INTRODUCTION
Chagas disease (ChD) is primarily a disease of poverty, disproportionately affecting the rural poor and immigrant populations predominantly in developing countries and is therefore considered a neglected disease 1 . Clinically, ChD exhibits a primary acute phase characterized by variable signs and symptoms ranging from mild to severe, and patent parasitemia is detected in direct blood tests. This phase progresses to the chronic phase, which persists throughout the patient’s life 2,3 . In this phase, detection of the parasite is rare due to subpatent and transient parasitemia. Approximately 60-70% of individuals will never develop apparent disease and are characterized as indeterminate; these patients present positive serologic reactivity for T. cruzi , which can sometimes be identified by hemoculture, xenodiagnoses, and polymerase chain reaction (PCR) 4–7 . Patients within this clinical form present normal 12-lead electrocardiogram (ECG) results, as well as normal radiological examinations of the chest, esophagus, and colon 8–11 . Moreover, 10-30 years after initial infection, some patients may develop the cardiac, digestive, and/or cardiodigestive clinical forms 12 . There are major variations in clinical manifestations and morbidities among patients with ChD 13,14 ; these differences can be attributed in part to the effectiveness of the immune response, genetic aspects of infected individuals, and the complex structure of the T. cruzi population 15,16 .
Trypanosoma cruzi exhibits a high degree of intraspecific variability, as detected by biological, biochemical, immunological, and genetic markers 17 . Based on different molecular markers, the Second Satellite Meeting recommended that T. cruzi should be classified into six discrete typing units (DTUs; T. cruzi I-VI) 18 . The biological characteristics of T. cruzi strains and clones, and particularly their tissue tropism, may play important roles as determinants of ChD and its clinical forms 19 . The diagnosis of T . cruzi infection should be directed based on the phase of infection. During the acute phase, parasitological methods should be prioritized due to the high number of trypomastigotes in peripheral circulation. In the chronic phase, T. cruzi can only be detected from a limited number of patients by parasitological methods, such as xenodiagnosis or blood culture 4,20–23 . Therefore, the development of additional molecular methods is necessary for parasite detection in patients with unclear serology and for better evaluation of the roles of specific trypanocidal treatments in patients with established Chagas’ cardiomyopathy 24–26 . In addition, these tools are important for studies involving biological and genetic characterization of the parasite.
Accordingly, the aim of this study was to detect circulating parasites by conventional PCR and blood culture (BC) in samples from patients with two defined and polar clinical forms of ChD, followed by evaluation of the genetic profiles of freshly isolated T. cruzi .
METHODS
Patients
This study included 91 patients in the chronic phase of ChD from endemic areas within the state of Minas Gerais, Brazil, who were identified and selected at the Referral Outpatient Center for Chagas Disease at the Clinical Hospital of the Universidade Federal de Minas Gerais (UFMG), Brazil, by Prof. Dr Manoel Otávio da Costa Rocha. All patients had at least two different reactive serological tests for T. cruzi and fulfilled eligibility criteria. Patients were subjected to a standard screening protocol that included medical history, physical examination, ECG, laboratory and chest X-ray examinations, and echocardiography evolution and characterized according the clinical classification of chronic Chagas cardiomyopathy, as previously reported 27 . None of the patients were undergoing etiological treatment nor had been previously treated for T. cruzi infection.
Ethical considerations
The study protocol was approved by the Research Ethic Committee of the Universidade Federal de Minas Gerais(protocol COEP-ETHIC 0559.0.203.000-11/2012/UFMG), and all participants provided written informed consent.
Blood culture
BC was performed with 30mL venous blood collected in heparinized vacuum tubes, and red cells were separated from the plasma by centrifugation at 300 × g for 10 min at 4°C 4 . The packed red blood cells were washed once, resuspended in 6mL liver infusion tryptose (LIT) medium, mixed, and distributed into six plastic tubes (Falcon, USA) containing 3mL LIT. The plasma supernatant was centrifuged at 900 × g for 20 min at 4°C, and 5mL LIT was added to the pellet. All tubes were maintained at 28°C, mixed gently twice a week, and examined monthly for up to 120 days. Microscopic examination was carried out in 10-µL aliquots of each preparation under 22-mm 2coverslips at a magnification of 400×.
