Sleeping sickness is endemic in 36 countries of sub Saharan Africa where approximately
60 million people are at risk (Aksoy, 2003; WHO, 1994). Glossina pallidipes
is a known vector for rhodesian sleeping sickness. Vector-related factors that
influence the epidemiology of sleeping sickness in Kenya are not well understood.
Intraspecific variations in vector competence have been reported among subpopulations
of Glossina sp. and are believed to contribute to the focal distribution
of rhodesian sleeping sickness (Welburn and Maudlin, 1999). According to Aksoy
(2003) and Aksoy et al. (1997), both intrinsic genetic factors and biological
factors such as presence of endosymbionts influence vector susceptibility to
infection by parasites and the subsequent transmission. The occurrence of endosymbiont
and gut molecules that modulates vector competence varies across tsetse subpopulations
(Aksoy, 2003). The intracellular endosymbiont are thought to act in diverse
ways (Scott et al., 1993; Aksoy et al., 1997), for example, Wigglesworthia
and Wolbachia have been reported to play roles in tsetse reproduction
and survival (Aksoy et al., 1997) and survival of tsetse population is
a key determinant of its vectorial capacity. In addition, midgut lectins (agglutinins),
trypanoagglutinins, trypanolysins (Ibrahim et al., 1984) and digestive
proteases (Stiles et al., 1991; Welburn and Maudlin, 1999) mediate establishment
and maturation of trypanosomes in tsetse flies. Earlier attempts to compare
the vectorial capacity of allopatric populations in Kenya used G. pallidipes
from non-sleeping sickness foci and non-human infective Trypanosomes (Moloo,
1993) and did not therefore reveal much with respect to epidemiology of rhodesian
sleeping sickness. In the present study, tsetse subpopulations from an endemic
and non-endemic areas with a human infective T. b. rhodesiense were used.
The role of vector-associated factors in the transmission of T. b. rhodesiense
and their possible contribution to the focal nature of rhodesiense sleeping
sickness is discussed.
Materials and Methods
This study was carried out in Busia and Nguruman. Busia study area lies
between latitude 0°136' South and 0°North and longitudes 33°54'
east and 340 25' 24' East (Fig. 1). The area is infested
with G. f. fuscipes along the riparian forest patches and G. pallidipes
which has patchy distribution, associated with woody hillside vegetation (Ford,
1971). Nguruman lies at latitude 1°55' S and longitude 35°25' E on the
floor of the rift valley in southern Kenya (Fig. 2). The area
is infested by G. pallidipes and G. longipennis within the woodlands
and G. swynnertoni on the adjoining escarpments (Brightwell et al.,
||A map of Western Kenya showing the study area
||Map of Nguruman showing the study area
Collection of Field Flies and Maintenance in the Insectary
Unbaited biconical traps (Challier and Laveissiere, 1973) were used for
collecting flies from the field. Caught flies were transported to the rearing
facilities, sorted and placed in rearing cages. The flies were maintained by
in vivo feeding from rabbit ears. Room conditions were maintained at
70±2% relative humidity and 23±2°C.
Pupae Production and Maintenance
Each day, pupae were collected from the rearing trays and placed in a 20
mL plastic tube lined with cotton wool. Every succeeding three-day collection
were grouped together to maintain uniform age groups and to synchronize emergences.
The pupae were transported by road to the experimental insectary at KARI-TRC,
Nairobi for incubated.
In vivo Infection, in vitro Maintenance and Probing of Flies
Emerged teneral flies were allowed to obtain their first blood meal from
a mouse infected with a T. b. rhodesiense clone, KETRI 3537. At peak
parasitaemia, three or four tsetse flies in a cage were held close to the mouse
belly to feed. Engorged flies were returned to the insectary where they were
maintained in vitro on bovine blood for 25 days. Day 23 post infective
feed, flies were starved for two days and probed on warmed slide.
