LGCmain
Indian Journal of Pathology and Microbiology
Home About us Instructions Submission Subscribe Advertise Contact e-Alerts Ahead Of Print Login 
Users Online: 1687
Print this page  Email this page Bookmark this page Small font sizeDefault font sizeIncrease font size


 
  Table of Contents    
REVIEW ARTICLE  
Year : 2022  |  Volume : 65  |  Issue : 5  |  Page : 164-175
Non invasive tests for diagnosis of parasitic infections of CNS


1 Department of Microbiology, Kidwai Memorial Institute of Oncology, Bangalore, Karnataka, India
2 Department of Neuropathology, National Institute of Mental Health and Neurosciences, Bangalore, Karnataka, India

Click here for correspondence address and email

Date of Submission17-Nov-2021
Date of Decision31-Jan-2022
Date of Acceptance06-Feb-2022
Date of Web Publication11-May-2022
 

   Abstract 


Central nervous system (CNS) infections are among the most devastating diseases with high mortality and morbidity. In the pre-human immunodeficiency virus (HIV) era, the occurrence of CNS infections was very infrequent. However, in the past four decades or so, with a global increase in the immunocompromised population, the incidence of opportunistic infections of the CNS has changed. This includes a global increase in the incidence of parasitic infections such as Toxoplasma gondii. Infections such as neurocysticercosis and cerebral malaria are quite prevalent in developing countries. Early diagnosis of these infections is crucial for instituting accurate therapy and preventing mortality and morbidity. Despite advances in neuroimaging techniques, laboratory diagnosis remains the mainstay for confirmation of diagnosis. We present an update on the noninvasive tests available for laboratory diagnosis of parasitic infections of the CNS.

Keywords: Cerebral malaria, CNS infections, immunodiagnosis, molecular diagnosis, neurocysticercosis, opportunistic infections, toxoplasmosis

How to cite this article:
Jayshree R S, Mahadevan A. Non invasive tests for diagnosis of parasitic infections of CNS. Indian J Pathol Microbiol 2022;65, Suppl S1:164-75

How to cite this URL:
Jayshree R S, Mahadevan A. Non invasive tests for diagnosis of parasitic infections of CNS. Indian J Pathol Microbiol [serial online] 2022 [cited 2022 May 24];65, Suppl S1:164-75. Available from: https://www.ijpmonline.org/text.asp?2022/65/5/164/345046





   Introduction Top


Over the past few decades, there has been a steady increase in the number of immunocompromised people due to various factors, including the spread of human immunodeficiency virus (HIV) infection in the community, widespread use of myeloablative therapies for management of hematological malignancies, and immunosuppressive therapy for organ transplantations and control of autoimmune diseases. This has led to a global increase in the incidence of opportunistic infections (OI) of the central nervous system (CNS). In the HIV-infected population, OIs of the CNS cause significant morbidity and mortality. While approximately 40%–70% of patients develop symptomatic neurological disease sometime during the course of their illnesses, 10%–20% of them present with infections at the time of diagnosis.[1] In general, parasitic infections of the CNS are more common than believed and affect the mental, cognitive, and neurological health of millions of adults and children in the developing world. Although sporadic, CNS infections also have an impact in the developed world.[2] Prompt diagnosis and treatment of these infections are essential to prevent devastating sequelae and to reduce neurological sequela and mortality. Herein we review recent advances in the laboratory diagnosis of three common neurotropic parasitic infections, namely toxoplasmosis, neurocysticercosis, and malaria, which are important both in the Indian context and globally.[2],[3]


   CNS Toxoplasmosis Top


Epidemiology

Toxoplasma gondii (T. gondii) is a coccidian parasite, which in the pre-HAART era was one of the most common causes of focal brain lesions in AIDS causing cerebral toxoplasmosis. The incidence of this disease differs in various countries and closely corroborates with seroprevalence in the normal population. In an autopsy study on 113 patients with focal brain lesions published from NIMHANS, cerebral toxoplasmosis was the single most common etiologic agent (89%) in HIV-positive patients with focal brain lesions.[4] In a study from Mumbai, CNS toxoplasmosis constituted 20% of the AIDS-associated CNS infections at postmortem.[5] A clinico-radiological study from Pune found 66% of HIV-infected patients to have CNS toxoplasmosis.[6] While presently, the seroprevalence of IgG antibodies to T. gondii among the general population in the United States is low (only 11%), it is higher in other geographic areas across the globe.[7] In India, the prevalence of antibodies to T. gondii in the general population shows a wide variation (1%–57%).[8],[9],[10],[11],[12],[13],[14] These differences in the prevalence and mean titers of IgG antibodies to the parasite observed across the world can be ascribed to several factors, including differences in hygiene, environmental factors, socio-cultural habits, and routes of transmission. The incidence of primary cerebral toxoplasmosis occurring in immunocompetent people is very low (<0.02%); contrastingly, in HIV-infected patients, it is as high as 30%–40%.[1] Before the advent of combination antiretroviral therapy (cART), cerebral toxoplasmosis was seen to develop in about a quarter of the AIDS patients seropositive for T. gondii within about 2 years after the onset of AIDS.[15] In this patient population, trimethoprim-sulfamethoxazole was proven to be effective as a primary prophylactic agent for preventing Pneumocystis pneumonia and Toxoplasma encephalitis infections.[15] Consequently, primary prophylaxis with cotrimoxazole has been recommended for Toxoplasma IgG seropositive HIV-infected patients with CD4+ T-cell counts of <100/μL.[16] Also, administration of antiretroviral therapy has been shown to reduce the risk of developing cerebral toxoplasmosis by nearly 50%.[15]

Diagnosis of cerebral toxoplasmosis is challenging as neuroimaging features of ring-enhancing lesions are not specific for toxoplasmosis and can be seen in tuberculosis, abscesses, metastases, and less commonly in lymphoma, with implications on treatment. Early diagnosis is mandatory for preventing mortality by initiating early treatment. Based on the clinical syndrome and neuroimaging features, a case definition of cerebral toxoplasmosis has been proposed involving definitive, probable, and possible cerebral toxoplasmosis.

