Open access peer-reviewed chapter
By Patrícia Sampaio Tavares Veras, Deborah Bittencourt Mothé Fraga, Manuela da Silva Solcà and Carlos Eduardo Sampaio Guedes
Submitted: September 19th 2013Reviewed: December 25th 2013Published: March 19th 2014
DOI: 10.5772/57573
Downloaded: 2051
Visceral leishmaniasis (VL) is a serious public health problem of great medical and veterinary importance. This disease is endemic in Brazil and in many other countries of Latin America, Asia, Africa and Europe (1). According to recent review (2), approximately 0.2 to 0.4 million cases of VL occur each year and although worldwide distributed, higher prevalence of the disease is concentrated in six countries, including India, Bangladesh, Sudan, South Sudan, Ethiopia and Brazil, that undertake for more than 90% of the cases. The clinical importance of VL resides in the severity of the disease that results in death of unrecognized cases and even for individuals with treatment access, death occurs in 10 to 20% of the cases [2-8].
Most of the VL cases are caused by the
The notion that dogs are the main urban domestic reservoir for this
Control strategies include performing accurate and early diagnosis of CVL to identify infected animals [19, 20]. CVL diagnosis is a difficult task since clinical signs of the disease in dogs can be confused with other diseases [19]. In endemic areas, a large percentage of infected animals are asymptomatic or present low number of discrete signs. The role these animals play in parasite transmission is still largely unknown. Several diagnostic strategies have been implemented based on parasitological, serological or molecular methods in association with clinical and epidemiological parameters [21]. Parasite culturing has been considered as gold standard for disease diagnosis [22, 23]. Although offering a high specificity since allows parasite identification, it offers very low sensitivity, besides it is laborious, time-consuming and largely dependent on the expertise of the observer [24, 25].
Serological tests are the most common diagnostic method employed for CVL diagnosis [3]. Several serological methods have been implemented for diagnosis of CVL, including direct agglutination assay (DAT), enzyme linked immunoassay (ELISA) and indirect immunofluorescent antibody test (IFI) [26]. However, most of these classical serological tests present important limitations for CVL diagnosis, including high consumption of time, and lack of sensitivity and specificity, mainly when animals present low antibody titers. This causes underestimation of disease, reflecting in failures in control measures, as well as the maintenance of infected untreated dogs in endemic areas [27, 28]. New methods based on immunochromatography have been implemented for serodiagnosis of CVL and have shown excellent results [29]. These techniques offer several advantages since they are rapid tests easily performed even in field areas, and more specific since they use recombinant DNA technology that additionally facilitates reproducibility and large-scale production. These advantages result in better identification of infected dogs. However, the efficacy of immunochromatographic techniques for CVL diagnosis needs to be improved [30]. In Brazil, a rapid test based in dual path platform (TR DPP®LVC - Biomanguinhos) had been recently implemented as screening test for CVL. This technique seems to be adequate to disease diagnosis in public health system. However, the TR DPP®LVC has shown an excellent performance identifying 98% of symptomatic dogs, it showed less efficacy for diagnosis of asymptomatic dogs (47%) [31]. Since there is evidence that asymptomatic dogs can participate in natural transmission cycle of VL, new strategies should be implemented in order to improve CVL diagnosis [16, 32-34]. For serological diagnosis one strategy can be the development of rapid tests based on impregnation of multi-antigen that would offer more sensitivity, as well specificity.
