Methods for the diagnosis of blood-borne parasitic infections have stagnated in the last 20–30 years. However, recently, there has been a tremendous effort to focus research on the development of newer diagnostic methods focusing on serological, molecular, and proteomic approaches. This article examines the various diagnostic tools that are being used in clinical laboratories, optimized in reference laboratories and employed in mass screening programmes.
by A. Ricciardi and Dr M. Ndao
Blood-borne protozoans are the causative agents of some of the world’s most devastating and prevalent parasitic infections. This group of pathogens includes members of the Trypanosoma, Leishmania, Plasmodium, Toxoplasma, and Babesia genera. Most of these infections, with the exception of toxoplasmosis and babesiosis, have always been described as being tropical or subtropical. However, the increase in international travel as well as the arrival of new immigrants has made some of these tropical diseases realities in developed countries as well. In addition, infection via contaminated blood (transfusions and organ transplants) has become a major problem. Clearly, the transmission of blood-borne protozoans is boundless and the actual number of cases is underestimated. Quick diagnosis has always been a priority in order to determine the appropriate treatment and prevent fatalities. In addition, now more than ever, advances in diagnostics can help prevent transmission and provide active surveillance. Currently, diagnostic and reference laboratories use an array of techniques including microscopy, serological assays, and molecular assays. Here, the advantages and disadvantages of the methods will be discussed.
Toxoplasmosis
Toxoplasmosis, caused by Toxoplasma gondii, has a worldwide distribution. In immunocompetent individuals, more than 80% of primary Toxoplasma infections are asymptomatic [1]. Toxoplasmosis becomes a problem when an individual is immunocompromised or during pregnancy. Diagnosis of toxoplasmosis varies according to the immune status of the patient.
Diagnosis of immunocompetent individuals relies on serology. Early antibody responses can be detected via methods such as the dye test, immunofluorescent assay, and agglutination test whereas later IgG titres are detected by enzyme-linked immunosorbent assay (ELISA). For many years, the Sabin-Feldman dye test was the gold standard diagnostic technique due to its sensitivity and specificity. In recent years, few laboratories have continued to use this method and rather focused on newer techniques such as indirect immunofluorescent antibody tests, hemagglutination tests, capture ELISAs, and immunosorbent agglutination assays (ISAGAs). Serological assays lack the capacity to differentiate between recent and older infections; IgM levels can persist for over two years [2]. In order to determine whether an infection is recent, avidity ELISA is performed. This assay verifies IgG avidity and is based on the concept that as the immune response progresses, an immunoglobulin’s affinity for a specific antigen will increase [3].
Diagnosis of Toxoplasma infection during pregnancy is crucial in order to prevent congenital toxoplasmosis. Prenatal diagnosis involves performing real-time polymerase chain reaction (PCR) using amniotic fluid. The PCRs used often target the B1 gene of the parasite [1]. Upon delivery, PCR is performed on either the placenta or the cord blood serum in order to detect parasites. ISAGAs are also often performed. If the tests are positive, cord blood samples at one week of life are sent to a reference laboratory [1]. Follow up serology is again performed at one month and then every two to three months. There have been recent advances in the field of toxoplasmosis post-natal diagnosis. An ELISA assay that measures interferon-gamma levels upon stimulation of whole blood cells with Toxoplasma crude antigens has been developed. This method has proven to be both sensitive and specific [4].
In the case of immunocompromised patients, a quick diagnosis is essential because the infection can be fatal. Diagnosis relies on detecting parasites either by PCR or microscopy. Microscopic examination of Giemsa-stained tissues or smears is the quickest and most inexpensive method for diagnosing toxoplasmosis. However, poor sensitivity is the major pitfall of this method. PCR can also be performed on blood or cerebral spinal fluid (CSF) samples in order to detect parasite DNA. However, the degree of sensitivity attained by the PCRs is questionable and requires further investigations [1].
Leishmaniasis
Protozoans of the Leishmania genus are transmitted to humans via sand fly bites. Visceral leishmaniasis (VL), which is a lethal infection if left untreated, can also be transmitted by blood transfusions, organ transplants, and sharing of needles among intravenous drug users.
Direct parasitological methods, such as microscopy and cultures, are the gold standard methods when diagnosing VL. These methods have high specificity, but varying sensitivity. Direct detection of parasites is performed by microscopic examination of aspirates from spleen, bone marrow, or lymph nodes [5]. Using spleen samples increases sensitivity, but the procedure to obtain the aspirates risks internal bleeding. Parasite culturing from aspirates is widely used by reference laboratories.
Extensive research on the development of Leishmania serological assays has uncovered a myriad of candidate diagnostic antigens. The most promising antigens were the kinesin-related proteins. From this group, rK39 was the most tested antigen [6–8]. The rK39 antigen has been used to develop an immunochromatographic strip test (ICT)-based rapid diagnostic test which is advantageous for mass screening in endemic areas. This test requires a drop of peripheral blood and can be completed in approximately fifteen minutes [7]. Although the rK39 ICT rapid test was quite successful in Asia, it was often unable to detect Leishmania infections in African patients [5]. Additionally, rapid diagnostic tests still need standardization in order to become a regular practice in clinical laboratories.
PCR is the main molecular tool for Leishmania diagnosis due to its high sensitivity and reliability. Different PCR target sequences that are commonly used include ribosomal RNA genes, kinetoplast DNA, mini-exon derived RNA, internal transcribed spacer regions, etc., [5]. Quantitative PCR is useful because it allows for the quantification of parasites as well as species typing. Furthermore, this technique can be used to monitor treatment efficiency. Unfortunately, equipment requirements as well as the high cost limit the use of PCR for mass screening purposes in the field. The introduction of loop-mediated isothermal amplification (LAMP) could facilitate the use of molecular techniques for diagnostics. LAMP is highly specific, carried out under isothermal conditions, quick, and requires less complicated equipment (5). Moreover, reagents can be kept at room temperature, and there are no post-PCR steps. Assessment of drug treatment can also be carried out through the use of nucleic acid sequence based amplification (NASBA) which amplifies RNA sequences under isothermal conditions. Coupled to oligochromatography, NASBA can be used to monitor the progression from active disease to cure [9].
Chagas Disease (American Trypanosomiasis)
Chagas disease is the result of an infection with the blood-borne protozoan Trypanosoma cruzi. The parasite is transmitted by the triatomine bug. The second most important mode of transmission is via contaminated blood. This includes blood transfusions, organ transplants, and congenital transmission.
