- Record: found
- Abstract: found
- Article: found
A pilot study on molecular diagnosis of Hapalotrema mistroides (Digenea: Spirorchiidae) infection in blood samples of live loggerhead turtles Caretta caretta
research-article
Author(s):
Erica Marchiori
1
,
,
Giorgia Dotto
1 ,
Cinzia Tessarin
1 ,
Mario Santoro
2 ,
Andrea Affuso
3 ,
Luciano Tarricone
4 ,
Ludovica Di Renzo
5
,
6 ,
Daniela Freggi
7 ,
Vincenzoleo Spoto
7 ,
Federica Marcer
1
Publication date (Electronic):
14 January 2020
Journal:
BMC Veterinary Research
Publisher:
BioMed Central
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Abstract
Background
Parasites of the family Spirorchiidae cause disease and mortality in marine and freshwater turtles; two species, Hapalotrema mistroides and Neospirorchis sp., are reported in the resident population of loggerhead turtles of the Mediterranean Sea, with the first being the most widespread. In vivo diagnosis of spirorchidiasis can represent a challenge in guaranteeing prompt control and treatment of the disease and is currently limited to copromicroscopy.
The aim of this study was the development of a real time PCR assay with TaqMan probe for the detection of H. mistroides infection in the blood of live loggerhead turtles, Caretta caretta, hospitalized in rehabilitation centres. Its potential use for in vivo diagnosis is explored.
Results
The developed real time PCR successfully detected H. mistroides DNA from both positive controls and experimental blood samples of live loggerhead sea turtles, showing good specificity, sensitivity and good reaction efficiency. Two out of three turtles which had demonstrated positivity at copromicroscopy also tested positive to this blood assay; DNA of H. mistroides was detected within the blood of one sea turtle, which tested negative for copromicroscopy.
Conclusions
This study describes a specific and rapid molecular assay to detect H. mistroides infection from live sea turtles and highlights for the first time the presence of DNA of this species in turtle blood samples. Since this assay is able to detect low amounts of the parasitic free DNA in blood samples, its application could be helpful for in vivo diagnosis of H. mistroides infection as well as for epidemiological purposes.
Related collections
Most cited references15
- Record: found
- Abstract: found
- Article: not found
Diagnosing Schistosomiasis by Detection of Cell-Free Parasite DNA in Human Plasma
Dominic Wichmann, Marcus Panning, Thomas Quack … (2009)
Introduction Schistosomiasis, also known as bilharzia, is caused by trematodes of the Schistosomatidae family. It is among the most important parasitic diseases worldwide, with a significant socio-economic impact [1]. More than 200 million people are infected, and about 200,000 may die from the disease each year. On a global scale, one of thirty individuals has schistosomiasis [2]. Movements of refugees, displacement of people, and mistakes in freshwater management promote the spread of schistosomiasis [3],[4]. Human disease is caused by S. haematobium, S. mansoni, S. japonicum, and less frequently, S. mekongi and S. intercalatum [5],[6]. Infection with cercariae occurs through intact skin via contact with infested water. Penetration of cercariae is followed by Katayama syndrome, an acute syndrome with fever, rash and eosinophilia. The syndrome is thought to be caused by antigen excess due to the presence of schistosomules in blood and the beginning of egg deposition [5],[6]. After maturation in the lung and liver sinusoids, adult male and female worms mate and actively migrate to their target organs [4],[5]. S. haematobium resides in walls of the bladder and sacral and pelvic blood vessels surrounding the urinary tract. The other mentioned species reside in mesenteric veins. After deposition of eggs in the capillary system, eggs penetrate the mucosa of target organs and are excreted in urine or feces. Sequelae of acute and chronic infection include hepato-splenic disease, portal hypertension with varices, pulmonary hypertension, squamous cell cancer of the bladder, liver fibrosis, and less common conditions such as myelo-radiculitis and female genital schistosomiasis. Co-infections with HCV and Schistosoma may also modify the course of hepatitis C [4], [7]–[14]. Anti-Schistosoma antibodies can be detected by enzyme immunoassay (EIA), immunofluorescence assay (IFA), and indirect hemagglutination assay [15]. Antibody detection is valuable in patients with rare exposure to Schistosoma, e.g., tourists. In patients with Katayama syndrome, a positive EIA antibody test is usually the earliest diagnostic laboratory result. Still, a large fraction of patients will initially test negative [16],[17]. False negative tests prevent timely treatment of schistosomiasis in travellers who present with fever of unknown origin. Moreover, the inability of serology to discriminate between active and past disease limits its clinical value for confirmation of the success of treatment [18]. Microscopic demonstration of eggs in stool or urine specimens is considered the diagnostic gold standard for confirmation of schistosomiasis in patients from endemic countries, as well as for the confirmation of the success of treatment. In field studies the rapid and inexpensive Katz-Kato thick smear technique is often used [19]. Because the shedding of eggs is highly variable, it is necessary to concentrate eggs from stool or urine prior to examination [20]. Even in concentrated samples the sample volume analysed in the microscope is limited. Due to random distribution effects, the analysed sample may not contain eggs even if the disease is active. It is thus very difficult to achieve a conclusive confirmation of successful therapy. In symptomatic patients with unsuccessful egg detection, it is often necessary to perform endoscopic biopsies of the bladder or rectal mucosa to increase the chance of detection [20]. Several groups have developed polymerase chain reaction (PCR) methods to improve the direct detection of Schistosoma. These tests are done on urine, stool, or organ biopsy samples, and involve the preparation of DNA from eggs prior to PCR amplification [21],[22]. Unfortunately, only a small volume of sample can be processed for DNA extraction, and it is dependent on chance whether the processed sample contains eggs or not. In this regard, PCR has the same limitations as microscopy and does not provide a significant clinical benefit. The detection of circulating cell-free DNA in human plasma has long been explored for the non-invasive diagnosis of a variety of clinical conditions (reviewed in [23] and [24]). It has been known since almost 20 years that patients with solid tumors have tumor-derived DNA circulating in plasma that can be used for diagnostic purposes [25]–[28]. Circulating fetal DNA in maternal plasma is used for diagnosing and monitoring of a range of fetal diseases and pregnancy-associated complications [29]–[33]. The normal concentration of cell-free DNA in plasma of adults is 10–100 ng/mL or 10e3 to 10e4 human genome equivalents per mL [34],[35]. It has been determined that the concentration of fetal DNA in maternal plasma is 3.4% of total serum DNA on average [16]. The presence of cell-free DNA in plasma may be a consequence of apoptosis, which is associated with physiological and pathological turnover of tissue, e.g., in tumor growth or embryonic development (reviewed in [36] and references therein). In parasitic diseases such as schistosomiasis, there is a huge turnover of parasites involving replication, maturation, and death of organisms. Multi-cellular parasites like Schistosoma contain DNA copies in stoichiometrical excess over parasite count. We reasoned that it might be possible to find cell-free parasite DNA (CFPD) circulating in plasma, and that this could be used to diagnose schistosomiasis. In contrast to eggs in stool or urine, CFPD would be equally distributed throughout the plasma volume of the patient, resolving the issue of random sampling that spoils clinical sensitivity of classical detection methods. As an extension of this rationale, we reasoned that it might also be possible to confirm the elimination of Schistosoma CFPD after successful treatment. To prove these concepts, Schistosoma-specific real-time PCR was established and optimised for detection of DNA from large volumes of plasma. A Balb/c mouse model of schistosomiasis was used to study the levels of CFPD in plasma during infection, as well as during and after therapy. The concept was then transferred to patients with different stages of infection, including