Supervisor: Dr. Terry W. Pearson
ABSTRACT
Procyclic culture forms of Trypanosoma brucei species and antibodies to these parasites were used in developing antibody-detection and antigen-detection assays for diagnosis of African human sleeping sickness. An agglutination assay using live procyclic
trypanosomes- the Procyclic Agglutination Trypanosomiasis Test (PATT) was developed for detecting anti-trypanosome antibodies in the sera of trypanosome-infected vervet
monkeys and humans. Antibodies to procyclic surface antigens were detected by the PATT in sera of vervet monkeys as early as 7 days post-infection with T. b. rhodesiense.
Positive agglutination titres were obtained with sera from monkeys with active, untreated infections and with sera taken soon after successful drag cure. Similar positive
agglutination results were also observed using the PATT with sera from T. b.gambiense- infected patients from Cote d'Ivoire and Sudan and with documented sera from T. b. rhodesiense—infected patients from Kenya. No agglutination reactions were observed with preinfection sera from vervet monk, ys, with sera from uninfected Canadians or with sera from Americans working in endemic areas. Together these results confirm the diagnostic value of using procyclic trypanosomes to detect anti-trypanosome antibodies in. human African sleeping sickness.
A double antibody sandwich ELISA using monoclonal antibodies and polyclonal rabbit antibodies to the surface membrane antigens of procyclic trypanosomes was developed. This assay detected circulating trypanosomal antigens in the sera of trypanosome-infected mice and in the sera from parasite-infected patients. However, limited success was obtained with this sandwich ELISA when tested on a larger repertoire of sera from infected humans. Rabbit antibodies made against whole lysates of T. b. rhodesiense procyclics were then employed in an antigen-trapping sandwich ELISA. The results demonstrated the effectiveness of this sandwich ELISA in revealing the infection status of vervet monkeys or humans infected with either T. b. rhodesiense or T. h.
gambiense. Trypanosomal antigens were detected in the sera of parasitologically confirmed monkeys and patients but not in preinfection sera nor in control sera from uninfected North Americans.
The PATT and the sandwich ELISA exhibited higher sensitivities than the currently employed diagnostic assay for human sleeping sickness, the Card Agglutination
Trypanosomiasis Test (CATT), when tested with sera of parasitologically-confirmed humans. The sandwich ELISA was superior to the antibody-detecting PATT and CATT in monitoring trypanocidal drug-treated patients. The overall sensitivity of the PATT and sandwich ELISA was 94.3% and 97.4% and the specificity was 84.5% and 95.5%, respectively. These results thus confirm the diagnostic value of these tests for the diagnosis of human African sleeping sickness.
Identification of diagnostically useful antigens was attempted in order to facilitate the adaptation of these diagnostic assays to a simpler format for field application. Pooled sera obtained from trypanosome-infected patients was used as a probe to detect
trypanosome antigens separated by high performance liquid chromatography,
immunoaffinity and immunoblotting techniques. Most of the antigens were detected in the higher molecular weight range (>62 Kd). Immunization of mice with the target antigens yielded six trypanosome-specific monoclonal antibodies. In a double antibody sandwich ELISA, these antibodies were successful in trapping circulating parasite antigens in sera from trypanosome-infected mice as early as 3 days post-infection. Some of these antigens have been partially biochemically characterized. Trypanosomal antigens were also detected by these antibodies in the urine of infected mice. The antigen-capture sandwich ELISA using either the selected monoclonal antibodies or the rabbit anti-procyclic whole lysate antibodies gave similar results with sera from trypanosome-infected mice, human sleeping sickness patients and uninfected humans from North America and Kenya. The results showed that these MAbs and their antigens were useful in the diagnosis of African human sleeping sickness.
Examiners:
Dr. T e rry ^ i Peaifs^, Supervisor lijche
(Department of Biochemistry and Microbiology)
Dr. William W.lCay, Departmental lum ber (Department of Biochemistry and Microbiology)
L
.
Dr.7Ro'SertWTOl^sm^ E&partmental Member (Department of Biophemis^and Microbiology]
Dr^Miphael J. Ashwood-Smith, Outside Member (Djepapment of Biology)
Dr. Robot D. Burice, Outside Member (Department of Biology)
Dr. Timothy Lee, External Examiner (University of Calgary)
TABLE O F CONTENTS
ABSTRACT... ii
TABLE OF CONTENTS ... v
LIST OF TABLES ... vii
LIST OF FIGURES... ix
ACKNOWLEDGEMENTS... xiii
FOREWORD... ... ... xiv
INTRODUCTION... 1
CHAPTER 1 Use of Procyclic trypanosomes in an antibody detection assay for African human sleeping sickness... 1
Introduction... ... 40
Materials and Methods... 42
R e su lts.,... 46
D iscussion... 60
CHAPTER 2 Detection of circulating trypanosomal antigens by double antibody sandwich ELISA using antibodies to procyclic trypanosom es... 62
Introduction... 62
Materials and Methods... 65
R esu lts... ... 73
D iscussion... 92
CHAPTER 3 Serodiagnosis of human African sleeping sickness by detection of anti-procyclic antibodies and trypanosome antigens 96 Introduction... 96
Materials and Methods ... 98
R esu lts ... 104
D iscussion... 143
CHAPTER 4 Identification of procyclic trypanosomal antigens that have serodiagnostic potential for human sleeping sickness... 149
Introduction... 149
Materials and Methods... 150
R esu lts... 166
DISCUSSION... 243
LIST OF TABLES
Table 1 Measurement of anti-procyclic surface antibodies in vervet monkey sera before and during infection with T. b. rhodesiense and at
various times after treatment with trypanocidal drugs... 52
Table 2 Total IgM and IgG levels in vervet monkey sera before and during
infection with T. b. rhodesiense and after drug treatment 56
Table 3 Cellular, biochemical, parasitological and serological
measurements on sera from African sleeping sickness patients
from Daloa, C6te d'Ivoire... 57
Table 4 Competitive solid-phase radioimmunometric assay of
anti-trypanosome monoclonal antibodies... 87
Table 5 Binding of biotin- or enzyme-labeled MAbs to trypanosomal antigens trapped by homologous or heterologous MAbs in double
antibody sandwich ELISA... 88
Table 6 Comparison of three different enzyme assay systems in indirect
E L ISA ... 89
Table 7 Stability of antibody coated microtiter plates and nitrocellulose paper in ELISA and dot-blot assays after storage at different time
intervals and temperatures... 90
Table 8 Detection of antigens or epitopes in bloodstream form and
procyclic culture form trypanosomes by ELISA... 91
Table 9 Measurement of anti-procyclic antibodies and circulating
trypanosomal antigens in vervet monkey sera before and during infection with T. b. rhodesiense and at various times after
treatment with trypanocidal drugs... 122
Table 10 Detection of anti-trypanosome antibodies and trypanosome antigens in sera of trypanosome-infected Kenyans before drug
treatment and at the time of relapse ... 126
Table 11 Measurement of anti-procyclic surface antibodies and circulating trypanosomal antigens in sera of trypanosome-infected patients
before and at various times after trypanocidal drug treatment 127
Table 12 Serological measurements on sera from African sleeping sickness patients from Daloa, Cote d'Ivoire, using the PATT, the CATT
and the double antibody sandwich ELISA... 129
Table 13 Cellular, biochemical, parasitological and serological
measurements on sera from trypanosome-infected Sudanese
Table 14 Cellular, biochemical, parasitological and serological
measurements c n sera from trypanocidal drug treated human
sleeping sickness patients from Sudan... 134
Table 15 Cellular, biochemical, parasitological and serological
measurements on sera from serologically positive (CATT) patients
from Sudan... 137
Table 16 Cellular, biochemical, parasitological and serological
measurements on sera from uninfected Sudanese, uninfected North Americans and human sleeping sickness patients from
Daloa, Cote d'Ivoire... 139
Table 17 Detection of anti-trypanosome antibodies and circulating
trypanosomal antigens in sera of trypanosome-confirmed,
trypanocidal drug-treated or serologically CATT positive Sudanese 141
Table 18 Measurement of antibodies and antigens in sera from patients with
different parasitic diseases... 142
Table 19 Binding characteristics of selected monoclonal antibodies to
detergent lysates of Trypanosoma species and Leishmania species. 228
Table 20 Detection of trypanosomal antigens by anti-PCF monoclonal
antibodies in trypanosome supernatants and pellets... 229
Table 21 The amino acid composition of immunoaffinity purified
trypanosomal antigen from MAb # 20 immunoadsorbent 230
Table 22 Amino terminal amino acid sequence of T. b. rhodesiense ViTat
1.1 PCF antigen recognized by monoclonal antibody # 2 0 ... 231
Table 23 The amino acid composition of immunoaffinity purified
trypanosomal antigens from MAb # 148 immunoadsorbent 232
Table 24 Amino terminal amino acid sequence of T. b. rhodesiense ViTat
1.1 PCF antigen recognized by monoclonal antibody # 148... 233
Table 25 Competitive solid-phase radioimmunometric assay of
anti-trypanosome monoclonal antibodies... 234
Table 26 Binding of biotin-labeled MAbs to trypanosomal antigens trapped
by homologous or heterologous MAbs in double antibody
LISTS OF FIGURES
Figure 1 The life cycle of Trypanosoma brucei species... 3
Figure 2 Transmission cycles involved in African human sleeping
sickness... 10
Figure 3 Maximum serum dilutions causing agglutination of
trypanosomes in the Procyclic Agglutination Trypanosomiasis
T est... 5 0
Figure 4 The effect of adding different biodnylated anti-7, h.
