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Blastocystis sp. are among the most commonly observed intestinal parasites in routine clinical parasitology. Blastocystis in humans consists of at least 9 genetic subtypes. Different subtypes of Blastocystis may be associated with differences in pathogenicity and symptomatology.
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Advanced microscopy on two samples and sequence-confirmed PCR on a third sample from the same individual were used for Blastocystis diagnosis and subtype analyses on routine clinical samples in a university hospital.
A high prevalence of Blastocystis infection was found with both advanced microscopy and sequence-confirmed PCR in our patient population. Most cases were caused by subtypes ST1, ST2, ST3 and ST4. A significantly higher prevalence was found among patients with a history of recent travel to tropical countries.
Permanent staining with Chlorazol Black was routinely performed on any SAF sample where further determination was needed, as is true for Dientamoeba fragilis and other vegetative stages of protozoa. Results of iodine wet smear and Chlorazol Black stain were compared on a selected series of positive and negative samples. Technicians performing microscopic diagnosis had extensive experience in parasitological diagnosis and were blinded for the results of PCR.
All non-fixed stools collected on the second day were mixed with Stool Transport and Recovery (S.T.A.R.) Buffer (Roche Applied Science) to an approximately 1:3 v:v suspension and vortexed and stored for DNA isolation and PCR. DNA was isolated from all faecal suspensions using the MagNA Pure LC Isolation instrument for nucleic acid extraction (Roche Applied Science). To monitor the process of extraction and amplification, a universal control consisting of phocine herpesvirus type 1 (PhHV-1) was added to the clinical specimens [36]. Briefly, 20 μl of faecal suspension was added to 500 μl lysis buffer with 2500 copies PhHV-1 and prelysed for 10 minutes at room temperature. After centrifugation, 500 μl of the lysate supernatant was used as input for the Total Nucleic Acid Isolation Kit (Roche Applied Science) as described by the manufacturer. Elution of nucleic acid was in a final volume of 50 μl. Blastocystis PCR was performed using primers and conditions described by Stensvold [33], aimed at a region of the small subunit rRNA gene of Blastocystis, using 15 μl of isolated DNA as input in a total volume of 50 μl. Fifteen microliters of PCR product were size-separated by agarose gel electrophoresis and visualized by UV illumination after ethidium bromide staining. Diluted amplicons were sequenced on both strands using amplification primers with BigDyeTerminator chemistry (Applied Biosystems) and analysed on an ABI 3900 sequencer. Subtypes following the nomenclature proposed by Stensvold et al [21] were determined by comparison of obtained sequences to Genbank reference sequences using CodonCode Aligner program (CodonCode Corporation), and MEGA 4.0 [37]. For sequences that seemed to contain a mixed signal of multiple subtypes, the predominant subtype was assigned. Only samples for which a Blastocystis sequence was obtained were considered PCR and sequencing positive. Samples for which no Blastocystis sequence was obtained or for which there was no PCR amplicon while the PhHV-1 control DNA was detected using real-time PCR [36], were considered negative. PCR and sequence interpretation were performed blinded to the microscopy results. Sequences were deposited in Genbank with accession numbers KF241991-KF242090.
Although microscopy on two samples proved very effective for diagnosis of Blastocystis, equal results with PCR diagnosis were obtained with examination of one sample only. In addition, determination of the subtype can be obtained, which is not possible by microscopy. We included sequence confirmation as a criterion for a positive PCR, to assure 100% specificity. Sequence-confirmed PCR was positive in 103 out of 107 (97%) positive patients. Discrepancies between sequence-confirmed PCR and microscopy in our study probably were related to very low numbers of Blastocystis shedded in the stools, resulting in insufficient amounts of PCR product to obtain a sequence.
The significantly higher prevalence of Blastocystis infection in patients presenting at the Department of Tropical Medicine as compared to other departments, suggests that travel to tropical countries can increase the risk of Blastocystis infection, as has been observed in earlier studies [7, 8, 53, 54]. High prevalence of Blastocystis has been reported in developing countries [1, 44, 55, 56]. Despite the higher prevalence, the Blastocystis subtype distribution among travelers and patients of other clinical departments was similar. This suggests that general poor hygiene and sanitation in many of the tropical countries rather than specific exposure to certain Blastocystis subtypes, causes the observed increase in prevalence. Unfortunately, no specific data about previous travel to tropical countries were available for the patients from departments other than the Department of Tropical Medicine.