Finally, it would be important to include more specific confirmatory tests for control strategies that can be advantageous to diagnose inconclusive cases. There is evidence that molecular diagnosis of
Since the discovery of canine visceral leishmaniasis (CVL) in Tunisia by Nicolle & Comte (1908), several reports have shown that dog and man share a common etiologic agent. The notion that dog is the main reservoir of visceral leishmaniasis (VL) in urban centers [38] is supported by several evidences including the high cutaneous parasitism observed in dogs infected by
Some studies have shown a correlation between the presence of clinical signs in infected animals and transmissibility of the parasite to the vector and, consequently, a correlation with the occurrence of human cases [16, 32, 51]. In accordance with these studies, Travi et al. (2001) and Verçosa et al. (2008) showed that asymptomatic dogs did not transmit the parasite to the vector [38, 51]. There is not a consensus about this idea, since there is a wide variation in the rates of infectivity (70 to 90%) between asymptomatic and symptomatic dogs. Studies show that, regardless of the clinical presentation, any dog has the ability to transmit
The diagnosis of VL in the dog must consider the association between clinical, laboratory and epidemiological data. As discussed above, clinical diagnosis is problematic and difficult for veterinarians to perform due to the great variability of clinical signs that
There are several laboratorial diagnosis methods for leishmaniasis: i) parasitological methods (detection of the parasite), ii) serological methods (detection of anti-
In spite of serological techniques such as enzyme-linked immunosorbent assay (ELISA) and indirect immunofluorescence assay (IFAT) being the most widely used methods for the diagnosis of CVL [60] parasitological methods, such as direct examination of slides and isolation from tissue cultures, allow the parasite to be detected and can be used as confirmatory diagnostic methods for CVL [61]. In recent decades, molecular techniques such as polymerase chain reaction (PCR) have been introduced for the diagnosis of CVL, exhibiting high sensitivity and specificity [21]. These techniques detect the genetic material of the parasite, which can be used as confirmatory methods in cases of recently infected or asymptomatic animals, which tend not to be diagnosed serologically, and in most cases, do not show seroconversion, having a low parasite load [4, 60]. In a study conducted in Belo Horizonte-MG, a VL-endemic area in Brazil, among 1,443 dogs evaluated, 15.3% of them were seropositive, while 84.7% showed negative serology. Interestingly, among serologically negative dogs, 24.4% showed up as positive using the molecular diagnostic technique, and most of these (97.6%) would not be diagnosed, since they consist of asymptomatic dogs with negative serology [19].
Dogs from endemic areas considered resistant remain clinically normal and asymptomatic without exhibiting clinical signs. There is evidence that the parasites in these animals are effectively eliminated at the infection site [62, 63]. However, in susceptible animals, a large number of parasites are detected in infected tissues. In these animals, the presence of the parasite may occur in multiple organs, accompanied by a granulomatous inflammatory reaction and production of immune-mediated phenomena, probably responsible for the appearance of various types of clinical signs [64].
Initial clinical signs of CVL include: hypertrophy of the lymph nodes, changes in skin appendages such as onychogryphosis, swelling of the footpad, localized alopecia, skin ulcers and nasal and periocular dermatitis. Alopecia and non-pruritic exfoliative dermatitis can spread to other parts of the animal's body. Weight loss may also be present, as well as cachexia, anorexia and conjunctivitis. Internal organs such as spleen, liver, kidney and lymph nodes may also be affected, when kidney injuries are present may lead to the dogs death [13, 65]. Fever, apathy, diarrhea, epistaxis, intestinal bleeding, hepatosplenomegaly, hyperkeratosis, keratoconjunctivitis are also found in affected animals [66-68]. Some clinical signs are more frequent than others; skin lesions are the most frequent manifestations affecting approximately 50 to 90% of symptomatic dogs [4, 67, 69, 70], including non-pruritic exfoliative dermatitis, with or without alopecia, which can be generalized or localized to the muzzle, ears and limbs [67, 71, 72]. Other very common signs are weight loss, observed in 25 to 80% of CVL cases, including onychogryphosis in 30 to 75%, and ocular abnormalities in 16 to 24% [28]. The most common clinical signs of VL in dogs are depicted in Figure 1.
Major clinical signs associated with CVL. A: alopecia on the muzzle, B: periocular dermatitis with keratoconjunctivitis and hyperkeratosis; C: hyperkeratosis of the nasal mucosa; D: generalized non-pruritic exfoliative dermatitis; E: ulcerated lesion in the ear; F: crust with vascular injury on the tip of the ear; G: lymphadenomegaly of the popliteal lymph node; H: cachexia; I: onychogryphosis. Photos of animals infected by
In dogs with CVL, clinical-pathological changes may occur such as intestinal lesions, renal and hepatic abnormalities [73]. The main biochemical laboratory findings from CVL are hyperglobulinemia, mainly due to increased production of antibodies, and hypoalbuminemia, attributed to chronic inflammation, as long as renal and hepatic failure [66]. The result of these changes is a reduction in the albumin/globulin ratio and hyperproteinemia [28]. Additionally, severe CVL is associated with changes in hematological parameters such as severe anemia and leukopenia, associated with lymphopenia, eosinopenia and monocytopenia [66, 74, 75]. Immune-mediated thrombocytopenia also occurs accounting for episodes of bleeding such as epistaxis, hematuria and hemorrhagic diarrhea [76].