Katayama syndrome, chronic disease with egg excretion, and patients treated for schistosomiasis in the past without current signs of disease. Materials and Methods Ethics statement Written informed consent was obtained from every patient. The study was approved by the ethics committee of the Board of Physicians of the City of Hamburg. Animal model of S. mansoni infection The Liberian isolate of S. mansoni [37], was maintained in Biomphalaria glabrata and Syrian hamsters. Maintenance of the life cycle was exactly performed as described elsewhere [38]. Adult female Balb/c mice (Charles Rivers Laboratories, Sulzfelden, Germany) were infected by intraperitoneal injection of 100 cercariae diluted in 200 µL sterile isotonic saline solution. Approval was obtained from the animal protection board of the City of Hamburg. Patients The study included patients with Katayama syndrome (n = 8) defined by fever, eosinophilia and a history of surface freshwater contact during a recent travel to a schistosomiasis endemic region. A second group had active, untreated disease defined by detection of eggs in stool or urine (n = 14). Most patients in this group were immigrants from endemic regions presenting to their primary care physician with acute manifestations like hematuria. Most of them where not aware of their disease. A third group of patients had treated schistosomiasis defined by prior anti-parasitic treatment and failure to detect viable eggs by microscopy (n = 30). Samples Serum and plasma samples were collected for antibody testing and DNA extraction, respectively. Serum was stored at +4°C and plasma was stored at −20°C prior to use. Stool samples were collected in Merthiolat-Iodine-Formol buffer and stored at +4°C until use. Serology All patient sera were tested for anti-Schistosoma antibodies by means of an extensively validated in-house EIA that has been described previously [15]. EIA used crude extracts from cercariae and adult worms of S. haematobium and S. mansoni, as well as extracts from adult worms of S. japonicum. Parasite detection Stool investigation was done essentially as described earlier [15]. For detection of S. haematobium eggs, urine was filtered as described by Peters et al. [39]. Microscopy was performed directly on untreated biopsies and on paraffin-embedded tissue. The latter was cut with a microtome into 5-µm sections. The sections were subsequently mounted on glass slides, stained with hematoxylin-eosin, periodic acid–Schiff and Trichrome stains and subsequently examined by an experienced pathologist for Schistosoma eggs. Plasma DNA preparation DNA from plasma was prepared by large volume phenol-chloroform extraction. In brief, up to 20 mL of plasma were mixed with an equal volume of phenol and centrifuged for 5 min at 1,200 g. The aqueous phase was transferred to a new tube, mixed with an equal volume of phenol∶chloroform 1∶1, and centrifuged 5 min at 3,500 rpm. Again the aqueous phase was transferred to a new tube, mixed with an equal volume of chloroform, and centrifuged 5 min at 3,500 rpm. DNA was precipitated by adding 1/10 volume of 3 M sodium acetate and 1 volume of 99% ethanol. After centrifugation for 1 h at 14,000 g the supernatant was discarded. To remove residual salt the pellet was washed with 1 mL ethanol 70% and centrifuged 20 min at 10,000 rpm. Supernatant was discharged. The DNA pellet was air-dried and dissolved in 50 µL of water and stored at −20°C. Real-time PCR In order to achieve high analytical sensitivity, the 121 bp tandem repeat sequence (GenBank accession number M61098) that contributes about 12% of the total Schistosoma mansoni genome sequence was chosen as the PCR target gene [40]. 20 µL reactions contained 3 µL of DNA, 2 µL 10X Platinum Taq PCR-Buffer (Invitrogen, Karlsruhe, Germany), 1.5 µL MgCl2 (50 µM), 200 µM of each dNTP, 0.8 µg bovine serum albumin, 500 nM of primers SRA1 (CCACGCTCTCGCAAATAATCT and SRS2 (CAACCGTTCTATGAAAATCGTTGT) each, 300 nM of probe SRP (FAM-TCCGAAACCACTGGACGGATTTTTATGAT-TAMRA), and 1.25 units of Platinum Taq polymerase (Invitrogen). Cycling in a Roche LightCycler® version 1.2 comprised: 95°C/5 min, 45 cycles of 58°C/30 s and 95°C/10 s. Fluorescence was measured once per cycle at the end of the 58°C segment. Quantification standard and technical sensitivity The PCR target fragment was cloned into plasmid by means of a pCR 2.1-TOPO TA cloning reagent set (Invitrogen, Carlsbad, California, USA). Plasmid purification was done with a QIAprep MiniPrep kit (Qiagen, Hilden, Germany). Plasmids were quantified by spectrophotometry. The standard plasmid was tested in 10-fold dilution series by PCR, showing a detection limit of 5.4 copies per reaction. If plasmids were inoculated into 200 µL of plasma prior to preparation, 68.8 copies per mL of plasma were detectable. Because the DNA contained in 200 µL was concentrated in 50 µL elution volume, of which 3 µL were tested by PCR, the PCR input at 68.8 copies per mL corresponded to a calculated 0.8 copies per PCR. Dilutions of the standard plasmid were also used as a quantification reference in real-time PCR. It should be mentioned that only approximate concentrations of Schistosoma DNA can be determined because the number of copies per genome of our target sequence varies between S.mansoni and hematobium and is unknown for S. japonicum [40]. Internal control Target gene nucleotides 39–79 bp were removed from the quantification standard plasmid, and replaced by an alternative probe binding site with techniques described earlier [41], using primers SRA-mut (ATCGTTCGTTGAGCGATTAGCAGTTTGTTT TAGATTATTTGCGGAGCGTGG) and SRS2-mut (CTGCTAATCGCTCAACGAAC GATTACAACGATTTTCATAGAACGGTTGG) for extension PCR, in combination with diagnostic PCR primers mentioned above. The resulting construct was cloned as described in the section “quantification standard”. Optimization of Schistosoma PCR for large plasma volumes One whole schistosome was ground in liquid nitrogen. Its nucleic acids were extracted and inoculated into human normal plasma. Different volumes of plasma were prepared by classical phenol-chloroform extraction, keeping the water volume in which DNA was resuspended at the end of the procedure constant at 50 µL. Parallel PCRs conducted on these nucleic acid solutions showed that an increase of detection signal was achieved up to an input volume of 10 mL of plasma, as evident by reduction of Ct values in real-time PCR. Above this input volume, no increase of signal was observed anymore, probably due to the introduction of interfering substances into PCR that were derived from large volumes of plasma. These experiments were repeated and confirmed with plasmid DNA spiked in human plasma. An input volume in humans of 10 mL of plasma was chosen as the volume to be analyzed in human diagnostic application in this study. It should be mentioned that outside this study, smaller volumes of plasma (down to 1 mL) were successfully used for CFPD detection. Quantitative correction factors Different input volumes of plasma were processed for mice or humans, respectively. For mice, 1 mL of plasma was extracted and the resulting DNA resuspended in 50 µL, of which 3 µL were tested in PCR. One DNA copy per PCR vial thus represented 16.7 copies per mL (50 / 3). For Humans, 10 mL of plasma were extracted and resuspended in 50 µL of water, of which 3 µL were tested in PCR. One DNA copy per PCR vial thus represented 1.67 copies per mL. Confirmation of PCR specificity Plasma from 30 blood donors and 35 patients examined for other conditions were tested by large-volume plasma extraction and CFPD real-time PCR. None yielded positive results. Statistical analysis The Statgraphics V 5.1 software package (Manugistics, Dresden, Germany) was use for all statistical analyses. T-tests were always two-tailed. Results In the case of tumors and pregnancy, cell-free DNA can be detected in plasma. Because the high turn-over rates of cells in these conditions resemble processes observed in parasitic infections, we reasoned that the detection of cell-free DNA from infecting parasites (CFPD) might be effective as a diagnostic approach in schistosomiasis. In preliminary experiments, stored serum samples from humans with confirmed schistosomiasis were processed with a method commonly used for detection of DNA viruses from cell-free plasma [42] and tested by Schistosoma PCR [21]. Plasma samples from mice infected with S. mansoni were also tested. In both cases, Schistosoma DNA was detectable in some but not all of the tested samples (data not shown). To determine systematically under which conditions and at what quantities CFPD was detectable in schistosomiasis, a quantitative real-time PCR assay for a Schistosoma multi-copy gene was established as described in the Materials and Methods section. A well-established mouse model of schistosomiasis was employed. In a first step it was tested whether CFPD circulated in plasma during the phase of chronic schistosomiasis. Four adult BALB/c mice were infected with 100 cercariae of S. mansoni and sacrificed after completion of the replication cycle on day 42 after infection. To enable testing of a large volume of mouse plasma, blood was pooled from four mice and one mL of pooled plasma was extracted. Quantitiative PCR with an absolute quantification standard (refer to Materials and Methods section) yielded a DNA concentration of 128.27 copies of CFPD target gene per mL of plasma (Figure 1, marked datum point). 