rhodesiense procyclic monoclonal antibodies on binding to
trypanosome procyclic water lysates in indirect ELISA 7 7
Figure 5 Optimization of double antibody sandwich ELISA for detection
of antigens in trypanosome water lysates or membranes 7 9
Figure 6 Detection of different solubilized extracts from Trypanosoma brucei rhodesiense bloodstream forms and procyclic culture forms using rabbit anti-PCF membrane polyclonal antibodies as 'capture' antibodies and biotinylated anti-TBRPl MAbs (247,346 and 477) as detector antibodies in a double antibody
sandwich ELISA... 8 1
Figure 7 Double antibody sandwich ELISA results and parasitemia data
for sera from 7. b. rhodesiense -infected mice... 8 3
Figure 8 Double antibody sandwich ELISA results and parasitemia data for the second set of sera from 7. b. rhodesiense -infected
m ice... 85
Figure 9 Detection of trypanosomal antigens in water lysates of
parasites by double antibody sandwich ELISA... 110
Figure 10 Summary of antibody and antigen detection tests with respect to the infection status of vervet monkeys infected with 7. b.
rh o d e sie n se ... 112 Figure 11 Anti-trypanosome antibodies and trypanosomal antigens in
sera of vervet monkeys No. 47 and No. 49 during 7. b.
rhodesiense infection... 114 Figure 12 Detection of anti-trypanosome antibodies and trypanosomal
antigens in sera from 10 Kenyan patients... 116
Figure 13 Measurement of anti-trypanosome antibodies and circulating trypanosomal antigens in sera from patients who relapsed
Figure 14 Detection of anti-trypanosome antibodies and circulating
trypanosomal antigens by the PATT and die sandwich ELISA in sera from eight patients who relapsed long after trypanocidal
drug treatm ent... 120
Figure 15 Separation of T. b. rhodesiense ViTat 1.1 PCF crude lysate by gel permeation chromatography using a Sephacryl S-200
colum n... 174
Figure 16 Ion-exchange-HPLC profiles of pooled fractions # 40-50 from
G PC ... 176
Figure 17 Separation of fractions # A-D from the ion-exchange HPLC
column by a reducing 5-15 % gradient SDS-PAGE gel 178
Figure 18 Reverse phase HPLC profiles of pooled fractions (A, B and C)
from the ion-exchange HPLC column... 179
Figure 19 Antigen profiles detected by pooled HSSS from Daloan patients (Cote d'Ivoire) in various parasite lysates using
immunoblotting ... 181
Figure 20 Antigen profiles detected by pooled HSSS from Kenyan
patients in various parasite lysates using immunoblotting 132
Figure 21 SDS-PAGE silver stain profiles of T. b. rhodesiense ViTat 1.1 PCF eluted from an immunoaffinity column coupled to pooled
immunoglobulins from HSSS from Kenyan patients 183
Figure 22 Silver stained SDS-PAGE gel patterns cf extracted gel
fractions used for immunization of BALB/c mice... i 84 Figure 23 Immunofluorescence patterns of formaldehyde-fixed or
acetone-permeabilized Trypanosoma and Leishmania species
using anti-PCF MAb # 20... 185
Figure 24 Immunofluorescence patterns of formaldehyde-fixed or acetone-permeabiiized Trypanosoma and Leishmania specier
using anti-PCF MAb # 236... 187
Figure 25 Immunofluorescence patterns of formaldehyde-fixed or acetone-permeabilized Trypanosoma and Leishmania species
using anti-PCF MAb # 65... ... 189
Figure 26 Immunofluorescence patterns of formaldehyde-fixed or acetone-permeabilized Trypanosoma and Leishmania species
using anti-PCF MAb # 401, 148 and 91... 191
Figure 27 The Trypanosoma specific antigen profile recognized by MAb
Figure 28 Detection of antigen in whole lysates of Trypanosoma and
Leishmania species by inrimunoblotting using MAb # 148... 194 Figure 29 Detection of antigen in whole iysates c f Trypanosoma and
Leishmania species by immunoblotting using MAb # 236... 195 Figure 30 Detection of antigen in whole lysates of Trypanosoma and
Leishmania species by immunoblotting using MAb # 6 5 ... 196 Figure 31 Detection of antigen in whole lysates of Trypanosoma and
Leishmania species by immunoblotting using MAb # 9 1 ... 197 Figure 32 Detection of antigen in whole lysates of Trypanosoma and
Leishmania species by immunoblotting using MAb #401... 198 Figure 33 Analysis of antigens Tecognized by the six selected and-FCF
monoclonal antibodies after heat, protease and chemical
treatm ent... *... 199
Figure 34 Enzyme-linked immunosorbent assay of T. b. rhodesiense ViTat 1.1 PCF after passing over Con-A agarose using MAb
# 20 , 201
Figure 35 Enzyme-linked immunosorbent assay of T. b. rhodesiense ViTat 1.1 PCF after passing over Con-A agarose using MAb #
65... ... 203
Figure 36 Extraction of trypanosomal antigens by various solubilization
b u ffers... 205
Figure 37 Enzyme-linked immunosorbent assay on T. b. rhodesiense ViTat 1.1 PCF lysates: eluted fractions from the MAb # 20
im m unoadsorbent... 208
Figure 38 Silver stained SDS-PAGE profile of ELISA positive, affinity
purified fractions from the MAb # 20 immunoadsorbent 210
Figure 39 Enzyme-linked immunosorbent assay on T. b. rhodesiense ViTat 1.1 PCF lysates: eluted fractions from the MAb # 148
im m unoadsorbent... 211
Figure 40 Silver stained SDS-PAGE profile of ELISA positive, affinity
purified fractions from the MAb #148 immunoadsorbent 213
Figure 41 Binding of different biotinylated anti-trypanosome monoclonal antibodies to trypanosome procyclic water lysates in indirect
E LISA ... 214
Figure 42 Detection of trypanosomal an'igens in water lysates of
Figure 43 ELIS measurement of trypanosomal antigens in sera and
urine from T. b. rhodesiense -infected mice... 218
Figure 44 ELISA measurement of trypanosomal antigens in sera and urine from T. b. rhodesiense -infected mice using rabbit ;ir.ti-
TBRP1 whole lysate antibodies... 220
Figure 45 Immunoblotting profiles of MAb # 148 on various T. brucei
lysates separated on a reducing 10% SDS-PAGE gel 27" Figure 46 Immunoblotting profiles of MAb # 148 on various T. brucei
species lysates separated on a non-reducing 10% SDS-PAGE
gel... 223
Figure 47 Detection of Trypanosomal antigens in parasitologically-confirmed sleeping sickness sera from Kenyan patients using rabbit anti-trypanosome whole lysate antibodies or MAb
m ix tu re... 224
F gure 48 Detection of Trypanosomal antigens in uninfected human sera from Kenyans (- ve controls) using rabbit anti-trypanosome
ACKNOWLEDGEMENTS
I am grateful to Dr. ferry Peaison for his inspiring supervision throughout this project. My thanks to the staff of the Department of Biochemistry and Microbiology, specifically to Robert Beecroft, Jennifer Duggan, Armando Jardim, Sandy Kielland, dbert Lubossiere and Scott Scholz, for their advice and assistance during my study. I would also like to the thank my Doctoral Dissertation C om m its: Drs. Michael Ashwood-Smith, William Kay, Robert Olafson and Robert Burke for their positive input.