Finally, nonspecific signs of illness that are mistaken for other diseases such as babesiosis, ehrlichiosis and canine trypanosomiasis also contribute to make CVL clinical diagnosis imprecise and difficult to perform [13].
The detection by optical microscopy of the parasite by direct observation of stained smears from spleen aspirate, lymph node and bone marrow tissues has high specificity, allowing confirmation of CVL diagnosis [3, 53, 61, 77]. However, the sensitivity of this method is less than 30%, since the direct parasite identification may be limited, especially in mildly and asymptomatic dogs that have low parasitic load, producing false negative results [3, 53, 61, 77].
Another method that can identify the parasite in tissues is the culturing of tissue fragments or aspirates, preferably in a biphasic medium [78], composed by Novy-MacNeal-Nicolle (NNN), or Tobie modified medium or United States Army Medical Research Units (USAMRU) as solid phase medium and, most often, Schneider as liquid phase medium. This parasitological diagnostic method offers high specificity allowing isolation and characterization of parasites, as well as determination of which species and/or variants are circulating in endemic areas [79]. However, the culturing consists of an indirect test, because when the parasites are isolated from various tissues, they are present in amastigote form and during cultivation they transform into the promastigote form. This process may be impaired as a result of parasite death due to a failure of temperature-control during transport of the tissue sample, or contamination during collection or cultivation [13]. Additionally, a culturing is time consuming and may take up to 4 weeks of observation for definitive diagnosis [13, 79]. Furthermore, specific media for promastigote isolation are not easily obtained, being a technique restricted to specialized laboratories [70, 80], in which the outcome also depends on the experience of the observer [24, 25]. Although culturing offers greater sensitivity compared to direct viewing of amastigotes in tissue [35], it still remains at very low levels.
In summary, parasitological techniques have high specificity but low sensitivity, especially for the detection of dogs, recently infected, asymptomatic or those presenting low parasite load. In addition, the need for skilled personnel and the long delays to obtain the results prevent parasitological techniques to be used in epidemiological surveys [4, 23, 61, 81-84].
Serological tests are based on the presence of specific humoral immune responses against the pathogen or purified fraction or recombinant proteins of the pathogen. These tests allow detection of immunoglobulin (IgG) levels, thus becoming an essential tool for the diagnosis of CVL. These methods are simple to carry out and therefore they are frequently used to determine the prevalence of leishmaniasis in epidemiological studies [85].
A wide variety of serological methods are available for CVL diagnosis, presenting variations in sensitivity and specificity. The performance of these diagnostic techniques varies depending on the type of antigen used and the detection of anti-
The most commonly employed serological tests for the diagnosis of CVL, including ELISA, indirect immunofluorescence test (IFAT), and direct agglutination test (DAT), uses parasite or crude extract of
Despite the practicality and simplicity of serological tests, they do not have 100% sensitivity because some dogs, especially those that are resistant or in the early stages of the disease, have negative results. Thus, the results of such tests should be evaluated carefully, always associating test results with epidemiological history, clinical state of the animal, and the result of a more specific diagnostic test [86]. In addition, since titers of anti-
IFAT
IFAT is a test in which anti-immunoglobulin antibodies labeled with fluorochromes react with parasites immobilized in a slide. IFAT is a laborious technique that presents difficulties for both standardization and interpretation of the results Therefore, detection of antigen-antibody reaction by fluorescence microscopy depends on the observer experience, compromising reproducibility of this test in different laboratories. Thus, it is not considered a simple and practical technique for evaluating a large number of canine sera [57]. In spite of these limitations, it is still being used as a diagnostic method for mass screening of infected dogs [87]. This method varies in its performance, with sensitivity ranging from 68 to 100% and specificity of 60 to 90% [5, 88-90].