10.1371/journal.pntd.0000422.g001 Figure 1 DNA copies per mL of pooled mouse plasma (y-axis, four mice per datum point) in mice infected intraperitoneally with 100 cercariae of S. mansoni. After completion of parasite maturation on day 42, mice were treated orally with praziquantel on day 45 (120 mg per kg). At the indicated times (x-axis), four mice were sacrificed, their blood pooled, and 1 mL of pooled plasma was tested as described in the Materials and Methods section for cell-free Schistosoma DNA. The untreated group is marked with an asterisk (*). It was next determined whether any associations might exist between the amount of living parasites in mice and the concentration of CFPD. Along with the four mice mentioned above, 16 more mice had been infected on the same day with the same dose of S. mansoni cercariae. On day 45 post infection all 16 mice were treated with a single oral dose of 120 µg praziquantel per gram body weight. This dose was known to eliminate Schistosoma in our model (own unpublished data). Groups of four mice were sacrificed on days 50, 80, 120, and 180 after infection, respectively, and from each group one mL of pooled plasma was tested. Figure 1 summarizes the CFPD target gene concentrations observed in all groups of mice, including the untreated group. Interestingly, in mice sacrificed five days after treatment the Schistosoma CFPD concentration in pooled plasma was considerably increased against the group that was sacrificed immediately before treatment (899.23 vs 128.27 target gene copies per mL). CFPD concentrations decreased to 182.93 and 70.97 target gene copies/mL on days 80 and 120, respectively, and became undetectable in the last group sampled on day 135 post treatment. It was concluded that the concentration of CFPD in plasma might be associated with the number of viable parasites or eggs in the mouse model, and the observed increase of CFPD immediately after treatment may have been due to parasite decay. To determine whether CFPD could also be detected in humans, fourteen patients with chronic disease were studied. These patients had been referred to our tropical medicine ward after being identified in routine screening for gastrointestinal conditions or other symptoms compatible with Schistosomiasis. It could not be reconstructed how long these patients had been infected, or how long ago they had been exposed. Diagnoses were initially made by EIA. Active infections were subsequently confirmed in all patients by microscopic detection of intact eggs in urine, stool, or organ biopsies. Either S. mansoni, or S. haematobium, or S. japonicum eggs were seen (Table 1). From each patient, 10 mL of plasma were extracted and tested for Schistosoma CFPD. All patients tested positive. The observed CFPD concentrations ranged from 1.22 to 27,930 target gene copies per mL of plasma. 10.1371/journal.pntd.0000422.t001 Table 1 Patients with chronic disease. Patient Suspected origin of infection Country of residence Residence status EIA Schistosoma species Sample in which eggs were detected Cell-free DNA cop/mLa 1 West Africa No data available Refugee + S. haematobium Bladder biopsy 27930.64 2 Mozambique Germany NGO worker + S. mansoni Rectum biopsy 27930.64 3 Nigeria Nigeria Immigrant + S. mansoni Rectum biopsy 3247.14 4 Egypt Egypt Immigrant + S. mansoni Rectum biopsy 1584.80 5 Egypt Egypt Immigrant + S. mansoni Rectum biopsy 1584.80 6 Philippines Germany Expatriate + S. japonicum Rectum biopsy 1584.80 7 Egypt Egypt Immigrant + S. mansoni Rectum biopsy 773.48 8 Zambia Germany NGO worker + S. mansoni Rectum biopsy 377.50 9 West Africa No data available Refugee + S. mansoni Rectum biopsy 184.24 10 Ghana Ghana Immigrant + S. mansoni Rectum biopsy 184.24 11 Gambia/Senegal Gambia/Senegal Immigrant + S. mansoni Rectum biopsy 21.42 12 West Africa West Africa Immigrant + S. haematobium Urine 21.42 13 Uganda Uganda Immigrant + S. mansoni Stool 2.49 14 Egypt Egypt Immigrant + S. mansoni Rectum biopsy 1.22 a Note that 10 mL of plasma were processed. 1 copy per mL = 1.67 copies per PCR vial. Because of the high detection rates in patients with active disease, it was tested whether CFPD might already be detectable in the early acute disease (Katayama syndrome). Eight patients were studied, as shown in Table 2. All of these patients had acute disease that was confirmed subsequently to be associated with Schistosoma infection. Although most patients were seen only in the third week of symptoms, two patients could be tested already on days 2 and 8 of symptoms, respectively. In three of eight patients, antibody EIA was still negative during the first visit. CFPD PCR was positive in all eight patients (Table 2). Target gene concentrations in the cohort seemed to increase with increasing times after exposure or after disease onset, as shown in Figure 2. Highest values were observed about six weeks from exposure or about 15 days from onset of symptoms. 10.1371/journal.pntd.0000422.g002 Figure 2 Cell-free Schistosoma DNA concentrations in plasma of patients with acute disease (Katayama syndrome) plotted against the days post exposure or post onset of symptoms when the tested samples were taken. 10 mL of plasma were tested for cell-free DNA. 10.1371/journal.pntd.0000422.t002 Table 2 Patients with Katayama syndrome. Patient Destination Purpose Visit DPEa DPOb DPTc LEUKd EOe EIAf Cell-free DNA copies per mLg 1 Mozambique Professional First 42 14 13.6 20.9 + 57227.85 Second 210 195 156 5.3 3.6 + 21.42 2 Ethiopia Professional First 42 15 10.2 26 − 27930.64 Second 135 120 79 7.3 4.1 + 1584.80 3 Uganda Professional First 56 18 10.1 19 + 21.42 Second 270 250 200 4.9 2.6 + 5.10 4 Uganda Professional First 12 20 7.3 22 + 10.45 Second 460 445 434 6.9 5.0 + 2.49 5 Malawi Tourist First 20 2 6.2 6.5 + 10.45 Second 750 740 716 5.2 0.8 + − 6 Mozambique Tourist First 56 21 50.7 65 − 13631.84 7 Jemen Tourist First 54 14 6.7 23 + 773.48 8 Malawi Tourist First 35 8 4.9 19.7 − 184.24 a Days post exposure with fresh water (most likely event). b Days post onset of symptoms. c Days post treatment for second visits. d Leukocyte count (n per nL). Average leukocyte count in patients 1 to 5: first visit, 9.48 cells/nl; second visit, 5.92 cells/nl (p<0.0017). e Percent eosinophiles in total leukocytes. Average eosinophile fraction in patients 1 to 5: first visit, 18.88%; second visit: 3.2% (p<0.033). f Enzyme immunoassay. g Note that 10 mL of plasma were processed. 1 copy per mL = 1.67 copies per PCR vial. It was next studied whether a decrease of CFPD concentration due to treatment could be confirmed. In the group of patients with Katayama syndrome, five of eight patients could be followed after treatment (Table 2). All five patients received praziquantel and prednisolon (1 mg/kg) within two weeks after initial diagnosis. A second treatment course (same dose of praziquantel, no prednisolon) was conducted in all patients 4 to 6 weeks later. Patients were appointed for control visits which took place 105 to 738 days after the initial visit (rows labelled “second visit” in Table 2). As expected, average leukocyte counts and levels of eosinophilia (% eosinophiles in leukocyte count) were significantly lower in second visits than in first visits. All patients had normal or only marginally increased eosinophile levels during their second visits (Table 2). Mean Schistosoma CFPD target gene concentrations in plasma were 17,040.20 copies/mL during first visits and 322.76 copies/mL during second visits. Means were significantly different (two-tailed T-test, p<0.05, Wilcoxon paired-sample test, p<0.04). Interestingly, only one patient had a completely negative CFPD PCR test during the second visit, and this was the patient with the longest interval between treatment and second visit. To obtain more data on Schistosoma CFPD concentrations after treatment, we tested 30 patients who had been treated for schistosomiasis during eight years in our institution, and who were available for a re-visit. These patients were in good clinical condition, had no eosinophilia, and had received between 1 and 6 treatment courses since their last exposure in endemic regions. Patient histories are summarized in Table 3. Ten of the 30 patients had positive CFPD PCR results. Intervals between treatment and PCR testing were significantly different between PCR-positive and PCR-negative patients (0.43 years vs. 3.4 years, p<0.0004, ANOVA f-test). The longest interval between treatment and a positive PCR result in any patient was 58 weeks. Interestingly, three of the ten patients with positive PCR showed dead eggs in histology. 