Polymerase chain reaction assays
PCR assays were performed according to previously optimized protocols 7 that were standardized and validated for the diagnosis of chronic ChD 28 . Blood samples (5mL) were collected from each patient and immediately transferred to sterile tubes containing guanidine/ethylenediaminetetraacetic acid buffer (pH 8.0, 5mL) 29 . The samples were stored for 5-7 days at room temperature, boiled at 100°C for 15 min, and stored at 4°C, after which 200-µL aliquots were collected for deoxyribonucleic acid (DNA) extraction 7,30 . Positive controls (ChD patients), negative controls (patients without ChD from nonendemic areas), T. cruzi DNA corresponding to TcI-VI, and reagents without DNA were included.
Trypanonoma cruzi isolation
Positive BCs in LIT medium were maintained in individual tubes for a short period of time without passage, and LIT medium was added every 10-15 days for a maximum of 8 weeks. To obtain epimastigote masses (1 × 10 6/mL), the cultures were washed twice in Krebs-Ringer-Tris buffer (pH 7.2). Cultures were then concentrated by centrifugation at 900 × g and 4°C for 10 min, and wet masses were stored at -20°C for DNA extraction 31 .
Analysis of discrete typing units of Trypanonoma cruzi isolates
Trypanonoma cruzi isolates were typed using three different parasite genomic sequences according to proposed protocols 32 , as follows: 24Sα ribosomal (rRNA) gene D7 domain 33 , mitochondrial cytochrome oxidase subunit 2 gene (COII) 34 , and the spliced leader genes intergenic region 35 , as markers of six DTUs 18 , TcI (Col1.7G2 Colombiana clone), TcII (JG), TcIII (222), TcIV (CAN III clone), TcV (3253 Lages-Silva et al.: unpublished data), and TcVI (CL) were used as reference strains and DTU controls 18,34 .
Statistical analysis
Fisher’s exact tests were used to determine the associations among BC, clinical classification, and the degree of cardiac involvement. Mann-Whitney-Wilcoxon tests were used to compare the numbers of positive tubes according to the BC results, patient age, and clinical form 36 . Kappa coefficient concordance and 95% confidence intervals were used to quantify the degree of agreement between the results of BCs and PCR 37,38 . To confirm or refute the evidence found by tests mentioned above, we used a 5% significance level.
RESULTS
Patient demographics
Approximately, 25.3% (23/91) of patients had the indeterminate clinical form, and 74.7% (68/91) of patients showed different degrees of cardiac involvement 27 . Among patients with the indeterminate form of ChD, 34.8% (8/23) were men, and 65.2% (15/23) were women [ages 33-70 years; 44 ± 10.3 years (mean ± standard deviation)]. For the cardiac form, 66.2% (45/68) of patients were men, and 33.8% (23/68) were women (ages 25-81; 54 ± 10.3 years).
Detection of Trypanonoma cruzi by blood culture
Fifty of the 91 (54.9%) patients with chronic ChD presented positive BCs. Patients with cardiac and indeterminate clinical forms showed 60.3% and 39.1% BC positivity, respectively. No significant difference in BC positivity could be verified between patients with cardiac and indeterminate forms of ChD ( Figure 1 a). Similarly, there was no consistent evidence of a relationship between BC positivity and cardiac morbidity ( Figure 1 b).
FIGURE 1 Distribution of blood culture results in patients with chronic Chagas disease versus clinical form (a) and blood culture versus chronic Chagasic cardiomyopathy levels (b) . CCC: chronic Chagasic cardiomyopathy vs: versus.