During the acute stage of Chagas disease, parasites can be observed in the blood. For this reason, diagnosis is carried out by direct microscopic viewing of Giemsa-stained thin and thick blood smears [10]. Parasites may also be detected through the use of hemocultures. In Chagas endemic areas, xenodiagnosis may be performed. This method involves allowing the naïve triatomine bug to take a blood-meal from the patient, and then analysing the bug for the presence of trypanosomes. It is believed that with continued research, molecular methods will eventually replace indirect diagnostic techniques such as blood cultures and xenodiagnosis [10]. However, molecular tests need to be standardized for routine clinical practice.
During the chronic stage of Chagas disease, diagnosis relies on serology; however, these tests often yield results that are difficult to interpret [10]. Commonly used, standardized serological assays include indirect immunofluorescence (IIF), indirect hemagglutination (IHA), and ELISA. IIF and IHA are commonly used due to their good sensitivity; however, their results are operator-dependent, and there is a lack of studies which analyse their reproducibility [10]. Currently, the immunoblot and radioimmunoprecipitation assays are in the process of being standardized. Both tests showed promise in early studies. A great deal of work is also being focused on the development and standardization of molecular methods such as PCR, which could be useful in monitoring chronic phase, reactivation, and treatment response.
As previously mentioned, disease transmission can also occur from mother to child, leading to congenital Chagas. Screening of neonates can be performed via direct methods, such as microscopy, or PCR using venous or cord blood samples from the newborn. These tests have very high sensitivity when performed during the first month of life [10]. Serological analysis may also be performed.
Sleeping Sickness (African Trypanosomiasis)
Trypanosoma brucei is the causative agent of African trypanosomiasis, and it is transmitted via the bite of the tsetse fly. During the first stage of the disease, parasites can be found circulating in the peripheral blood. The second stage is marked by parasites crossing the blood-brain barrier and infecting the central nervous system (CNS). The parasitic subspecies dictates geographic distribution, prognosis, and diagnosis.
T. b. gambiense causes West African trypanosomiasis, which is a slow progressing disease and is characterized by low parasite loads [11]. Definite diagnosis is carried out by microscopic observation of blood, lymph node aspirate, or CSF for the presence of parasites. In the field, the card agglutination test for trypanosomiasis (CATT/T. b. gambiense) has been widely used since its development in 1978 (12). Whole blood is used, and the assay directly detects T. b. gambiense specific antibodies. CATT/T. b. gambiense is cheap, quick, and highly sensitive. However, the test can give rise to false positives in individuals who are co-infected with malaria [12]. Although CATT/T. b. gambiense is the most sensitive, similar tests such as micro-CATT and LATEX/ T. b. gambiense can also be used. If these assays generate positive results, they need to be confirmed by microscopy or other molecular methods.
T. b. rhodesiense causes East African trypanosomiasis, which progresses quickly and is characterized by high parasite loads (11). For this subspecies, there is no diagnostic equivalent to the CATT/T. b. gambiense. However, diagnosis by microscopic observations of thick and thin smears is simple due to the elevated parasite load associated with T. b. rhodesiense.
Microscopy is the most practical technique to be used in rural areas. However, microscopy requires adequately qualified personnel in order to prevent misdiagnosis. Molecular methods would substantially improve the diagnosis of African trypanosomiasis. PCR techniques have been developed to screen the CSF of patients. The discovery of the SRA gene in T. b. rhodesiense has proven to be a breakthrough for the promotion of PCR techniques. Reactions targeting this gene have the potential to identify a single trypanosome [11]. There has also been the introduction of fluorescence in-situ hybridization in combination with peptide nucleic acid probes aimed towards ribosomal RNA. However, these tools for diagnosis are new and require further optimization. Extensive research is being focused on standardizing molecular techniques and rendering them more accessible. The use of LAMP is a step forward in improving molecular
approaches [11].
Future research needs to focus on the improvement of molecular diagnostic techniques. Currently, second stage infections are diagnosed by microscopic observation of CSF. Research is being conducted to test various cytokines and antibodies as biomarkers for CNS infection [11].
Malaria
Malaria is the most important parasitic infection in the world due to its high mortality. The causative agents, parasites of the Plasmodium genus, are transmitted by Anopheles mosquitoes. Quick diagnosis is essential in order to determine the appropriate treatment as well as to prevent further transmission.
Microscopy is the gold standard for laboratory diagnosis. This method involves detecting parasites in Giemsa-stained thick and thin blood smears. However, microscopic results are operator-dependent, thereby causing the sensitivity to vary. A great deal of effort has been focused on developing rapid diagnostic tests (RDTs) which can be used in the field. These tests can supplement microscopy, but they cannot replace it yet. Current RDTs are serology based and use three different Plasmodium antigens: Plasmodium histidine-rich protein, Plasmodium lactate dehydrogenase, or Plasmodium aldolase [13]. These tests are quick, easy to perform, and require minimal patient samples. However, they are not specific for species such as P. malariae, P. ovale, and P. knowlesi. Furthermore, false positives may be observed due to cross-reactions in patients with Schistosoma mekongi or rheumatoid factor [14]. In addition, the tests inefficiently detect P. falciparum infections from South America, as this species does not produce the common histidine-rich proteins [15].
Currently, there are no commercially available molecular assays. Although some reference and government laboratories have developed their own molecular assays, their availability is limited. LAMP is currently in the spotlight. Poon et al. developed a LAMP test which detected the target sequence of P. falciparum 18S ribosomal RNA gene [16]. They stated that the price of this test was one tenth that of a conventional PCR. Recently, LAMP was further simplified in the form of a card test. It was used in combination with DNA filter paper and melting curve analysis. This system was shown to be highly specific and sensitive [17]. Improvement of the LAMP technique should be geared towards the development of rapid diagnostic tests which could potentially be used in the field.
Babesiosis
Babesiosis is caused by parasites belonging to the Babesia genus that are spread by certain ticks commonly found in North America. The parasites infect red blood cells (RBCs), and consequently cause hemolytic anemia. The disease can be fatal in splenectomy patients, immunocompromised individuals, and the elderly. Diagnosis is complicated by the symptoms’ resemblance to other tick-borne illnesses.
The gold standard of babesiosis diagnosis relies on detecting the parasites in the patients’ RBCs. This is achieved by microscopic observation of thick and thin blood smears. Babesia infections can be easily mistaken for P. falciparum infections [18]. Additionally, false negatives are common in immunocompetent individuals whose parasitemia can be lower than 1% [18]. Samples are often sent to reference laboratories in order to confirm ambiguous results. IFFs are used to detect anti-babesial IgM and IgG [18]. They are sensitive, specific, and reliable. ELISAs and immunoblots, although not standardized, can be performed to confirm the IFF results. However, compared to IFFs, Babesia detecting ELISAs require higher concentrations of antigen and have varying sensitivity [18]. Future research on babesiosis diagnosis is aimed at developing multiplex PCR assays that will be able to detect several tick-borne infections. PCR assays have the potential to yield positive results from 100µl blood samples containing as little as three parasites; demonstrating the incredible advantage that molecular techniques could contribute to diagnosis of this parasitic disease [18].