Definitive: When there is a histopathological demonstration of the protozoa in brain biopsy/autopsy specimens. Biopsies can be obtained by stereotactic computed tomography (CT)/MRI-guided biopsy, and T. gondii tachyzoites are demonstrated along the periphery of the lesions by immunohistochemistry using tachyzoite specific antibodies to T. gondii [Figure 1]. Although these tests yield good sensitivity (67%–87%), it is not without significant morbidity and mortality.[17],[18] Alternatively, detection of T. gondii DNA in cerebrospinal fluid (CSF) can be accomplished by PCR.
Figure 1: Biopsy from cerebral toxoplasmosis lesion shows characteristic central zones of coagulative necrosis with thrombosed vessels walled in by zone of chronic inflammation (a). Numerous ruptured tachyzoites are demonstrated along the wall by immunohistochemistry (b). Inset (b) shows an intact bradyzoite [a: H and E ×Obj. 20; b: Immunoperoxidase, p30 antigen of T. gondii ×Obj 20; b, inset: immunoperoxidase ×Obj40]

Click here to view


Probable: Definite radiological response to empirical anti-toxoplasma therapy comprising sulfadiazine and pyrimethamine for 10–14 days. Cases wherein invasive procedures are not advocated or when lumbar puncture is contraindicated or molecular tests are not available as in low resource centers would fall under this category.

Possible: Presence of anti-T. gondii antibodies (IgG) in the serum with no other alternative diagnosis. This group would also comprise cases that cannot be followed up: those who succumb early after hospitalization or refuse treatment and get discharged against medical advice.[15]

Laboratory Diagnosis of Toxoplasmosis

Various approaches are used to diagnose acute primary toxoplasmosis:

  1. Direct evidence comprises direct histological demonstration of the parasite [Figure 1], culturing the organism from clinical samples using cell lines, and detection of the parasite DNA using molecular tests.
  2. Indirect evidence includes serological diagnosis by demonstration of specific antibody isotypes toward parasitic antigens, particularly IgM or low-avidity IgG, both of which help in discriminating acute from chronic infections.[19] As against crude lysates of T. gondii tachyzoites, which are currently used for ELISA-based serodiagnosis of toxoplasmosis, a mixture of carefully selected tachyzoite/bradyzoite stage-specific recombinant antigens of the parasite appear promising in differentiating acute from chronic infections. Numerous T. gondii antigens have been cloned in Escherichia coli, such as surface antigens: SAG1 (P30), SAG2 (P22), SAG3 (P43), and P35; microneme antigens: MIC2, MIC3, MIC4, and MIC5; dense granule antigens: GRA1 (P24), GRA2 (P28), GRA4, GRA5, GRA6 (P32), and GRA7 (P29); rhoptry antigens: ROP1 (P66) and ROP2 (P54); and matrix antigens: MAG1.[20]


Laboratory diagnosis of cerebral toxoplasmosis

Serology

While serodiagnosis of acute primary T. gondii infection can be established using the tests detailed above, serological diagnosis of cerebral toxoplasmosis is often riddled with difficulties. ELISA using recombinant antigens listed above appear to be very promising for diagnosing congenital toxoplasmosis; however, its efficacy in diagnosing cerebral toxoplasmosis remains to be proven.[20] The majority of cases of cerebral toxoplasmosis are not acute acquired primary infections; they represent reactivation of latent infections in the brain and hence serum IgM antibodies have been reported only in a small percentage (0%–9%) of cases.[8],[21],[22],[23],[24],[25],[26] In our own experience, an autopsy-confirmed cohort of cerebral toxoplasmosis in the HIV-infected revealed IgM antibodies in 61% of the serum samples.[27] Considering that seroprevalence of toxoplasmosis among voluntary blood donors in that area was ~20% and the fact that IgM antibodies to the parasite have been recognized to persist for about 2 years following primary infection, these antibodies may not be indicative of recent T. gondii infection.[13],[28] Thus, anti-T. gondii IgM antibody test does not serve any diagnostic role in cerebral toxoplasmosis unless accompanied by low-avidity IgG antibodies to the parasite.[27] On the contrary, detection of high titers of high-avidity serum T. gondii IgG antibodies in HIV-positive individuals with clinical signs and imaging results suggestive of toxoplasmosis can be an adjunct to arrive at a diagnosis, especially where lumbar puncture is contraindicated and where molecular tests are not available as in low-resource centers.[7],[21],[27],[29],[30],[31],[32] Although the local presence of parasite-specific antibodies in the CSF indicates toxoplasmic encephalitis, the sensitivity of its detection is variable. Of the many factors that influence the appearance of antibodies in the CSF, the most crucial is the proximity of the lesions to the meninges and the duration since reactivation.[21] Additionally, intrathecal synthesis of IgG antibodies has been observed in 54.5%–70% of patients.[29],[30] Further, using affinity-mediated immunoblotting, the oligoclonal nature of the intrathecally synthesized antibodies was shown in 100% of the TE patients, and bands specific to 29kDa protein of T. gondii corresponding to tachyzoite antigen p30 were uniformly observed.[30] Detection of CSF anti-toxoplasma IgG antibodies in AIDS-associated cerebral toxoplasmosis has been reported to be 100% sensitive.[27],[33] Duration, severity of illness, and institution of anti-toxoplasma therapy may, however, modulate CSF IgG positivity and titers.