No difference was observed between results obtained for BC in patients of different ages, regardless of the clinical form ( Figure 2 a). Moreover, there were no correlations between age and BC results after adjustment for clinical form ( Figure 2 b and Figure 2 c), i.e., there was no evidence that the age associated with the clinical form may influence BC results. The number of positive tubes for BC was compared in patients with indeterminate and cardiac clinical forms. There was no evidence that patients with the indeterminate form showed a greater frequency of positive tubes than those with the cardiac form. Moreover, in samples from both clinical forms, a high frequency of negative tubes was observed ( Figure 3 ).
FIGURE 2 Boxplots for the age of patients with chronic Chagas disease according to clinical form and blood culture results, and pvalues of Mann-Whitney-Wilcoxon tests. (a) Patients with indeterminate or cardiac Chagas disease; (b) patients with the indeterminate form; (c) patients with the cardiac form. From bottom to top in the boxplot: lower quartile-1.5 IQlower quartile (25%), median (50%), upper quartile (75%), and upper quartile-1.5 IQ. IQ (interquartile range): difference between the upper and lower quartile.
Associations between blood cultures and PCR methods
Trypanosoma cruzi was amplified by PCR in 98.9% (90/91) of samples from patients with chronic ChD. Of these, 54.9% (50/91) were positive by both PCR and BC; 43.9% (40/91) were only positive by PCR, and 1.1% (1/91) of samples was negative for both methods. All concordance coefficients were negative, indicating no evidence of agreement between BC and PCR results. In addition, all 95% confidence intervals possessed negative lower and upper limits or contained zero ( Table 1 ).
TABLE 1: Percentage of concordances, point and interval estimates of the kappa coefficient according to the clinical form and the total.
| Clinical form | Blood culture | PCR | Patients % (no.) | Agreement coefficient | Type of agreement |
|---|---|---|---|---|---|
| Indeterminate | Positive | Positive | 39.1 (9/23) | -0.394 [-0.803; 0.0147] | none |
| Negative | Positive | 56.5 (13/23) | |||
| Positive | Negative | 0.0 (0/23) | |||
| Negative | Negative | 4.3 (1/23) | |||
| Cardiac | Positive | Positive | 60.3 (41/68) | -0.248 [-0.485; -0.0100] | none |
| Negative | Positive | 39.7 (27/68) | |||
| Positive | Negative | 0.0 (0/68) | |||
| Negative | Negative | 0.0 (0/68) | |||
| Total | Positive | Positive | 54.9 (50/91) | -0.282 [-0.487; -0.0762] | none |
| Negative | Positive | 44.0 (40/91) | |||
| Positive | Negative | 0.0 (0/91) | |||
| Negative | Negative | 1.1 (1/91) |
Genotyping of Trypanonoma cruzi isolates
A total of 42 T. cruzi stocks were isolated from 91 patients with ChD using the BC technique and were analyzed by 24Sα rRNA and mitochondrial COII genes. Approximately, 95.2% (40/42) of the analyzed T . cruzi isolates belonged to rDNA group 1 [fragments of 125 base pairs (bp)] and COII haplotype C (fragments of 212 and 81bp), corresponding to TcII. Among these patients, 34 had the cardiac form of ChD, and eight had the indeterminate form of the disease. However, two isolates (50 and 54) from patients with cardiac and indeterminate clinical forms, respectively, were classified as rDNA½ amplified fragments of 110 and 125bp ( Figure 4 a). Genotyping of the mitochondrial COII genes of these isolates amplified restriction fragments of 264, 81, and 30bp for isolate 50 (mitochondrial haplotype A, T. cruzi I) and restriction fragments of 294 and 81bp for isolate 54 (mitochondrial haplotype B), which suggests of TcIII, TcIV, TcV, or TcVI ( Figure 4 b). In order to distinguish populations belonging to TcIII and TcIV (amplicons of 200bp) from those with TcI, TcII, TcV, or TcVI, which present fragments of approximately 150-157bp, we used the SL-IR gene marker. Most of the analyzed T. cruzi populations (40/42) showed amplification of 150-/157-bp fragments corresponding to TcII, as previously identified by rDNA and COII markers. In contrast, analysis of the SL-IR gene showed that isolate 50 corresponded to TcIII or TcIV (amplicons of 200bp), whereas isolate 54 was associated with TcV or TcVI (amplicons of 150-157bp; Figure 4 c).