Proteomics
Dr Momar Ndao’s laboratory focuses on the improvement and advancement of diagnosis. Through our work, we hope to encourage the development of proteomic strategies for the diagnosis of parasitic infections. Mass-spectrometry platforms are the future of proteomics, and they can be used to identify biomarkers from biological fluids. Some techniques that can be used to analyse protein expression include matrix-assisted laser desorption ionization time-of-flight mass-spectrometry (MALDI-TOF MS), surface-enhanced laser desorption ionization time-of-flight mass-spectrometry (SELDI-TOF MS), liquid chromatography combined with mass-spectrometry, isotope-coded affinity tags, and isobaric tags for relative and absolute quantification [19]. When SELDI is used, samples are directly spotted onto chemically active ProteinChip Array surfaces which can be chosen based on specific chemical and biological properties. With MALDI, samples are mixed with the matrix component prior to loading on a chip. These proteomic platforms can be useful in identifying biomarkers that are indicative of a specific pathophysiological state. Currently, members of our laboratory are using both SELDI and MALDI techniques extensively to identify biomarkers of blood borne parasites.
Summary
Quick and correct diagnosis of parasitic infections is crucial to avoid deaths and further disease transmission. Diagnostic methods include parasitological techniques, such as microscopy and culturing, serological assays, and molecular tests [Table 1]. Although several serological and molecular diagnostic tools are being tested and used by certain reference laboratories, results are always confirmed by microscopy which remains the gold standard. Many newer assays have not been standardized yet, thus, forcing diagnosticians to rely on microscopic observations. Unfortunately, the evolution of diagnosis in the field of parasitology has been slow to progress. Fortunately, in recent years, several groups have focused their research on the improvement of diagnostics. Current research emphasizes the development and optimization of molecular techniques such as PCR and LAMP. Additional work must concentrate on rendering molecular diagnostics more accessible. Although relatively new at the moment, proteomic platforms seem to be the future of diagnosis. These new techniques can identify biomarkers which can categorize susceptible individuals, distinguish between the different stages of an infection, and monitor whether treatments lead to cure. Diagnostic research has made much progression, however, there is still a lot of work to be done and improvements to be made. In order to better the diagnosis of blood-borne parasitic infections, research plus communication is the answer.
References
1. Robert-Gangneux F and Darde ML. Clin Microbiol Rev 2012; 25: 264–96.
2. Gras L, et al. Epidemiol Infect 2004; 132: 541–8.
3. Lefevre-Pettazzoni M, et al. I Clin Vaccine Immunol 2007; 14: 239–43.
4. Chapey E, et al. J Clin Microbiol 2010; 48: 41–5.
5. Srividya G, et al. Parasitol Res 2012; 110: 1065–78.
6. Badaro R, et al. J Infect Dis 1996; 173: 758–61.
7. Chappuis F, et al. Trop Med Int Health 2006; 11: 31–40.
8. Singh S, et al. Clin Diagn Lab Immunol 2002; 9: 568–72.
9. Saad AA, et al. PLoS Negl Trop Dis 2010; 4: e776.
10. Lescure FX, et al. Lancet Infect Dis 2010; 10: 556–70.
11. Welburn SC, et al.. Adv Parasitol 2012;79: 299–337.
12. Magnus E, et al. Ann Soc Belg Med Trop 1978; 58: 169–76.
13. Wilson ML. Clin Infect Dis 2012; 54: 1637–41.
14. Leshem E, et al. J Clin Microbiol 2011; 49: 2331–2.
15. Gamboa D, et al. PLoS One 2010; 5: e8091.
16. Poon LL, et al. Clin Chem 2006; 52: 303–6.
17. Yamamura M, et al. Jpn J Infect Dis 2009; 62: 20–5.
18. Hunfeld KP, et al. Int J Parasitol 2008; 38: 1219–37.
19. Ndao M. Interdiscip Perspect Infect Dis 2009; 2009: 278246.
The authors
Alessandra Ricciardi, BSc
National Reference Centre for Parasitology, Research Institute of the McGill University Health Center, Montreal, Canada
Momar Ndao, DVM, MSc, PhD
National Reference Centre for Parasitology at the Montreal General Hospital, Montreal, Quebec, Canada
E-mail: momar.ndao@mcgill.ca
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, /in Featured Articles /by 3wmediaPertussis (whooping cough) has been a significant cause of morbidity and mortality in young children since the first epidemic was described in 1578. Currently in the West even when infants suffering from the disease are hospitalized and appropriately treated, around 1% still die, and in less developed countries the mortality rate in infants is as high as 4%. However, following the isolation of the causative organism Bordetella pertussis over a century ago, years of research and development resulted in the introduction of an effective vaccine in the 1940s.
The whole cell vaccine used heat-killed bacteria combined with diphtheria and tetanus toxoids to give the classical DPT vaccine, usually given to infants three times during their first year of life with further booster doses twice during childhood. The advent of this vaccine did not prevent the three to five year pertussis epidemic cycle, but it elicited a strong immune response and the total number of cases plummeted in immunized populations. There were some common side-effects, including swelling, mild fever and pain, but these were trivial compared with the high risk of children contracting pertussis if they were not immunized. Sadly, though, very dubious research linked cases of SIDS and encephalopathy with use of whole cell pertussis vaccine, and the popular press eagerly disseminated this dangerously misleading information. Parents began to exercise their so-called ‘freedom of choice’ based on a dearth of unbiased information and stopped having their children immunized, so in the 1990s a new acellular vaccine (DPaT) with fewer side effects gradually replaced the classical DPT.
Now cases of pertussis have more than tripled in the last five years in much of the globe, and the resulting whooping cough epidemic is the worst for 50 years. While it is possible that a more virulent strain of bacterium has evolved, the most likely explanation is that the ‘new’ vaccine is not as effective as its predecessor. Indeed a recent robust study from Australia compared incidence of pertussis in 40,694 children who were immunized in 1998 with either DPT or DPaT (both vaccines were still in use at that time). Significantly higher rates of pertussis were found in the children who had received the latter vaccine.
The suggested solution to the pertussis epidemic is to extend immunisation programmes to cover pregnant women as well as all those who come in contact with young infants. Wouldn’t reintroducing the old vaccine be simpler?