My special thanks to Dr. Michael Clarke for his support and encouragement at the commencement of my graduate studies as well as to Chrystal McNabb for her assistance during one 'long' summer. I am indebted to Dr. Jennifer Richardson for kindly providing the anti-TBRPl hybridomas; to my collaborators: Drs. Paul Sayer, Tony Vervoort, Bruce Wellde and Pierre Cattand for providing sera from vervet monkeys and humans used in this study; to the B. C. government and the University of Victoria for financial aid in the form of scholarships; and to the International Development Research Center of Canada and the
»
UNDP/World Bank/WHO Special Programme for Research and Training in Tropical Diseases for funding tins project.
I am most grateful for the unfailing support of my family: Bill Conconi, Ellie Conconi, Flo Conconi and especially Diarmaid 6 Foighil.
is a widely misunderstood branch of secret operations. Its pur pose is not to apprehend enemy agents; that is an aim of the security forces. It is the word 'counter' which causes the trouble, since it is generally interpreted to mean 'against'; a defensive operation against the enemy's intelligence
operations. Quite to the contrary, CE is an offensive operation by using- or, more usually, attempting to use- the opposition's operations The ultimate goal of all CE operations is to penetrate the opposition's own secret operations apparatus; io become, obviously without the opposition's knowledge, an integral and functioning part of their calculations so far as intelligence is concerned, you know what he knows. You have thereby annulled, in one stroke, the value to him of his secret intelligence about you you are iu position to control his action.*
Introduction
African trypanosomiasis is a complex of diseases caused by trypanosomes, a taxon of flagellated protozoan parasites. Trypanosomiasis occurs throughout the tropical regions of Africa and the parasites are cyclically transmitted by tsetse flies (Glossina spp.; ILRAD, 1986) to a variety c f vertebrate hosts. Different species of trypanosomes infect a great variety of feral mammals and also domesticated species including cattle, sheep, goats, pigs, horses and camels. Some also infect humans. Trypanosomiasis in domesticated African animals is caused by Trypanosoma congolense, T. vivax, T. brucei brucei, T. simiae and T. evansi. In humans, African trypanosomiasis is also known as "sleeping sickness" and the infective organisms are T. b. rhodesiense and T.b. gambiense (Hoare, 1970).
Collectively, these diseases have severely hindered the economic and social development of Sub-Saharan Africa (WHO, 1979).
Presently, sleeping sickness is endemic in 36 African countries, coinciding with the geographic range of the tsetse fly vector. Recent estimates suggest that 50 million people are at risk of infection (Maurice and Pearce, 1987). Until 1979, there were approximately 10,000 new cases recorded per annum, but recent serious outbreaks in Cameroon, Sudan i- id Uganda have at least doubled the infection rate (Goodwin, 1985). Present estimates are probably low because of inadequate reporting (WHO, 1986). Sleeping sickness is often fatal if left untreated and has led to the depopulation of many parts of Africa since the beginning of the century (WHO, 1979). The recent Ugandan epidemic and outbreaks in the southern Sudan since the 1970's result from the breakdown in implementation of diagnostic and treatment facilities brought about by civil unrest (Goodwin, 1985). In the past 20 years, however, increased research funding has led to significant progress in the development of diagnostic and therapeutic technology and in our understanding of the complex molecular biology of trypanosome-mammalian host interaction. Nevertheless, no permanent solution, such as the development of a vaccine or the elimination of the vector, can be expected in the near future (WHO, 1986). Because of this, the development of effective diagnostic tests and curative therapies continue to hold the most short-term promise in controlling sleeping sickness in Africa (Maurice and Pearce, 1987).
Historical Perspective
Human sleeping sickness was first described in the 14th century by the Arab physician A1 Qualquashaudi (Hoeppli, 1959). However, it was not until the late 19th century that the importance of trypanosomes in this disease were recognized by Lewis and Evans in India and Bruce in Africa, all of whom gave their names to species of
trypanosomes (Hoare, 1972; Goodwin, 1985). The causative agents of human African sleeping sickness, T. b. rhodesiense and T.b. gambiense, were first described in 1902 and 1903, respectively ( Hoare, 1972). Endemic foci of sleeping sickness in Western Africa and the Congo River Basin were mapped by European colonizers in the late 19th century. Sleeping sickness has been documented to the east of the African Rift Valley and in the Zambesi Basin only within the last 70 years. This does not necessarily imply that the disease originated in West Africa. The East African form of sleeping sickness caused by T. b. rhodesiense is distinct from the West African form and its relatively diffuse distribution may have prevented early observers from pinpointing endemic foci (Duggan, 1970). More recent human population migrations caused by climatic or socio-economic factors have facilitated the spread of sleeping sickness from its ancient foci to other regions of Africa (Duggan, 1970).
Developmental Cycles Of African Trypanosomes
Human-infecting Trypanosoma brucei are cyclically transmitted by tsetse fly vectors (Glossina spp.) to their mammalian hosts (Ormerod, 1976) (see Fig. 1). The parasites adapt to these distinct environments (fly and mammal) by unde going a series of
morphological and metabolic metamorphoses. Three main developmental stages occur in the parasite life cycle. In the mammalian host the parasites occur predominantly in the blood, hence this form is termed the bloodstream stage. The parasite resides in two sites in the tsetse fly: the midgut and the salivary glands, each with a distinct developmental stage, respectively termed procyclic and metacyclic forms. During developmental
metamorphoses, the most dramatic changes occur in the mitochondrial system and the surface membrane of the parasite (Vickerman, 1971). Both can be related to the survival mechanisms of the parasite in its different hosts. The parasite switches from a total reliance on glycolysis for its source of metabolic energy in the mammalian host to a cytochrome- mediated metabolism in the insect midgut (Vickerman et al., 1988). Alterations in
MET ACYCLIC
LONG SLENDER BLOODSTREAM FORMS
EPIMASTIGOTE
SHORT STUMPY BLOODSTREAM FORMS PROCYCLIC
molecules of the plasma membrane allow the flagellate to evade the mammalian host's specific and non-specific defense mechanisms (Vickerman and Barry, 1982). Other developmental changes occur in the parasite's endocytotic apparatus (Steiger, 1973) and in its giycosomes (Opperdoes and Borst, 1977). During its life cycle the parasite alternates between proliferative phases when it undergoes multiplication and non-proliferative periods in which it is incapable of cell division. The latter are associated with major transitions in environment, the former with the establishment of the parasite in a newly acquired environment (Vickerman, 1985).