10.1371/journal.pntd.0000422.t003 Table 3 Patients seen after treatment. Patient Country of origin Residence status Schistosoma speciesa Histologyb YPEc NTd TPTe Cell-free DNA cop/mLf 1 Egypt Immigrant S. haematobium Negative 3 1 2 wk 6653.16 2 Sierra Leone Refugee Unclassified Negative 6 1 2 wk 89.92 3 Guinea Immigrant S. mansoni DE 8 1 4 wk 89.92 4 Sierra Leone Refugee Unclassified Negative 7 2 13 wk 43.88 5 Zimbabwe/Botswana Immigrant Unclassified Negative 4 1 2 wk 21.41 6 German Tourist Unclassified Negative 3 3 24 wk 10.45 7 Ghana Immigrant S. mansoni EO 2 4 54 wk 10.45 8 Data not available Immigrant Unclassified DE 2 2 58 wk 2.49 9 Germany Expatriate Unclassified DE 3 2 9 wk 2.49 10 Egypt Immigrant Unclassified Negative 4 5 52 wk 2.49 11 Cameroon Immigrant S. haematobium Negative 2 3 2 y - 12 Uganda Immigrant Unclassified Negative 3 1 0 y - 13 Germany Tourist S. haematobium Negative 3 3 1 y - 14 Germany Tourist S. mansoni Negative 3 3 2 y - 15 Philippines Immigrant S. japonicum DE 4 6 2 y - 16 Ghana Immigrant Unclassified EO 4 2 3 y - 17 Germany Tourist S. mansoni Negative 4 3 3 y - 18 Egypt Immigrant S. mansoni EO 5 1 0 y - 19 Egypt Immigrant Unclassified Negative 5 4 3 y - 20 Cameroon Immigrant Unclassified EO 6 2 5 y - 21 Egypt Immigrant S. mansoni Negative 6 4 4 y - 22 Ghana Immigrant S. haematobium DE/EO 10 1 0 y - 23 Germany Tourist Unclassified Negative 10 3 8 y - 24 Egypt Immigrant S. haematobium Negative 6 2 6 y - 25 Germany Expatriate Unclassified Negative 5 3 4 y - 26 Cameroon Immigrant Data not available Negative 5 3 4 y - 27 Germany Tourist Unclassified Negative 5 3 4 y - 28 Cameroon Immigrant Unclassified Negative 6 3 5 y - 29 Egypt Immigrant Unclassified Negative 7 4 4 y - 30 Data not available Immigrant Unclassified DE 8 2 8 y - a Identified by microscopy during earlier active disease episode (recorded data). b Microscopic findings in colon or bladder biopsy upon re-visit. DE = degenerated eggs; EO = eosinophilic infiltrates; Negative = normal histology. c Years post exposure = time (years) between last exposure in endemic country and PCR testing. d Number of earlier treatment courses since last exposure. e Time post treatment = time (wk = weeks; y = years) between completion of last treatment course and PCR testing. f Note that 10 mL plasma were processed. 1 copy per PCR mL = 1.67 copies per PCR vial. To obtain an estimate of the approximate duration of CFPD detection after therapy, the CFPD target gene concentrations were plotted against time for all patients in this study who provided positive PCR results after treatment (patients from the Katayama syndrome ever group and post-treatment group). As shown in Figure 3, linear regression or exponential curve fitting suggested that negative results could be expected by weeks 82 or 120 after treatment, respectively. 10.1371/journal.pntd.0000422.g003 Figure 3 Cell-free Schistosoma DNA concentrations after treatment. DNA concentrations were plotted only for those patients still showing cell-free Schistosoma DNA in plasma after treatment. These data were pooled from patients who had been followed prospectively after being diagnosed with Katayama syndrome, as well as from patients examined retrospectively after concluded treatment. Linear regression analysis yielded the graph equation Y = 2.03−0.02 X. Exponential regression yielded the graph equation Y = e ∧ −0.02 (X−30.4). Discussion Schistosomiasis involves a wide range of symptoms and is difficult to diagnose. In this study we have explored the utility of detecting cell-free parasite DNA (CFPD) in serum as an alternative to detecting eggs in stool, urine, or organ biopsies. The concept of using cell-free DNA for diagnostic purposes has been proven in oncology and prenatal diagnostics [25]–[33]. It was our rationale that schistosomiasis involves parasite turnover, liberating DNA from decaying parasites that would reach the blood. Unlike eggs in stool or urine, CFPD in plasma would not undergo random sampling effects that complicate diagnostics. By means of a well-established murine model of schistosomiasis, it was confirmed that DNA could be detected in plasma during active disease, and that praziquantel treatment led to clearance of Schistosoma CFPD from plasma. Consistent with the hypothesis that circulating Schistosoma DNA stemmed from decaying parasites, a marked increase of CFPD concentration was observed in plasma of mice sampled short after initiation of therapy. Because of the large differences in plasma volume between mice and humans, we have not undertaken any further mouse experimentation but continued a proof-of-concept study on available patients with schistosomiasis in various clinical stages. In a first approach, we showed that CFPD could be detected in all of 14 patients with active disease. Due to the small number of available patients, this finding clearly awaits confirmation in larger studies. It should also be mentioned that the sensitivity of our assay may vary between Schistosoma species, as the target gene has not been formally evaluated in S. japonicum (e.g., our whole study contained only one patient with S. japonicum), and it has been shown that S. hematobium contains less copies of it than S. mansoni [40]. More recent PCR protocols (e.g., [21]) may be better suited to detect all species with the same sensitivity. This study therefore does clearly not provide a protocol intended for direct transfer into clinical application. Nevertheless, it is an interesting perspective that CFPD PCR might reach a clinical sensitivity of 100% for active schistosomiasis. In industrialized countries, it may be easier to find well-equipped molecular diagnostic laboratories than experienced microscopists with sufficient expertise in Schistosoma egg detection. Because of the ease of taking blood samples, and in view of the risk contributed by undiagnosed Schistosomiasis, it could become a realistic option to integrate Schistosoma CFPD PCR in routine diagnostic regimens for the clarification of gastrointestinal or urological conditions. Katayama syndrome caused by acute Schistosoma infection is a major differential diagnosis in returning travellers presenting with fever of unknown origin [6]. Although eosinophilia is a helpful criterion to distinguish Katayama syndrome from other conditions such as malaria or dengue fever, it is difficult to make a distinctive diagnosis due the shortcomings of serology and the inability of demonstrating Schistosoma infection before egg production [14]. We have demonstrated here that CFPD can be detected very early after onset of symptoms in patients with Katayama syndrome. Despite the limited number of patients studied, the concentrations of CFPD observed in our patients were well above the detection limit of the PCR assay. Based on experiments on limiting dilution series and quantitative correction factors as described in the Materials and Methods section, it could be assumed that the technical sensitivity limit of our assay was ca. 1.67 CFPD target gene copies per mL of plasma. The earliest patient with Katayama syndrome sampled on day 2 of symptoms already had a plasma concentration of ca. 10 copies per mL. If larger studies can confirm the high clinical sensitivity seen in our study, the detection of CFPD in plasma might become an accepted way of ruling out Katayama syndrome. It should be mentioned here that we have meanwhile modified our protocol by testing smaller volumes of plasma (in the order of 1–2 mL) and using a larger input volume of DNA in PCR. This modification makes the method easier to handle in routine laboratories, and still seems to provide sufficient sensitivity to diagnose patients with Katayama syndrome. A third field of application is the monitoring of therapy. In