Diagnosis of blood-borne parasitic infections: an overview
, /in Featured Articles /by 3wmediaMethods for the diagnosis of blood-borne parasitic infections have stagnated in the last 20–30 years. However, recently, there has been a tremendous effort to focus research on the development of newer diagnostic methods focusing on serological, molecular, and proteomic approaches. This article examines the various diagnostic tools that are being used in clinical laboratories, optimized in reference laboratories and employed in mass screening programmes.
by A. Ricciardi and Dr M. Ndao
Blood-borne protozoans are the causative agents of some of the world’s most devastating and prevalent parasitic infections. This group of pathogens includes members of the Trypanosoma, Leishmania, Plasmodium, Toxoplasma, and Babesia genera. Most of these infections, with the exception of toxoplasmosis and babesiosis, have always been described as being tropical or subtropical. However, the increase in international travel as well as the arrival of new immigrants has made some of these tropical diseases realities in developed countries as well. In addition, infection via contaminated blood (transfusions and organ transplants) has become a major problem. Clearly, the transmission of blood-borne protozoans is boundless and the actual number of cases is underestimated. Quick diagnosis has always been a priority in order to determine the appropriate treatment and prevent fatalities. In addition, now more than ever, advances in diagnostics can help prevent transmission and provide active surveillance. Currently, diagnostic and reference laboratories use an array of techniques including microscopy, serological assays, and molecular assays. Here, the advantages and disadvantages of the methods will be discussed.
Toxoplasmosis
Toxoplasmosis, caused by Toxoplasma gondii, has a worldwide distribution. In immunocompetent individuals, more than 80% of primary Toxoplasma infections are asymptomatic [1]. Toxoplasmosis becomes a problem when an individual is immunocompromised or during pregnancy. Diagnosis of toxoplasmosis varies according to the immune status of the patient.
Diagnosis of immunocompetent individuals relies on serology. Early antibody responses can be detected via methods such as the dye test, immunofluorescent assay, and agglutination test whereas later IgG titres are detected by enzyme-linked immunosorbent assay (ELISA). For many years, the Sabin-Feldman dye test was the gold standard diagnostic technique due to its sensitivity and specificity. In recent years, few laboratories have continued to use this method and rather focused on newer techniques such as indirect immunofluorescent antibody tests, hemagglutination tests, capture ELISAs, and immunosorbent agglutination assays (ISAGAs). Serological assays lack the capacity to differentiate between recent and older infections; IgM levels can persist for over two years [2]. In order to determine whether an infection is recent, avidity ELISA is performed. This assay verifies IgG avidity and is based on the concept that as the immune response progresses, an immunoglobulin’s affinity for a specific antigen will increase [3].
Diagnosis of Toxoplasma infection during pregnancy is crucial in order to prevent congenital toxoplasmosis. Prenatal diagnosis involves performing real-time polymerase chain reaction (PCR) using amniotic fluid. The PCRs used often target the B1 gene of the parasite [1]. Upon delivery, PCR is performed on either the placenta or the cord blood serum in order to detect parasites. ISAGAs are also often performed. If the tests are positive, cord blood samples at one week of life are sent to a reference laboratory [1]. Follow up serology is again performed at one month and then every two to three months. There have been recent advances in the field of toxoplasmosis post-natal diagnosis. An ELISA assay that measures interferon-gamma levels upon stimulation of whole blood cells with Toxoplasma crude antigens has been developed. This method has proven to be both sensitive and specific [4].
In the case of immunocompromised patients, a quick diagnosis is essential because the infection can be fatal. Diagnosis relies on detecting parasites either by PCR or microscopy. Microscopic examination of Giemsa-stained tissues or smears is the quickest and most inexpensive method for diagnosing toxoplasmosis. However, poor sensitivity is the major pitfall of this method. PCR can also be performed on blood or cerebral spinal fluid (CSF) samples in order to detect parasite DNA. However, the degree of sensitivity attained by the PCRs is questionable and requires further investigations [1].
Leishmaniasis
Protozoans of the Leishmania genus are transmitted to humans via sand fly bites. Visceral leishmaniasis (VL), which is a lethal infection if left untreated, can also be transmitted by blood transfusions, organ transplants, and sharing of needles among intravenous drug users.
Direct parasitological methods, such as microscopy and cultures, are the gold standard methods when diagnosing VL. These methods have high specificity, but varying sensitivity. Direct detection of parasites is performed by microscopic examination of aspirates from spleen, bone marrow, or lymph nodes [5]. Using spleen samples increases sensitivity, but the procedure to obtain the aspirates risks internal bleeding. Parasite culturing from aspirates is widely used by reference laboratories.
Extensive research on the development of Leishmania serological assays has uncovered a myriad of candidate diagnostic antigens. The most promising antigens were the kinesin-related proteins. From this group, rK39 was the most tested antigen [6–8]. The rK39 antigen has been used to develop an immunochromatographic strip test (ICT)-based rapid diagnostic test which is advantageous for mass screening in endemic areas. This test requires a drop of peripheral blood and can be completed in approximately fifteen minutes [7]. Although the rK39 ICT rapid test was quite successful in Asia, it was often unable to detect Leishmania infections in African patients [5]. Additionally, rapid diagnostic tests still need standardization in order to become a regular practice in clinical laboratories.
PCR is the main molecular tool for Leishmania diagnosis due to its high sensitivity and reliability. Different PCR target sequences that are commonly used include ribosomal RNA genes, kinetoplast DNA, mini-exon derived RNA, internal transcribed spacer regions, etc., [5]. Quantitative PCR is useful because it allows for the quantification of parasites as well as species typing. Furthermore, this technique can be used to monitor treatment efficiency. Unfortunately, equipment requirements as well as the high cost limit the use of PCR for mass screening purposes in the field. The introduction of loop-mediated isothermal amplification (LAMP) could facilitate the use of molecular techniques for diagnostics. LAMP is highly specific, carried out under isothermal conditions, quick, and requires less complicated equipment (5). Moreover, reagents can be kept at room temperature, and there are no post-PCR steps. Assessment of drug treatment can also be carried out through the use of nucleic acid sequence based amplification (NASBA) which amplifies RNA sequences under isothermal conditions. Coupled to oligochromatography, NASBA can be used to monitor the progression from active disease to cure [9].
Chagas Disease (American Trypanosomiasis)
Chagas disease is the result of an infection with the blood-borne protozoan Trypanosoma cruzi. The parasite is transmitted by the triatomine bug. The second most important mode of transmission is via contaminated blood. This includes blood transfusions, organ transplants, and congenital transmission.
