continuously for three
nights per week during the same time as the CDC light traps
from 30 April to 6 September (1984). Mosquitoes collected
in the traps were removed, brought to the laboratory,
counted, identified, and used for colonization. (See
section on colonization of Cj<. errati cus).
CDC light traps, operated from 1800 to 0800 hrs the
following morning, were used to determine seasonal changes
in the biting fly populations and to determine which species
were present at the Hatchet Creek study site. Mosquitoes
collected in the light traps were returned to the
laboratory. Those that were dead were removed from the


-35-
The progression of the acute infection can be explained as a
exponential linear relationship for parasitemia levels of
fewer than 500 parasites/10,000 RBC (5% parasitemia). In
general, the slopes of the acute infections of the lizards
with less than 5% parasitemia were similar. Except for
lizard S-47 (patent infection on day 23 post-exposure), S-51
patent infection on day 40 post-exposure), and S-43, the
slopes of parasitemia increase were not significantly
different (p=.01). Lizards S-42, S-43 and S-51 that
survived the longest and which had terminal parasitemias of
>70% (71.2, 87.7, and 119.6%, respectively) had the lowest
slopes (rate of increase in parasitemia) (Table 2).
However, subsequent to parasitiemias of >500 parasites per
10,000 RBC, the parasitemia increases at a reduced rate and
is better explained as a quadratic relationship. When
considering all lizards that died or were killed (and
probably would have died within a few days), there were
significant differences (p =.01) in the curves of the
quadratic equation. However, the curves are similar enough
to average over all lizards (Figure 7). A regression of
parasitemia over the course of the infection for all lizards
2
was performed (R =.88). The initial positive bloodfilm
(patent infection) was adjusted to begin on the mean day of
patent infection (28.6) since we were interested in the
average course of infection. The predicted parasitemia over
the course of the infection is shown by the dotted line
(Figure 7 ) .


-77-
Sporozoite formation. Figures 25 to 28 illustrate
differentiation of the sporoblastoid into sporoblasts and
the initial development of sporozoites. Differentiation of
the sporoblastoid and development of sporozoites is a
dynamic process within the oocyst and occur concurrently.
Differentiation of the oocyst is initiated by the formation
of irregularly-shaped vacuoles in the sporoblastoid body and
immediately beneath the oocyst capsule (Figure 25). A
subcapsular space is formed along the periphery of the
sporoblastoid as the sporoblastoid plasma membrane contracts
away from the oocyst capsule. Linear to convex dense
membranes are sometimes present along the sporoblastoid
membrane during vacuolization, but are more numerous
following complete separation of the sporoblastoid from the
oocsyt capsule (Figures 26 and 29). Large vacuoles form
within the sporoblastoid body (Figure 27). Coelescense of
the internal cytoplasmic vacuoles and their extension to the
sporoblastoid surface produces clefts which subdivide the
oocyst cytoplasm into several sporoblasts (Figure 28).
Figure 29 illustrates the initial sporozoite formation
which is characterized by dense inner thickened convex
membranes (= linear dense areas) which develops beneath the
sporoblast plasma membrane. Immediately inside and along
the newly condensed inner membrane, a single row of
microtubules is formed. The outer sporoblast plasma
membrane continues to evaginate and forms the outer
sporozoite membrane, while the denser inner membrane forms


-174-
which bloodfed on lizards infected with S. golvani were
dissected at daily intervals and observed for parasites.
Midguts with Schel1ackia parasites were teased apart, air
dried, fixed with absolute methanol, and stained with
Giemsa .
Since hatchling lizards were not available, wild caught
lizards that did not demonstrate either S_. occidental is or
. go 1 vani parasites in bloodfilms, and which were collected
from areas where < 2 % of the wild caught lizards had positive
bloodfilms for Schellackia parasites were used. Sand flies
and mosquitoes were force fed to Sc_. undul atus and A.
carolinensis from <1-29 days subsequent to bloodfeeding on
infected lizards. Lizards which were force fed one or more
infected sand flies or mosquitoes, were maintained in a
screened cage with a 40 watt lamp in the laboratory as
previously described, or else placed in a temperature
controlled environmental chamber at 32C (90F). Lizards,
serving as control groups, were collected from the same
locality as those which were force fed infected arthropods
and maintained under the same conditions as the
experimentally infected lizards. Bloodfilms were prepared
from all lizards which ingested infected flies at 3-4 day
intervals.
Mites (G_. texana), which were bloodfeeding on one Sc.
undul atus infected with S_. occidentals, were removed and
forcefed to other non infected Sc. undulatus. The bodies of
some of these mites were also teased apart, fixed in