Molecular tests

There is substantial published literature addressing PCR-based diagnosis of toxoplasmosis.[15] However, despite the vast experience from many centers across the globe, consensus on the steps to be employed for the molecular diagnosis of this infection is lacking. Standard operating procedures regarding the ideal protocol for sample preparation, gene target to be selected, molecular technique to be employed, etc., are yet to be prescribed for the molecular diagnosis of this infection. Moreover, as standard commercial DNA-based diagnostics for T. gondii are lacking, scientists are forced to resort to in-house protocols and procedures for making a diagnosis.

There are several factors that can influence sensitivity of the test: suboptimal transport and storage of clinical samples before testing; nonavailability of standardized reagents and methods of extraction of nucleic acids from clinical samples, for example, simple boiling/inhouse extraction method using proteinase K and phenol chloroform/extraction using commercial kits; sample testing done singly/in replicates; establishing the limit of detection of an assay before testing clinical samples; sets of controls used; variation in laboratory conditions: highly proficient accredited laboratory/nonaccredited laboratory lacking proper facilities to contain laboratory contamination of PCR products; target gene selected: single copy gene, for example, gene coding for p30 antigen (SAG1), genes SAG2, SAG3, SAG4, and GRA4, gene coding for alpha tubulin, beta tubulin, multi copy or repetitive gene, for example, B1 gene: 35-fold repetitive, rRNA gene: 110-fold repetitive, 529bp coding gene: 200–300-fold repetitive—TGR1A, TGR1E, TGR2, TGR4, etc.; usage of varied primer sets for different target genes, for example, within the B1 gene itself, various primers have been designed toward different locations in the gene; conventional PCR/hemi-nested PCR/nested PCR/real time PCR/loop-mediated isothermal amplification (LAMP); time lapsed after initiation of anti-toxoplasma therapy; etc.[34],[35] Considering all these factors, it is not surprising to find a wide variation in the sensitivity of T. gondii detection in clinical samples, which also makes the comparison of results between various studies rather difficult. In-house nB1 PCR has been widely used in most diagnostic clinical microbiology laboratories. Although both nPCR and real-time qPCR have been reported to have similar performance characteristics, qPCR appears to be better as it is rapid, reproducible, and there is less chance of laboratory contamination.

Detecting parasitic DNAemia comes into use when lumbar puncture is contraindicated. HIV-associated cerebral toxoplasmosis is usually attributed to local reactivation of the latent brain infection and hence parasitemia is very rarely observed. This perhaps explains the poor sensitivity of methods detecting protozoal DNA in the blood even with tests amplifying highly repetitive targets.[26] A wide variation in the sensitivity of blood T. gondii PCR in HIV+ patients has been reported ranging from 2% to 87% with an average of 30%–50%.[15] The sensitivity of PCR for the detection of parasitic DNA in the CSF in cases of cerebral toxoplasmosis also ranges from 11.5%[15] to 100%[27],[30],[33] (mean: 50%–60%), with a specificity of 96%–100%, a positive predictive value of 100%, and negative predictive values of 71%–92%.[15] In addition to the factors listed above, the sensitivity of lumbar CSF PCR has been shown to be affected by the duration and severity of illness, the number and extent of brain lesions, the time since institution of anti-toxoplasma therapy, and the closeness of the lesions to the meninges.[15] The latter point has been substantiated by the higher sensitivity of ventricular versus lumbar CSF in B1 nPCR (100% vs. 76.5%) collected at autopsy.[27] Notwithstanding its modest sensitivity, considering its high specificity and positive predictive value (100%), B1 PCR should become a part of the panel of diagnostics for cerebral toxoplasmosis.[36] In other words, while the detection of T. gondii DNA in the CSF confirms the diagnosis of cerebral toxoplasmosis, a negative result does not rule it out, and accordingly, initiation or continuation of specific anti-toxoplasma treatment is recommended.[15] Specific antiprotozoal therapy greatly affects the sensitivity of PCR and hence the first week following treatment initiation is the ideal time to sample CSF for PCR, and as a corollary, therapeutic effectiveness can be monitored by CSF PCR.[15] REP529 being a highly repetitive gene appears to be an attractive target for diagnosing cerebral toxoplasmosis,[37] However, the sensitivity of amplification of this target in the blood of patients with cerebral toxoplasmosis was dismal (25%).[26] In contrast, real-time PCR amplification of this gene in the CSF samples from cases of cerebral toxoplasmosis yielded a sensitivity of 69% but a specificity and a positive predictive value of 100%.[38] In the 21st century, metagenomic next-generation sequencing (NGS) is radically transforming the diagnosis of infections in a noninvasive manner. In this technique, cell-free nucleic acids are sequenced and has the advantage of being a universal pathogen detector in a single run; this technique can also discover possible novel pathogens in clinical specimens. The entire process is a target-independent assay without a priori knowledge of the possible causative agents. A recent report describes diagnosing an atypical case of TE by NGS using the CSF.[39] While the specificity of the assay is very high, its sensitivity on clinical samples for diagnosing cerebral toxoplasmosis is yet to be evaluated. A panel of noninvasive tests, namely B1 nPCR and T. gondii IgG serology both on the blood and CSF (whenever lumbar puncture is not contraindicated), are useful diagnostic adjuncts to imaging, especially in the HIV-infected population with a clinical suspicion of cerebral toxoplasmosis.