During the acute stage of Chagas disease, parasites can be observed in the blood. For this reason, diagnosis is carried out by direct microscopic viewing of Giemsa-stained thin and thick blood smears [10]. Parasites may also be detected through the use of hemocultures. In Chagas endemic areas, xenodiagnosis may be performed. This method involves allowing the naïve triatomine bug to take a blood-meal from the patient, and then analysing the bug for the presence of trypanosomes. It is believed that with continued research, molecular methods will eventually replace indirect diagnostic techniques such as blood cultures and xenodiagnosis [10]. However, molecular tests need to be standardized for routine clinical practice.
During the chronic stage of Chagas disease, diagnosis relies on serology; however, these tests often yield results that are difficult to interpret [10]. Commonly used, standardized serological assays include indirect immunofluorescence (IIF), indirect hemagglutination (IHA), and ELISA. IIF and IHA are commonly used due to their good sensitivity; however, their results are operator-dependent, and there is a lack of studies which analyse their reproducibility [10]. Currently, the immunoblot and radioimmunoprecipitation assays are in the process of being standardized. Both tests showed promise in early studies. A great deal of work is also being focused on the development and standardization of molecular methods such as PCR, which could be useful in monitoring chronic phase, reactivation, and treatment response.
As previously mentioned, disease transmission can also occur from mother to child, leading to congenital Chagas. Screening of neonates can be performed via direct methods, such as microscopy, or PCR using venous or cord blood samples from the newborn. These tests have very high sensitivity when performed during the first month of life [10]. Serological analysis may also be performed.
Sleeping Sickness (African Trypanosomiasis)
Trypanosoma brucei is the causative agent of African trypanosomiasis, and it is transmitted via the bite of the tsetse fly. During the first stage of the disease, parasites can be found circulating in the peripheral blood. The second stage is marked by parasites crossing the blood-brain barrier and infecting the central nervous system (CNS). The parasitic subspecies dictates geographic distribution, prognosis, and diagnosis.
T. b. gambiense causes West African trypanosomiasis, which is a slow progressing disease and is characterized by low parasite loads [11]. Definite diagnosis is carried out by microscopic observation of blood, lymph node aspirate, or CSF for the presence of parasites. In the field, the card agglutination test for trypanosomiasis (CATT/T. b. gambiense) has been widely used since its development in 1978 (12). Whole blood is used, and the assay directly detects T. b. gambiense specific antibodies. CATT/T. b. gambiense is cheap, quick, and highly sensitive. However, the test can give rise to false positives in individuals who are co-infected with malaria [12]. Although CATT/T. b. gambiense is the most sensitive, similar tests such as micro-CATT and LATEX/ T. b. gambiense can also be used. If these assays generate positive results, they need to be confirmed by microscopy or other molecular methods.
T. b. rhodesiense causes East African trypanosomiasis, which progresses quickly and is characterized by high parasite loads (11). For this subspecies, there is no diagnostic equivalent to the CATT/T. b. gambiense. However, diagnosis by microscopic observations of thick and thin smears is simple due to the elevated parasite load associated with T. b. rhodesiense.
Microscopy is the most practical technique to be used in rural areas. However, microscopy requires adequately qualified personnel in order to prevent misdiagnosis. Molecular methods would substantially improve the diagnosis of African trypanosomiasis. PCR techniques have been developed to screen the CSF of patients. The discovery of the SRA gene in T. b. rhodesiense has proven to be a breakthrough for the promotion of PCR techniques. Reactions targeting this gene have the potential to identify a single trypanosome [11]. There has also been the introduction of fluorescence in-situ hybridization in combination with peptide nucleic acid probes aimed towards ribosomal RNA. However, these tools for diagnosis are new and require further optimization. Extensive research is being focused on standardizing molecular techniques and rendering them more accessible. The use of LAMP is a step forward in improving molecular
approaches [11].
Future research needs to focus on the improvement of molecular diagnostic techniques. Currently, second stage infections are diagnosed by microscopic observation of CSF. Research is being conducted to test various cytokines and antibodies as biomarkers for CNS infection [11].
Malaria
Malaria is the most important parasitic infection in the world due to its high mortality. The causative agents, parasites of the Plasmodium genus, are transmitted by Anopheles mosquitoes. Quick diagnosis is essential in order to determine the appropriate treatment as well as to prevent further transmission.
Microscopy is the gold standard for laboratory diagnosis. This method involves detecting parasites in Giemsa-stained thick and thin blood smears. However, microscopic results are operator-dependent, thereby causing the sensitivity to vary. A great deal of effort has been focused on developing rapid diagnostic tests (RDTs) which can be used in the field. These tests can supplement microscopy, but they cannot replace it yet. Current RDTs are serology based and use three different Plasmodium antigens: Plasmodium histidine-rich protein, Plasmodium lactate dehydrogenase, or Plasmodium aldolase [13]. These tests are quick, easy to perform, and require minimal patient samples. However, they are not specific for species such as P. malariae, P. ovale, and P. knowlesi. Furthermore, false positives may be observed due to cross-reactions in patients with Schistosoma mekongi or rheumatoid factor [14]. In addition, the tests inefficiently detect P. falciparum infections from South America, as this species does not produce the common histidine-rich proteins [15].
Currently, there are no commercially available molecular assays. Although some reference and government laboratories have developed their own molecular assays, their availability is limited. LAMP is currently in the spotlight. Poon et al. developed a LAMP test which detected the target sequence of P. falciparum 18S ribosomal RNA gene [16]. They stated that the price of this test was one tenth that of a conventional PCR. Recently, LAMP was further simplified in the form of a card test. It was used in combination with DNA filter paper and melting curve analysis. This system was shown to be highly specific and sensitive [17]. Improvement of the LAMP technique should be geared towards the development of rapid diagnostic tests which could potentially be used in the field.
Babesiosis
Babesiosis is caused by parasites belonging to the Babesia genus that are spread by certain ticks commonly found in North America. The parasites infect red blood cells (RBCs), and consequently cause hemolytic anemia. The disease can be fatal in splenectomy patients, immunocompromised individuals, and the elderly. Diagnosis is complicated by the symptoms’ resemblance to other tick-borne illnesses.
The gold standard of babesiosis diagnosis relies on detecting the parasites in the patients’ RBCs. This is achieved by microscopic observation of thick and thin blood smears. Babesia infections can be easily mistaken for P. falciparum infections [18]. Additionally, false negatives are common in immunocompetent individuals whose parasitemia can be lower than 1% [18]. Samples are often sent to reference laboratories in order to confirm ambiguous results. IFFs are used to detect anti-babesial IgM and IgG [18]. They are sensitive, specific, and reliable. ELISAs and immunoblots, although not standardized, can be performed to confirm the IFF results. However, compared to IFFs, Babesia detecting ELISAs require higher concentrations of antigen and have varying sensitivity [18]. Future research on babesiosis diagnosis is aimed at developing multiplex PCR assays that will be able to detect several tick-borne infections. PCR assays have the potential to yield positive results from 100µl blood samples containing as little as three parasites; demonstrating the incredible advantage that molecular techniques could contribute to diagnosis of this parasitic disease [18].