FIGURE 4 A: Representative profiles of 24Sα rRNA of T. cruzi isolates from patients with chronic Chagas disease and controls obtained by polyacrylamide gel electrophoresis. Lane 1: MW marker; Lane 2: Col1.7G2 clone (~110bp), TcI; Lane 3: JG (~125bp), TcII; Lane 4: 222 strain (~110bp), TcIII; Lane 5: CAN III clone (~117/119bp), Tc IV; Lane 6: 3253 (~110 and 125bp), TcV; Lane 7: CL strain (~125bp); Lanes 8-29: DNA samples of patients; Lane 30: NC: negative PCR control (reagents without DNA). B: RFLP analysis of the mitochondrial COII gene in the T. cruzi isolates belonging to different haplotypes, obtained by polyacrylamide gel electrophoresis. Digestion of the DNA with AluI generated three RFLP patterns for the T. cruzi strains: restriction fragments of 264, 81, and 30bp are classified as TcI (mitochondrial haplotype A; control, Col1.7G2 clone); restriction fragments of 212 and 81bp are classified as TcII (mitochondrial haplotype C; control, JG strain); restriction fragments of 294 and 81bp are classified as TcIII, TcIV, TcV, and TcVI (mitochondrial haplotype B; control, 222, CAN III, 3253, and CL strains). Lanes 2-7: TcI-VI controls; Lanes 8-29: DNA samples of patients; Lane 30: NC, negative PCR control (reagents without DNA). C: Genetic profiles of T. cruzi isolates obtained by SL-IR genes used to separate T. cruzi III from other DTUs. Lanes 2-7: TcI – VI controls, Col1.7G2 clone, and JG strain (fragments of 150 – 157bp corresponding to TcI and TcII), 222 and CAN III strains (fragment of 200bp, associated with strains TcIII and TcIV), 3253 and CL strains (fragments of 150/157bp, belonging to TcV and TcVI). Lanes 8-29: DNA samples of patients; Lane 30: NC, negative PCR control (reagents without DNA). 24Sα rRNA: 24Sα ribosomal; T. : Trypanosoma; MW: molecular weight; bp: base pairs; Tc: T. cruzi ; DNA: deoxyribonucleic acid; NC: negative control; PCR: polymerase chain reaction; RFLP: restriction fragment length polymorphism; CO: cytochrome oxidase; SL-IR: spliced leader ingergenic regions; DTUs: discrete typing units.
DISCUSSION
In this study, we evaluated the detection of circulating parasites by conventional PCR and BC in samples from patients with ChD. Notably, our findings showed that more than half of the samples from patients with chronic ChD with different clinical forms and not subjected to specific treatment were positive by BC analysis. This positivity could be related to the high number of patients with the cardiac form, most of whom were 50 or more years old; however, there was no evidence of the degree of cardiac involvement and BC and/or PCR positivity. Despite these findings, we cannot rule out the possibility that BC variability may be directly related to the rate of T. cruzi infection in our specific patient group 22,39 . Higher positivity by BC has been shown in seroreactive patients between 4 and 20 years old when submitted to blood culture 40 . The period of cultivation also affects BC positivity; in previous studies, BC positivity was increased at 6 months after inoculation 41,21 . Additionally, increased positivity when BCs were repeated in the same patient using a larger blood volume and centrifugation at 4°C 21,22 .
In our study, no correlation was observed when we compared the number of BC-positive tubes with the clinical form of ChD, indicating that this parameter may be useful for assessing parasitemia in patients with chronic ChD; the number of positive tubes from a single BC could be an indicator of parasitemia. T. cruzi parasitemia in patients with chronic ChD varies considerably in different endemic regions. Moreover, clinical forms of the disease, morbidity, and mortality can also vary from one endemic area to another, and the intensity of infection and the immune response of patients also differs 19 . ChD progression may result from changes in the patient’s immune response during the course of T. cruzi infection 42 . Moreover, age influences the degree of parasitemia in patients 43,44 , and the decline in trypomastigote circulating forms is related to age 45 . However, we did not observe significant differences among BC results, patient age, and clinical form.