   Neurocysticercosis Top


CNS infection by the larval stage (Cysticercus cellulosae) of the cestode Taenia solium (T. solium) results in neurocysticercosis (NCC). It is a zoonotic infection wherein humans accidentally become the intermediate host of the parasite. Infection occurs by the feco-oral route by ingesting eggs of the helminth in contaminated water or food. Thus, vegetarians too can acquire NCC. The activated oncospheres liberated in the lumen of the human gut penetrate the mucosa and migrate to various organs, including the CNS. The brain parenchyma is the most common site where the larvae (cysticerci) lodge and remain encysted but they can also be found extra-parenchymally in the ventricles or subarachnoid spaces. Indian patients with NCC are often young with solitary cysticercal granuloma.[40],[41],[42],[43],[44],[45] While parenchymal cysts are less aggressive, manifesting as headache and seizures, extraparenchymal cysts can lead to raised intracranial pressure due to blockage of CSF circulation, thereby causing mortality and morbidity.[46] Parenchymal cysts are initially quiescent, inducing little or no inflammation. Later, as the host immune response is stimulated, resultant inflammation induces cyst degeneration and resolution with calcification of the cyst.[47] It is this degeneration or calcification of the encysted parenchymal parasites that gives rise to symptoms of the disease, with seizures being the most frequent manifestation.[43],[44] In the developing world, NCC is considered the most common preventable etiology of acquired epilepsy.[48] Estimates of the World Health Organization (WHO) indicate an annual occurrence of 2.5–8.3 million cases of NCC worldwide, which may be an underestimate.[49] The WHO has labeled cysticercosis as endemic in South-East Asia, Mexico, and sub-Saharan Africa, and its prevalence represents the economic and social development in a community.[50] However, an increase in the incidence in nonendemic regions has also been observed, perhaps attributable to international travel and globalization.[51] An autopsy-based study has reported NCC as the most common infective cyst in India, occurring in 7.8% of the cases (42/538).[52] A systematic meta-analysis found 29% of epilepsy cases to be attributable to NCC, while 78% of NCC present with seizures. In India, NCC manifesting as epilepsy has been projected to be 1 per 1000 population.[53] On the same lines, about 25% of patients with epilepsy harbored antibodies to T. solium.[50]

Although the etiology is known, considering the nonspecific nature of clinical signs and symptoms, the diagnosis of NCC is not straightforward. Definitive evidence requires histological diagnosis, which involves invasive procedures added to which are inaccessible locations in the CNS. Classification of NCC was first described in 1996 and has undergone a series of periodic revisions. Current recommendations comprise clinical, neuroimaging, immunological, and epidemiological criteria, and depending on the degree of evidence, cases are classified into three categories: definitive, probable, and possible.[46]

Immunodiagnosis

Antibody detection

Presently, none of the immunological tests is a stand-alone tool for the diagnosis of NCC. Nevertheless, they are useful adjuncts to neuroimaging. A Western Blot assay called enzyme-linked immunoelectrotransfer blot assay (EITB) uses lentil lectin purified parasite glycoprotein antigens (LLGP) for serological diagnosis of cysticercosis. Although considered as a benchmark for diagnosing cysticercosis, the assay is very demanding in terms of availability of antigens and expertise, which may not be possible in rural areas. The strip contains seven parasite-specific antigenic bands that belong to three major protein families, namely GP50, T24/42, and 8kDa.[51] Various banding patterns observed in the patient's sera indicate the possibility of severe infection and the viability of the encysted parasite. Immunoreactivity to all families of proteins correlated with severe extraparenchymal and intraparenchymal infections. The probability of a positive EITB assay was higher in cases with perilesional edema.[54] Antibodies to GP50 are the first to appear and the last to disappear; subsequently, antibodies to the other two protein families emerge. The performance characteristics of this assay varies depending on the form of NCC. It works best in extraparenchymal NCC and in cases with more than one parenchymal lesion, demonstrating a sensitivity and specificity of 98% and 100%, respectively. EITB can detect antibodies in the serum by about 5 weeks after infection, irrespective of the general immune status of the patient and is useful for monitoring therapy. The sensitivity using CSF is similar to that of serum. However, in cases with a single cyst or with calcified cysts, its sensitivity was seen to be only 50%.[48] An alternative immunoblot assay has been developed using recombinant and synthetic peptides antigens with rGP50, rT24, and 8kDa proteins, which performed well in comparison with the conventional EITB assay in terms of sensitivity and specificity, circumventing the need for handling live cysticerci and simultaneously, the quality and supply of antigens can be maintained.[47] In contrast, the performance characteristics of ELISA and dot blot assays using crude lysates of cysticerci as antigen were inferior: their sensitivity/specificity was 89%/81% and 89%/73%, respectively. However, in cases with multiple lesions, a specificity of 100% was observed by ELISA. Other commercial ELISA kits fared badly, demonstrating very poor performance characteristics: the sensitivity for detecting antibodies in cases with viable parasites was low, varying from 22% to 44%. Also, these antibodies were nonspecific—cross-reacting with sera of patients with hydatid disease, other helminthic infections, and extra-neural cysticercosis. Adding to the challenge was the observation that in neurocysticercosis, the antibodies continued to remain positive for a year post anti-helminthic therapy.[47] A recent study described the performance of another three new recombinant antigens of the parasite, two of which, namely TsF78 and TsP43, corresponding to filamina and peroxidase, respectively, appeared promising. The antigens were reported to have high sensitivities (~94% and 92%) in ELISA and did not cross-react with sera from other helminthic infections (specificities: ~95% and 93%), making them good diagnostic candidates suitable for commercialization.[55] Circulating antibodies to excretory/secretory antigens were detected with moderate sensitivity in NCC, raising the possibility of using serum samples for diagnosing this infection.[56]