In a study evaluating IFAT for the diagnosis of CVL, the efficacy of the test was evaluated using 254 sera from infected and uninfected dogs and sera from animals with other parasitic diseases. The authors observed low sensitivity (72%) and specificity (52%), as well as cross-reactions when sera from dogs infected with other pathologies, such as
DAT
The direct agglutination test (DAT) is an alternative method for the diagnosis of VL, first described in 1975 and adapted for the diagnosis of human and canine infection in the late 1980s [93, 94]. DAT is a method that uses whole stained promastigotes as antigen, either in suspension or freeze-dried [35]. The advantage of this test lies in its low cost when compared with other tests [93]. However, this test is not desirable for screening large numbers of samples, since it is a laborious procedure, due to the production process for crude
Changes to the DAT protocol have been proposed by Gómez-Ochoa
ELISA
For various reasons, ELISA tests based on whole parasites or crude lysate of parasite antigens for the diagnosis of CVL do not provide satisfactory results, as follows: i) it is a laborious technique, which leads to a delay in the delivery of results and, consequently, the implementation of treatment or the removal of infected dogs from endemic areas [68, 99]; ii) leads to the appearance of cross-reactions with sera from individuals infected with other
A study using 234 domesticated dogs in an endemic area for CVL assessed the efficacy of ELISA, IFAT and DAT for the diagnosis of CVL. In this study, dogs were also parasitologically evaluated for identification of
Using sera from dogs with CVL, a comparison of an ELISA test using crude soluble antigen of
Thus, the search for tests with higher sensitivity and specificity for dogs with a variety of conditions became necessary for control of CVL, which would lead to a reduction of errors in actions taken for treatment or control. In countries that adopt culling of seropositive dogs as a control measure, low sensitivity of diagnostic tests can lead to the maintenance of dogs that transmit disease and lack of specificity can result in unnecessary culling of healthy dogs. The identification of new proteins of
Another way to overcome the obstacles of ELISA based on whole parasites or crude parasite antigen was the development of ELISA tests based on parasite fractions such as that using the parasite surface molecule, fucose-mannose ligand antigen (FML). The FML-based ELISA showed a high sensitivity, which was similar in detecting either oligosymptomatic (90%) or symptomatic (90%) dogs. Regarding specificity, ELISA using crude parasite antigen for the diagnosis of oligosymptomatic dogs was superior, achieving 100% in comparison to FML-based ELISA that was 93.3%. However, for symptomatic dogs the specificity of the FML-based ELISA showed similar results of 96.7% compared to that obtained by ELISA based on crude parasite antigen (93.3%) [101].
Other ELISA assays based on recombinant antigens such as rA2 from
Interestingly, the association of the recombinant proteins enhanced test performance both for detection of symptomatic and asymptomatic infected dogs. Indeed, using IFAT as the gold standard, ELISA based on the mix of rK9, rK26 and rK39 from
The combination of these findings reinforces the notion that the use of multiple antigens in diagnostic tests enhances test performance and the need to search for new antigens that may compose a diagnostic test able to better diagnose asymptomatic dogs.
New recombinant proteins are being evaluated. Faria
Another study evaluated the performance of the ELISA based on another recombinant antigens of
In summary, most studies using ELISA suggest that in comparison to tests based on crude antigen, those based on recombinant antigens improves accuracy, increasing sensitivity and specificity for the diagnosis of symptomatic dogs. Although improved, test accuracy is still low for the detection of asymptomatic animals.
Rapid tests
