Latex agglutination test for the detection of urinary antigens in visceral leishmaniasis

Acta Tropica 78 (2001) 11 – 16
www.parasitology-online.com
Latex agglutination test for the detection of urinary antigens
in visceral leishmaniasis
Zamil J. Attar a, Michael L. Chance a, Sayda el-Safi b, James Carney c,
Ahmed Azazy d, Maha El-Hadi b, Cibele Dourado a, Marcel Hommel a,*
a
Molecular Biology and Immunology Di6ision, Li6erpool School ofTropical Medicine, Li6erpool, L3 5QA, UK
b
Department of Microbiology and Parasitology, Uni6ersity of Khartoum, Khartoum, Sudan
c
Kalon Biological Ltd, Ash Vale, GU12 5QJ, UK
d
Faculty of Medicine & Health Sciences, Sana’a Uni6ersity, Sana’a, Republic of Yemen
Received 4 April 2000; received in revised form 21 August 2000; accepted 25 August 2000
Abstract
This paper describes a new latex agglutination test (‘KATEX’) for the detection of leishmanial antigen in the urine
of patients with visceral leishmaniasis. In preliminary laboratory trials, using urine collected from well-defined cases
and controls from Brazil, Yemen and Nepal, the test had 100% specificity and a sensitivity between 68 and 100%.
When used in a time-course experiment in cotton rats infected with Leishmania dono6ani, the test became positive 1
week after inoculation and antigen levels in urine declined rapidly after chemotherapy (the test was negative before
the end of the course of treatment). Finally, in an integrated study performed in Sudan, KATEX was compared to
microscopy and four different serological tests in a group of 73 patients having presented with clinical manifestations
suggestive of visceral leishmaniasis. Compared to microscopy, KATEX performed better than any single serological
test in predicting positivity and a particularly good result was obtained by combining KATEX and the direct
agglutination test (DAT). © 2001 Published by Elsevier Science B.V.
Keywords: Kala azar; Visceral leishmaniasis; Diagnostic; KATEX; Latex; Urine antigen
1. Introduction
Visceral leishmaniasis (VL), is a disease with an
annual incidence of 500 000 cases worldwide and
an estimated 200 million individuals at risk of
contracting the infection (Ashford et al., 1992);
the fact that 90% of clinical cases occur in the
* Corresponding author. Tel.: +44-151-708-9393.
E-mail address: mhommel@liv.ac.uk (M. Hommel).
poorest communities in developing countries is an
important consideration with regard to diagnosis
and treatment. Clinical diagnosis relies on noncharacteristic symptoms (cachexia, anaemia,
chronic fever with hepato-splenomegaly) and is
only reliable in advanced cases and in epidemic
situations. Mortality of VL is high in the absence
of treatment (using mostly injectable pentavalent
antimony) which is lengthy (20–28 days in most
regimes), very expensive and rather toxic. The
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Z.J. Attar et al. / Acta Tropica 78 (2001) 11–16
finding of leishmanial amastigotes in stained
smears from lymph nodes, bone marrow or
splenic aspirates is generally accepted as the ‘gold
standard’ for diagnosis, but the method is invasive, generally not feasible where is it most needed
and it has poor sensitivity; while the in vitro
cultivation of parasites from any of the above
samples will improve sensitivity, this requires sophisticated laboratory facilities. Serological tests
for the detection of antileishmanial antibodies are
a well-developed diagnostic tool and a number of
methods have been described, including indirect
immunofluorescence (Badaro et al., 1983), ELISA
(Hommel et al., 1978) and direct agglutination
test (El-Harith et al., 1986). The use of recombinant leishmanial antigens (for example rK39;
Houghton et al., 1998; Sundar et al., 1998) or
synthetic peptide antigens (Fargeas et al., 1996)
has recently been introduced for serology both in
an ELISA and a dipstick format.
There are a number of problems with serological assays, including the possible cross-reaction
with other pathogens including other Leishmania
spp (responsible for cutaneous or mucocutaneous
infections), Trypanosoma cruzi and mycobacteria
and the fact that most serological tests cannot
readily distinguish between current infection, subclinical infections or past infections. In an infection which remains asymptomatic in the majority
of cases and where only an estimated 5 – 20% of
infections ever become clinically patent (Hommel,
1999), this is a significant drawback to the use of
serological tests in areas where infections are common, since seroconversion does not necessarily
signify VL.