Antigen detection: Antigen detection assays act as a complementary tool in the diagnostic armamentarium for cysticercosis. Antibodies have been used for capturing cestodic antigens both in the CSF and serum. Other biological samples such as urine, tears, and saliva have also been tested. The sensitivity of antigen detection in the CSF is ~90%. Although the sensitivity of these tests is lower than EITB, when positive, it has the advantage of indicating the presence of viable cysticerci, being negative in calcified disease and hence is useful for monitoring antiparasitic therapy. Crude antigenic extracts used to raise antibodies have shown poor sensitivity and specificity in antigen detection assays. Detection of excretory/secretory antigens such as the 32kDa, 43kDa, and 67kDa peptides in the serum and CSF are better targets than somatic antigens of the vesicular stage of the cestode. Monoclonal antibody-based sandwich ELISA is another step forward toward improving antigen detection.[50] A novel approach from a Brazilian laboratory has been to use fractionated and purified antibodies (IgY) made in hens immunized with a hydrophobic fraction of Taenia crassiceps metacestodes for antigen capture in sandwich ELISA. The test was found to be ~ 94% specific and 93% sensitive in detecting circulating immune complexes in NCC.[57] Considering the simplicity of performing an ELISA, the test has the potential to become a noninvasive tool for diagnosing NCC even in smaller and resource-limited centers.

Molecular tests

PCR-based amplification of the highly repetitive region Tsol19 of T. solium DNA in the CSF of patients with extraparenchymal lesions has been higher than in cases of intraparenchymal NCC (90.9% vs. 42.9%).[58] On the contrary, using the same set of primers, CSF PCR detected 100% of the intraparenchymal NCC.[59] Larger studies with a higher statistical power are required to understand this discrepancy. Amplification of Tsol19 DNA has also been documented in the blood and urine of both extra and intraparenchymal lesions of NCC although at moderate sensitivity (57%/60% and 60%/88%, respectively).[60] A recent study targeting another highly repetitive DNA sequence Tsol13 documented a good performance characteristic for diagnosing and follow-up of subarachnoid and ventricular NCC by qPCR on CSF.[61] The overall sensitivity and specificity of blood LAMP directed toward the cox 1 gene of T. solium in NCC patients was 74% (72% in intraparenchymal and 87% in extraparenchymal) and 90%, respectively. Thus, LAMP on blood is a simple test that can serve to be a useful diagnostic tool for NCC diagnosis in resource-poor settings.[62] A proof of principle study detected cell-free parasitic DNA in the CSF by NGS wherein genome detection was found to be more sensitive than detection of parasite-specific antibodies and has been used for prognostication and also correlated well with symptomatic relief.[63],[64] Moreover, metagenomic sequencing has also been shown to detect unsuspecting cases of NCC. However, these pilot results need to be verified and their performance characteristics determined in larger studies including different forms of NCC, location, and number of lesions, particularly intraparenchymal NCC.[51] Another attractive approach is proteomic profiling of sera and CSF from NCC cases, which may give useful leads for the diagnosis as well.


   Cerebral Malaria Top


The etiological agent of malaria is a protozoan parasite of the genus Plasmodium. Malaria even today remains a fatal disease and impacts health worldwide. Of the 229 million cases of malaria and 409,000 deaths reported in 2019 globally,[65] ~90% were caused by Plasmodium falciparum (P. falciparum). It is endemic to many countries; globally, more than half of the population residing in tropical and subtropical regions is at risk of infection.[65] Five species of the organism, namely vivax, falciparum, malariae, ovale, and knowlesii are known to infect humans. Cerebral Malaria occurs in ~2% of the patients infected with P. falciparum. It is the most severe neurological complication of P. falciparum parasitemia and is associated with high morbidity and mortality. While optimal treatment decreases mortality by 15%–25%, about a third of the treated are not without neurological deficits. Although rare, P. vivax and P. knowlesii infections can also cause cerebral malaria.[66]

Laboratory diagnosis

Conventional method

Prompt diagnosis of cerebral malaria is crucial for prompt institution of antimalarial treatment and thereby prevention of mortality. The gold standard diagnostic test is the detection of asexual forms of P. falciparum in the Giemsa-stained peripheral blood smears (thin and thick). While a veteran microscopist can detect about 5 parasites/μL, an average one can pick up 30–50 parasites/μL in a thick smear. Thus, in cases having lower parasitemia, alternative sensitive modes of diagnosis are needed. Also, considering that P. falciparum erythrocytic schizogony occurs within the postcapillary venules of internal organs, the highly pathogenic mature parasitized erythrocytes are sequestered there. Thus, a low peripheral parasitemia in a patient with severe disease may not be a good prognostic sign. Due to the sequestration of parasites in the internal vessels, a 100-fold difference in the total parasite biomass can be seen in early malaria versus late disease at an identical peripheral percentage parasitemia.[65],[67] Nevertheless, peripheral parasitemia of >10% is one of the diagnostic criteria of severe infection with a poor prognosis.[68]

Quantitative Buffy Coat (QBC) test

This test adds some objectivity to smear examination and simplifies detection. Briefly, about 55–65-μL sample of blood is centrifuged in a micro-hematocrit tube having a fluorescent dye, acridine orange, following which the tube is viewed under a fluorescent microscope with epi-illumination. All the DNA in the blood sample including leukocytic and parasitic DNA exhibit yellow fluorescence. While QBC is a simple and objective tool, it cannot speciate or quantify plasmodia; in addition, fluorescent staining of leucocytes makes detection of plasmodia difficult.

