The Value of Molecular Studies for the Diagnosis of Congenital Chagas Disease in Northeastern Argentina

Chagas disease caused by the hemoflagellate Trypanosoma cruzi is an endemic trypanosomiasis in Latin America, with 2.3 million people being infected in Argentina.¹

Although the infection is mostly acquired through vectorial transmission, congenital transmission is epidemiologically relevant as well. Recent studies in Argentina have indicated that the probability of vertical transmission of the infection ranges from 2.6% to 6.7%; congenital Chagas disease represents an important health problem in this country.² At the level of public hospitals, the number of pregnant chagasic women receiving treatment in Argentina is 9%.³

Because premature treatment in newborns leads to serologic and parasitologic conversion to negativity, early diagnosis of the infection will help to prevent further lesions.⁴ Current methods for parasitologic diagnosis show low sensitivity, and longitudinal evaluations by means of serologic procedures have many follow-up losses. As such, complementary methods are needed. Previous studies have pointed out the substantial value of polymerase chain reaction (PCR), in terms of sensitivity and specificity, in the direct parasitologic diagnosis of congenital Chagas disease, when using nuclear or kinetoplastic DNA from T. cruzi.^5–8 It follows that the use of PCR at the time of birth will allow for a faster infection diagnosis, avoiding repeated examinations in the ensuing months.

In this study, we evaluated the value of PCR for the detection of congenitally transmitted Chagas disease in relation to the routine diagnostic criteria based on the identification of bloodstream forms and/or seroconversion after 9 months of age.⁹ In parallel, a study comparing the sensitivity of this parasite DNA-detecting procedure, with respect to the traditional microhematocrit identifying whole parasite forms, was performed. This study was carried out in newborns from mothers with positive T. cruzi serology attending maternity services at the Hospital Central de Reconquista. This health center is situated in a high endemicity area of northern Santa Fe province in Argentina. Informed consent was obtained from all human adult participants and parents or legal guardians of minors. The protocol was approved by the hospital review board, as requested by the ethical committee of the School of Biochemistry and Biologic Sciences from the Universidad Nacional del Litoral.

Two milliliters of venous blood was taken from each patient: 1 mL was immediately mixed with an equal volume of a solution of 6 mol/L guanidine hydrochloride and 0.2 mol/L EDTA in sterile polypropylene tubes for PCR, with the sample being stored at 4°C for further processing. The remaining amount was used for routine parasitologic and serologic methods. The former consisted of the microhematocrit technique,¹⁰ and results were expressed in a qualitative form (presence or absence of parasites), whereas indirect hemagglutination (IHA) and enzyme immunoassay (ELISA) were used for serologic diagnosis. IHA was performed by means of a commercially available kit (Chagatest; Wiener Laboratories, Rosario, Argentina), according to the manufacturer’s recommendations, with a 1/16 dilution being regarded as positive sera. A commercial kit was also used for the ELISA method (ELISA kit; Wiener Laboratory). As recommended by the manufacturer, the sera were analyzed by a 1/20 dilution, and the cut-off value was established as the average of two optical density (OD) readings from negative control sera plus 0.200 OD units. Samples kept in polypropylene tubes containing guanidine/EDTA lysates were immersed in a boiling water bath for 15 minutes to shear the DNA molecules, as described by Britto and others.¹¹ Each sample was processed in duplicate, using two 100-μL aliquots each for the phenol–chloroform extraction, ethanol precipitation, and DNA resuspension in 100 μL sterile distilled water. Purifications were performed in six sample series, plus the respective controls, using sterile distilled water as a negative control and DNA from a patient with acute Chagas disease as a positive control. PCR was performed according to the protocol of Wincker and others.¹² Amplification was performed in 50-μL reaction mixtures containing 1/10 of the total DNA isolated, 250 mmol/L of each deoxynucleotide triphosphate, 100 pmol of each specific oligonucleotide for kinetoplast minicircle DNA S121 (5′-AAATAATGTACGGG[T/G]GAGATGCATGA-3′) and S122 (5′-GGTTCGATTGGGGTTGGTGTAATATA-3′), and 1.25 IU of Taq polymerase. The reaction mixtures were overlaid with 50 μL mineral oil and subjected to 30 cycles of amplification in a thermocycler. Denaturation, annealing, and extension steps were performed for 60 seconds each at 94°C, 60°C, and 72°C, respectively, with an initial denaturation at 94°C for 180 seconds, and a final extension at 72°C for 5 minutes.