Infection of the mammalian host is initiated by the bite of a parasitized fly vector which results in the deposition of tsetse saliva containing metncyclic trypanosomes into the dermal connective tissue of the mammal (Hoare, 1972). Parasites then enter the draining lymphatic vessels and from there access the host bloodstream where they multiply with a doubling time of circa 6 hours (Seed, 1978) as long slender bloodstream forms. These bloodstream trypanosomes display two significant specializations to their mammalian habitat. Each individual's single mitochondrion is non-functional (Vickerman, 1965) and the parasites obtain their energy solely from glycolysis which occurs in a special organelle called the glycosome (Opperdoes, 1985, ±987). Bloodstream trypanosomes survive in the mammalian host by utilizing r unique immune evasion tactic, antigenic variation (Gray,
1965; Tanner et al., 1980). Each parasite is ensheathed in a 12-15 nm thick surface coat composed of a single form of glycoprotein, the variant surface glycoprotein (VSG), (Vickerman, 1969; Cross, 1975). Upon infection, the host's immune system responds primarily to this surface antigen. However, trypanosomes, as a population, can evade the host's immune surveillance mechanism by successively changing to different and unique surface coats, a phenomenon known as antigenic variation (Turner, 1984, Haduk e t al.,
1984). This antigenic switching results in a fluctuating parasitemia that often characterizes a trypanosome infection. Dividing slender parasites predominate in the ascending
parasitemia and a particular antigenic type, called the homotype, forms the major part of the population (Van Meirvenne e t al., 1975). The parasitemia goes into remission as
trypanosomes of the predominent variable antigenic type (VAT) are eventually removed by the specific host immune response to that homotype (Seed, 1977, Hajduk and Vickerman, 1981a). However, the few slender trypanosomes that undergo antigenic variation
temporarily evade the host immune response. Some of these cells transform into a non dividing morphologically distinct "stumpy" form and the parasitemia declines. This change from slender to stumpy bloodstream trypanosome coincides with the activation of
mitochondrial enzymatic pathways (Vickerman, 1965). Stumpy forms may be preadapted for further differentiation into the procyclic stage in the tsetse midgut once they are imbibed when a fly feeds on an infected mammalian host. Repeated syringe passage of bloodstream form T. brucei among mammalian hosts results in the formation of slender forms
throughout the infection period (Ashcroft, 1960), and such trypanosome populations are described as being monomorphic. Tsetse flies feeding on rodents infected with
monomoiphic strains of T. brucei are not capable of reinfecting other rodents, suggesting that the slender to stumpy differentiation is necessary for trypanosome survival in the insect and, hence, completion of the life cycle (Ashcroft, 1960; Hadjuk and Vickerman, 1981).
Trypanosome-infected blood is ingested by the tsetse into its crop and from there passes to the lumen of the midgut, which is the site of the transformation from stumpy bloodstream to procyclic insect form (Vickerman, 1985). Most ingested slender
bloodstream forms are probably unable to develop into procyclics and thus die (Wijers and Willett, 1960). Transformation takes place in the posterior part of the midgut in the
endoperitrophic space, i.e., inside the peritrophic membrane that separates the bloodmeal from the midgut epithelium (Evans and Ellis, 1983). Procyclic development is associated with a loss of the VSG coat (Turner et al., 1988), cessation of endocytosis (Steiger, 1973) replication of the kinetoplast-DNA network (Hajduk etal., 1984), a switch from the utilization of glucose to ptoline as a principal energy source (Evans and Brown, 1972; Bowman and Flynn, 1976; Bienen et al., 1981), complete activation of the mitochondrion (Brown et al., 1973) and a change in glycosome morphology (Hart et al., 1984). These changes occur over a 48-72 hour period in the tsetse gut and are accompanied by active division of parasites. Comparable transformations occur in vitro when bloodstream forms are grown in various culture media at 26 °C (Brun and Jenni, 1985), but the time course is often shorter (Brown et al., 1973). Cultivated procyclic trypanosomes resemble their midgut counterparts in their morphology (Steiger, 1973), metabolism (Brown et al., 1973), antigenicity (Honigberg et al., 1976; Richardson et al., 1986), protein patterns (Pearson et al., 1987) and by il.rir capacity to produce mature infections in the tsetse fly (Evans,
1979).
Procyclic trypanosomes penetrate the peritrophic membrane of the tsetse gut to access the ectoperitrophic space (Evans and Ellis, 1983). As the parasites migrate to the proventriculus, they cease cell division and develop into elongate mesocyclic forms. The parasites again cross the peritrophic membrane and migrate via the oesophagus, proboscis
lumen and hypopharynx to the tsetse's salivary glands. Mesocyclic trypanosomes then attach to the microvilli of the salivary gland cells using punctate junctional complexes and resume cell division as attached epimastigote forms (Tetley and Vickerman, 1985). These events are followed by a series of morphological transformations: epimastigotes (lacking the VSG coat) lead to uncoated premetacyclics, which give rise to VSG-coated nascent metacyclics and eventually produce VSG-coated, free-swimming metacyclic trypanosomes in the salivary gland lumen (Tetley et al., 1987). Production of the VSG coat in
metacyclics is accompanied by the repression of mitochondrial metabolic activity, reversion of glycosomes to form spherical organelles and the cessation of cell division (Vickerman et al., 1988).
The developmental cycle within the tsetse vector takes 3-5 weeks to complete (Vickerman et al., 1988). When innoculateti into the new environment of the mammalian host by the feeding tsetse fly, metacyclic trypanosomes transform into proliferating long slender bloodstream forms and reestablish the mammalian infection (Evans and Ellis, 1983). Whether or not the trypanosome life cycle is at all stages programmed in one direction, or if certain stages can be omitted, are still open questions. However,
accumulating evidence on the transcription of the maxicircle component of kinetoplast DNA during the slender > stumpy > procyclic form transformations suggest that this
developmental sequence is unidirectional. Stuart (1987) and Michelotti and Hajduk (1987) found that the 9S and 12S mitochondrial ribosomal RNA levels are 30-fold lower in slender forms than in stumpy bloodstream parasites. Transcripts from 3 other mitochondrial genes: cytochrome B and subunits I and II of cytochrome oxidase are undetectable in slender bloodstream trypanosomes but are present in stumpy forms at a similar level to that of procyclics. However, the stumpy transcripts are not translated until metamorphosis to the procyclic stage occurs. These results imply that stumpy forms are preadapted to the tsetse midgut. Recent identification of developmentally regulated proteins that appear during the course of differentiation from bloodstream to procyclic stages
provide additional evidence that the trypanosomal developmental cycle may be indeed unidirectional (Richardson et al., 1988; Roditi etal., 1989b; Varner et al., 1989).
There is considerable interest in identifying the signals that trigger the large number of developmental stage transformations in the trypanosome life cycle. According to
Czichos and co-workers (1986), a 10 *C temperature drop (37 *C to 27 *C) and the addition of 3 mM cis-aconitate to minimal essential medium (plus 15% heat-inactivated
transformation of bloodstream to procyclic trypanosomes. A role has been implicated for lectins in inducing developmental changes in the life cycle stages that parasitize the tsetse vector (Maudlin and Welbum, 1987,1988). Black et al. (1985) suggested that mammalian host factors may prompt the slender to stumpy transformation of bloodstream
trypanosomes. The recent identification of an epidermal growth factor receptor homologue in T. brucei has raised the possibility that growth factor interactions similar to those found in m am m alian cells are involved in cell growth regulation in these parasites (Hide et al.,
1989).
It was long believed that trypanosomes replicated solely by binary fission (Vickerman, 1985). However, the existence of a sexual stage in the T. brucei life cycle was implicated by isoenzyme studies of natural parasite populations (Gibson et al., 1980; Tait, 1980,1983), by direct analysis of DNA complexity vborst et al., 1980) and content (Borst et al., 1982), by restriction site polymorphisms (Gibson et al., 1985) and by general protein gene product polymorphisms (Anderson et al., 1985). Genetic recombination in T. brucei was confirmed by the production of hybrid progeny when two genetically marked clones were co-transmitted through the tsetse vector (Jenni et. al., 1986; Paindavoine etal.,
1986; Wells et al., 1987; Sternberg et al., 1988; Pearson and Jenni, 1989). In contrast to the malaria parasite, sexual fusion does not appear to be an obligate part of the
trypanosome's development in the insect vector (Sternberg et al., 1989).