Rapid diagnostic tests

There are about 86 rapid diagnostic tests (RDTs) available commercially; these are immunochromatographic assays that are designed to capture various circulating malaria antigens on a membrane containing specific antimalaria antibodies. The popular ones are those targeting detection of P. falciparum-specific antigens: histidine-rich protein 2 (HRP2)/lactate dehydrogenase (LDH)/aldolase. Other tests are directed against the detection of genus-specific malaria antigens pLDH, P. vivax and are capable of diagnosing non-falciparum infections in mixed infections. HRP2 is a falciparum-specific protein produced by the asexual and sexual forms and is expressed on the surface of infected erythrocytes. It is liberated into the blood upon maturation of the schizont and serves as a good diagnostic and prognostic marker as it represents parasite burden. The limit of detection of this test is >40 parasites/μL of blood, with a sensitivity of >90% for falciparum malaria. The specificity of the test, however, is low (~40%). This may be due to delayed clearance of plasma HRP2; thus, another confirmatory test is needed to verify the findings especially in endemic regions. Alternatively, false-positive results observed in RDT may be because the test may be more accurate in detecting submicroscopic parasitemia. Thus, with molecular techniques being routinely used for diagnosis, an alternative “gold standard” other than estimating parasitemia based on peripheral blood microscopy may be needed. Histidine-rich protein gene deletion mutants of P. falciparum, specifically of pfhrp2 and pfhrp3 genes, have been reported from many countries such as South America, Africa, and India.[69] In view of this discovery, the WHO has recommended the use of alternative targets such as LDH/aldolase in RDT assays and microscopy for diagnosing malaria in such regions.[69] Besides, tests such as real-time quantitative PCR targeting conserved regions of the parasite DNA can detect infection with PfHRP2 gene deletion mutants. Another crucial cause for false-negative RDT is high parasitemia (parasite density >1000/μL), which gives rise to the prozone effect. This is a well-known phenomenon in immunological reactions; in RDT, it occurs when there is a high level of antigen circulating in the blood. Testing samples in dilution may help overcome such false-negative results.[70] In a high malaria transmission region in Tanzania, sandwich ELISA-based quantification of plasma concentrations of PfHRP2 (a measure of total parasite biomass) was proven to be an exceptional biomarker and of prognostic value in severe malaria.[71] Increased CSF levels of PfHRP2 and lowered ratios of plasma/CSF PfHRP2 indicated poor prognosis due to egress of the antigen across a compromised blood–brain barrier.[72] Polymorphisms of the PfHRP2 gene across various geographical regions also need to be kept in mind while designing antibodies to the protein. Parasitic LDH and aldolase are intracellular products produced by both sexual and asexual stages; they are enzymes belonging to the glycolytic pathway and are useful diagnostic biomarkers. While the latter is a pan malaria antigen (PMA), the former had a limit of detection of >100 parasites/μL of blood and a sensitivity of >95%.[69] It is recommended that RDT results be confirmed by microscopy, both to rule out false negativity due to low parasitemia and in RDT-positive cases to verify the species and to check the parasite burden.

Serological tests to detect intrathecal synthesis of malaria-specific antibodies are not useful for the diagnosis of cerebral malaria. In blood samples too, malarial serology does not serve any diagnostic purpose as antibodies to the parasite persist for years after exposure, especially in patients residing in malaria-endemic regions.

Molecular diagnosis: Considering the emergence of gene deletion mutants of the parasite, molecular tools have been developed particularly for bedside diagnosis of malaria. PCR-based detection of the nucleic acid of P. falciparum in the blood samples can help arrive at a diagnosis of cerebral malaria in a patient with clinical signs and symptoms suspicious of malaria. Loop-mediated isothermal amplification of the conserved gene of 18S ribosomal RNA of P. falciparum and nucleic acid amplification technique (NAT) on saponin lysed erythrocytes are simple, low-cost automated tests that offer both high sensitivity and specificity. The advantage is that submicroscopic parasitemia can be detected with ten-fold higher sensitivity compared to microscopy. An added advantage is that detection of end products in LAMP can even be simplified to naked eye visualization of color change in visible light.[73] Similarly, recently, Taqman-based detection of Plasmodia-specific DNA in CSF samples has been reported in a large surveillance study on children below 5 years with suspected meningitis from West Africa.[74] Also, a high mortality was observed in HIV-positive children in whom Plasmodium were detected in the CSF. Alternative targets for the LAMP assay are mitochondrial DNA and apicoplast genome.[73] Further, LAMP assays blended with electronic detection—lab-on-chip platform is an attractive and economical step further providing diagnosis at the point of care in low resource countries.[75] Also, other methods for the detection of plasmodia using isothermal amplification such as nucleic acid sequence-based amplification (NASBA), recombinase polymerase amplification (RPA), and thermophilic helicase dependent amplification (tHDA) have been published.[73] Thus, although molecular tests are technically demanding, added advantages are higher sensitivity, quantification of parasite burden, speciation, and detection of drug-resistant mutants.

Hemozoin or malarial pigment, a metabolic byproduct of cellular hemoglobin in plasmodial infected red blood corpuscles is an attractive tool for diagnosing malaria. Various techniques have been developed with a merger of biophysics, biochemistry, and engineering fields for the detection of these biocrystals in finger-prick blood samples by using handheld devices. The methodologies capitalize on the peculiar paramagnetic and optical properties of hemozoin crystals, which are different from hemoglobin.[73] Flow cytometry and laser desorption mass spectrometry have been shown to be useful for detecting hemozoin even within leukocytes.[76] Hemozoin has been shown to have photoacoustic properties as well, which can be exploited for economical diagnosis of malaria.[77] Also stated is the use of lasers for transdermal detection of hemozoin-induced nanobubble vapor.[73] In malaria-endemic areas, it is not uncommon to find incidental peripheral smear positivity, and clinical presentations are often common to many infectious conditions, with severe malaria being one of the etiologies. Malarial parasites do not synthesize miRNA, but the pathogen's mRNA can bind the host miRNA, influencing its action and can thus regulate the function of various organs. Hence, differential miRNA expression profile in combination with P. falciparum parasitemia can become a critical tool for the differential diagnosis of severe malaria, for example, cerebral malaria in patients presenting to the hospital with a clinical picture depicting severe infection. Also, an exaggerated host immune response represented by heightened levels of several biomarkers such as interferon-gamma inducible protein 10, tumor necrosis factor-α, C-reactive protein, procalcitonin, angiopoetin2, and intercellular adhesion molecule-1 has been associated with mortality in cerebral malaria.[76]