Recently, rapid immunodiagnostic tests have begun to be employed as routine laboratory tests for detection of diseases such as leishmaniasis. The recombinant antigens of the parasite are impregnated onto nitrocellulose membranes and serum samples are applied in the rapid test platform. Antigens impregnated in nitrocellulose membranes are recognized by specific immunoglobulin present in the serum of infected individuals. This reaction is revealed by the interaction of protein A coupled to colloidal gold particles, with the Fc portion of the immunoglobulins associated with the recombinant antigens. The use of immunochromatographic assays as diagnostic methods has the main advantages of being rapid, completed in around 15 minutes, easy to carry out and can dispense with the need for equipment to read the results [110]. Furthermore, these tests are easily stored, and test supplies and samples do not need to be maintained at low temperatures and can it even be performed at the place of collection. These tests are already widely used to detect HIV [111] and H1N1 [112] infection. For the diagnosis of CVL and human VL, among the tested and commercially available recombinant proteins, the most widely used for composing immunochromatographic tests is the recombinant protein rK39. This protein contains repetitive sequences of 39 amino acids from a protein related to kinesin of kinetoplast from
Recently, a meta-analysis was performed in order to broadly assess the performance of rapid tests using rK39 as the antigen in the diagnosis of CVL. The combined analysis of 16 studies using rapid tests based on rK39 offered a sensitivity of 86.7% (95% CI: 76.9–92.8%) for the detection of clinical disease and 59.3% (95% CI: 37.9–77.6%) for identification of
| Harith | DAT | 44 | 6 | 176 | 98.9 | 100 | |
| Barbosa-de-Deus | ELISA | LMS | 188 | 1582 | 55 | 98.0 | 95.0 |
| Scalone | ELISA | K39 | 209 | 81 | 62 | 97.1 | 98.8 |
| Schallig | DAT | 79 | 67 | 24 | 88.6 | 96.7 | |
| FAST | 79 | 67 | 24 | 93.6 | 89.0 | ||
| Rosati | ELISA | K26 | 202 | 20 | 0 | 100 | 100 |
| ELISA | K9 | 202 | 20 | 0 | 95 | 95 | |
| ELISA | K39 | 202 | 20 | 0 | 95 | 95 | |
| Mohebali, | Dipstick | rK39 | 268* | 0 | 0 | 70.9 | 84.9 |
| Boarino | ELISA | K9-K39-K26 chimera | 232 | 362 | 0 | 95.8 | 99.1 |
| Mettler | Rapid test | rK39 | 47 | 50 | 26 | A: 52.9 S: 96.7 | 94 |
| IFAT | 47 | 50 | 26 | A: 29.4 S: 90.0 | 100 | ||
| Lira | EIE® - LVC | 25 | 16 | 11 | 72.0 | 87.5 | |
| IFI® - LVC | 25 | 16 | 11 | 68.0 | 87.5 | ||
| Ferreira | EIE® - LVC | 234* | 20 | 20 | 96.0 | 100 | |
| IFI® - LVC | 234* | 20 | 20 | 72.0 | 100 | ||
| DAT | 234* | 20 | 20 | 93.0 | 100 | ||
| Ferroglio | SNAP® CLATK | CTA | 59 | 124 | 0 | 91.1 | 99.0 |
| Porrozzi | ELISA | rK26 | 100 | 25 | 14 | A: 66.0 S: 94.0 | 90.0 |
| ELISA | rK39 | 100 | 25 | 14 | A: 66.0 S: 100 | 85.0 | |
| ELISA | rA2 | 100 | 25 | 14 | A: 88.0 S: 70.0 | 96.0 | |
| ELISA | CTA | 100 | 25 | 14 | A: 30.0 S: 88.0 | 87.0 | |
| Cândido | ELISA | CTA | 60 | 30 | 0 | O: 86.7 P: 90.0 | O: 100 P: 93.3 |
| ELISA | FML | 60 | 30 | 0 | O: 90.0 P: 86.7 | O: 93.3 P: 96.7 | |
| Lemos | RDTs | rK39 | 76 | 33 | 0 | 83 | 100 |
| ELISA | 76 | 33 | 0 | 95 | 100 | ||
| Babakhan | FAST | 73 | 74 | 0 | 98.6 | 78.7 | |
| Coelho | ELISA | LRP | 111 | 47 | 14 | 100 | 98.2 |
| ELISA | CTA | 111 | 47 | 14 | 96.0 | 100 | |
| Troncarelli | IFAT | 51 | 0 | 0 | 83.0 | 92.5 | |
| Figueiredo | EIE® - LVC | 305* | 0 | 0 | 100 | 96.6 | |
| IFI® - LVC | 305* | 0 | 0 | 22.2 | 97.0 | ||
| de Lima | ELISA | CTA | 52 | 52 | 0 | 91.5 | 94.7 |
| RDTs | rK39 | 52 | 52 | 0 | 100 | 91.2 | |
| Marcondes | SNAP® CLATK | CTA | 283 | 86 | 31 | 94.7 | 90.6 |
| Alves | EIE® - LVC | 39 | 39 | 39 | 100 | 68.0 | |
| ELISA | 39 | 39 | 39 | 100 | 93.6 | ||
| IFI® - LVC | 39 | 39 | 39 | 100 | 70.5 | ||
| IFAT | 39 | 39 | 39 | 100 | 61.5 | ||
| DPP® - LVC | rK28 | 39 | 39 | 39 | 100 | 97.5 | |
| Grimaldi | DPP® - LVC | rK28 | 120 | 59 | 11 | A: 47.0 S: 98.0 | 96.0 |
| Souza | ELISA | rLci1A | 138 | 119 | 86 | 96.0 | 92.0 |
| ELISA | rLci2B | 138 | 119 | 86 | 100 | 95.0 | |
| Barral-Veloso | ELISA | 31 | 37 | 45 | 93.5 | 97.6 | |
| ELISA | 31 | 37 | 45 | 87.1 | 100 | ||
| Quinnell | RDTs | rK39 | 322 | 59 | 0 | 46.0 | 98.7 |
Sensitivity and specificity of some serological techniques by type of antigen, and evaluated population
* Dogs from an endemic area for CVL without confirmed diagnosis.