An antigen detection test would, in principle,
provide better means for diagnosis since antigen
levels are expected to broadly correlate with the
parasite load. Antigen detection systems are also
an ideal alternative to the antibody detection systems in immunocompromised patients and more
particularly with the growing number of HIV
co-infected cases, especially in advanced cases
where the immune response is impaired (De-Gorgolas and Miles, 1994; Rosenthal et al., 1995). No
satisfactory antigen detection test is currently
commercially available and attempts to develop
such tests have been unconvincing (Kohanteb et
al., 1987; Senaldi et al., 1996).
This paper describes the development of an
immunoassay to detect antigens in the urine of
patients with visceral leishmaniasis and a preliminary field trial of its effectiveness.
2. Materials and methods
2.1. Maintenance of Leishmania parasites
Leishmania dono6ani (MHOM/ET/67/HU3;
LV9) were maintained in vivo in cotton rats (Sigmodon hispidus) by intraperitoneal sub-inoculation of amastigotes from infected cotton rats.
Cotton rats are very susceptible to Leishmania
infection and develop a heavy infection approximately 3 months after infection (Fulton and
Joyner, 1948).
L. dono6ani promastigotes were maintained in
vitro at 26°C in liquid HOMEM culture medium
(Berens et al., 1976) containing 10% heat inactivated foetal calf serum (FCS).
2.2. Urine samples
The initial panel of human urine samples was
collected in Jacobina, Brazil. It included confirmed VL patients (25) and endemic controls (34),
which included patients with cutaneous leishmaniasis patients (12) and patients with Chagas disease (seven). Further urine samples from five
parasitologically confirmed cases were collected in
Nepal and 29 in Yemen; a set of 23 endemic
control were also collected in Yemen, including
patients with proteinuria, malaria, brucellosis,
schistosomiasis, cutaneous leishmaniasis and typhoid fever. As non-endemic negative controls,
312 fresh urine samples, taken randomly from a
single day admission to the Royal Liverpool Hospital, were used. All these samples were used in
the laboratory to perform initial sensitivity and
specific studies.
In a study performed in the Immunology Unit,
Department of Microbiology and Parasitology of
the University of Khartoum (Sudan), urine and
serum samples were collected from 73 patients
having presented with suggestive clinical signs of
VL (fever which did not respond to antimalarial
Z.J. Attar et al. / Acta Tropica 78 (2001) 11–16
therapy, with splenomegaly and/or lymphadenopathies); parasitological confirmation was performed in 60/73 patients by microscopical examination of Giemsa-stained smears from bone
marrow or lymph node aspirates.
2.3. Urine samples from a timecourse experiment
Urine samples from experimentally infected
cotton rats followed in a time course experiment
(Azazy, 1993) were used. Briefly, a group of cotton rats were infected with L. dono6ani amastigotes, and urine samples were collected weekly by
placing the rats into metabolic cages overnight. At
week 12 post-infection, all animals were treated
with Pentostam (20 mg Sbv/kg i.p. on alternate
days) for 20 days. Urine was collected from week
1 to week 26 (when the experiment was ended).
2.4. Preparation of the polyclonal antibodies
New Zealand White rabbits were repeatedly
immunised with L. dono6ani promastigotes collected from stationary phase cultures. IgG fractions of the anti-Leishmania hyperimmune sera
were prepared using Protein A-Sepharose chromatography (CL4B; Pharmacia). The anti-Leishmania IgG was labelled with horseradish
peroxidase (HRP) using the periodate method
described by Wilson and Nakane (1978).
2.5. Capture ELISA
Immulon-2 ELISA Plates (Nunc) were coated
overnight with optimal dilutions of rabbit antiLeishmania IgG (5 mg/ml) in coating buffer. After
washing, the undiluted urine specimens were incubated for 2 h followed, after further washing, by
HRP-labelled anti-Leishmania IgG at 1:400 dilution for 1 h. ABTS and H2O2 were used as the
substrate. Absorbance was measured at 410 nm
on a Dynatech MR5000 ELISA reader.
2.6. Latex agglutination test
Latex beads were coated with antibodies using
the method described by Hudson and Hay (1980)
with some modifications. One millilitre of opti-
13
mum IgG concentration was mixed with an equal
volume of 1% latex beads (Prolabo 0.8 mm) for 2
h. The optimum IgG concentration was determined by coating the latex beads with different
IgG concentrations ranging between 0.1–1 g/ml.