Although human tryy .osomiasis is a vector-borne disease, the transmission cycles are subject to a variety of interacting factors (biotic as well as abiotic) which influence the dynamics of transmission. Owing to the multiplicity of determinants that apply to the host, vector, and parasite, transmission cycles are inevitably complex and subject to
environmentally imposed changes. In addition to the trypanosome-infected person that can serve as parasite reservoir, domestic and wild animals can also be reservoir hosts of both T. b. gambiense and T. b. rhodesiense The zoonotic nature of endemic T. b. rhodesiense. disease has long been recognized. Heisch et al. (1958), in an ethically questionable
experiment, isolated T. b. rhodesiense from a naturally infected bushbuck which on inoculation into humans produced the sleeping sickness disease. Human pathogenic T. b. rhodesiense has also been isolated and identified from domestic cattle, sheep, dogs and goats (Rickman and Robson, 1970; Hawkin, 1975; Herbert etal., 1980). With the advent of new methods of characterizing trypanosomes, such as isoenzyme electrophoresis and
DNA analysis, parasites similar or identical to T. b. gambiense have been demonstrated in domestic animals (dogs, cattle, sheep and pigs) and game animals (kob and hartebeest) in areas endemic for human sleeping sickness (Maurice and Pearce, 1987). The prevalence of these animal trypanosome infections is difficult to estimate, since animal parasitemaia can be low with long parasitemic phases. Wild animals and some domestic animals such as cattle and pigs (Mehlitz et al., 1982) are likely important animal reservoirs for human pathogenic trypanosomes because they are usually said to be 'trypanotolerant', i.e. they do not develop the clinical signs of trypanosomiasis (Olubayo, 1978). The transmission cycles in T. b. gambiense and T. b. rhodesiense diseases thus involve man-fly-man, game- fly-man and domestic animal-fly-man cycles (Fig. 2), further complicating the
epidemiological control of African human sleeping sickness.
Pathology and Immunology of African Sleeping Sickness
There are two distinct forms of African sleeping sickness in humans, each due to a subspecies of Trypanosoma brucei (Apted, 1970a). The Gambian form is caused by T.b. gambiense and the Rhodesian or East African form is caused by T.b. rhodesiense. The subspecies are morphologically indistinguishable. However, their presence in an infection can be inferred by the geographic location of the disease and by its clinical course (De Raadt and Seed, 1977). Gambian sleeping sickness is distributed mainly in Western and Central Africa and is a chronic disease which patients normally survive for 2-3 years without treatment. By contrast, the Rhodesian form is found predominantly in Eastern and Southern Africa and is an acute disease usually ending in death within 2-6 months of infection (Brown, 1983). In the absence of chemotherapy both forms of African sleeping sickness are generally fatal in humans (Poltera, 1985).
An infection can be divided into three stages according to its clinical manifestations. Initially, the parasites remain localized at the site of the tsetse bite (initial stage) producing a chancre. Subsequently, the trypanosomes become widely distributed by the host's
bloodstream circulation (systemic stage) and the host displays symptoms of recurrent parasitemia. Finally the parasite invades the central nervous system and other organs (advanced stage) inducing heart and nervous system irregularities in the host (Molyneux et al., 1984; Shapiro and Pearson, 1986).
Fig. 2
Transmission cycles involved in human African Sleeping sickness (Adapted from Maurice find Pearce, 1987).
Hranan-tishu; oaa cycle % ' $ U ' r * * ^ 0 ' * - r ' V '‘ ’S' , * ' H '"" " a . , v < v V * ' ' V j t J
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The first sign of a trypanosome infection is often a chancre or lesion at the site of the tsetse bite (Fairban and Godfrey, 1957: Basson et al., 1977). The chancre represents a combination of an acute inflammatory response and an immune reaction to the locally proliferating trypanosomes. It exhibits marked oedema and an intense cellular reaction which appear 5 15 days after the tsetse bite and lasts for approximately 2 weeks. Infiltration of polymorphonuclear leukocytes, small lymphocytes, lymphoblasts and macropha ges into the infection focus comprise the initial host reaction (Shapiro and Pearson, 1986). The frequency with which this chancre occurs in infections has been the subject of some controversy. It is more commonly observed in infected Europeans than in Africans and also in T.b. rhodesiense infections than in T.b. gambiense infections
(Duggan and Hutchinson, 1966; Gelfand, 1966).
Trypanosomes that escape the initial host response at the chancre will migrate via local draining lymph nodes and the lymphatic system into the bloodstream (Ssenyonga and Adam, 1975). Antigenic variation is used by the bloodstream trypanosomes to evade the host's immune surveillance. This results in fluctuating parasitemia accompanied by symptoms such as intermittent fevers, headaches, joint pains, splenomegaly and
lymphadenopathy (Apted, 1970a; Greenwood and Whittle, 1980). Although swelling of the posterior cervical lymph nodes is fairly characteristic of a T.b. gambiense infection, most other symptoms occur only periodically, coinciding with the increase of parasite numbeis in the circulation (Molyneux et al., 1984).
As the infection progresses, par asites migrate from the vascular system into the interstitial fluid of many organs, especially the heart and the central nervous system (W€ry et al., 1982). Dysfunction of hean, liver, lungs, kidneys and the endocrine system is frequently observed at this, the advanced stage of the infection (Apted, 1970a, 1980; Poltera etal., 1976,1977; Bafort and Schmidt, 1983; Wellde etal., 1989a). Typical histopathological features include vasculitis of the capillary vessels (Poltera and Sayer,
1983) and organ lesions which are associated with granular immunoglobulin deposits and with a marked cellular infiltration (Poltera and Cox, 1977). Lesions in the central nervous system (CNS) progressively lead to a variety of clinical manifestations: headaches,
irritability', tremors, ataxia, convulsions, personality changes, somnolence, pronounced wasting and coma (Poltera, 1985). Once the trypanosomes access the CNS, death is inevitable if the patients do not receive prompt chemotherapy, which at this late stage is not always effective.
During the progress of the disease a state of immunodepression deveiops. As a result, the cause of death is sometimes a secondary infection such as pneumonia (Brown,
1983; Shapiro and Pearson, 1986). Profound immune system changes occur in trypanosome-infected patients. The immune system plays important roles in both the attempted control of the parasite and the pathogenesis of the disease state. An intense immunoproliferative reaction develops in most lymphoid organs during the initial and systemic stages. This is followed by, in the murine system at least, a general
immunosuppression in the advanced stage which affects antibody responses as well as T- cell mediated immune responses to both the parasite and non-parasite-relaied antigens (Mansfield, 1981; Bancroft and Askonas, 1982). Hosts succumb in the long term to either very heavy parasite loads or else to secondary infections to which they become more susceptible due to their weakened irrunune function. The extent of immune disruption varies with the virulence of the trypanosome clone (Sacks et al., 1980).
Trypanosomes stimulate an intense proliferation of B- and T-cells, null cells and macrophages during the early course of an infection. B-cell proliferation occurs in the lymph nodes, bone marrow and spleen, resulting in a striking increase in levels of
circulating IgM (Clarkson, 1976). Trypanosome whole lysates or membrane fractions can also elicit this reaction (Clayton et al., 1979). Some of the IgM production is non-parasite specific and may result from activated host macrophages producing factor(s) mitogenic for lymphocytes (Sacks et al., 1982). Polyclonal activation and the parasite-specific antibody response have been implicated in some aspects of the pathogenesis of trypanosomiasis. Antibodies (Abs) may play a role in causing the haemolytic anemia during the systemic stage of infection and in the immune-complex dysfunction and autoimmunity characteristic of advanced infections (Mansfield and Kreier, 1972; Lambert et a., 1981). Studies with nude mice, incapable of IgG production, indicate that the IgM isotype alone is effective in controlling parasitemia (Campbell et al., 1978). Antibodies produced against the surface antigen coat of trypanosomes are responsible for the elimination of these parasites by a combination of complement-mediated lysis (Murray and Urquhart, 1977) and antibody- dependent phagocytosis (Greenblatt et al., 1983; Ngaira et al., 1983).