-191-


-109-


Figure 13. Number of Plasmodium mexicanum trophozoites,
schizonts, gametocytes and percent of
infected white blood cells per 10,000 red
blood cells during the course of the
infection of Seeloporus undulatus (S-14)
infected by bite of Lutzomyia vexator.


collected in the trap. Culex territans is an early spring
mosquito, while Cx^. erraticus is abundant in the summer and
fall months when transmission of P_. flor i dense occurs.
Transmission of P_. f 1 or i dense was accomplished by bite of
Cx. erraticus and intraperitoneal injection of sporozoites.
Sporogony of P_. f 1 ori dense was demonstrated in L_. vexato r,
but sporozoites were rarely seen, and never in the salivary
glands. The prepatent period for P_. f 1 ori dense and P_.
mexicanum is relatively long (>20 days) at 18-24C.
PIasmodiurn mexicanum is very pathogenic, invading the spleen
and endothelial cells of the brain, in Sc_. undul atus, while
^P. floridense rarely kills A. carol i nensi s .
Sporogony of both saurian malarias is similar to that
described for other malarias. The gross morphology and
ultrastructure of P_. floridense sporozoites is similar to
other mammalian and avian malarias. However, P_. mexi canum
sporozoites are short, stout, and have a subpel1icular
microtubular arrangement which is different from other
malarias, but similar to P_. agamae, another saurian malaria.
Transmission of Schel1ackia parasites was accomplished
by ingestion of infected mosquitoes, sand flies, and mites
that previously fed on conspecific hosts, i. e., S_. gol vani
to A. carol i nensi s and S^. occi dental i s to Sc undul atus .
The prepatent period was reduced significantly by high
temperatures. At room temperatures, the prepatent period
for both Schel1ackia parasites was >20 days. However, at
32C, the prepatent period was decreased to as few as 7
days .
XVII


-26-
The F ^ progeny of two species of mosquitoes, Cu 1 ex
erraticus and Cu 1 ex territans collected in lizard baited
traps were also provided bloodmeals on S£. occidentals
infected with P_. mexi can um during the same time as l^.
vexator. None of the mosquitoes developed oocysts while all
of the vexator dissected had oocysts (range 9-54, mean
22.1) .
Patent JP. mexi canum infections were first observed in
nine of the experimentally infected Sc_. undul atus from days
23-40 post-exposure (mean 28.6 days) (Table 1 and Figure 5).
Because lizards were bled only every third day, infections
may have been patent as early as two days previous to the
positive bloodfilm. The acute infection was allowed to run
its course in each of six lizards. The remaining three
lizards were killed when they became anorexic and lethargic
and probably would have survived only a few days longer.
The six lizards which were not killed died of fulminating
infections by day 96 post-exposure and became lethargic and
anorexic several days prior to death. Force feeding two of
the lizards during this critical period did not appear to
increase the survivability of the lizards. The period of
survival for these lizards varied from 13-56 (mean 27.0)
days following the detection of parasites in the bloodfilm
and 39-96 (mean 57.8) days post-exposure. Two of the
longest living lizards, S-42 and S-51, were adult females
that deposited abnormal infertile eggs during the course of
the infection.


-94-
several days later. Since only a few sand flies were
dissected, it is unclear whether the low temperatures
destroyed the parasites. However, this may indicate that
the sporogonic temperature range of P_. mexicanum may be from
about 20C to 30C. It is believed that temperatures above
30C, even if it would not detrimentally affect the
parasite, would be detrimental to the longevity of the sand
fly. Although there were differences in the rate of
sporogonic development of P_. mexi canum maintained at 24 and
27C, there appeared to be no other differences.
Sporozoites from sand flies maintained at 27C were highly
infective to S_c. undul atus (Chapter 2).
Ultrastructure of Extrinsic Stages of Plasmodium mexicanum
The ultrastructure of the oocyst, formation of
sporozoites from the sporoblast, and sporozoites of several
species of mammalian and avian malaria have been described
by a number of authors (Garnham et al., 1960, 1961, and
1963; Sinden and Garnham, 1973; Duncan et al., 1960;
Terzarkis, 1971; Terzarkis et al., 1967; Vanderberg et al.,
1967). The first and only reported ultrastructure of the
oocyst, sporozoite development, and sporozoite of a saurian
malaria, P_. agamae in C u 1 i c o i d e s nubecul osus was described
by Boulard et al ( 1983). While there are many similarities
among the transformation of the oocyst and development of
sporozoites of £. mexicanum and other malaria species, there
are also some differences.


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