Proteomics
Dr Momar Ndao’s laboratory focuses on the improvement and advancement of diagnosis. Through our work, we hope to encourage the development of proteomic strategies for the diagnosis of parasitic infections. Mass-spectrometry platforms are the future of proteomics, and they can be used to identify biomarkers from biological fluids. Some techniques that can be used to analyse protein expression include matrix-assisted laser desorption ionization time-of-flight mass-spectrometry (MALDI-TOF MS), surface-enhanced laser desorption ionization time-of-flight mass-spectrometry (SELDI-TOF MS), liquid chromatography combined with mass-spectrometry, isotope-coded affinity tags, and isobaric tags for relative and absolute quantification [19]. When SELDI is used, samples are directly spotted onto chemically active ProteinChip Array surfaces which can be chosen based on specific chemical and biological properties. With MALDI, samples are mixed with the matrix component prior to loading on a chip. These proteomic platforms can be useful in identifying biomarkers that are indicative of a specific pathophysiological state. Currently, members of our laboratory are using both SELDI and MALDI techniques extensively to identify biomarkers of blood borne parasites.
Summary
Quick and correct diagnosis of parasitic infections is crucial to avoid deaths and further disease transmission. Diagnostic methods include parasitological techniques, such as microscopy and culturing, serological assays, and molecular tests [Table 1]. Although several serological and molecular diagnostic tools are being tested and used by certain reference laboratories, results are always confirmed by microscopy which remains the gold standard. Many newer assays have not been standardized yet, thus, forcing diagnosticians to rely on microscopic observations. Unfortunately, the evolution of diagnosis in the field of parasitology has been slow to progress. Fortunately, in recent years, several groups have focused their research on the improvement of diagnostics. Current research emphasizes the development and optimization of molecular techniques such as PCR and LAMP. Additional work must concentrate on rendering molecular diagnostics more accessible. Although relatively new at the moment, proteomic platforms seem to be the future of diagnosis. These new techniques can identify biomarkers which can categorize susceptible individuals, distinguish between the different stages of an infection, and monitor whether treatments lead to cure. Diagnostic research has made much progression, however, there is still a lot of work to be done and improvements to be made. In order to better the diagnosis of blood-borne parasitic infections, research plus communication is the answer.
References
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2. Gras L, et al. Epidemiol Infect 2004; 132: 541–8.
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18. Hunfeld KP, et al. Int J Parasitol 2008; 38: 1219–37.
19. Ndao M. Interdiscip Perspect Infect Dis 2009; 2009: 278246.
The authors
Alessandra Ricciardi, BSc
National Reference Centre for Parasitology, Research Institute of the McGill University Health Center, Montreal, Canada
Momar Ndao, DVM, MSc, PhD
National Reference Centre for Parasitology at the Montreal General Hospital, Montreal, Quebec, Canada
E-mail: momar.ndao@mcgill.ca
Molecular diagnosis and sub-speciation of cutaneous leishmaniasis
, /in Featured Articles /by 3wmediaDiagnosing cutaneous leishmaniasis histologically depends on the identification of the amastigotes, which is inconclusive and leads to cases of missed diagnosis or misdiagnosis. In this article, we describe a rapid diagnostic molecular method for Leishmania species identification and differentiation using DNA extracted from formalin-fixed paraffin-embedded (FFPE) skin tissue biopsies.
by L. Yehia and Dr I. Khalifeh
Clinical background
Cutaneous leishmaniasis is a chronic disease caused by Leishmania protozoan parasites that is on the increase in endemic and non-endemic regions because of environmental changes triggered by humans [1, 2]. It is most prevalent in the Middle East and North Africa. With changes in vector (sandfly), habitat and increased travel among populations, the incidence of leishmaniasis is showing a clear increase [3].
There are more than 20 strains of Leishmania that are pathogenic to humans [4], and these are partially responsible for its clinical diversity. The diagnosis of cutaneous leishmaniasis rests on the pathological identification of the amastigotes, which may be inconclusive [5]. This is dependent on the strain type, host response and the disease stage. Accurate microscopic diagnosis is essential to permit appropriate targeted therapy [6].
Clinically, cutaneous leishmaniasis may be asymptomatic and self-limiting. However, cases progressing to mutilating ulceration and disfiguring scarring have also been reported [7]. As the disease progresses, the number of amastigotes decreases to the point where none can be detected microscopically. The absence of amastigotes is a common problem encountered in up to 47% of cases [8]. In such instances, the diagnosis of cutaneous leishmaniasis must not be excluded [4].
Materials and methods
Skin biopsies embedded into FFPE tissue blocks were collected for 122 patients diagnosed clinically with cutaneous leishmaniasis. Cases included in the study were restricted to cutaneous lesions of patients who did not receive treatment prior to the biopsy. Cases with visceral or mucocutaneous involvement and with material insufficient for PCR or histopathological examination were excluded. Clinical information pertaining to the lesion was also collected including: number, duration, location and dermatologic appearance. In addition, the patient’s age, gender and country of residency were tabulated.
Cases were classified according to the modified Ridley’s parasitic index, a traditionally used pathological scoring system based on microscopic analysis of hematoxylin and eosin stained slides. DNA was then extracted from FFPE tissue blocks of each patient. Polymerase chain reaction (PCR) was performed using Leishmania-specific ribosomal internal transcribed spacer 1 (ITS1-PCR). Nested ITS1-PCR was performed on cases negative for conventional ITS1-PCR. ITS1-PCR amplicons were then digested with HaeIII for subsequent restriction fragment length polymorphism (RFLP) subspeciation.
Results
Of the 122 skin biopsies, microscopic evaluation of stained slides identified 54 cases (44.3%) labeled as histologically negative (with no unequivocal amastigotes detected). Of these negative cases, 9 (17%) were shave biopsies and 45 (83%) were punch biopsies.
DNA extracted from FFPE tissue blocks collected for all cases ranged from 4 to 1672 ng/μl (mean=213 ng/μl, SD=289 ng/μl). The oldest blocks were 19 years of age, whereas the newest were less than 1 year old. The quantity of the extracted DNA dating back to 1992 was 166 ng/μl (SD=128 ng/μl), whereas that for specimen from the year 2010 was 272 ng/μl (SD=161 ng/μl) indicating that a good quantity of DNA could be extracted from archival well-preserved FFPE tissues, even when they were old.