In vivo Transmission of Trypanosomes to Mice
In vivo transmission was used to transfer infection from the tsetse fly
to mice through natural feeding process. Tsetse flies that had been fed on infectious
mice as tenerals were allowed to feed on clean Swiss white mice, one fly per
mouse. Each mouse was monitored for parasitaemia and Packed Cell Volume (PCV)
every day for 20 days. Trypanosomes were detected in the whole blood by dark-ground
buffy coat phase contrast technique (Murray et al., 1977). PCV from heparinised
whole blood samples was measured after haematocrit centrifugation using the
method described by Schalm et al. (1975).
Determination of Infection Rate and Infection Load in Tsetse Flies
Flies were starved for two days then dissected at the mouthparts, midgut
and salivary gland. The fly parts found infected with trypanosomes were cut-off
and transferred into a 0.2 mL conical ampoule. Zero point one mL of 20% glycerine
Phosphate-Buffer Saline Glucose (PBS), was used to make a homogenous suspension
from which a wet smear was made and mean trypanosome load determined underx400
magnification. Sample of each fly found parasitologically negative in all its
three dissected parts were pooled into an ampoule and stored at -4°C for
Preparation of Template for PCR
Ampoules were thawed and the contents expelled into single 1.5 mL micro
centrifuge tubes containing 0.5 mL lysis buffer (1%v/v Triton-100 in 10 mM Tris-HCL,
pH 7.5) in which they were vortexed briefly. The material was pelleted by centrifugation
at 13,000 g for 10 min and the supernatant discarded. This step was repeated
until the material was free of haemoglobin. The final pellet was resuspended
in 100 μL of PCR buffer (10 mM Tris-HCl, pH 8.3, 50 mM KCl, 1.5 mM MgCl2)
with 60 μg mL-1 proteinase K. The mixture was incubated at 55°C
for 1 h and at 95°C for 10 min to denature the protainase K. Two microliters
of this extract was used as a template for PCR amplification.
DNA Amplification, Electrophoresis and Filming
Amplification of trypanosome DNA was performed as a 20 μL reaction
mix in a micro-centrifuge tube. The reaction mixture consisting of 10 mM Tris-HCl,
pH 8.3, 1.5 mM MgCl2, 50 mM KCl, 150 μM of each deoxynucleotide
triphosphate, 0.8 μM of each pair of trypanozoon specific primers (TBR1
and TBR 2) (PE Applied Biosystems, USA), 2 μL of the DNA extract and 0.5
units of Taq DNA polymerase (Fisher). The reaction mixture was placed in a thermo-
cycler and incubated at 94°C for 1 min, followed by 30 cycles of denaturation
at 92°C for 30 sec, annealing at 60°C for 45 sec, extension at 72°C
for 45 sec and a final extension at 72°C for 4 min. Fifteen microliters
of each PCR product was analyzed by gel electrophoresis through 1.5% agarose
gels stained with ethidium bromide and photographed using a Polaroid camera.
Tsetse Rearing and Pupae Production in Field Insectaries
Pupae production per day averaged 22 in Nguruman and 35 in Busia subpopulation
colonies. A total of 879 and 1,256 pupae were collected from Nguruman and Busia
respectively. Out of the 2135 pupae collected, 564 (26.42%) emerged, 303 (24.12%)
from Busia and 261 (29.69%) from Nguruman subpopulation groups.
|| Mean daily survival and mean percent survival of tsetse flies
to day 25
|| Mean (%) trypanosome infection rate of tsetse flies from
Busia and Nguruman
|Figures followed by the same letters within a column are not
significantly different at 0.05% LSD
|| Mean trypanosome infection load in Glossina pallidipes
25 days post infection
|Figures followed by the same letters within a column are not
significantly different at 0.05% by LSD
|| Transmission success of trypanosomes to mice by tsetse flies
detected with infection by microscopy
|| Survival curves and exponential trend line for male and female
tsetse flies from Nguruman
Survivals of Experimental Flies
Analysis of variance showed that the daily survival rates of flies from
Nguruman and Busia subpopulations did not differ significantly (t = 0.420, df
= 7, p = 0.687). Paired sample t-test for mean percent survival upto day 25
showed no significant difference (t = 0.989, df = 8, p = 0.395) between the
two-subpopulation groups (Table 1).