A: asymptomatic dogs; S: dogs with clinical signs related to CVL; O: oligosymptomatic dogs (presenting one to three clinical signs of CVL); P: polysymptomatic dogs (presenting more than three clinical signs of CVL).
RDTs = Kalazar Detect - Canine Rapid Test
LMS =
CTA = Crude Total Antigen - antigenic fractions of
FML = Fucose-Mannose Ligand antigen - glycoprotein complex isolated from the surface of
LRP =
Efforts have been made to improve the efficacy of rapid tests by developing more sensitive and specific method that could be used in mass screening for the diagnosis of CVL. An alternative proposal is to use a mixture of recombinant proteins or chimeric proteins. The protein rK28 chimeric for the relevant epitopes of three antigens, rK9, rK26 and rK39 [87, 108] that showed promising efficient results in an ELISA based test [124], was recently used to compose a new rapid test in DPP format. This format consists of a double track platform that offers greater sensitivity and specificity [125]. In addition, this rapid test has advantages over previously used serological methods due to greater precision, simplified interpretation of the data, minimal use of sample volumes, and compatibility with different types of body fluids such as blood, serum, saliva, plasma and urine. In contrast to these advantages, recently Grimaldi et al (2012) showed that rK28-based DPP despite its high sensitivity (98%) and specificity (96%) towards sera from symptomatic dogs, showed low sensitivity of only 47% towards sera from dogs with no signs [31]. With regard to sera from dogs with other diseases, the observed specificity was 96%, with false-positive reactions mainly for some sera of dogs infected with
In recent decades, due to advances in molecular biology techniques and reduced implementation costs, the polymerase chain reaction (PCR) began to be used in VL diagnosis [23, 126]. Its use has demonstrated superior results to those obtained by ELISA, IFA and culture in detecting animals infected with
PCR is a technique based on the principle of complementary bases pairing of the DNA molecule, allowing amplification and detection of a particular region of the target genome using a pair of specific oligonucleotide primers. The reaction can produce tens of billions of DNA fragments from a single molecule, and has high sensitivity small quantities of samples to be used. This type of PCR, hereafter referred as "conventional PCR" (cPCR) needs electrophoresis in agarose or polyacrylamide gels along with dyes such as ethidium bromide, SYBR Green or silver nitrate to view the amplified product. This approach is usually qualitative, with analysis of the presence or absence of bands, or semi-quantitative, when densitometry of bands is used in comparison with known standards. Since it uses qualitative or semi-quantitative analysis, it is imprecise and generates false negatives with some frequency.
A variant of cPCR called "quantitative real-time PCR" (qPCR) became popular in the 2000s. It uses a quantitative approach that allows real-time monitoring of the amplification of the target PCR fragment using fluorophores that bind to double stranded DNA or linked to probes. The most commonly used method is SYBR Green: fluorophore binds to double stranded DNA molecules produced during amplification of the target fragment, leading to the emission of fluorescence during the PCR. This method has the disadvantage of not being able to directly discriminate the amplification of nonspecific DNA fragments, which is usually solved by analyzing the dissociation curve. In contrast, the TaqMan method uses a probe containing between 13 and 30 nucleotides, specifically for the target sequence and combined with a fluorophore and a fluorescence inhibitor. During polymerization of the target fragment, DNA polymerase degrades the probe and fluorescence is emitted. The use of this technique enables an increase in the specificity of this method.
Various PCR-based protocols have been developed for the detection of parasite's DNA and CVL diagnosis. However, the methods used may vary with respect to several parameters, such as fluorophores, probes, target regions and tissue used for detection of target DNA (Table 2), making it difficult to do a comparative analysis between the different protocols. It is known that the sensitivity and specificity of PCR for detection of
The PCR protocol sensitivity is also affected by the type of tissue used in the detection of
The selection of target region in the parasite genome is important because the variation in the number of copies, depending on the region, influences the sensitivity for detecting the parasite's DNA and for quantification of parasite load. The highly conserved and repetitive regions are the most commonly employed, such as the gene for subunit ribosomal RNA (rRNA) or minicircle kinetoplast DNA (kDNA) [21, 23, 127, 140, 141], that has 40-200 copies per cell, while the kDNA minicircles have about 10,000 copies distributed among 10 different classes of sequences. Using this as a target region confers high sensitivity to PCR [142]. For quantification of the parasitic load is recommended to normalize the amount of parasite gene amplification in relation to a constitutive gene derived from the host genome in order to correct distortions caused by errors in the DNA used in the PCR reaction [127].