ITS1-PCR was performed on DNA extracted from all cases. Initially, and regardless of the histopathological analysis, 55 (45%) cases were positive and showed a band of between 300 and 350 base pairs indicative of Leishmania by agarose gel electrophoresis. The remaining 67 (55%) were negative (Fig. 1A, B). The negative cases were subjected to nested ITS1-PCR and 100% of these cases actually turned out to be positive for Leishmania (Fig. 1C).
Comparing the resultant ITS1-PCR bands to the DNA pattern of normal skin tissues, we identified 54 cases – that had been shown as negative by histopathology according to Ridley’s parasitic index – that amplified DNA with Leishmania-specific primers by conventional or nested ITS1-PCR, and that failed to show the normal skin profile seen in the negative controls tested. RFLP analysis identified L. tropica subspecies in all cases, identified by the presence of a 200 and 60 base pairs restriction fragments (Fig. 2) [9].
Clinical and diagnostic significance
Cutaneous leishmaniasis is a disease that is endemic in many regions of the world. With the ease of travel in the world, human and animal reservoirs of Leishmania parasites have been established in regions that previously were not known to harbour the sandfly vector because of habitat incompatibility. Thus, novel endemic areas have emerged in regions across the world. Therefore, a high index of suspicion becomes crucial for early diagnosis and control of leishmaniasis. With the advent of molecular diagnostic techniques and their high sensitivity and specificity, it has become easier to detect and control many infectious diseases, including leishmaniasis, as shown in this and other studies.
Traditionally, direct detection of parasites is performed by microscopic examination of clinical specimens or by cultivation, but either approach may be diagnostically problematic [1, 4, 10]. Cultures may take long periods, possibly weeks, for sufficient parasites to grow for species characterization. In addition, success in microscopic identification of amastigotes in stained preparations varies depending on the number of parasites present and/or the experience of the person examining the slide [11]. This is mainly due to the fact that all Leishmania species are morphologically similar and may present with a variable number of amastigotes. As the disease progresses, the number of amastigotes decreases to the point where none can be detected histopathologically.
Despite these drawbacks, microscopic identification and parasite cultivation are still the primary diagnostic tools used in most regions where leishmaniasis is endemic. However, it is stressed that accurate and rapid species identification is not possible using either technique. In the last decade, polymerase chain reaction (PCR) analysis has been successfully introduced and has been proven to be the most sensitive molecular tool for direct detection and parasite characterization of Leishmania species in clinical samples [1, 5, 12].
Accurate Leishmania species identification and subspeciation in clinical specimens is now possible by subjecting the extracted DNA to PCR, followed by enzymatic digestion to identify restriction fragments indicative of the subspecies. Such amplification using Leishmania-specific primers allows the indirect yet conclusive detection of the amastigotes, when present in a given clinical specimen. A highly sensitive method is valuable especially in chronic cases where the parasitic index is low and potentially undetectable by conventional microscopy.
Conclusion
This study successfully identified L. tropica in 54 skin biopsies from patients clinically suspected of having cutaneous leishmaniasis with negative biopsies. The importance of this result is manifested in the need for diagnostic tools that are sensitive, specific, rapid and capable of identifying all clinically significant Leishmania species from FFPE tissue blocks (Fig. 3).
Therefore, ITS1-PCR carried out on DNA extracted from FFPE tissue specimens, followed by HaeIII RFLP analysis, is a valuable method for the rapid and reliable diagnosis of cutaneous leishmaniasis. In chronic cases where the parasite load is low, or when insufficient tissue is available, nested ITS1-PCR can be performed to increase sensitivity. The advantages of this method are also highlighted with the possibility of using different biological specimens, and the ability to detect both Old World and New World leishmaniasis.
The work summarized here was first published as Yehia L. et al., 2012 [13].
References
1. Schonian G, et al. Diagn Microbiol Infect Dis 2003; 47: 349.
2. Goto H, Lindoso JA. Expert Rev Anti Infect Ther 2010; 8: 419.
3. Scarisbrick JJ, et al. Travel Med Infect Dis 2006; 4: 14.
4. Ameen M. Clin Exp Dermatol 2010; 35: 699.
5. Singh S, et al. Expert Rev Mol Diagn 2005; 5: 251.
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13. Yehia L, et al. J Cutan Pathol 2012; 39: 347–355.
The authors
Lamis Yehia, BSc
Biomedical Sciences Training Program, Case Western Reserve University in Cleveland, Ohio, USA
Ibrahim Khalifeh, MD
Department of Pathology and Laboratory Medicine, American University of Beirut, Beirut, Lebanon
E-mail: ik08@aub.edu.lb
Real time RT-PCR is the gold standard for laboratory diagnosis
, /in Featured Articles /by 3wmediaNoroviruses are the most common cause of viral gastroenteritis in humans. In recent years diagnostic methods for Noroviruses, especially real-time reverse transcription-polymerase chain reaction (RT-PCR) for the detection of Norovirus-RNA, have been improved and become more widely available.
by Dr Christoph Metzger-Boddien
Noroviruses are transmitted by fecally contaminated food or water, by person-to-person contact, and via aerosolization of the virus and subsequent contamination of surfaces. They are the most common cause of viral gastroenteritis in humans [15]. Symptoms include nausea, vomiting, diarrhea, and stomach cramping. Additional symptoms are fever, chills, headache, muscle aches and a general sense of tiredness. The onset of symptoms can begin quickly and an infected person may feel sick after a very short period of time. In most people, the illness lasts for about one or two days. People with Norovirus illness are contagious from onset of symptoms until at least three days after recovery. Some people may be contagious for even longer. Noroviruses are highly contagious. The estimated dose is as low as 18 viral particles. Approximately 5 billion infectious doses can be present in each gram of feces during peak shedding [16]. Infection can be more severe in young children and elderly people. Dehydration can occur rapidly and may require medical treatment or hospitalization [10].
Sporadic disease
In recent years diagnostic methods for Noroviruses, especially real-time reverse transcription-polymerase chain reaction (RT-PCR) for the detection of Norovirus-RNA, were improved and became more widely available. Subsequently, it became obvious that Noroviruses are the leading cause of sporadic gastroenteritis in all age groups. In Germany, since the implementation of the notification requirement according to §§6 and 7 of the infection protection act (Infektionsschutzgesetz, IfSG) a rise of reported cases can be observed with a seasonal accumulation during the winter months from October to March (2001: 9,223 cases, 2004: 64,973 cases, 2007: 201,242 and 2008: 212,769 cases, source: Robert-Koch-Institute, RKI, Berlin), but still a high estimated number of unreported cases remain.