order to prevent relapse and long term sequelae from insufficient treatment, it is important to achieve a laboratory confirmation of the success of treatment [18], [43]–[45]. Unfortunately, patients after therapy as well as patients after a long course of disease with spontaneous healing (“burnt out bilharzia”) are difficult to judge based on clinical or laboratory findings [16],[18]. Several repetitive, parallel samplings are necessary to increase the statistical chance of detection of eggs by microscope, and thus to increase the clinical sensitivity of laboratory diagnostics [20],[22],[46]. This problem applies not only to microscopy, but also to conventional PCR on stool or urine samples [21],[47]. In the latter tests, there are additional issues such as PCR inhibition in stool samples. We have shown here that the concentration of CFPD in plasma was significantly reduced after therapy. The average CFPD concentration in those patients who still had detectable DNA after treatment (25.1 copies per mL) was significantly lower than in patients with Katayama syndrome (first visits, 537 copies per mL) or active disease (323.6 copies per mL), as determined by ANOVA (F-test, p<0.035; refer to Figure 4 for a Box Plot diagram). The decline of CFPD concentration in patients before and after treatment may thus become an effective parameter for monitoring patients under therapy. On the contrary, we were surprised to see that it took considerably longer in humans than in mice for CFPD PCR to become entirely negative after treatment. Lo et al. have determined that the half-life of fetal DNA in mother's plasma after birth ranges between 4 and 30 minutes [48]. In our study, pooled data from patients followed prospectively and patients re-examined retrospectively after treatment suggest that it may take more than one year until CFPD becomes entirely undetectable. Although we have no experimental evidence, it can be speculated that inactive eggs may release DNA with very slow kinetics. The greater number of eggs in humans with chronic disease as opposed to mice in our experiments may be responsible for a considerably longer duration until CFPD is totally eliminated in humans. Future studies should address the utility of paired CFPD determinations in individual patients before and after treatment, rather than insisting on negative CFPD results for a confirmation of treatment success. 10.1371/journal.pntd.0000422.g004 Figure 4 Box plot analysis of cell-free DNA concentrations in patients with Katayama syndrome (first visits only), patients with chronic disease, and all patients who had positive plasma PCR after treatment (pooled from treated Katayama syndrome patients and patients examined retrospectively after treatment). Boxes represent the innermost two quartiles (25%–75% percentiles = interquartile range, IQR) of data. The whiskers represent an extension of the 25th or 75th percentiles by 1.5 times the IQR. The notches represent the median +/−1.57 IQR √n. If the notches of two boxes do not overlap, the medians ( = notch centers) are significantly different (true for the active disease vs. the treated group). In summary, the detection and quantification of CFPD from plasma might carry the potential of becoming a novel diagnostic tool for any stage of schistosomiasis. With increased automation and better instrumentation for molecular diagnostics, the cost efficiency and quality of results in clinical laboratories can exceed that of repetitive diagnostic determinations by microscopy. The cost of reagents and consumables for our method range around 3 USD per determination, which is probably too expensive in many endemic countries. However, this price is compatible with application in funded surveillance and control programmes, and should be affordable for individualized application in emerging countries. Instrumentation and expertise for proper PCR diagnostics has considerably improved in many countries due to the demands created by HIV and TBC treatment programmes. If future studies can prove the clinical benefits suggested here, Schistosoma CFPD PCR may become a new priority in molecular diagnostics in developing and emerging countries.
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- Abstract: found
- Article: found
A Schistosoma haematobium-Specific Real-Time PCR for Diagnosis of Urogenital Schistosomiasis in Serum Samples of International Travelers and Migrants
Lieselotte Cnops, Patrick Soentjens, Jan Clerinx … (2013)
Introduction Urogenital schistosomiasis due to Schistosoma haematobium is a serious underestimated public health problem. It is endemic in 53 countries of the African continent and of the Middle East [1], [2]. Adult worms live in the capillary plexus of the bladder and other parts of the urino-genital system and eggs are excreted in the urine and occasionally found in feces. Diagnosis of S. haematobium infections is traditionally done by microscopy but is often unreliable due to the circadian and day-to-day variations in egg excretion, and to low parasite load, especially in the traveler. Antibody-based assays are useful to confirm infection, but do not distinguish active infection from past exposure, and false-negative results occur, mainly in S. haematobium infections. Antibody tests are usually negative during acute symptomatic schistosomiasis. Assays that detect circulating antigens seem very promising in the early phase of infection but still lack sensitivity in the diagnosis of light infections [3], [4], [5], [6]. Recently, we developed a genus-specific real-time PCR (further called ‘genusPCR’) that sensitively detect all human infectious Schistosoma species in feces and urine [7]. The genusPCR was not able to detect schistosome DNA in serum although molecular analysis of serum is of interest in acute schistosomiasis before detectable levels of eggs are excreted [8]–[12]. In 2009, Wichmann and colleagues [10] described a real-time PCR, targeting a highly repeated 121-bp sequence of S. mansoni (named Sm1-7) to detect cell-free schistosome DNA in serum. This was proven successful in acute and chronic S. mansoni infection, but not so much in S. haematobium infection. To fill that gap, we developed a real-time PCR specific for the diagnosis of S. haematobium in serum samples. The real-time PCR targets Dra1, a S. haematobium-specific 121-bp repeat sequence originally described by Hamburger et al. and present in hundreds to thousands of copies and representing at least 15% of its genome [13]. We first tested this PCR (further called ‘draPCR’) on urine and feces samples to evaluate its species-specificity and its performance in comparison with microscopy, and then on serum samples to determine its potential as diagnostic tool for acute phase schistosomiasis. Methods Ethics statement The diagnostic procedures described in this manuscript are part of the standard diagnostic work-up of patients suspected of schistosomiasis. All samples were routine diagnostic samples from patients presenting at the Institute of Tropical Medicine (ITM, Antwerp, Belgium) policlinic and were stored after completion of the routine tests. The ITM has the policy that sample left-overs of patients presenting at the ITM policlinic can be used for research unless the patients explicitly state their objection. The Institutional Review Board of ITM approved the institutional policy of this presumed consent as long as patients' identity is not disclosed to third parties. All data have been analysed anonymously. Clinical samples PCR analysis was retrospectively performed between January and October 2012 on samples that were stored at 0.45×109/L), a positive serology (IHA titer ≥1/160 or positive ELISA), the presence of Charcot-leyden crystals in feces or hematuria (>7 RBC/µL) or who travelled together with a confirmed case. DNA extraction As previously described [7], [14], DNA was extracted with the QIAamp DNA stool mini kit (Qiagen Benelux, Venlo, The Netherlands) from 1 gram of feces that was dissolved into 5 mL ASL buffer (Qiagen). An 200 µl urine sediment was processed for DNA extraction with the QIAamp DNA mini kit (Qiagen) after centrifugation of 10 mL of urine and three wash/centrifugation steps. For DNA extraction of serum with the QIAamp DNA MIDI kit (Qiagen) or by phenol/chloroform, 1 to 2 mL of serum was used [7]. Control samples Positive control DNA of S. mansoni and S. haematobium were kindly provided by Dr. T. Huyse (ITM/KUL, Belgium). Positive control DNA of S. intercalatum and S. guineensis were kindly provided by Dr. F. Allan from SCAN at the Natural History Museum (London, UK) [15]. DNA