| Ferreira et al. 2012 | Syber α pol DNA | NI | Yes | Yes | ß - canine actin | (80) Infected dogs | Conjunctival swab, blood, bone marrow and skin | Comparative1,2 | Skin > Bone marrow > Conjunctival swab > Blood | |
| Solcà | TaqMan kDNA | 0.01 parasites/ reaction | Yes | Yes | 18S eukaryotic rRNA | (51) Dogs | Bone marrow, conjunctival swab, lymph node, skin and spleen | Comparative1,2 | Spleen > Blood > Lymph node > Skin > Bone marrow > Conjunctival swab | |
| Belinchón-Lorenzo et al. 2013 | TaqMan kDNA | 0.0079 parasites/ reaction | Yes | Yes | 18S eukaryotic rRNA | (28) Dogs | Blood, hair and lymph node | Comparative 2 | Lymph node > Hair = Blood | |
| Ferreira et al. 2013 | Syber α pol DNA | NI | Yes | Yes | ß - canine actin | (62) CVL positive dogs | Conjunctival, nasal and ear swab, blood, Bone marrow and skin | Comparative 1,2 | Skin = Nasal swab and bone marrow > Conjunctival swab > Oral swab > Ear swab | |
| Geisweid et al. 2013 | Syber kDNA | NI | Yes | No | Canine NCX1 | (74) CVL suspected dogs | Conjunctival swab, blood, bone marrow and lymph node | Comparative 2 | Bone marrow > Conjunctival Swab | |
| Reis | Syber α pol DNA | NI | Yes | No | G3PDH | (60) Seropositive dogs | Skin and spleen | Comparative 1,2 | Spleen > Skin | |
| Pennisi | NI kDNA | NI | No | No | --- | (6) Treated dogs | Blood, lymph node and skin | Not comparative | --- | |
| Francino | TaqMan kDNA | 0.001 parasites/ reaction | Yes | No | 18S eukaryotic rRNA | (15) Dogs with clinical signs suggestive of CVL | Blood and bone marrow | Comparative1, 2 | Bone marrow > Blood | |
| Rodriguez-Cortez | TaqMan kDNA | 0.001 parasites/ reaction | Yes | Yes | 18S eukaryotic rRNA | (6) Experimentally infected dogs | Blood, bone marrow, liver, lymph node, skin and spleen | Not comparative | --- | |
| Solano-Gallego | Syber kDNA | 7 parasites/ml | Yes | No | Canine GAPDH | (10) Symptomatic dogs naturally infected | Blood, bone marrow and urine | Comparative 2 | Bone marrow > Blood > Urine | |
| Manna | TaqMan kDNA | 0.001 parasites/ml | Yes | Yes | ß - actin | (18) Naturally infected treated dogs | Blood, lymph node and skin | Comparative 2 | Lymph node > Skin > Blood | |
| Manna | TaqMan kDNA | NI | Yes | Yes | ß - actin | (56) Dogs | Blood and lymph node | Not comparative | --- | |
| Quaresma | Syber kDNA | 0.1pg DNA/ml | Yes | Yes | ß -canine globin | (35) Dogs | Blood and bone marrow | Comparative2 | Blood = Bone marrow | |
| Maia | TaqMan kDNA | 1 parasite /reaction | Yes | No | ß - canine actin | (12) Experimentally infected dogs | Blood, bone marrow, buffy coat, liver, lymph node, skin and spleen | Comparative1, 2 | Spleen / Buffy coat / Liver / Lymph node / Bone marrow / Skin > Blood | |
| Galletti | TaqMan kDNA | 0.03 parasite/ reaction | No | No | --- | (88) Dogs | Conjunctival swab, Lymph node, bone marrow and blood | Comparative1 | --- | |
| Lombardo | TaqMan kDNA | NI | No | No | --- | (138) Dogs | Blood, conjunctival and oral swabs and lymph node | Comparative1 | --- | |
| Naranjo | TaqMan kDNA | NI | Yes | No | 18S eukaryotic rRNA | (22) Sick dogs | Main lacrimal gland, tarsal gland and nictitating membrane gland | Comparative1 | --- | |
List of papers using the qPCR technique for
* Study design: Comparative1: aim to compare different diagnostic techniques results; Comparative2: aim to compare different canine tissue sensitivity; ** Sensitivity: evaluation of
In a cytological study, Reis
Splenic collection, bone marrow and lymph node aspirates are considered invasive procedures [153] in addition to having an elevated cost compared to blood collection. Thus, it can be recommended to use samples obtained less invasively, such as blood and conjunctival swabs [136, 154, 155]. These samples are quick and easy to obtain, and it is low-cost compared to more invasive procedures, in addition to their higher acceptance by animal owners [132, 154, 155].