The optimum concentration is defined as the most
sensitive, economical concentration with no autoagglutination. One millilitre washing buffer (1%
BSA in PBS) was then added to the IgG-latex
beads mixture and mixed for another 30 min. The
beads were then washed twice and resuspended in
1 ml of washing buffer and stored at 4°C.
The ‘Kala azar latex agglutination test’ (named
‘KATEX’ for convenience) was performed by
mixing 50 ml of the prepared latex reagent with 50
ml of the neat urine sample on a ceramic slide. The
slide was rotated and rocked consistently for 2
min. Any agglutination was recorded as follows:
+ + + + , Most of latex agglutinates and
moves to the edges;
+ + + , Resembles chalk dust scattered onto
surface;
+ + , Clear agglutinated particles against background of granular latex; and
+ , Agglutination can just be noted as compared to the negative control.
2.7. Serology
In the Sudanese study, serology was performed
on serum samples from all 73 patients using the
direct agglutination test (DAT), indirect immunofluorescence test (IFAT), enzyme-linked immunosorbent assay (ELISA kit, Novum
Germany) and rK39-impregnated test strips
(kindly provided by Dr P. Desjeux, WHO). Standard protocols were used for each test and a rate
of positivity was determined for each test: rK39
by the presence of a red line on the upper end of
the dipstick (Sundar et al., 1998), DAT by agglutination at serum dilution of 1/1600 or above,
ELISA by an absorbance at two standard deviations above the cut-off, and an IFAT titre above
1/80. The combined results of all four serological
tests (rather than the results of individual serological tests), together with the results of microscopy,
was compared to the KATEX results.
Z.J. Attar et al. / Acta Tropica 78 (2001) 11–16
14
Table 1
Preliminary tests of KATEX in confirmed cases of visceral leishmaniasis (VL) compared to non-endemic controls (NEC) and
endemic controls (EC)
Samples
VL (Brazil)
VL (Nepal)
VL (Yemen)
NEC (Liverpool)
EC (Yemen)
EC (Brazil)
Number
25
5
29
312
23
34
LAT score
Sensitivity
+++/++++
0/+
16
5
25
9
0
4
312
23
34
3. Results
All urine samples from both endemic and European controls as well as VL patients from
Brazil, Nepal and Yemen were tested using KATEX. Using this set of well-defined samples, the
test had 100% specificity, since there were no
cross-reaction with urine samples from endemic
controls (zero positive out of a total of 57 samples
tested). When the specificity was further evaluated
by testing another 312 fresh urine samples from
European negative controls from the Royal Liverpool Hospital, the initial results gave 74 false
positives (23.7%). However, when the false positive urine samples were boiled for 5 min and
re-tested with KATEX, they all turned negative,
while all genuinely positive VL urine samples
from endemic areas remained positive after boiling. Consequently, systematic boiling of urine
specimens was introduced as a routine procedure
in the KATEX protocol. The results are summarised in Table 1. Overall, 48/59 VL patients
had a KATEX positive test (+ + or above),
which represents a sensitivity of 81.4%.
Following the preliminary screening of human
urine samples, urine samples from experimentally
infected cotton rats taken during the time-course
experiment were tested with KATEX. The urine
antigen was detected as early as 1 week post-infection, and more importantly, the antigen level
started to decline very quickly after treatment at
week 12 (i.e. KATEX became negative before the
end of the course of treatment; Fig. 1).
Specificity
68
100
86
100
100
100
Finally, urine samples were collected in the
Gedarif State of Sudan, as part of a larger field
trial of a variety of VL diagnostic methodologies.
The results obtained in 73 patients in whom KATEX and at least four serological tests had performed are summarized on Fig. 2. A combined
serological score had to be used since the performance of each individual serological test was variable and qualitative analysis of their respective
performance was outside the scope of this study.
When compared to microscopy (performed in 62
of the 73 patients), 41/47 smear negative patients
were also KATEX negative (specificity of 87.2%),
while 15/15 smear positive patients were KATEX
positive (indicating a sensitivity of 100%). In this
comparative trial, latex agglutination performed
Fig. 1. Follow-up of urine antigens by capture ELISA and
KATEX in a time course experiment in cotton rats.