BC is considered positive if at least one blood sample contains T. cruzi trypomastigote forms that can differentiate into epimastigotes and multiply in culture medium. Interestingly, regardless of the clinical form, most cases were positive between 30 and 60 days, corroborating with previous data 4,21,22,40. BC may be wrongly considered positive due to in vitro production of motionless amastigotes for extended periods until the detection of moving forms, since a low number of amastigotes may be unnoticed 46,47. Variabilities in BC positivity have been reported and can be explained by various factors, such as higher volume of collected blood, which increases the chance to obtain intact forms of the parasite; maintenance of blood samples on an ice bath or at 4°C after collection; immediate removal of the plasma; quickness in the processing of samples; quality of culture medium; and correctness of the technique 4,21,48. Another important factor to be considered is the technical expertise to examine BCs, since T. cruzi can grow in different morphologies 46,47.
Trypanosoma cruzi k-DNA was detected by conventional PCR in most patients with ChD. Regardless of the protocol used, PCR is considered superior to BC and/or xenodiagnoses 5,7,22,23,49,50. This can be explained by parasitemia levels in infected individuals living in different endemic areas and may be related to the complexity of the T. cruzi life cycle. Patients living in Virgem da Lapa, Minas Gerais showed high levels of parasitemia based on positive xenodiagnosis when compared with individuals from other areas 51. The presence of T. cruzi in the peripheral circulation at any given time of blood collection depends on the parasite’s life cycle, the immune balance between parasite and host 52, and the time between the blood collection and sample processing 4,21. In this study, the volume of blood collected did not influence the positivity of PCR, consistent with a previous study of patients from the Triângulo Mineiro 50. Another hypothesis is that the infection may have been caused by Trypanosoma rangeli ; however, this is unlikely because infection by this protozoan has been detected only in animals in the State of Minas Gerais 53.
Studies in different endemic areas have shown that genetic differences between parasite strains can influence parasitemia and PCR positivity 54. This positivity may still be related to several factors, such as the number of transient trypomastigotes in the peripheral circulation at the time of blood collection, appropriate storage conditions of the samples, DNA extraction procedures, amplification targets, the use of the same thermal cycler, and the ability to detect minimal quantities of parasite DNA 7,22,28. PCR is more convenient than BC since it requires the collection of a lower blood volume, has a shorter processing time, and allows analysis of multiple samples simultaneously 55. Nevertheless, it is not possible to use PCR for T. cruzi isolation and subsequent biological, biochemical, and/or molecular studies. Thus, BC is the most efficient, particularly if repeated, because it allows the isolation and multiplication of the parasite in culture medium 22. BC and PCR will not achieve higher sensitivity because they are under the influence of various intrinsic factors of vertebrate hosts and the heterogeneity of parasite populations circulating in different endemic regions. However, PCR suitability for diagnostic purposes in laboratory routine is important for the detection of chagasic infection in patients with low levels of parasitemia or those with inconclusive serology 22. Importantly, recent data have demonstrated the standardization and validation of PCR for the diagnosis of chronic ChD. Despite this, the absence of a reliable method to detect and quantify parasitemia is still an obstacle for improving our understanding of the impact of persistent parasitemia in the natural history of ChD and to characterize the parasite load in order to evaluate prognosis and therapy 28. Recent findings have demonstrated a reliable protocol based on real-time PCR for validation and quantification of T. cruzi DNA in human blood samples, aiming to provide an accurate surrogate biomarker for diagnosis and treatment in patients with ChD 56.
Our findings showed that association of BC and PCR did not increase T. cruzi detection since PCR was positive in 40/91 patients with negative BC results. These results showed that the combination of the two methods improved the chance of detecting the parasite and/or its genomic fragments, corroborating with previous studies 22,50.