During the infection, antibodies are also produced against the, trypanosome invariant plasma membrane components, nuclear and cytoplasmic constituents. A pathological effect is suspected for these Abs. Infected athymic mice do not produce Abs to trypanosome
non-variant molecules and do not exhibit the tissue damage and immune-complex symptoms produced in thymic-intact controls (Mansfield, 1990). In addition to parasite- specific Abs, auto-antibodies against the host's self-antigens such as complement C3, fibrinogen-fibrin degradation products (Boreham and Facer, 1974) and tissue antigens such as liver, thymus and brain (Mansfield and Kreier, 1972; Poitera et al., 1980) are also generated during polyclonal B-cell activation. Heterophile, or anti-eiythrocyte Abs and anti-DNA Abs have also been reported in cases of human trypanosomiasis (Parratt and Herbert, 1976). Although the precise contribution of tnese diverse autoantibodies to pathogenesis is unclear, they undoubtedly play some role in part of the disease syndrome, including haemolytic anemia and coagulation impairment (Parratt and Herbert, 1976).
Infection of the central nervous system (CNS) during the advanced stage of ;he disease is usually associated with a significant increase in the cerebrospinal fluid of IgM levels, lymphocytes and immune complexes formed by parasite variant antigens and Abs produced against them (Green wooc und Whittle, 1973; Lambert et al., 1981). If deposited in the brain, the immune complexes can potentially damage capillaries and lead to localized oedema (Lambert et al., 1981). Autoimmune responses in the CNS have been suggested to occur during trypanosome infections, but the inconsistent evidence for anti-neuronal
anti) xxiies in experimentally infected mice (Pentreath,1989) suggests that other factors are involved in neuronal damage. Astrocytes are one of the most numerous cell types in the CNS and are closely associated with the CNS tissue/blood system interface (McCarron et al., 1985). Expressionof astrocyte surface-exposed Class II major histocompatibility antigens (Ia-determinants) is induced by activated T-cells that cross the blood/brain barrier during an infection (Wekerle et al., 1987). Pentreath (1989) hypothesizes that la-induced astrocytes can serve as antigen-presenting cells. Once trypanosomes invade the CNS, astrocytes may interact with the activated T-cells to produce cytokines. These cytokines may, in turn, induce the proliferation of lympho cytes, astrocytes or microglia, and provoke a concomitant R-cel1 response and antibody secretion. The cerebral manifestations of the disease may depend on the inflammation sites, the inteiaetions between parasites,
antibodies and astrocytes, a id on the levels and types of mediator release at these sites (f entreath, 1989).
Although levels of circulating IgM remains high auring a chronic infection and some IgM against novel trypanosome antigens continues to be produced (Luckins and Mehlitz, 1976; Sacks and Askonas, 1980), the ability to mount effective, specific antibody
responses to new trypanosomal antigens decreases with time (Oka et al., 1984). Various mechanisms ha /e been proposed to explain the immunosuppression phenomenon in
advanced stages of infection. Evidence for the involvement of suppressor cells (Pearson et al., 1978; Jayawardena et al., 1978; Kar etal., 1981, Yamamoto etal., 1985), soluble suppressor substances (Tizard et al., 1978) and differential macrophage activity (Paulnock et at., 1988) have been presented. Theories of "clonal exhaustion" (Askonas et al., 1985) or "down-regulation" via an anti-idiotypic network (Sacks, 1984) have also been proposed to account for the immune system hyporesponsiveness observed during trypanosome infections. Interestingly, drug-cure of an infected immunosuppressed host does lead to rapid recovery of immune responsiveness (Roelants et al., 1979; Clayton et al., 1980). Indeed, immunosuppression in these hosts can be reversed within a few days post treatment (Askonas, 1985).
Trypanosomal Antigens
The antigenic profile of African trypanosomes varies throughout their complex life cycle and involves many different parasite molecules. One group of antigens, the VSGs have received much research attention due to their central role in antigenic variation in the mammalian host. A considerable amount of knowledge has accumulated about VSG structure, synthesis and molecular genetics (Turner, 1982; Donelson, 1988). In contrast, non-VSG antigens in these parasites have been less well studied. Although their existence has long been known, only a few non-VSG antigens have recently been biochemically characterized. For example, there are a Trypanosoma brucei species-specific 30-40 kDa glycoprotein, procyclin (Roditi et al., 1987; Richardson et al., 1988) and a giycosyl- phosphatidylinositol-specific phospholipase C of Trypanosoma brucei with an apparent molecular mass of 37-40 kDa (Bulow and Overath, 1986; Hereld et al., 1986; Fox et al.,
1986). Expression of these molecules in the bloodstream and procyclic stages of trypanosomes is developmentally regulated (Richardson et al., 1986; Roditi et al., 1989; Carrington et al., 1989).
a) Variant Surface Glycoproteins
Franke (1905) was the first to infer immunological shifts in African trypanosomes parasitizing mammalian hosts. He observed that monkey blood, removed 2 weeks post infection, contained a substance that was capable of lysing the initial trypanosome
innoculum but was ineffective against parasites collected later in the course of the infection. It was subsequently demonstrated that this variation was an inherent property of individual trypanosomes and not a reflection of innoculum heterogeneity (Ritz, 1916). We now know that the systemic stage of a trypanosome infection is characterized by a relapsing
parasitemia in which each parasitemia peak consists predominantly of parasites with a distinct VSG coat (Vickerman and Luckins, 1969). The potential number of different VSGs expressible by individual trypanosome clones is >100 (Capbem et al., 1977) and, based on gene-counting estimates, the total repertoire may approach 1,000 (Van der Ploeg et al., 1982).
The identity of a particular bloodstream trypanosome, as determined by it's VSG, is called the "variable antigen type" (VAT). A cloned trypanosome can potentially produce a great number of VATs which constitute its VAT repertoire (WHO, 1986). In the first parasitemic wave, the mammalian host is exposed to VSG coats synthesized by the metacyclic trypanosomes in the tsetse salivary glands (Hajduk and Vickerman, 1981a). Monoclonal antibody screening of metacyclic trypanosomes in single tsetse flies revealed that they are antigenically heterogeneous, but exhibit a more limited antigenic range than bloodstream stages (Esser et al., 1982). A cloned T. b. rhodesiense stock may contain up to 16 different metacyclic VATs (Esser and Schoenbechler, 1985), all of which are also expressible as bloodstream form surface antigens (Barry, 1986).
In the mammalian host, the entire cell surface of bloodstream trypanosomes is covered by a 12-15 nm-thick coat (Vickerman, 1969; Vickerman and Luckins, 1969) consisting of circa 107 molecules of the membrane form of a variant surface glycoprotein (mf VSG; Cross, 1975; Cardoso de Almeida and Turner, 1983; Ferguson et al., 1988). Cross (1975,1984) developed techniques for the large ssale isolation of pure VSGs , allowing their biochemical and structure-function analyses. VSGs have a molecular mass of circa 60,000, the carbohydrate content ranges from 7-10% by weight (Johnson and Cross, 1977) and the polypeptide chains contain 450-500 amino acids (Turner, 1985). Although the carbohydrates are heterogeneous in structure, it is the diversity in amino acid
sequence that is responsible for antigenic variation (Turner, 1988). The N-terminal 350 amino acids of distinct VSG sequences differ markedly from each other (Rice-Ficht et al., 1981). However, some conservation in cysteine placement and some conservative amino acid changes within the N-terminal 30 residues have been observed (Olafson et al., 1984). Sequence homologies are also present at the C-terminal 50-100 VSG residues (Rice-Ficht et al., 1981; Holder and Cross, 1981). Two different forms of carbohydrate are present in all VSGs; a N-linked oligosaccharide(s) attaching to the nascent polypeptide chain
(Strickler and Patton, 1980; McConnell et al., 1983) and a glycan that is covalently attached to the C-terminus of the protein via an ethanolamine residue (Holder, 1983). In addition, the C-terminal glycan is attached to a dimyristoyl phosphatidylinositol moiety which anchors the VSG to the plasma membrane (Ferguson et al., 1985,1988). The diacyl glycerol of the membrane-attached VSG (mf VSG) may be cleaved by a
glycophosphatidylinositol-specific phospholipase C (Hereld et al., 1986). This results in the release of soluble form VSG from the plasma membrane (Gumett et al., 1986) and the exposure of an epitope at the C-terminus conser ed carbohydrate portion, also called the cross-reacting determinant (CRD), that is responsible for the immunological cross- reactivity observed among most VSGs (Holder and Cross, 1981).