In conclusion, fast and reliable diagnosis of Blastocystis infection in routine clinical parasitology is possible by advanced microscopy on multiple fixed stool samples with immediate fixation after production of stools. In terms of sensitivity, specificity and subtype determination, direct PCR on a single stool sample is also an excellent diagnostic tool. The prevalence of Blastocystis infection and the ongoing debate about pathogenicity warrant the use of good diagnostic methods in routine clinical laboratories. Other subjects of interest are patients with gastrointestinal complaints returning from the tropics without pathogenic microorganisms but only Blastocystis in stools. Further studies on pathogenicity, and especially the importance of subtypes of Blastocystis, are needed, as well as identification of an effective treatment. For many of these studies excellent laboratory diagnosis is a prerequisite.
The Zika virus outbreaks brought to light the lack of globally applicable guidelines for the diagnosis and management of GBS. Such guidelines are necessary because the diagnosis of GBS can be challenging owing to heterogeneity in clinical presentation, an extensive differential diagnosis, and the lack of highly sensitive and specific diagnostic tools or biomarkers. Guidance for the treatment and care of patients with GBS is also needed because disease progression can vary greatly between patients, which complicates an entirely prescriptive approach to management. In addition, treatment options are limited and costly, and many patients experience residual disability and complaints that can be difficult to manage.
Availability of globally applicable clinical guidelines for GBS is especially important as new outbreaks of pathogens that trigger GBS are likely to occur in the future. To generate this globally applicable clinical guideline for GBS, the ten most important steps in the management of GBS, covering diagnosis, treatment, monitoring, prognosis and long-term management, were identified by a group of international experts on GBS (Fig. 1). For each step, recommendations were provided on the basis of evidence from the literature and/or expert opinion, and consensus was sought for each recommendation to finalize the guideline. These recommendations are intended to assist providers in clinical decision-making; however, the use of the information in this article is voluntary. The authors assume no responsibility for any injury or damage to persons or property arising out of or related to any use of this information, or for any errors or omissions.
Following the outbreak of Zika virus and its association with an increase in the incidence of GBS, the European Union-funded Zika Preparedness Latin American Network (ZikaPLAN) was established22. Our new guideline was initially prepared by participants of the ZikaPLAN network, comprising experts on GBS from the Netherlands (S.E.L., M.R.M. and B.C.J.), Brazil (F.d.A.A.G. and M.E.D.) and the United Kingdom (H.J.W.). These members brought specific clinical and research expertise to the guideline from their leading roles in large international projects on GBS (such as the International GBS Outcome Study (IGOS)), along with direct experience in managing the large increases in GBS cases in Zika virus-affected regions of Latin America23. To develop the preliminary guidelines, a series of in-person meetings were held between lead authors on the writing committee (S.E.L., M.R.M., B.C.J. and H.J.W.), along with smaller individual meetings with colleagues in Latin America (S.E.L., F.d.A.A.G. and M.E.D.) and continuous e-mail correspondence to review drafts and receive input. On the basis of their expert opinion and through consensus, this group identified ten of the most important steps in the diagnosis and management of GBS.
In the absence of sufficiently sensitive and specific disease biomarkers, the diagnosis of GBS is based on clinical history and examination, and is supported by ancillary investigations such as CSF examination and electrodiagnostic studies. The two most commonly used sets of diagnostic criteria for GBS were developed by the National Institute of Neurological Disorders and Stroke (NINDS) in 1978 (revised in 1990)2,3 (Box 1) and the Brighton Collaboration in 2011 (ref4) (Supplementary Table 1). Both sets of criteria were designed to investigate the epidemiological association between GBS and vaccinations but have since been used in other clinical studies and trials. We consider the NINDS criteria to be more suited to the clinician as they present the clinical features of typical and atypical forms of GBS, although the criteria from the Brighton Collaboration are also important, widely used, and can help the clinician to classify cases with (typical) GBS or MFS according to diagnostic certainty. Various differential diagnoses must also be kept in mind when GBS is suspected, and some symptoms should raise suspicion of alternative diagnoses (Boxes 1 and 2). The role of ancillary investigations in confirming a GBS diagnosis is described in more detail in the following section. 041b061a72