Controls for each PCR assay included positive and negative controls for DNA purifications corresponding to samples of such reaction and a reaction mixture without DNA as a control of the reaction itself. All reactions were performed in duplicate. Amplification products were analyzed in agarose gel electrophoresis stained with ethidium bromide, and results were qualitatively expressed (presence or absence of the specific 330-bp band). The sensitivity of PCR was determined according to Virreira and others¹³ with slight modifications. Briefly, serial dilutions of parasite DNA were performed to obtain 100–0.001 equivalents of parasites in 5 μL water (1 equivalent = 300 fg of DNA). PCR assays were carried out with all dilutions, with an amplification band being obtained using the 0.05 equivalent parasite per tube.

As stated earlier, this study was made up of two parts. The first one compared PCR diagnosis in relation to standard methods (presence of bloodstream parasite and/or positive serology at 9 months of age). For this purpose, 17 neonates were studied in the follow-up investigation, and their venous blood samples were taken at birth and at 2–4 and 9 months of age. Concomitantly, a comparison between microhematocrit and PCR in 121 newborns from chagasic mothers, including the former 17 children, was carried out. For this comparative study of sensitivity between both parasitologic methods (microhematocrit and PCR), blood samples were taken from the neonates at the time of birth.

Comparisons for qualitative variables, within and between groups, were performed by non-parametric tests for > 2 dependent or independent samples, respectively.

The framework of the overall study is presented in Figure 1. Among children completing parasitologic and serologic follow-up after 9 months of age, five of them were diagnosed as having acute Chagas disease (29.4%), one by microhematocrit and four by specific serology (Table 1). Two of the five children had a negative PCR result in the sample taken at birth but had a positive PCR in the second sample, which remained positive in the third sample (9 months of age), whereas the other children were PCR positive since delivery (representative PCR gels obtained during the follow-up are shown in Figure 2). As such, it seems that PCR sensitivity to detect infection at the time of delivery is lower than that seen at 2 months of age. Comparisons by the McNemar test did not reach the level of statistical significance mainly because of the reduced sample size, for which future studies are needed to confirm this trend. The lower sensitivity coexisted with a reduced intensity in the specific T. cruzi DNA band observed in the agarose gel of amplified products from the first sample compared with the second and third samples (Figure 2). Increased sensitivity of PCR after the second month of age may be related to several non–mutually exclusive possibilities such as 1) a gradual decay of maternally transferred IgG anti-T. cruzi antibodies, reducing the control of parasitemia; 2) a recent infection with placental transmission occurring near the time of delivery; and 3) the individual immunologic ability of the fetus to control the infection by developing a CD8 T-cell response toward parasites.¹⁴ Among the 12 remaining children completing follow-up, all of them had a negative microhematocrit in the three samples and serology converted to negative on 9 months of age. The three samples yielded negative results for PCR. Such findings are indicative of a correlation between PCR and the follow-up of seroconversion after 9 months of age but not with the microhematocrit currently used for the diagnosis of congenital Chagas disease. Nevertheless, PCR did not detect the total amount of infected children at birth.

All cases with confirmed infection were subjected to treatment with benznidazol 7 mg/kg/day for 60 days.