All the tests reviewed above for diagnosis of CNS parasitic infections of importance are summarized in [Table 1].
Table 1: Overview of list of tests for diagnosis of CNS protozoal infections

Click here to view



   Conclusion Top


CNS infections, particularly in the immunocompromised, are on the rise due to advances in immunomodulatory therapies. Clinical diagnosis of infections in the immunocompromised state is a challenge as suppression of immune responses alters clinical presentation and neuroimaging features, delaying accurate diagnosis. In addition, laboratory diagnosis is a challenge as immunological tests often have very low sensitivity and specificity. Interpretation of molecular tests, despite its exquisite sensitivity, is fraught with difficulties, especially in the context of immunocompromised host, to differentiate latent from active infections. As diagnosis is delayed, treatment is also not instituted in time, leading to increased morbidity and mortality. The role of the laboratory in patient management is emerging as a crucial one for providing precise, diagnosis prompt institution of antimicrobial treatment and thereby preventing mortality and morbidity. In this update, we reviewed the existing tests and provide a ready reference of the utility, advantages, and disadvantages of tests available in three parasitic infections prevalent in our country: cerebral toxoplasmosis, NCC, and cerebral malaria. With advances in technology, the ultimate goal of finding the ideal test for early diagnosis and successful treatment of these infections may soon become a reality.

Financial support and sponsorship

Nil.

Conflicts of interest

There are no conflicts of interest.



 
   References Top

1.
Dzikowiec M, Góralska K, Błaszkowska J. Neuroinvasions caused by parasites. Ann Parasitol 2017;63:243–53.  Back to cited text no. 1
    
2.
Garcia HH, Nath A, Del Brutto OH. Parasitic infections of the nervous system. Semin Neurol 2019;39:358-68.  Back to cited text no. 2
    
3.
Kumar D, Pannu AK, Dhibar DP, Singh R, Kumari S. The epidemiology and clinical spectrum of infections of the central nervous system in adults in north India. Trop Doct 2021;51:48-57.  Back to cited text no. 3
    
4.
Shankar SK, Mahadevan A, Satishchandra P, Kumar RU, Yasha TC, Santosh V, et al. Neuropathology of HIV/AIDS with an overview of the Indian scene. Indian J Med Res 2005;121:468-88.  Back to cited text no. 4
    
5.
Lanjewar DN, Surve KV, Maheshwari MB, Shenoy BP, Hira SK. Toxoplasmosis of the central nervous system in the acquired immunodeficiency syndrome. Indian J Pathol Microbiol 1998;41:147-51.  Back to cited text no. 5
[PUBMED]  [Full text]  
6.
Wadia RS, Pujari SN, Kothari S, Udhar M, Kulkarni S, Bhagat S, et al. Neurological manifestations of HIV disease. J Assoc Physicians India 2001;49:343-8.  Back to cited text no. 6
    
7.
Marra CM. Central nervous system infection with Toxoplasma gondii. Handb Clin Neurol 2018;152:117-22.  Back to cited text no. 7
    
8.
Meisheri YV, Mehta S, Patel U. A prospective study of seroprevalence of toxoplasmosis in general population and in HIV/AIDS patients in Bombay, India. J Postgrad Med 1997;43:93-7.  Back to cited text no. 8
[PUBMED]  [Full text]  
9.
Mittal V, Bhatia R, Sehgal S. Prevalence of toxoplasma antibodies among women with BOH and general population in Delhi. J Commun Dis 1990;22:223-6.  Back to cited text no. 9
    
10.
Singh S, Nautiyal BL. Seroprevalence of toxoplasmosis in Kumaon region of India. Indian J Med Res 1991;93:247-9.  Back to cited text no. 10
    
11.
Joshi YR, Vyas S, Joshi KR. Seroprevalence of toxoplasmosis in Jodhpur, India. J Commun Dis 1998;30:32-7.  Back to cited text no. 11
    
12.
Mohan B, Dubey ML, Malla N, Kumar R. Seroepidemiological study of toxoplasmosis in different sections of population of Union Territory of Chandigarh. J Commun Dis 2002;34:15-22.  Back to cited text no. 12
    
13.
Sundar P, Mahadevan A, Jayshree RS, Subbakrishna DK, Shankar SK. Toxoplasma seroprevalence in healthy voluntary blood donors from urban Karnataka. Indian J Med Res 2007;126:50-5.  Back to cited text no. 13
[PUBMED]  [Full text]  
14.
Stephen S, Pradeep J, Anitharaj V, Janarthanam V. Seroprevalence of toxoplasmosis in voluntary blood donors of Puducherry and surrounding districts of Tamil Nadu. J Parasit Dis 2017;41:1158-61.  Back to cited text no. 14
    
15.
Vidal JE. HIV-related cerebral toxoplasmosis revisited: Current concepts and controversies of an old disease. J Int Assoc Provid AIDS Care 2019;18:2325958219867315.  Back to cited text no. 15
    
16.
Panel on Opportunistic Infections in HIV-Infected Adults and Adolescents. Guidelines for the prevention and treatment of opportunistic infections in HIV-infected adults and adolescents: Recommendations from the Centers for Disease Control and Prevention, the National Institutes of Health, and the HIV Medicine Association of the Infectious Diseases Society of America. Available from: https://clinicalinfo.hiv.gov/sites/default/files/guidelines/documents/Adult_OI.pdf. [Last accessed on 2021 Oct 09].  Back to cited text no. 16
    