Z.J. Attar et al. / Acta Tropica 78 (2001) 11–16
Fig. 2. Comparison between the results of KATEX, the combined serology score and microscopy in a field trial in Sudan.
better than any serological test in predicting
smear positivity and a particularly good result
was obtained by combining KATEX and DAT,
since the 18 samples which were found to be
positive in both tests included all 15 smear positive samples. Overall, the results were most reliable when KATEX positivity was ] + + (this
category included 12/15 smear positive samples
and only two smear negative samples); worst results were amongst the borderline positives (the
+ category included three smear positive and
four smear negative samples).
4. Discussion
In chronic infections, such as visceral leishmaniasis, the detection of antigens in patient serum is
complicated by the presence of high levels of
antibodies, circulating immune complexes (CIC),
serum amyloid, rheumatoid factors and auto-antibodies, all of which may mask immunologically
important antigenic determinants or competitively
inhibit the binding of antibodies to free antigen.
This may explain why no antigen detection assay
for VL is routinely in use to-date, despite a number of reports describing the existence of circulating antigens and immune complexes in VL
(Sehgal et al., 1982; Galvao-Castro et al., 1984;
15
Azazy et al., 1994, 1997). By looking at the presence of antigen in the urine, many of the problems
related to immune complexes may be avoided
(Kohanteb et al., 1987; Senaldi et al., 1996;
Urnovitz et al., 1996).
Although the capture ELISA format gave satisfactory results in the laboratory, the latex format
was chosen for development because this format
is better suited for use in remote rural areas in the
tropics, where VL is mostly endemic. In its
present format, the test is simple to use, economical and robust: its two main advantages are the
fact that the performance of the test does not
require any electric appliance and that most technicians are familiar with the latex format (which is
routinely used for pregnancy tests) and did not
experience any difficulty in scoring the test. The
need for boiling the urine is a step which complicates the performance of the test, but is indispensable to ensure specificity. The presence of such
non-specific reactions in un-boiled urine is a feature of latex agglutination tests applied to urine
and is not unique to KATEX (the non-specific
reactions do not exist in other test formats, e.g. in
the capture ELISA format).
The results obtained with KATEX using samples collected from different foci of VL (Brazil,
Nepal, Sudan and Yemen) indicate that the test
works well regardless of the geographical origin of
samples. The specificity of the test is high, particularly when using a high reading threshold (e.g.
only considering as positive samples that are ]
+ + ), even if this reduces sensitivity; in these
conditions and in the present format of the test,
sensitivity is no more than 70–80%. When used
together with a robust serological test (e.g. DAT)
and in patients with clinical suspicion of VL,
KATEX performs as well as microscopy, the accepted gold standard for the diagnosis of leishmaniasis. Whether the test has applications for the
detection of asymptomatic or pre-patent cases of
VL has not yet been evaluated in population
surveys. It is theoretically conceivable that KATEX may cross-react with other trypanosomatid
infections (e.g. sleeping sickness or Chagas disease) in areas where these infections are co-endemic, and this possibility will need to be
examined in future epidemiological surveys (the
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Z.J. Attar et al. / Acta Tropica 78 (2001) 11–16
problem does not arise when KATEX is used for
individual patient diagnosis since these diseases
are clinically very different). Further clinical evaluation of KATEX is also required to confirm the
rapid return to negative after treatment, which
was observed during the experimental time-course
in cotton rats; this feature would be of particular
interest to detect early treatment failure and antimony-resistance, which cannot readily be
achieved with existing methodologies.
References
Azazy, A.A., 1993. The development of circulating antigen
system for the diagnosis of visceral leishmaniasis. PhD
Thesis, University of Liverpool.
Azazy, A.A., Devaney, E., Chance, M.L., 1994. A PEGELISA for the detection of Leishmania dono6ani antigen in
circulating immune complexes. Trans. R. Soc. Trop. Med.
Hyg. 88, 62 – 66.
Azazy, A.A., Chance, M.L., Devaney, E., 1997. A time-course
study of circulating antigen and parasite-specific antibody
in cotton rats infected with Leishmania dono6ani. Ann.
Trop. Med. Parasit. 91, 153–162.
Ashford, R.W., Desjeux, P., de Raadt, P., 1992. Estimation of
population at risk and number of cases of leishmaniasis.
Parasitol. Today 8, 104–105.
Badaro, R., Reed, S.G., Carvalho, E.M., 1983. Immunofluorescent antibody test in American visceral leishmaniasis: sensitivity and specificity of different morphological
forms of two Leishmania species. Am. J. Trop. Med. Hyg.
32, 480 – 484.