|| Survival curves and exponential trend lines for male and
female tsetse flies from Busia
||Survival curves and exponential trend line for tsetse flies
from Busia and Nguruman subpopulations
||Percent infection of flies from Nguruman and Busia subpopulation
flies as detected by microscopy and PCR
|| Mean parasitaemia in mice infected with T. b. rhodesiense
using flies from Busia sub population
Both sexes had similar survival in the Nguruman subpopulation (Fig.
3). Males had higher survival rates than females in the Busia subpopulation
(Fig. 4). On average, Nguruman subpopulation recorded lower
survival than the Busia subpopulation (Fig. 5).
Susceptibility of Experimental Flies to Trypanosome Infection
Infection rates between the Nguruman and Busia subpopulation flies were
not significantly different (χ2 = 0.387, df = 1, p = 0.534).
However, Busia subpopulation flies had higher infection rate (58.9%; N = 63)
than Nguruman subpopulation flies (47.6%; N = 39). Figure 6
shows that higher infection was detected by microscopy among the Busia subpopulation
(19%) than the Nguruman subpopulation (17.9%) flies. Contrastingly, PCR detected
higher proportion of trypanosome infections among the Nguruman (41.0%) than
the Busia (28.6%) subpopulation flies. Table 2 shows that
all the infected flies had midgut infection, 19% for Busia and 17.9% for Nguruman
subpopulation. Table 3 shows that infection load was highest
in the midgut (8.2) followed by the salivary gland (1.95) and proboscis (1.18)
in the Busia subpopulation flies.
Infection rates in the proboscis were 2.6% (N = 39) and 3.2% (N = 63) in the Nguruman and Busia subpopulations flies, respectively. Salivary gland infection was 2.6% (N = 39) in the Nguruman subpopulation and 4.8% (N = 63) in the Busia subpopulation flies. Chi-square test showed no significant difference among infected parts (χ2 = 0.024, df = 2, p = 0.988).
Analysis of variance of the Busia flies showed significant difference in infection load among the three parts dissected (F = 26.167; df = 2; p<0.001). Separation of means by Least Significance Difference (LSD) showed that infection load in the midgut was significantly higher than in the proboscis and salivary gland. Analysis of variance of the Nguruman subpopulation showed that there were significant differences in infection load among the three parts dissected ((F = 20.043, df = 2, p<0.001). Separation of means by LSD showed that midgut had significantly higher infection load than proboscis and salivary gland. Comparison of mean infection loads between the Busia and Nguruman subpopulation groups using chi square test showed no significant difference (χ2 = 0.646, df = 2, p = 0.439).
In vivo Transmission of Trypanosomes to Mice by Infected Tsetse Flies
Forty one point six percent (N = 12) of flies from the Busia subpopulation
successfully transmitted trypanosomes to mice, while none (0%, N = 7) of the
flies from the Nguruman subpopulation group transmitted the infection to mice
(Table 4). Chi square test of association between subpopulation
group and transmission success indicated insignificant difference (χ2
= 2.601, df = 1, p = 0.107).
Parasitaemia and Packed Cell Volume in T. b. Rhodesiense Infected Mice
Five flies successfully transmitted trypanosomes to mice in the Busia subpopulation.
The mean prepatent period in mice was 5 days. The first parasitaemia wave peaked
on day 10 followed by the second and third on days 13 and 17, respectively (Fig.