Some studies have shown that detection of parasites in the peripheral blood is less sensitive compared to other tissue samples such as spleen, bone marrow, lymph nodes and skin and tends to have variable parasitic load in accordance with the stage of infection [129, 141, 156]. However, depending on the technique and the target, blood can be used for detection of
According to Solano-Gallego
Among other less invasive sample types investigated, Solano-Gallego et al (2007) evaluated urine samples with qPCR technique, but the results described showed positivity only in dogs with severe renal injury [160]. Naranjo et al. (2012) identified the presence of
| Elleviti – Torino, Italy | 26.80* | --- | 63.00* | --- |
| Scanelis - Toulouse, France | --- | --- | 60.30* | --- |
| Laboratoire d'Anatomie Pathologique Vétérinaire du Sud-Ouest – Toulouse, France | --- | --- | --- | 127.30* |
| Laboratório Veterinário INNO – Braga, Portugal | 20.60* | 54.40* | --- | --- |
| Instituto Nacional de Investigação Agrária e Veterinária, I.P. – Lisbõa, Portugal | 28.00* | 41.20* | --- | --- |
| Centro de Investigación y Análisis Biológicos – Madrid, Spain | 13.60* | 60.30* | 73.70* | --- |
| Texas Veterinary Medical Diagnostic Laboratory – San Antonio TX, USA | 19.20 | --- | --- | --- |
| Cornell University - Ithaca NY, USA | 22.50 | 60.00 | --- | --- |
| Hermes Pardini - Belo Horizonte MG, Brazil | 17.20* | 60.20* | --- | --- |
| Análisis Biológicos– Chapecó SC, Brazil | 9.40* | 42.15* | 72.25* | -- |
| Laborlife - Rio de Janeiro RJ, Brazil | 30.10* | 77.40* | --- | --- |
Cost of the main diagnostic test for CVL in different laboratories consulted in the second semester of 2013 in U$
*Prices in Euros (€) and Brazilian Real (R$) converted in US dollars (U$), quotation of November 14th of 2013. €1 = aprox. U$ 1.34 - €1 = aprox.R$ 0.43
Despite the high sensitivity and specificity, the use of molecular methods for the CVL diagnosis presents some limitations to its use in epidemiological surveys: i) it has higher costs than other techniques (Table 3) used in the CVL diagnosis, including reagent and equipment costs; ii) it presents relative complexity in its implementation, requiring personnel with training in the execution of PCR reactions. However, this method has advantages in terms of sensitivity and specificity when compared to other diagnostic techniques, which justify its use in confirming cases screened by serology [24, 132]. Particularly due to the possibility of quantifying target DNA, qPCR may be used to monitor the parasitic load of the animal during the experimental infection, or during and after treatment in countries where it is permitted [35-37, 162]. Compared with cPCR, qPCR enables a reduction in the probability of false positives resulting from amplification artifacts and greater speed in obtaining results, once electrophoresis is no longer performed [163].
In summary, detailed clinical evaluation complemented with highly sensitive test allows proper identification of infected dogs in an endemic area. Evidence shows that the use of a rapid serological test associated with a molecular diagnostic test with high specificity, such as qPCR, is required for identification of all infected dogs, both asymptomatic and symptomatic. On the other hand, for sick dogs a correct diagnosis is necessary either to perform dog culling in countries where this measure is used as a control strategy of VL or to define treatment. In this case, a detailed clinical evaluation should be associated with biochemistry and hematological tests to identify signs of renal and hepatic failure, in conjunction with a serological test to confirm animal clinical condition.
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