Berens, R.L., Brun, R., Krassner, S.M., 1976. A simple
monophasic medium for axenic culture of hemoflagellates.
J. Parasitol. 62, 360 –365.
De-Gorgolas, M., Miles, M.A., 1994. Visceral leishmaniasis
and AIDS. Nature 372, 734.
El-Harith, A.E., Kolk, A.H.J., Kager, P.A., Leeuwenburg, J.,
Muigai, R., Kiugu, S., Laarman, J.J., 1986. A simple and
economical direct agglutination test for serodiagnosis and
sero-epidemiological studies of visceral leishmaniasis.
Trans. R. Soc. Trop. Med. Hyg. 80, 583–587.
Fargeas, C., Hommel, M., Maingon, R., Dourado, C., Monsigny, M., Mayer, R., 1996. Synthetic peptide-based enzyme-linked immunosorbent assay for the serodiagnosis of
visceral leishmaniasis. J. Clin. Microbiol. 34, 241–248.
.
Fulton, J., Joyner, L., 1948. Infection by Leishmania dono6ani
in the cotton rat. J. Gen. Microbiol. 2, 103 – 109.
Galvao-Castro, B., Ferreira, J.A., Marzochi, K.F., Marzochi,
M.C., Coutinho, S.G., Lambert, P.H., 1984. Polyclonal B
cell activation, circulating immune complexes and autoimmunity in human American visceral leishmaniasis. Clin.
Exp. Immunol. 56, 58 – 66.
Hommel, M., 1999. Visceral leishmaniasis: biology of the
parasite. J. Infect. 39, 101 – 111.
Hommel, M., Peters, W., Ranque, J., Quilici, M., Lanotte, G.,
1978. Micro ELISA techniques in the serodiagnosis of
visceral leishmaniasis. Ann. Trop. Med. Parasitol. 72, 213 –
218.
Houghton, R.L., Petrescu, M., Benson, D.R., Skeiky, Y.A.W.,
Scalone, A., Badaro, R.T., Reed, S.G., Gradoni, L., 1998.
A cloned antigen (recombinant K39) of Leishmania chagasi
diagnostic for visceral leishmaniasis in human immunodeficiency virus type 1 patients and a prognostic indicator for
monitoring patients undergoing drug therapy. J. Infect.
Dis. 177, 1339 – 1344.
Hudson, L., Hay, F.C., 1980. Practical Immunology, 2nd ed.
Blackwell, Oxford.
Kohanteb, J., Ardehali, S.M., Rezai, H.R., 1987. Detection of
Leishmania dono6ani soluble antigen and antibody in the
urine of visceral leishmaniasis patients. Trans. R. Soc.
Trop. Med. Hyg. 81, 578 – 580.
Rosenthal, E., Marty, P., Poizot-Martin, I., Reynes, J., Pratlong, F., Lafeuillade, A., Jaubert, D., Boulat, O., Dereure,
J., Gambarelli, F., Gastaut, J.A., Dujardin, P., Dellamonica, P., Cassuto, J.P., 1995. Visceral leishmaniasis and
HIV-1 co-infection in southern France. Trans. R. Soc.
Trop. Med. Hyg. 89, 159 – 162.
Sehgal, S., Aikat, B.K., Pathania, A.G.S., 1982. Immune
complexes in Indian kala-azar. Bull. Wld. Hlth. Org. 60,
945 – 949.
Senaldi, G., Xiao, S.H., Hoessli, D.C., Bordier, C., 1996.
Serological diagnosis of visceral leishmaniasis by a dot-enzyme immunoassay for the detection of a Leishmania dono6ani-related circulating antigen. J. Immunol. Methods 193,
9 – 15.
Sundar, S., Reed, S.G., Singh, V.P., Kumar, P.C., Murray,
H.W., 1998. Rapid accurate field diagnosis of Indian visceral leishmaniasis. Lancet 351, 563 – 565.
Urnovitz, H.ll., Murphy, W.H., Gottfried, T.D., FriedmanKien, A.E., 1996. Urine-based diagnostic technologies.
Trends Biotech. 14, 361 – 364.
Wilson, M.B., Nakane, P.R., 1978. Recent development in the
periodate method of conjugating horseradish peroxidase
(HRPO) to antibodies. In: Knap, W., Halubar, R., Wicks,
G. (Eds.), Immunofluorescence and Related Staining Techniques. Elsevier, Amsterdam, pp. 421 – 426.