Outbreaks
Noroviruses are the predominant cause of gastroenteritis outbreaks worldwide. Data from the United States and European countries show that Norovirus is responsible for approximately 50% of all reported gastroenteritis outbreaks (range: 36%–59%) [12]. Periodic increases in Norovirus outbreaks are associated with the emergence of new GII.4 strains. These emergent GII.4 strains are rapidly replacing existing strains predominating in circulation and sometimes cause seasons with high Norovirus activity, as in 2002–2003 and 2006—2007 [17, 20]. Genetic drift successfully promotes the re-emergence of GII-4 variants in the population [13]. Because the virus can be transmitted by food, water and contaminated environmental surfaces as well as directly from person to person, and because there is no long-lasting immunity to Noroviruses, outbreaks can occur in a variety of institutional settings (e.g. nursing homes, hospitals, and schools) and affect people of all ages. Multiple routes of transmission can occur within an outbreak; for example, point-source outbreaks from a food exposure often result in secondary person-to-person spread within an institution or community [4]. Of the 1,518 Norovirus outbreaks in the USA, during 2010 – 2011, laboratory confirmed by the CDC, 59% were from long-term care facilities (889 outbreaks); 8% were from restaurants (123 Outbreaks); 7% were from parties & events 7% (99 outbreaks); 4% were from hospitals (65 outbreaks); 4% were from schools (64 outbreaks); 4% were from cruise ships (55 outbreaks); and 14% were from other and unknown events (223 outbreaks) [10].
Foods that are commonly involved in outbreaks of Norovirus infection are e.g. leafy greens, fresh fruits, and shellfish. However, any food that is served raw or is being handled after cooking can get contaminated.
In Germany, according to data published by the RKI, the number of Norovirus outbreaks has increased by 20% between 2009 and 2010. Recently, the RKI published the final report of a huge outbreak of acute gastroenteritis in five Eastern German federal states. The source of the outbreak was a batch of deep-frozen strawberries. In total, over 11,000 cases of disease occurred. It was Germany’s largest foodborne outbreak of gastroenteritis, with several hundred institutions affected. In a considerable proportion of tested patients, Noroviruses were found [4].
Analysis of outbreak costs
In fact there is a huge socio-economic impact of Norovirus-associated diseases. A study of Johnston et al. 2007, showed the costs of an outbreak including the estimated loss of revenue because of unit closures, sick leave and cleaning expenses [7]. Because of the high contagiousness of Noroviruses early diagnosis in order to set up appropriate hygiene interventions is the most useful measure. In 2004, Lopman et al. showed, that diagnosis of the first case within three days instead of four reduces the duration of an outbreak by seven days [5, 8].
Diagnostic methods
The clinical specimens used for Norovirus diagnosis in most cases are stool and vomit samples. There is no cell culture method for the isolation of Noroviruses from clinical specimens available. Therefore, the majority of clinical virology laboratories perform RT-PCR assays for Norovirus detection. Additionally, for preliminary identification of Norovirus as the cause of gastroenteritis outbreaks, there are enzyme immunoassays (EIA) and rapid tests available. However, these kits are not recommended for individual diagnosis.
Real-time RT-PCR assays
The region between ORF1-ORF2 is the most conserved region of the Norovirus genome, with a high level of nucleotide sequence identity across strains within a genogroup [6]. This region is ideal for designing broadly reactive primers and probes for real-time RT-PCR (RT-qPCR) assays for high throughput screening in clinical diagnostic laboratories and for the detection of Norovirus RNA in
environmental samples (e.g. food and water).
The quality of the real time RT-PCR results is dependent on the quality of template RNA-extraction from clinical and environmental samples. The implementation of extraction controls in commercial RT-PCR duplex assays (e.g. Control-RNA in MutaREX Norovirus Kit, Immundiagnostik AG, Bensheim, Germany) minimizes the risk of false negative results due to inhibition or partial inhibition of the reverse transcription step and/or the PCR and due to processing errors during the extraction of RNA. Control RNA is added to a sample before RNA extraction with a commercial kit (e.g. High pure viral RNA Kit, Roche Diagnostics GmbH, Mannheim, Germany; or intron viral gene spin, gerbion, Kornwestheim, Germany) and its recovery is measured subsequently in the duplex real time RT-PCR. The latest generation of commercially available Norovirus real time RT-PCR Kits is extremely sensitive and specific [18]. Therefore such tests have become the gold standard for Norovirus laboratory diagnosis in the past few years.
Enzyme immunoassays
For detection of Norovirus antigen in clinical samples, rapid assays (e.g. EIA) offer an alternative to real time RT-PCR assays. However, the development of a broadly reactive EIA for Noroviruses has been challenging because of the number of antigenically distinct Norovirus strains and the high viral load required for a positive signal in these assays. Commercial kits include pools of cross-reactive monoclonal and polyclonal antibodies. In evaluation studies, the sensitivity of these kits ranged from 36% to 80%, and specificity has ranged from 47% to 100% compared with real time RT-PCR [1, 2, 3, 9, 11, 14, 19].
Summary
Norovirus real time RT-PCR Kits offer a sensitive, specific, fast and cost effective diagnosis. Results can be generated within one hour. But clearly only real time RT-PCR Kits containing control RNA used as extraction control for process monitoring produce feasible and reliable results. RNA extraction from clinical specimens and the reverse transcription of RNA to cDNA are the most crucial steps in Norovirus RT-PCR procedures. Errors in sample preparation and/or RT-reaction can lead to false negative results in conventional RT-PCRs as well as real time RT-PCRs when internal controls (RNA or DNA) are already added to the PCR master-mix. Laboratories performing in-house RT-PCR for Noroviruses should critically evaluate their tests with regard to these high quality standards. Because of the modest performance of Norovirus Enzyme Immunoassays, particularly their poor sensitivity, they are not recommended for clinical diagnosis of Norovirus infection in sporadic cases of gastroenteritis. Negative samples will have to be confirmed by real time RT-PCR in outbreaks as well as in sporadic cases.
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4. Großer Gastroenteritis-Ausbruch durch eine Charge mit Noroviren kontaminierter Tiefkühlerdbeeren in Betreuungseinrichtungen und Schulen in Ostdeutschland, 09-10/2012. Epidemiologisches Bulletin Nr. 41/12: 414-417, Oct 15th, 2012
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The author
Christoph Metzger-Boddien, PhD
gerbion GmbH & Co. KG
Remsstr. 1, D-70806 Kornwestheim, Germany