Understanding structure of the T. cruzi population is essential due to the links between parasite transmission cycles and the infection/disease. Herein, 42 T. cruzi isolates from untreated patients with chronic ChD and with well-defined clinical forms of ChD were identified. Most isolates from these patients were analyzed by rDNA 24Sα, COII, and SL-IR molecular markers and were found to be associated with DTU II. We also identified one isolate (50) associated with DTU III or IV from a patient with the cardiac form of the disease, and another (54) corresponding to DTU V or VI from a patient with the indeterminate form of the disease. We believe that isolate 50 is an excellent candidate for microsatellite analysis, a technique sensitive enough to detect small differences between clones within a single isolate.
Importantly, our data were consistent with previous studies showing that DTU II was associated with human infection in the State of Minas Gerais 32,57,58. The methodologies applied for genetic characterization of isolates of different clinical forms may be a limiting factor for typing T. cruzi populations since the analysis was based on a single marker, leading to potential misinterpretations 59. Our results showed that using two molecular markers was not sufficient for typing of T. cruzi isolates. These limitations were also observed by other authors 32,60–63.However, the use of three genetic markers with different evolutionary rates was able to ensure the correct typing of T. cruzi isolates. In contrast, it was not possible to correlate the parasite genotype with the clinical form. Findings using low-stringency single specific primer PCR and rDNA 24Sα markers showed TcII in all isolates from patients with indeterminate, cardiac, digestive, and cardiodigestive forms 58. Although TcII was identified in most T. cruzi isolates, no correlation with clinical forms was observed using rDNA 24Sα, COII, and SL-IR markers and polymorphisms in microsatellite analysis 32. TcII has been reported as the primary cause of acute ChD and is the typical genotype detected in patients with cardiac and digestive forms of ChD, confirming the association of this DTU with human infection in Brazil and Argentina 57,63. TcII has also been detected using the same markers with high homology among different hosts and locations in northeastern semiarid 62 . TcI is associated with human infection in the Amazon, the Andean region, Central America, Mexico, and endemic areas of Southeastern and Northeastern Brazil 63–67. Few cases of human infection caused by TcI have been described in Brazil 63,68.However, recent findings have detected TcI in patients with the indeterminate form of ChD in the northern region of Minas Gerais State 32 and in patients with cardiac, digestive, and indeterminate forms of ChD from Rio Grande do Norte State 69. Interestingly, TcIII (formerly known as TcIIc) is associated with terrestrial ecotopes from different reservoirs and vectors 70,71 and is able to coexist with other DTUs in the environment, supporting an overlap between sylvatic and peridomestic transmission cycles of T. cruzi69. In Colombia, TcIII has been detected in patients with chronic ChD in whom mixed infection with TcI and TcII was identified 72. Therefore, previous studies have shown how difficult it is to distinguish the role of each DTU in the disease. DTUs may be under-reported from domiciliary and sylvatic transmission cycles because some genotyping methodologies fail to distinguish between TcIV (TcIIa) and TcIII 73. TcIII was identified in patients with chronic ChD from Minas Gerais State 32 and in patients with the indeterminate form of ChD from Rio Grande do Norte State 69. Therefore, we cannot rule out the possibility of other parasite strains triggering the disease in humans in Minas Gerais. Further studies are needed to fully characterize the epidemiological and clinical aspects of TcI and TcIII in Brazil.
In summary, in this study, we demonstrated that BC detected the parasite in approximately half of the patients, whereas PCR was more effective for detecting T. cruzi in almost all samples, with the exception of one patient. Therefore, PCR and BC were efficient for detecting T . cruzi following collection of only a single blood sample, and the combined use of both methods did not increase positivity in the diagnosis of chronic Chagas disease. PCR does not allow the isolation of T. cruzi , emphasizing the importance of BC for further biological, biochemical, immunological, and genetic studies of parasite populations. Considering genotyping data, it was not possible to establish a correlation between the clinical form of ChD and the genetic profile of T. cruzi isolates.
