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Am. J. Trop. Med. Hyg., 69(3), 2003, pp. 309-313
Copyright © 2003 by The American Society of Tropical Medicine and Hygiene

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SHORT REPORT: QUANTIFICATION OF LEISHMANIAVIRUS RNA IN CLINICAL SAMPLES AND ITS POSSIBLE ROLE IN PATHOGENESIS

MONICA M. OGG, RICARDO CARRION, JR., ANA CRISTINA DE CARVALHO BOTELHO, WILSON MAYRINK, RODRIGO CORREA-OLIVEIRA, AND JEAN L. PATTERSON
Department of Microbiology University of Texas Health Science Center San Antonio, San Antonio, Texas; Department of Virology and Immunology, Southwest Foundation for Biomedical Research San Antonio, Texas; Laboratorio de Imunologia Celular e Molecular, Centro de Pesquisas Rene Rachou, FIOCRUZ, Belo Horizonte, Minas Gerais, Brazil; Departamento de Parasitologia Instituto de Ciencias Biologicas, Universidade Federal de Minas Gerais, Belo Horizonte, Minas Gerais, Brazil

 

ABSTRACT

Leishmaniavirus (LRV) is a double-stranded RNA virus that infects the protozoa Leishmania and has been identified in numerous strains of Leishmania braziliensis and L. braziliensis guyanensis. In general, the species of Leishmania dictates disease manifestation except in the case of L. braziliensis, which is capable of causing either cutaneous or mucocutaneous leishmaniasis. We wanted to determine 1) the quantity of LRV RNA present in a clinical sample and 2) if infection with LRV was associated with a specific disease manifestation. A real-time reverse transcriptase–polymerase chain reaction assay was used to assay clinical samples for the presence of LRV. Of 47 samples tested, 12 positive samples were obtained from patients with cutaneous lesions, lesions in the process of scarring, and cutaneous scars. This is the first study to examine the prevalence of LRV RNA within a small cohort from Brazil.

 

INTRODUCTION

Leishmaniasis is an infection caused by an intracellular parasite transmitted by sand flies. The human infection can display a range of manifestations from cutaneous involvement, to the destruction of mucous membranes, to a visceral disease with a fatal outcome. Each year, 1.5 million people worldwide develop cutaneous leishmaniasis and 500,000 develop visceral leishmaniasis.1 Within the 82 countries with endemic leishmaniasis, approximately 12 million are currently infected with leishmaniasis and 350 million people are at risk of acquiring this parasitic infection.2

Typically, the species of Leishmania responsible for the infection dictates the disease manifestation, except in the case of L. braziliensis, which can cause either cutaneous or mucocutaneous leishmaniasis. From 1% to 3% of those patients infected with L. braziliensis go on to develop the mucocutaneous form of leishmaniasis.3 Host factors such as gender, age, immune status, and area of anatomic exposure are associated with disease progression. However, another factor, the presence of Leishmaniavirus (LRV) within the Leishmania parasite, could be responsible for the variation of disease manifestation.3,4 Thus, the ability to predict the clinical outcome by identifying whether LRV is present is of great importance.

Leishmaniavirus is a double-stranded RNA virus with a genome approximately 5.3 kilobases in length that encodes two large open reading frames on the plus-strand.5 The first large open reading frame, ORF 2, encodes the 82-kD capsid protein.6 The second large open reading frame, ORF 3, encodes a 98-kD protein containing conserved RNA-dependent RNA polymerase motifs and therefore is predicted to be the viral polymerase.7–9 Leishmaniavirus has been detected in cultures of L. braziliensis, L. b. guyanensis, and Leishmania major.10 Viral RNA has been isolated from the cytoplasm in all three cultures, and shows no hybridization with nuclear extracts.5 Leishmania parasites are persistently infected with LRV. The virus can be eradicated using hygromycin B selection, which permanently removes the virus even after selection is discontinued.11 Leishmania cultures that are free of LRV infection are capable of being transiently infected with RNA from LRV.12

A reverse transcriptase–polymerase chain reaction (RT-PCR) assay was previously developed to detect LRV RNA in cutaneous leishmaniasis lesions from human biopsy tissues collected in Peru.13 The fact that viral RNA could be detected in human samples led us to develop a quantitative assay system that could detect the presence of viral RNA in clinical samples. A real-time RT-PCR assay system was used with primers and probe directed toward the projected third stem loop located within the 5' untranslated region (UTR). This region/sequence was conserved across all nine viral isolates whose sequences are known and therefore provide a region that may be preserved in other viral isolates that have not been identified or sequenced.