7). Mice survival was 100% to day 15. Mortality 60% was recorded between
on day 16, while 40% survived to day 20 (Fig. 8). There was
however no successful transmission of trypanosomes to mice to Nguruman subpopulation
flies. Mean Packed Cell Volume (PCV) of infected mice was 36.96±1.1 SE
while that of control mice was 38.95±1.0 SE. Two-sample t- test for mean
PCV indicated no significant difference (t = 1.35, df = 31, p = 0.187). Linear
trend lines indicated stable PCV level for the control mice and a gradual decline
in PCV for the infected mice with time (Fig. 9). Mean PCV
and mean antilog parasitaemia showed a weak and insignificant inverse correlation
(r = -0.390, p = 0.135).
|| Survival of mice infected with T. b. rhodesiense using
flies from Busia
||Changes in mean packed cell volume and linear trend lines
in mice infected naturally with T. b. rhodesiense by flies from Busia
The present study demonstrated remarkable differences in vector density between Western Kenya (Busia) and Southeastern Kenya (Nguruman). Vector density plays a central role in transmission of vector borne parasites to susceptible hosts, as the average host-biting rate is dependent upon the per capita vector load. Although Weir and Davidson (1965) reported that other factors such as host abundance, host preference and human-vector contact also influence transmission, higher human biting rate would be expected at Nguruman. Human biting rate is a measure of risk of transmission of humans and therefore human population at Nguruman stand at higher risk of contracting the infection should the human infective T. b. rhodesiense circulate in the vector population. The present study also showed that G. pallidipes from Western Kenya had higher survival and longevity than those from Southeastern Kenya. The competence of a vector population depends on the average longevity of individuals in the population (Macdonald, 1957). For T. b. rhodesiense, which requires considerably longer time to undergo its reproductive cycle in the invertebrate host, a vector population with high survival and longevity would be more suitable and this therefore, indicate that Busia subpopulation would be a better vector.
The present study showed that G. pallidipes subpopulation from Western Kenya was more susceptibility to trypanosomes infection than those from Southeastern Kenya. Susceptibility to trypanosome infection is regulated by several factors such as endosymbionts and genetic predisposition (Aksoy, 2003). Influence of heritability on vectorial capacity in insects has been reported (Gooding, 1988). Wakelin (1978) insinuated that susceptibility to infection depend on the genotype of the insect and often on the inheritance of a single gene. Comparative genetic studies based on microsatellite loci between the two subpopulations showed significant variability (Nei, 1987). Part of the variation in susceptibility to trypanosome infection between the two subpopulations may therefore be attributed to genetic variability, although this needs to be quantified. In an earlier study (Moloo, 1993) found that G. pallidipes from Nguruman were more susceptible to T. congolense infection than those from Shimba hills. This finding is consistent with the genetic data (Nei, 1987) where Nguruman and Shimba hills subpopulations were found to be closer and Shimba hills was genetically further from Busia subpopulation than Nguruman subpopulation. Data on tsetse endosymbionts in the two areas is however lacking and it would be interesting to study the occurrence and possible contribution of endosymbionts in the observed variations in susceptibility. Another striking difference between the two tsetse subpopulations was in their ability to transmit infection to susceptible host. The reason Nguruman subpopulation flies which showed mature transmissible metacyclics could not transmit the parasites to mice is not clear. However, it could have been due to the low metacyclic load detected in the salivary gland or other factors peculiar to the subpopulation.
These findings reveal the underlying differences between the two subpopulations and, by extension, other G. pallidipes subpopulations with respect to tsetse longevity and refractoriness to trypanosome infection and transmission capacities, which are important factors in vectorial capacity and overall epidemiology of sleeping sickness. In general, the present study revealed that G. pallidipes subpopulation from western Kenya was more competent in acquiring, establishing and transmitting T. b. rhodesiense than that from the southeastern Kenya, an observation which partially explains the localised distribution of rhodesian sleeping sickness to specific foci in Kenya.
I am grateful to WHO/TDR, Swiss Tropical Institute (STI) and KARI-TRC for funding this work. Thank also to Godfrey Emase, Joseph Etyang, Charles Nambiro Sam Guya, Kariuki Ndungu and Purity Gitonga for their invaluable effort in making the bench work a success.