Monoclonal antibodies (MAbs) have been produced against purified VSGs since 1980 and have become valuable tools in investigating VSG epitopes and their relationship to the trypanosome cell surface (Pearson et al., 1980; Miller et al., 1984a,b; Hall and Esser, 1984; Pinder etal., 1987; Clarke etal., 1987). Although all of the MAbs were specific for their immunizing VSGs, only a few of the MAbs produced bound to living trypanosomes or, more critically, neutralized infectivity (Turner, 1988). Many of the MAbs can bind to trypanosomes only after chemical fixation (Turner,1988). These data suggest the presence of cryptic antigenic determinants with variant specific sequences that are not surface-exposed (Pearson et al., 1981). The non-cryptic, surface-exposed epitopes are located within the N-terminal domain and are topographically assembled (Clarice et al.,
1987).
The molecular mechanisms involved in trypanosome antigenic variation have been studied extensively using recombinant DNA technology. It is now known that the process of antigenic variation is regulated at the transcriptional level (Donelson and Rice-Ficht, 1985; Boothioyd, 1985; Borst, 1986). Genes encoding VSGs are scattered within the T. brucei genome at both intrachromosomal (Van der Ploeg et al., 1982) and telomeric
positions (DeLange and Borst, 1982; Williams et al., 1982). Only one VSG gene is transcribed at one time and this expressed VSG gene is usually located at one of several telomeric expression sites (Donelson and Rice-Ficht, 1985). Taree mechanisms have been proposed for the antigenic switch: telomeric activation, reciprocal telomeric exchange and gene conversion (Donelson, 1987). In the telomeric activation model, transcription is switched from one expression site to another with no apparent change in the genomic environment of these VSG genes (Williams et al., 1979; Myler et al., 1984a; Bernards,
1984). Telomeric exchange involves reciprocal recombination occurring between two telomeric VSG genes resulting in the placement of a previously inactive VSG gene into an active expression site (Pays et al., 1985). The third proposed mechanism, gene
conversion, involves duplication of a telomeric or intrachromosomal basic copy (BC) VSG gene which is then reciprocally translocated with the p reviously expressed VSG gene at the active expression site (Myler et al., 1984b; Pays, 1985,*.
Despite the recent advances in our understanding of V SG genes, very little is known about the molecular mechanisms that control transcription initiation of a VSG gene. Nor do we know what triggers the switch from the tra” "cription of one VSG gene to another (Donelson, 1988). It was hypothesized that host antibodies might induce the antigenic switches in bloodstream trypanosomes (Dtvle, 1977). This was dismissed when antigenic variation was observed in the absence of specific antibodies during in vitro culture (Doyle et al., 1980). The release of soluble form VSGs from their membrane-attached anchor is catalyzed by a glycosylphosphatidylinositoi (GPI) - specific phospholipase C (PLC). This VSG lipase is a membrane-associated enzyme which cleaves the GPI- membrane anchor of the VSG molecule, forming diacylglycerol and a 1,2-cyclic phosphate on the inositol ring (Cardoso de Almeida and Turner, 1983; Fer^ -son and Cross, 1984; Ferguson et al., 1985,1988). In cell lysates of bloodstream trypanosomes the lipase is recovered in the particulate fraction and detergents are required for its solubilization (Cardoso de Almeida and Turner, 1983). Purification of this enzyme has allowed its further biochemical characterization (Hereld et al., 1986; Biilow and Overath, 1986; Fox et al., 1986). It is now known that the lipase is a non-glycosylated protein of 37-40 kDa which is highly specific for the GPI-moiety; phosphatidylinositol is only slowly cleaved and other common phospholipids do not serve as substrates. In contrast to other
phospholipase C's, this GPI-specific lipase does not require Ca++ for its activity (Biilow and Overath, 1986). About 30,000 copies of this VSG lipase are present per cell, each molecule can convert circa 100 molecules of membrane form VSG to soluble form VSG
per minute. The GPI-specific lipase is undoubtedly responsible for the rapid release of VSG from the plasma membrane of bloodstream trypanosomes under certain circumstances (England et al., 1988). However, it is still unclear if the enzyme plays an obligatory role in the differentiation of bloodstream to procyclic forms (Biilow et al., 1989a). Using
monoclonal antibodies to undetermined epitopes, Biilow et al. (1989b) have localized the enzyme, not on the plasma membrane, but predominantly on the peripheral face of intracellular vesicles of obscure function. The expression of the VSG lipase’s
developmenially regulated, with bloodstream trypanosomes containing high levels of both GPI-specific mRNA and VSG lipase activity (Biilow and Overath, 1985). Procyclic forms, however, yield no detectable GPI-specific lipase activity and contain significantly reduced mRNA levels for this enzyme (Carrington et al., 1989).
b) Procyclin
Procyclin is an immunodominant species-specific glycoprotein found on the surface of Trypanosoma brucei (Richardson et al., 1986,1988; Roditi et al., 1989b). It is an unusual molecule of which approximately 40 % of the protein sequence is a glutamic acid- proline (glu-pro) dipeptide repeat (Roditi et al., 1987; Mowatt and Clayton, 1987;
Richardson etal., 1988). Eight procyclin genes have now been identified in the T. brucei genome. These genes are arranged as four unlinked pairs of tandem repeats (Mowatt and Clayton, 1987). At least two closely related versions of procyclin are expressed (Roditi et al., 1987; Mowatt and Clayton, 1987; Richardson et al., 1988; Mowatt and Clayton,
1988). Both forms consist of a 31-amino acid N-terminal domain which contains a
glycosylation site, followed by a (Asp-Pro)2 (Glu-Pro)22-29 sequence and a 23-amino acid C-terminal hydrophobic domain which might serve as a membrane anchor (Clayton and Mowatt, 1989). Although affinity-purified procyclin from T.b. rhodesiense procyclic culture forms has an apparent molecular mass of 30-40 Kda when run on SDS-PAGE gels, the gene nucleotide sequences predict a 11-15 Kda protein (Roditi et al., 1987; Mowatt and Clayton, 1987; Richardson et al., 1988). The discrepancy between these estimates is probably due in part to the anomalous migration of proline-rich proteins on polyacrylamide gels (Ferguson et al., 1984; Young et al., 1985) and in part to glycosylation of the
trypanosomes is estimated to be 0.7% of the total cell protein, or 6X106 molecules per cell (Mowatt and Clayton, 1989).
All ten monoclonal antibodies that have been produced against live T. b.
rhodesiense procyclic trypanosomes can bind to procyclin (Richardson et al., 1986,1988; Roditi et al., 1989a). Some of the epitopes recognized by these antibodies have been localized using synthetic peptides which correspond to three different regions of procyclin. Four of these monoclonals bind to the glutamic acid-proline repeats while one antibody recognizes the N-terminal fragment containing amino acids 1-20 fragment and another binds to fragments having amino acids 21-35 (Richardson et al., 1988). Interestingly, although procyclin could be used to raise antibodies which bound to synthetic dipeptide repeats, the converse was not achievable (Roditi et al., 1989a). Both Roditi et a/.(1987) and Mowatt and Clayton (1987) were unable to induce antibodies to dipeptide repeats even when the dipeptides were coupled to several different protein carriers. Improper antigen presenation of the synthetic repeats may account for the failure to induce antibodies
specific for the procyclin molecule.
Expression of procyclin in trypanosomes is developmentally regulated.