Epidemiologic data on congenital Chagas disease are mostly based on parasitologic studies performed at birth by the microhematocrit method, which may bias its actual occurrence. Therefore, a comparative study between PCR and microhematocrit in samples taken at birth was carried out. Analysis of these samples (children with and without follow-up) showed that 3 of 121 neonates were diagnosed as infected using microhematocrit (2.5%), whereas 12 of them yielded a positive PCR result (9.9%), emphasizing the greater sensitivity of this molecular method as determined by the Fisher exact test (P < 0.0008). The present rate of congenital Chagas disease detected by microhematocrit was similar to results recorded in other studies using the same method in newborn children from Santa Fe city¹⁵ and from endemic regions of Salta province,⁸ being 2.9% and 2.6%, respectively. Our findings are in line with data obtained in Paraguay, which reported a 3.0% occurrence under similar study conditions,⁷ whereas a recent survey in Bolivia showed a 5.7% rate of congenital transmission in microhematocrit studies from cord blood samples.¹³ In agreement with other reports, our results point to an underevaluation of the occurrence of congenital transmission by classic parasitologic methods compared with other confirmatory methods or PCR. For example, a study in Paraguay showed a 10.0% or 3.0% incidence depending on whether PCR or microhematocrit was used, respectively.⁷ A survey in 302 children from northwestern Argentina detected 8.9% of congenital cases when applying different methods, with microhematocrit and PCR yielding a 2.9% and 6.4% of detection, respectively.⁸ Less marked differences were reported by Virreira and others,¹³ who studied 311 Bolivian children born to chagasic mothers and found 18 positive newborns by microhematocrit and PCR methods, with only 1 of the remaining 293 newborns being negative by microhematocrit but positive by PCR. In line with this study, Schijman and others¹⁶ also showed a similar sensitivity of PCR and microhematocrit, although, in their study, both procedures were performed in duplicate with a 20-day difference between blood samples.

Beyond these comments, these data suggest that there would be a higher transmission rate with many cases left untreated or given late treatment. Introduction of PCR will not only improve our current knowledge on congenital disease epidemiology, but also favors a better diagnosis, and the ensuing specific treatment. Nevertheless, the lower positivity rate obtained at the time of birth, with respect to the ones recorded in subsequent samples, raises the need for improving its sensitivity.

This study was done in a highly endemic region of northeastern Argentina. To the best of our knowledge, it constitutes the first analysis of neonatal transmission of Chagas disease by PCR in this area, extending the epidemiologic information provided by surveys carried out in the northwestern region of our country.^8,17

Study framework and results.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 78, 4; 10.4269/ajtmh.2008.78.624

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Study framework and results.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 78, 4; 10.4269/ajtmh.2008.78.624

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Study framework and results.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 78, 4; 10.4269/ajtmh.2008.78.624

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PCR products amplified from neonate peripheral blood using primers S121 and S122, separated on a 2% ethidium bromide–stained agarose gel. Lane 1, amplification reaction without added DNA; Lanes 2–4 and 5–7, samples taken at birth and at 2–4 and 9 months of age from two confirmed congenital Chagas infection cases by routine diagnosis, respectively; Lanes 8–10, samples taken at birth and at 2–4 and 9 months of age from a serologic-confirmed negative baby; Lane 11, positive control sample; Lane 12, 100-bp ladder. Arrow indicates the expected 330-bp product of the minicircle DNA amplification.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 78, 4; 10.4269/ajtmh.2008.78.624

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PCR products amplified from neonate peripheral blood using primers S121 and S122, separated on a 2% ethidium bromide–stained agarose gel. Lane 1, amplification reaction without added DNA; Lanes 2–4 and 5–7, samples taken at birth and at 2–4 and 9 months of age from two confirmed congenital Chagas infection cases by routine diagnosis, respectively; Lanes 8–10, samples taken at birth and at 2–4 and 9 months of age from a serologic-confirmed negative baby; Lane 11, positive control sample; Lane 12, 100-bp ladder. Arrow indicates the expected 330-bp product of the minicircle DNA amplification.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 78, 4; 10.4269/ajtmh.2008.78.624

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PCR products amplified from neonate peripheral blood using primers S121 and S122, separated on a 2% ethidium bromide–stained agarose gel. Lane 1, amplification reaction without added DNA; Lanes 2–4 and 5–7, samples taken at birth and at 2–4 and 9 months of age from two confirmed congenital Chagas infection cases by routine diagnosis, respectively; Lanes 8–10, samples taken at birth and at 2–4 and 9 months of age from a serologic-confirmed negative baby; Lane 11, positive control sample; Lane 12, 100-bp ladder. Arrow indicates the expected 330-bp product of the minicircle DNA amplification.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 78, 4; 10.4269/ajtmh.2008.78.624

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