was extracted from one adult worm and used in a 1/100 dilution (∼0.1 ng/µL). Positive control DNA of S. mekongi was obtained from a stool sample of a patient seen at ITM, containing 50 eggs per gram (EPG). Positive control DNA of S. japonicum derived from cercariae spotted on FTA filter paper kindly provided by Dr. J.P. Webster (London, UK). Primer and probe design The highly repetitive Dra1 sequence of S. haematobium (Accession number DQ157698.1) was selected as target and primers were identical to those described [13] (Sh-FW 5-gatctcacctatcagacgaaac-3′; Sh-RV 5′-tcacaacgatacgaccaac-3′). An additional fluorescent labeled hydrolysis probe was developed (Sh-probe 5′-tgttggtggaagtgcctgtttcgcaa-3′) for real-time monitoring of the PCR signal and was labeled with a 5′-FAM reporter, an internal ZEN quencher and a IowaBlack Fluorescent Quencher at the 3′-end (IDT, Leuven, Belgium). The amplicon size was 96 base pairs. Real-time PCR The draPCR was performed with a 25 µL reaction mix containing 5 µL DNA, 1× Perfecta qPCR Supermix (Quanta Biosciences), 500 nM of Sh-FW and Sh-RV primer, 250 nM of Sh-probe and 0.1 mg/mL bovine serum albumin. The program consisted of an initial step of 2 min at 95°C followed by 50 cycles of 15 s at 95°C and 60 s at 60°C. The reaction was run on the SmartCycler II (Cepheid Benelux, Belgium). DNA detection was expressed by Cycle threshold (Ct)-values. In every run, the non-template control was negative (Ct = 0) and the S. haematobium control was positive. To detect DNA of schistosome species other than S. haematobium, the Sm1-7PCR and genusPCR were performed as described before [7], [10]. PCR validation The primer and probe design was verified with Integrated DNA Technology (IDT) Oligo Analyzer software (v3.1) (http://eu.idtdna.com/analyzer/Applications/OligoAnalyzer/). Primer and probe specificity was checked in silico by BLAST analysis (http://blast.ncbi.nlm.nih.gov/Blast.cgi) and by 2% agarose gel electrophoresis at 100 V for 35 minutes. The analytical specificity of the PCR was tested on a panel of clinical control samples containing 23 different intestinal or blood parasites. The panel included stool samples (n = 14) from patients infected with protozoa (Giardia lamblia, Entamoeba dispar, E. histolytica, Blastocystis hominis, Enterocytozoon bieneusi, Encephalitozoon spp), nematodes (Ascaris lumbricoides, Strongyloides stercoralis, Trichostrongylus spp., Trichuris trichiura, or Ancylostomidae), trematodes (Clonorchis spp., Fasciola hepatica) or a cestode (Taenia saginata) and blood samples (n = 9) from patients infected with Plasmodium falciparum, P. vivax, P. ovale, P. malariae, Leishmania donovani, Loa loa, Onchocerca volvulus, Dirofilaria repens or Trypanosoma brucei rhodesiense. The detection limit was determined on a 10-fold dilution series of a stool sample containing 580 EPG of S. haematobium. It was diluted in a negative stool sample which was dissolved in ASL buffer (Qiagen). DNA was extracted from each dilution and the highest dilution with a positive signal indicated the detection limit. The variation in Ct-values was determined in a serum sample that was processed 8 times within the same run (repeatability) or 5 reactions that were run at different days (reproducibility). The coefficient of variation (CV, expressed as %) of the Ct-values was calculated. Microscopy Microscopy was performed on a single urine and/or fecal sample per patient at the time of presentation and in some cases, on a single follow-up sample one month after treatment. Diagnosis of schistosomiasis was confirmed when S. haematobium or S. mansoni eggs were detected in urine and/or feces. Microscopic examination of urine samples was performed on the sediment of at least 20 mL end-stream urine and of stool samples following a concentration method on 3 grams of feces that had been homogenized in 42 mL of 10% formaldehyde-saline solution [16]. The infection intensity in stool was expressed by the number of EPG. The limit of detection was 10 EPG. Serology A combination of an in-house enzyme-linked immunosorbent assay (ELISA) using S. mansoni antigen (mixture of egg and adult worm extract) and an indirect hemagglutination inhibition assay (IHA), using a S. mansoni adult worm extract (ELI.H.A. Schistosoma, EliTech MICROBIO, France) with a cut-off at 1/160, were used to detect anti-schistosome antibodies. Results Primer and probe design IDT Oligo analysis approved no self- or heterodimerization between the primers and the probe. BLAST analysis with probe and primers indicated 100% query coverage and maximum identity with S. haematobium. No species other than Schistosoma were in silico recognised by the primers and probe. Gel electrophoresis obtained a single band of expected length for the amplicon of S. haematobium and no signal for the non-template control. PCR validation To determine the species-specificity, schistosome species of the three complexes were tested with the draPCR, the Sm1-7PCR and the genusPCR. The DNA controls of human species of the S. haematobium complex revealed a strong signal with the draPCR (Ct-values ranging from 15.21 to 16.65) and the cattle species S. bovis revealed a signal of medium intensity (Ct 29.68). DNA controls of other Schistosoma species gave no (S. japonicum, S. mekongi) or a very weak signal (S. mansoni, Ct 41.93) (Table 1). In comparison, the genusPCR easily recognized all species of the three complexes while the Sm1-7PCR detected a strong signal for S. mansoni and S. bovis, a medium to weak signal for the human species of the S. haematobium complex and no signal for species of the S. japonicum complex (Table 1). 10.1371/journal.pntd.0002413.t001 Table 1 Species-specificity of the draPCR in comparison to the Sm1-7PCR and the genusPCR. complex species host extract from draPCR Sm1-7PCR genusPCR S. mansoni complex S. mansoni human adult worm 41.93* 15.08 21.35 S. haematobium complex S. haematobium human adult worm 15.21 29.44 20.67 S. intercalatum human adult worm 16.65 39.16 23.64 S. guineensis human adult worm 15.12 38.97 23.26 S. bovis cattle adult worm 29.68 13.49 18.92 S. japonicum complex S. japonicum human cercariae 0.00 0.00 23.52 S. mekongi human clinical sample 0.00 0.00 27.98 Ct-values indicate a strong (Ct 38) or no (Ct = 0) recognition of the species. * a 5-fold dilution series on this DNA demonstrated that the weak signal resulted from cross-amplification and not from a-specific background noise. Of interest is the difference in Ct-values measured for S. haematobium (Ct 15.21) and S. mansoni (Ct 41.93) by the draPCR. Since the amount of amplicon doubles every PCR cycle (i.e. increase by one log2), the difference of 26 Ct's is equivalent to a more than 67 million times lower sensitivity to detect S. mansoni in comparison to S. haematobium. The same counts for the detection of the S. bovis species in comparison to the other S. haematobium complex species by the draPCR with a difference of 14 Ct's that accounts for a 16,000 times lower sensitivity (Table 1). No cross-reaction was seen with the draPCR in the 23 control samples with intestinal and blood parasites other than Schistosoma. The analytical sensitivity demonstrated a detection limit of 0.5 EPG. Repeatability and reproducibility testing revealed a CV of 1.03% and 1.04% respectively. PCR analysis on urine samples and biopsy A panel of 110 urine samples and one bladder wall biopsy was analysed with the draPCR. A positive PCR signal was obtained in 14 urine samples (Ct-values ranging from 16.77 to 32.40) of which seven were positive for S. haematobium ova by microscopy (Table 2). The other seven urine samples (Ct-values ranging from 30.74 to 46.63) were from patients treated for schistosomiasis two weeks or one month before (n = 2) or from patients without previous treatment but with anti-schistosome antibodies (n = 3) or with S. haematobium eggs in feces (n = 1) or in a urine sample obtained three days earlier (n = 1). The latter urine sample also contained Trichomonas vaginalis. The 96 urine samples that were negative for S. haematobium or any other parasite by microscopy, were also negative by PCR (Table 2). 10.1371/journal.pntd.0002413.t002 Table 2 Evaluation of the draPCR on urine samples. Microscopy (urine) Dra PCR S. haematobium negative Total positive 7 7 14 negative 96 96 Total 7 103 110 The biopsy containing S. haematobium eggs was positive by PCR with a Ct-value of 17.08. PCR analysis on stool samples The draPCR was evaluated on a panel of stool samples in which eggs of S. haematobium (n = 11), of S. mansoni (n = 21) or no schistosome eggs (n = 52) were microscopically detected. All samples with eggs of S. haematobium were positive (Ct-values ranging from 20.35 to 37.87) and all samples with eggs of S. mansoni were negative (Table 3). In addition, the draPCR revealed a positive signal in three samples without eggs (Ct-values varied from 36.78 to 45.33), two of which were follow-up samples of confirmed patients one month after treatment and one from a clinically suspected patient with eosinophilia. 