 

MATERIALS AND METHODS

The study was a blind study, with the samples being tagged only with a number until after all the samples were processed. Samples were collected from patients at the reference medical service for cutaneous leishmaniasis, Ambulatory Paulo de Magalhaes, located in the city of Caratinga in Minas Gerais, Brazil. The patients were from the rural regions of Vale do Rio Doce, an area endemic for cutaneous leishmaniasis. The patients were diagnosed by clinical examinations, Montenegro test, and/or direct observation for the presence of the parasite using imprint of tissue fragments obtained from the skin biopsies stained with Giemsa, and in some cases by staining of the tissue with hematoxylin and eosin. The results of the Montenegro test and parasitologic examination are shown in Table 1Go. Written informed consent was obtained from each subject prior to enrollment in the study. Ethical approval for this study was obtained from the Universidade Federal de Minas Gerais Ethics Committee on Studies with Humans. Samples were collected on cotton swabs that were passed over the scar/lesion, placed into a plastic bag, and then sent to the United States, where they were kept frozen at -20°C until processed. As shown in Table 2Go, samples included 8 cutaneous scars, 5 cutaneous lesions in the process of scarring, 28 cutaneous lesions (one of which is not due to leishmaniasis), 2 mucocutaneous lesions (one of which is not due to leishmaniasis), and 4 water controls. The samples came from a variety of subjects, both male and female, ranging in age from 4 to 75 years.


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TABLE 1
Results of tests confirming a diagnosis of leishmaniasis
 

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TABLE 2
Summary of samples examined for the presence of Leishmaniavirus (LRV) RNA
 
The initial step in our extraction process was to place the swab into 200 µL of RNAzol B (Leedo Medical Laboratories, Houston TX), followed by incubation at room temperature for 30 minutes. The liquid squeezed from the swab was subjected to RNA extraction following the manufacturer’s recommended protocol. Isolated RNA was resuspended in 50 µL of nuclease-free water. Positive control samples from a culture of L. b. guyanensis strain M4147, which is known to contain LRV, were processed in an identical manner to that of the clinical samples.

Nucleotides 171 to 251 from the LRV 1-4 sequence contain the region that house the primer and probe set for LRV detection. The following primers and probe were used: forward primer sequence: 5'-GAG TGG GAG TCC CCC ACA T-3'; reverse primer sequence: 5'-TGG ATA CAA CCA GAC GAT TGC T-3'; and LRV probe sequence: 5'-FAM-CTG CAT TTC ATG CAG TTC CTC ACG TTC C-TAMARA-3'.

Conditions for the real-time PCR assay are essentially the same as those described for the TaqMan Gold RT-PCR Kit (Applied Bioystems, Foster City, CA, and modified by R. Carrion and others, unpublished data). Before being placed into the reaction mixture, the samples were heated to 75°C for five minutes and then quickly chilled on ice to destabilize any RNA secondary structure. The reaction mixture (25 µL) contained 1x final TaqMan buffer A, 5.5 mM MgCl2, 200 µM dATP, 200 µM dCTP, 200 µM GTP, 400 µM dUTP, 0.2 pmol/µL of forward primer, 0.2 pmol/µL of reverse primer, 0.12 pmol/µL of probe, 0.4 units/µL of RNase inhibitor, 1.25 units/µL of MultiScribe reverse transcriptase (50 units/µL), 0.025 units/µL of AmpliTaq Gold DNA polymerase (5.0 units/µL), RNase-free water, and 5 µL of the sample. The real-time PCR conditions used were similar to those suggested by the manufacturer with one exception: due to the fact that there were mismatches between the probe sequence and some of the isolates’ sequences, specifically LRV 1-4, which was used to generate the standard curve, the annealing temperature was 50°C to ensure that the probe annealed to the sequence. The PCR conditions are as follows: one cycle reverse transcription at 48°C for 30 minutes, one cycle AmpliTaq Gold Activation and RT inactivation at 95°C for 10 minutes, 40 cycles of PCR denaturation at 95°C for 15 seconds, and annealing/extension at 50°C for one minute. This routinely allowed us to detect from 5 x 101 to 5 x 107 copies of LRV RNA per sample with a detection limit of 5–50 copies.