Transformation of bloodstream trypanosomes into procyclic forms is accompanied by a rapid increase in procyclic-specific mRNA soon after cell differentiation commences. Within a few hours of these developments, a rapid increase in procyclin occurs (Roditi et al., 1989b). Two-color flow cytometry has been utilized to investigate the temporal, cell- surface expression of both procyclin and VSG. Results show that during bloodstream > procyclic transformation, loss of the VSG coat is followed by the gradual appearance of procyclin at the cell surface (Roditi et al., 1989b). The biological function of the procyclin molecule is presently unclear, although it could conceivably play a role in the
trypanosome's survival in the tsetse midgut (Richardson et al., 1988) or in determining tropism within the tsetse fly (Roditi and Pearson, 1990).
c) Other Non-YSG Antigens
It has long been recognized that infected mammalian hosts produce antibodies against both VSG and non-variant, common parasite antigens (Gray, 1960; De Raadt,
1974b), the identities of most of these common antigens are still poorly established. Methods such as complement fixation (Schoenaers et al., 1953), agglutination (Binz,
1972), ELISA (Luckins, 1977) and immunofluorescence techniques (Sadun, 1963; Mehlitz, 1979; Shapiro and Murray, 1982) have determined the presence of antibodies to non-variant trypanosome antigens in infected mammals. Other techniques, including double immuno-diffusion (Gray, 1961; Shapiro and Murray, 1982),
immunoelectrophoresis (Le Ray, 1975), counter-electroimmunophoresis (Poupin et al., 1976; Taylor and Smith, 1983), and immunoprecipitation (Shapiro and Murray, 1982; Gardiner et al., 1983), allow a crude enumeration of the different trypanosomal antigens recognized by host antibodies. These studies identified 5-7 different non-variant antigens. However, few of these were further characterized by immunoprecipitation or
immunoblotting (Shapiro and Murray, 1982; Gardiner et al., 1983; Burgess and Jeirells, 1985). In one study, serum from a mouse chronically infected with T.b. gambiense recognized 7-8 antigens with molecular weights of 50-120,175 and 300 kDa, all of which could be iodinated on the surface of procyclic trypanosomes (Gardiner et al., 1983).
Burgess and Jerrells (1985) also identified a low molecular weight doublet invariant antigen of approximately 22 kDa, using sera from 12 Kenyan patients infected with T.b.
rhodesiense . However, none of these antigens were characterized.
Several investigators have employed mouse infection-cure regimens to identify the common antigens that elicit the host's immune responses (Campbell et al., 1981; Parish et al., 1985). In most of these experiments, VSG-specific antibodies were obtained (Pearson et al., 1980; Miller et al., 1984a, b; Hall and Esser, 1984; Pinder et al., 1987; Clarke et al., 1987), although monoclonal antibodies (MAbs) to internal non-VSG antigens were selected by screening hybridoma supernatants with acetone-fixed, or air-dried trypanosomes
(Campbell et al., 1981). None of the non-variant antigens recognized by the MAbs were characterized. A 31 kDa protein, shared among bloodstream and procyclic trypanosomes of T. congolense, has been identified using MAbs produced using these immunization protocols (Parish et al., 1985). Shapiro and Murray (1982) attempted to correlate the type of trypanosomal antigens recognized with the clinical course of the disease. Sera from 18 cattle actively infected with T. brucei and from 14 drug-cured cattle were used to identify
the trypanosomal antigens by immunoprecipitation and SDS-pclyacrylamide gel
techniques. Antibodies from the infected cattle bound to 8 protein antigens of molecular weights 20,40, 45,100,110,150,180 and 300 kDa. Only three of these antigens: 110,
150 and 180 kDa, were recognized by sera from all the recovered cattle. The role of the immune response to these antigens in host control of the disease remains obscure. Recently, a trypanosome peptidase of 60 kDa molecular weight has been found in the plasma of mice infected with T. brucei (Knowles et al., 1987) and of heifers infected with T. congolense (Knowles et al., 1989). Whether hosts respond immunologically to this trypanosome peptidase is unknown. However, it is thought that the peptidase could be involved in the pathology of the disease.
Immunization with homogenates of African trypanosomes or with purified
trypanosomal components has also been used to identify non-variant antigens. While some of these studies only demonstrate the presence/absence of antigens commonly shared between the bloodstream and procyclic developmental stages (Weitz, 1960; Seed, 1963; Clarkson and Awan, 1969; Stanley et al., 1978, Beat et al., 1984), others have attempted to identify the antigens present in different trypanosomal species using
immunoelectrophoresis (Le Ray et al., 1973; Le Ray, 1975; Marcus and Schwarting, 1976), lectin-blotting (Frommel et al., 1987; Balber and Ho, 1988) and 2-dimensional gel electrophoresis (Anderson et al., 1985). Immunoelectrophoresis studies have revealed that the antigenic contents of T. b. brucei and T.b. rkodesiense stocks were indistinguishable. Immunization with procyclic culture forms elicited antibodies to 34 detectable components, 5 of which were not present in bloodstream forms. Conversely, immunization with bloodstream trypanosomes elicited antibodies to 23 detectable components and only the VSGs appeared to be bloodstream-specific (Le Ray et al., 1973; Le Ray, 1975; Marcus and Schwarting, 1976). Lectin-blotting studies detected 21 concanavalin A (Con A)-binding glycoproteins that are shared among the procyclic and bloodstream forms of T.b. brucei and T.b. gambiense. Additionally, 2 bloodstream-specific, con A-binding glycoproteins (one of which was a VSG, the other being a 81 kDa non-variant glycoprotein) were detected, together with an 84 kDa, procyclic-specific glycoprotein (Frommel et al., 1987). These various glycoproteins have differential distributions within the two phases of the detergent Triton X-114 (Balber and Ho, 1988). Biochemical characterization and cellular localization of these antigens have not yet been accomplished.
Other investigators have attempted to characterize immunogenic trypanosomal components by establishing their location in the parasite cell body. Early studies used hyperimmune sera prepared in rabbits innoculated wi th T.b. brucei homogenates and localized trypanosomal antigens in different subcellular fractions (Brown and Williams,
1962,1964). Two antigens were detected in trypanosomal nuclei using
immunoelectrophuresis. Recently McLaughlin (1982) has analyzed the immunogenic components of semi-purified African trypanosomal particle fractions. About 7-10 different antigens were detected by cross-immunoelectrophoresis using hyperimmune rabbit sera against different cellular fractions. Most of the antigens appeared to occur at the parasite flagellar pocket (McLaughlin, 1982). Of these, two principal glycoproteins (60 and 66 kDa) and four minor antigens (35-50 kDa) were distinguished (McLaughlin, 1984,1987). Immunization of mice with flagellar pocket membrane fragments yielded up to 60% protection against a challenge infection with T.b. rhodesiense bloodstream forms
(McLaughlin, 1987; Olenick et al., 1988). Gardiner et al., (1983) analyzed 14 procyclic trypanosome membrane antigens using hyperimmune rabbit sera against a crude parasite membrane fraction. An 83 kDa surface membrane disposed, non-variant glycoprotein was identified in both T. brucei species and T. vivax, but despite stimulating the production of high titres of specific antibodies, no protection was afforded against tsetse-transmitted challenge with either parasite (Rovis et al., 1984).
Some African trypanosome enzymes are sufficiently different from host enzymes to be immunogenic. Antisera have been prepared against a few purified trypanosome
enzymes (Risby et al., 1969). Monoclonal antibodies have also been produced that bind to cytoskeletal components of Trypanosoma brucei. These include a 55 kDa tubulin (Gallo and Anderson, 1983; Gallo et al., 1988); both high (320 kDa) and low (41 kDa) molecular weight microtubule-associated proteins (Schneider etal., 1988a, 1988b); spectrin-like proteins (Schneider et al., 1988c); 68 and 72 kDa paraflagellar rod structural proteins (Gallo and Schrevel, 1985) and a 60 kDa protein that interacts with microtubules and membranes (Stieger and Seebeck, 1986; Seebeck etal., 1988). More detailed analyses of trypanosomal cytoskeletal antigens may provide new leads for chemotherapeutic attack on African trypanosomiasis.