10.1371/journal.pntd.0002413.t003 Table 3 Evaluation of the draPCR on feces. Microscopy (feces) DraPCR S. haematobium S. mansoni negative Total positive 11 3 14 negative 21 49 70 Total 11 21 52 84 PCR analysis on serum There is no reference method for schistosome DNA detection in serum. We therefore used the level of evidence of infection (confirmed S. haematobium (n = 12) or S. mansoni infection (n = 20) or suspected cases (n = 64)) as a reference (Table 4). Of all suspected cases, 8/64 were migrants and 56/64 travelers of whom serum was collected within 12 weeks upon return in 39 travelers and after more than 12 weeks (varying from 91 to 336 days) in 17 travelers. 10.1371/journal.pntd.0002413.t004 Table 4 Evaluation of the draPCR on serum. Infection status Number of serum samples (number of patients) DraPCR confirmed S. haematobium confirmed S. mansoni Suspected Total positive 22 (11) $ 5 (2) ○ 27(13) negative 1 (1) * 22 (20) 85 (62) 108 (83) Total 23 (12) 22 (20) 90 (64) 135 (96) $ Of the 22 samples, 11 serum samples were obtained at the moment of egg detection, one sample before egg detection, and 10 samples after treatment. Of the 11 patients with a confirmed S. haematobium infection, 7 patients demonstrated eggs in urine (see Table 2 ) and 4 patients had eggs in feces (part of the group described in Table 3 ). * only 1 mL of serum was available for PCR analysis and it was obtained 514 days after treatment. ○ 2 samples were collected before treatment, and 3 samples were taken 21 to 75 days after treatment. In total, 135 serum and blood samples were analysed from 96 patients of whom 64 patients with a single serum sample and 32 patients with one or more follow-up samples. No discordant results were obtained in different samples from the same patient. The draPCR was positive in 27 samples from 13 patients of which 11 patients (22 samples) with a microscopic confirmed S. haematobium infection and two patients (5 samples) with a clinical suspicion based on the presence of eosinophilia (0.88 and 2.13 10*9/L) and anti-schistosome antibodies (IHA 1/160 and 1/640) (Table 4). Of all 22 PCR positive samples of individuals with a confirmed S. haematobium infection, all serum samples collected at the same date of the parasitological confirmation (n = 11), were positive. All follow-up serum samples obtained 14 to 96 days after treatment (n = 10 from 8 patients), were also positive by PCR but demonstrated higher Ct-values. In two of the 8 patients, both serological tests remained negative 1 month and 2 months after treatment. PCR was additionally positive on one serum collected about 5 weeks after exposure (n = 1) while at that moment the urine and feces were microscopically negative and no antibodies could be detected. Detection of ova 42 days later, confirmed the S. haematobium infection. In one serum sample from a patient for which a single S. haematobium egg was detected in urine 514 days after treatment, no PCR signal was observed but the analysis was performed on an insufficient volume of serum (1 mL instead of 2 mL). All other serum samples with a negative PCR signal were from patients with a confirmed S. mansoni infection (20 patients, 22 samples) or from clinically suspected patients without schistosome eggs in urine and feces (62 patients, 85 samples) (Table 4). Four of these suspected patients had microscopically confirmed infections with Ascaris lumbricoides (n = 1), Ancylostomidae (n = 1), Strongyloides stercoralis (n = 1) or Trichuris trichuria (n = 1) and nine had been treated 6 months to 3 year prior to serum collection. The 85 PCR negative samples showed schistosomal antibodies by ELISA (n = 4), IHA (n = 10) or both (n = 15) and/or belonged to patients suspected because of recent freshwater exposure or eosinophilia. Discussion Urogenital schistosomiasis remains an important public health problem affecting approximately 112 million people, about half of the worldwide schistosome infections [2]. Schistosomiasis imported by travelers, expatriates and migrants is often caused by S. haematobium, with a frequency in the same range to that of S. mansoni [6], [17]–[19]. The presently developed PCR was designed to be used as a highly sensitive diagnostic tool for urogenital schistosomiasis in travelers returning from endemic regions. We opted for a real-time PCR format which has a short turn-over time and is preferred over conventional PCR methods due to its lower risk of contamination and higher sensitivity [20], [21]. The latter is of particular interest in travelers as they have often low parasite loads in the acute phase [6], rendering confirmation by microscopy erratic. The draPCR was able to detect all microscopy-confirmed S. haematobium infections in urine, bladder wall biopsy and feces and demonstrated no cross-reaction in clinical samples with microscopy-confirmed S. mansoni and other intestinal or blood parasites. Moreover, the draPCR detected ten extra S. haematobium-positive samples (7 urine and 3 stool samples) from 8 non-egg excretor patients that were highly suspected for urogenital schistosomiasis based on recent freshwater exposure, a strong antibody response (IHA≥1/1280) and/or the presence of eosinophilia. This confirms previous findings that PCR is highly sensitive in urogenital schistosomiasis diagnosis [22]–[24]. The draPCR can be of value on urine and stool samples of suspected patients when no eggs can be demonstrated by microscopy, especially as sampling is not invasive. Alternatively, the genus-specific PCR [7] could be used on urine and feces enabling the detection of schistosome DNA, regardless the causal species. Besides the excellent performance of the draPCR on urine and stool samples, the most striking result of this study is the specific detection of S. haematobium in serum. All but one of the serum samples from patients with a confirmed S. haematobium infection, and none of the serum samples from patients with a confirmed S. mansoni infection, were positive with the draPCR. Schistosomal DNA could additionally be detected in the serum of one patient about 5 weeks after freshwater exposure and 42 days before confirmation of the S. haematobium infection by microscopy. This clearly demonstrates the diagnostic potential of the draPCR to detect S. haematobium in serum during the acute phase of the infection. One serum sample of a patient with a confirmed S. haematobium infection was negative, which could be explained by previous treatment or the rather low volume (1 mL) of serum analysed. Due to the retrospective design of this study, PCR could not always be performed on an adequate volume of serum. We consider 2 mL as the optimal volume required for analysis. Two extra S. haematobium infections were detected by the draPCR in serum samples of suspected patients. False-positivity by PCR seems very unlikely as both patients had recent exposure to freshwater in Mali, developed typical severe symptoms related to Katayama syndrome with hypereosinophilia and had a positive serological response five to eight weeks post-exposure. Moreover, S. haematobium DNA was also detected in the follow-up serum samples while urine and feces remained negative after treatment. The decreasing PCR signal in follow-up samples demonstrates the PCR's potential to semi-quantitatively monitor treatment. Extra studies are required to confirm this. Further research could additionally compare the persistence of detectable levels of parasite DNA in serum with levels of circulating antigen that are more related to the actual worm burden and rapidly decrease after treatment [25]. Also, more scientific data is needed to assure when parasite DNA is cleared from the blood stream after treatment in order to determine the discriminating power of PCR between active or past present infections with the same Schistosoma species. PCR positivity was seen until at least 3 months after treatment in the present study, and up to more than one year after treatment in patients with S. mansoni infections [10]. A drawback of this study is that we had no genital samples available to test with the draPCR. S. haematobium parasites can cause genital