 

RESULTS

Table 1Go serves as the basis for how the samples were diagnosed as leishmaniasis. All but five samples were positive for the Montenegro test. Sample 5-1-a /5-1-b was positive when biopsy samples were examined for the presence of parasites. Samples 11, 19, and 40 were negative in both the Montenegro test and by parasitologic examination, but were positive based upon physician examination. (In an area where leishmaniasis is endemic, diagnosis based upon clinical examination is routine and definitive.) Sample 8 was diagnosed with leishmaniasis in 1991 by both a positive Montenegro test result and parasitologic examination. The subject then returned to the clinic in 2001 with nasal lesions where an earlier diagnosis of mucocutaneous leishmaniasis was made. Both the Montenegro test result and parasitologic examination in 2001 were negative, as well as the result of the PCR assay, which indicated that the destruction was not due to mucosal leishmaniasis but to carcinoma.

Forty-seven samples were processed for the presence of LRV (Table 2Go). Twelve (25.5%) of the 47 samples tested positive for the presence of LRV. The 12 samples that were positive contained cutaneous scars, lesions in the process of scarring, and cutaneous lesions; the one mucocutaneous scar was not positive. The number of viral genomes found in each sample ranged from 58 to 610,000 copies (Figure 1Go). There did not seem to be a correlation between the genome equivalents and type of sample in which they were found.



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  		    FIGURE 1. Samples that were positive for the presence of Leishmaniavirus (LRV) RNA and the quantification of that RNA. The sample data represent only the minimum genome equivalent for a single reaction of each sample. Samples show a range of types including scars, lesions in the process of scarring, and lesions, but not from the mucocutaneous sample. The detection limit (DL) for this assay was 5–50 copies/sample and is shown as the shaded region.

 
A natural infection model for leishmaniasis continues to be more clearly defined as different models for disease progression are used. A current low-dose challenge model revealed that there was a "silent" phase of infection, which is characterized by an absence of pathology but an increase in parasite number.14,15 Not only is there a "silent" phase that shows no pathology, but there is also the development of a chronic or persistent phase where parasites are maintained either in the lesion after it heals, in the lymphatic system, or in the spleen at nearly undetectable levels.14,16,17 Since there is a precedent for the development of a persistent phase during Leishmania infection, determining that each subject did indeed have (or had) leishmaniasis was a factor that was taken into consideration. As stated earlier, all patients who were determined to be positive for leishmaniasis were diagnosed by clinical examination, including direct observation of the parasite using imprints of tissues obtained from skin biopsies, which were stained with Giemsa to determine the presence of Leishmania parasites. Since parasite load is very difficult to determine by skin biopsy and is not a truly quantitative measure, a PCR assay to detect kinetoplast DNA was set up in an attempt to ascertain if Leishmania was present in the clinical samples. The DNA was reverse extracted from both the interphase and also the organic phase left at the end of the RNA extraction step of the RNAzol B protocol to determine if kinetoplast DNA could also be recovered and quantitated from these samples. The recovered DNA underwent PCR amplification. Controls for DNA extraction included a Leishmania culture sample extracted specifically for DNA and a sample reverse extracted in a manner identical to the extraction of the clinical samples. Both positive controls produced the expected 120-basepair fragment upon PCR amplification; however, the reverse extracted samples generated less product. None of the clinical samples were positive for kinetoplast DNA. Since LRV can be isolated from and visualized in infected parasites,18 the fact that kinetoplast DNA was not amplified would suggest that while the extraction method allows us to detect LRV RNA, the methodology is not optimal for DNA extraction and therefore determining parasite load.

 

DISCUSSION

This study further corroborates the results reported by Saiz and others13 in their study to detect LRV from Peruvian clinical samples. Their results indicated that two (18%) of 11 samples were positive for LRV RNA, which is similar to our value of 25.5%. The variance in this percentage could be due to the fact that the number of samples screened in our study was four times that of the previous study. The variability could also be due to an assay system that now uses primers and probes based upon a consensus sequence from nine viral isolates versus a system that was based upon a single viral sequence for the plus-sense primer and two viral sequences for the minus-sense primer. The use of the consensus sequence could increase the binding of the probe to a potentially diverse pool of viral isolates. Another factor is that the clinical samples previously surveyed were originally used for determining Leishmania species, whereas our sample types did not have such a restriction. This allowed us to include not only cutaneous lesions but also lesions in the process of healing, scars, and mucocutaneous samples as well. Our means of obtaining samples are much different from the punch biopsy method that Saiz and others used. Our procedure is certainly less invasive to the patient since only a swab is needed to collect samples. In addition, our method allows us to sample different stages within the disease progression such as active Leishmania infection, infections beginning to heal, and the scars that are the result of healing. The ability to look at samples at various stages of infection will allow us not only to follow disease progression of the parasite, but also to evaluate the presence of virus at each stage as well.

The results of the study indicate that there is no correlation between genome equivalents and type of lesion. The amounts of LRV genome equivalent detected in scars, lesions in the process of scarring, and active lesions were as variable as the sample types from which they were derived. Leishmaniavirus was even detected in a sample from a patient who was undergoing treatment. Although lesions in the process of scarring seem to have the lowest genome equivalents, this trend is not consistent across all samples. The sampling method may be partly responsible for some of the variability in genome equivalents detected. The only invariable tendency is that LRV is found in all types of samples except for the one mucocutaneous sample.

This study not only demonstrates the effectiveness of this new assay, but also allows us to further define the role LRV plays in disease manifestation by quantitating the number of genome equivalents present, as well as allowing us to analyze various types of clinical sample. Since the species of parasite determines clinical manifestation, and host factors such as age, gender, and immune status are able to affect disease progression, it is possible that LRV is another factor that is affecting disease progression. Although detectable virus levels, greater than 50 genome equivalents per sample, indicate that 25% of the samples are positive for virus, this does not take into account the species of parasite that caused the initial infection and that some species of Leishmania are not susceptible to viral infection. However, it is known that the prevalent species in Caratinga is L. braziliensis,19–21 which can be infected with LRV. It is also unlikely that the parasites carry viral isolates that do not share a similar sequence with identified LRV isolates, since the identity of the sequence is more than 90% conserved among known isolates. Due to the fact that only one available sample arose from mucocutaneous leishmaniasis, it is premature at this point to suggest that parasites causing the mucocutaneous form of leishmaniasis harbor very low numbers of LRV or are free of virus. More studies will need to be done to draw a more definite causal relationship between LRV and disease manifestation.

Received October 21, 2002. Accepted for publication June 16, 2003.

Acknowledgment: We thank Dr. Raymond Cologna for his reviews and suggestions.

Financial support: This work was supported by National Institutes of Health grant AI28473. Monica M. Ogg received financial support through a virology training grant (T32 AI07522).

Authors’ addresses: Monica M. Ogg and Ricardo Carrion, Jr., Department of Microbiology, University of Texas Health Science Center San Antonio, San Antonio, TX 78229, Telephone: 210-258-9865, Fax: 210-670-3329, E-mails: ogg{at}uthscsa.edu and carrion{at}sfbr.org. Ana Cristina de Carvalho Botelho and Rodrigo Correa-Oliveira, Laboratorio de Imunologia Celular e Molecular, Centro de Pesquisas Rene Rachou, FIOCRUZ, Av. Augusto de Lima 1715, Belo Horizonte, Minas Gerais, Brazil, 30190-002, Telephone: 55-31-3295-3566, Fax: 55-31-3295-3115, E-mail: correa{at}cpqrr.fiocruz.br. Wilson Mayrink, Departamento de Parasitologia, Instituto de Ciencias Biologicas, Universidade Federal de Minas Gerais, Av. Antonio Carlos 6627, Belo Horizonte, Minas Gerais, Brazil. Jean L. Patterson, Department of Virology and Immunology, Southwest Foundation for Biomedical Research, PO Box 760549, San Antonio, TX 78245-0549, Telephone: 210-258-9431, Fax: 210-670-3329, E-mail: jpatters{at}sfbr.org.

Reprint requests: Jean L. Patterson, Department of Virology and Immunology, Southwest Foundation for Biomedical Research, PO Box 760549, San Antonio, TX 78245-0549.

 

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