schistosomiasis [1], [26] but in travelers, only few cases were reported [26], [27]. Only one bladder wall biopsy was tested, and although this does not allow drawing conclusions, it is worth to mention that examination of tissue samples by PCR might be helpful to diagnose schistosomiasis in non-egg excretor individuals. Another drawback is that due to the retrospective sample collection, early acute phase samples were not frequently available because antibody or egg detection followed weeks later. A prospective study is needed with selection of patients during the (early) acute phase in order to compare the performance of different tools for urogenital schistosomiasis diagnosis [24]. The PCR targets Dra1, a sequence specific for S. haematobium [13] and previously successfully used in a conventional PCR on cercaria and infested snails [28] and on urine [24]. Similar to the Sm1-7 121-bp tandem repeat sequence that comprises 11% of the S. mansoni genome [8], [10], [29] and to the multicopy retrotransposon gene representing 14% of the S. japonicum genome [30], Dra1 also has a highly repetitive nature and represents 15% of the S. haematobium genome [13]. The presence of multiple copies of this target sequence enables the highly sensitive detection of S. haematobium DNA in serum and probably explains why the single copy 28S gene used in the genusPCR [7] was not successful to detect schistosome DNA in this matrix. The findings of this study demonstrate that the draPCR for detection of S. haematobium infections in serum is complementary to the Sm1-7PCR that is most sensitive to detect S. mansoni infections [9]–[10]. Furthermore, we demonstrated the species-specificity of both PCRs in control DNA of adult worm extracts. Apart from the strong signal for the human species of the S. haematobium complex group, we also observed a very weak signal for S. mansoni with the draPCR and a weak signal for S. haematobium with the Sm1-7PCR. This can be explained by the fact that the highly repetitive sequences of S. haematobium or S. mansoni respectively, are most likely present in at least a single copy in the genome of other Schistosoma species [13] and are only detectable when a huge amount of parasite DNA is present as in the case of the adult worm extracts. Since we did not detect a signal with the draPCR in all 43 clinical samples of patients with confirmed S. mansoni infections (egg load varying between 10 and 120 EPG), we conclude that the draPCR and Sm1-7PCR are suitable for analysis of serum of patients suspected for urogenital and intestinal schistosomiasis, respectively. What do we detect by the draPCR in serum, urine and feces? In analogy with prenatal diagnostics and oncology [31], [32], Wichmann et al [10] used the term ‘cell-free parasite DNA’ (CFPD) to comprise the DNA that was detected in serum. Indeed, due to the high parasite turnover, diverse stadia of the parasite might be present in the blood circulation and are detectable depending on the phase of the infection. Once penetrated through the human skin, schistosomules travel with the venous circulation to the lungs within 7 to 10 days and thereafter to the liver region for maturation [1]. In the acute phase of urogenital infections, the draPCR probably detects DNA of degrading schistosomules or juvenile worms that did not survive or mate. Schistosomes are complex multicellular eukaryotes, and the schistosome DNA in serum might also originate from rapid turn-over of the tegument during maturation of the worm [33]. In addition, after the upstream migration of mature worms to the venous plexus of the bladder and deposition of eggs, DNA of eggs that circulate into the systemic circulation due to retrograde venous flow could be detected. In chronic infections, DNA from desintegrated eggs, or from killed worms after treatment could also be a target for PCR in serum. In urine samples, the PCR primarily detects DNA from S. haematobium eggs. It is not unlikely that also transrenal nucleic acids of breakdown products of the parasite are detectable in the urine as demonstrated before for S. mansoni [34], [35] and other parasitic infections [36], [37]. In feces, DNA of S. haematobium eggs can be found due to the atypical location of the worm in the colon or rectal wall, or due to contamination of the stools with urine in case of a high-intensity infection [1], [38]. So far, no S. haematobium-specific PCR has been described before to be used in human serum of recently infected travelers. Our findings suggest that the draPCR in serum is suitable for diagnosis of urogenital schistosomiasis in a non-endemic setting and might be of value in diagnosing travelers during the acute phase of infection (4 to 6 weeks after exposure to infested water) before eggs excretion and seroconversion, and in light infections. Serology tests turn positive only about 6 to 12 weeks after exposure [6]. In addition, weak positive serological reactions are difficult to interpret and false-negative tests occur, especially with S. haematobium [4], [39], [40]. Further prospective evaluation of the draPCR on serum samples is needed, to demonstrate its diagnostic role during the early acute phase of the infection. Supporting Information Supporting Information S1 STARD checklist. (DOC) Click here for additional data file. Figure S1 Flowchart for evaluation of the draPCR. (DOC) Click here for additional data file.
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- Abstract: found
- Article: not found
Postmortem diagnostic investigation of disease in free-ranging marine turtle populations: a review of common pathologic findings and protocols.
Mark Flint, Janet C Patterson-Kane, Colin Limpus … (2009)
Over the past few decades, there have been increasing numbers of reports of diseases in marine turtles. Furthermore, in recent years, there have been documented instances of apparently new diseases emerging in these species of which the etiology and/or pathogenesis remain unknown. These instances i) raise concern for the survival of marine turtles, and ii) question the health and stability of the benthic marine environments in which turtles live. Knowledge of common disease processes and pathologic changes in lesions, along with a standardized approach to postmortem and sample collection are required to document and understand the host-agent-environment interactions in marine turtle health. This review combines, for the first time, a standardized approach to the postmortem of marine turtles for veterinary clinicians, with a concurrent descriptive review of the gross and microscopic pathologic changes in lesions commonly seen.
Author and article information
Contributors
Erica Marchiori:
ORCID: http://orcid.org/0000-0002-4401-0636
erica.marchiori@unipd.it
Journal
Journal ID (nlm-ta): BMC Vet Res
Journal ID (iso-abbrev): BMC Vet. Res
Title:
BMC Veterinary Research
Publisher:
BioMed Central
(London
)
ISSN
(Electronic):
1746-6148
Publication date
(Electronic):
14
January
2020
Publication date PMC-release: 14
January
2020
Publication date Collection: 2020
Volume: 16
Electronic Location Identifier: 16
Affiliations
[1
]ISNI 0000 0004 1757 3470, GRID grid.5608.b, Department of Animal Medicine, Production and Health, , University of Padova, ; Viale dell’Università 16, 35020 Legnaro, PD Italy
[2
]ISNI 0000 0004 1758 0806, GRID grid.6401.3, Department of Integrative Marine Ecology, , Stazione Zoologica Anton Dohrn, ; Villa Comunale 1, 80121 Naples, Italy
[3
]ISNI 0000 0004 1758 0806, GRID grid.6401.3, Centro Ricerche Tartarughe Marine, , Stazione Zoologica Anton Dohrn, ; Via Nuova Macello 16, 80055 Portici, NA Italy
[4
]Veterinary practitioner, Centro Recupero Il Benvenuto, S.S. 16 2287/C, 45038 Polesella,
RO Italy
[5
]Istituto Zooprofilattico Sperimentale Abruzzo e Molise “G. Caporale”, Via Campo Boario,
64100 Teramo, Italy
Author information
Erica Marchiori
http://orcid.org/0000-0002-4401-0636
Article
Publisher ID: 2232DOI: 10.1186/s12917-020-2232-y
PMC ID: 6961338
PubMed ID: 31937305
SO-VID: 026f21c0-31f9-4458-9c53-6af649d800b1
Copyright © © The Author(s). 2020
License:
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License ( http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver ( http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
History
Date
received
: 9
October
2019
Date
accepted
: 3
January
2020
Funding
Funded by: Società Italiana di Parassitologia
Award ID: Young Researcher Grant 2017
Award Recipient
:
Erica Marchiori
Categories
Subject:
Research Article
Custom metadata
issue-copyright-statement © The Author(s) 2020
ScienceOpen disciplines: Veterinary medicine