17.
Shyam babu C, Satishchandra P, Mahadevan A, Pillai Shibu V, Ravishankar S, Sidappa N, et al. Usefulness of stereotactic biopsy and neuroimaging in management of HIV-1 Clade C associated focal brain lesions with special focus on cerebral toxoplasmosis. Clin Neurol Neurosurg 2013;115:995-1002.  Back to cited text no. 17
    
18.
Antinori A, Ammassari A, Luzzati R, Castagna A, Maserati R, Rizzardini G, et al. Role of brain biopsy in the management of focal brain lesions in HIV-infected patients. Gruppo Italiano Cooperativo AIDS & Tumori. Neurology 2000;54:993-7.  Back to cited text no. 18
    
19.
Zeehaida M, Khalid H. Effective diagnostic marker for serodiagnosis of toxoplasma gondii infection: New developments and perspectives. In: Akyar I, editor Toxoplasmosis. Chapter 7. ISBN: 978-953-51-3270-7. London, UK: IntechOpen Limited; 2017. p. 105-17.  Back to cited text no. 19
    
20.
Ybañez RH, Ybañez AP, Nishikawa Y. Review on the current trends of toxoplasmosis serodiagnosis in humans. Front Cell Infect Microbiol 2020;10:204.  Back to cited text no. 20
    
21.
Luft BJ, Remington JS. Toxoplasmic encephalitis in AIDS. Clin Infect Dis 1992;15:211–22.  Back to cited text no. 21
    
22.
del Rio-Chiriboga C, Orzechowski-Rallo A, Sanchez-Mejorada G. Toxoplasmosis of the central nervous system in patients with AIDS in Mexico. Arch Med Res 1997;28:527-30.  Back to cited text no. 22
    
23.
Singh S, Dubey JP. Toxoplasmosis in India. Ghaziabad, New Delhi: Pragati Publishing Co.; 1999.  Back to cited text no. 23
    
24.
Chandramukhi A. Diagnosis of neurotoxoplasmosis by antibody detection in cerebrospinal (CSF) fluid using Latex Agglutination Test and ELISA. J Commun Dis 2004;36:153-8.  Back to cited text no. 24
    
25.
Prandota J, Gryglas A, Fuglewicz A, Zesławska-Faleńczyk A, Ujma-Czapska B, Szenborn L, et al. Recurrent headaches may be caused by cerebral toxoplasmosis. World J Clin Pediatr 2014;3:59-68.  Back to cited text no. 25
    
26.
Ajzenberg D, Lamaury I, Demar M, Vautrin C, Cabié A, Simon S, et al. Performance testing of PCR assay in blood samples for the diagnosis of toxoplasmic encephalitis in AIDS patients from the French departments of America and genetic diversity of toxoplasma gondii: A prospective and multicentric study. PLoS Negl Trop Dis 2016;10:e0004790.  Back to cited text no. 26
    
27.
Adurthi S, Mahadevan A, Bantwal R, Satishchandra P, Ramprasad S, Sridhar H, et al. Utility of molecular and serodiagnostic tools in cerebral toxoplasmosis with and without tuberculous meningitis in AIDS patients: A study from South India. Ann Indian Acad Neurol 2010;13:263-70.  Back to cited text no. 27
[PUBMED]  [Full text]  
28.
Paul M. Immunoglobulin G avidity in diagnosis of toxoplasmic lymphadenopathy and ocular toxoplasmosis. Clin Diagn Lab Immunol 1999;6:514-8.  Back to cited text no. 28
    
29.
Potasman I, Resnick L, Luft BJ, Remington JS. Intrathecal production of antibodies against Toxoplasma gondii in patients with Toxoplasmic encephalitis and the acquired immunodeficiency syndrome (AIDS). Ann Intern Med 1988;108:49-51.  Back to cited text no. 29
    
30.
Contini C, Fainardi E, Cultrera R, Canipari R, Peyron F, Delia S, et al. Advanced laboratory techniques for diagnosing Toxoplasma gondii encephalitis in AIDS patients: Significance of intrathecal production and comparison with PCR and ECL-western blotting. J Neuroimmunol 1998;92:29-37.  Back to cited text no. 30
    
31.
Colombo FA, Vidal JE, Oliveira de AC, Hernandez AV, Bonasser-Filho F, Nogueira RS, et al. Diagnosis of cerebral toxoplasmosis in AIDS patients in Brazil: Importance of molecular and immunological methods using peripheral blood samples. J Clin Microbiol 2005;43:5044-7.  Back to cited text no. 31
    
32.
Adurthi S, Mahadevan A, Bantwal R, Satishchandra P, Ramprasad S, Sridhar H, et al. Diagnosis of cerebral toxoplasmosis. Ann Indian Acad Neurol 2011;14:145-6.  Back to cited text no. 32
[PUBMED]  [Full text]  
33.
Vidal JE, Colombo FA, Oliveira de AC, Focaccia R, Chioccola VL. PCR assay using cerebrospinal fluid for the diagnosis of cerebral toxoplasmosis in Brazilian AIDS patients. J Clin Microbiol 2004;42:4765-8.  Back to cited text no. 33
    
34.
Bastien P. Molecular diagnosis of toxoplasmosis. Trans R Soc Trop Med Hyg 2002;96 (Suppl 1):S205-15.  Back to cited text no. 34
    
35.
Contini C. Clinical and diagnostic management of toxoplasmosis in the immunocompromised patient. Parasitologia 2008;50:45-50.  Back to cited text no. 35
    
36.
Cingolani A, De Luca A, Ammassari A, Murri R, Linzalone A, Grillo R, et al. PCR detection of Toxoplasma gondii DNA in CSF for the differential diagnosis of AIDS-related focal brain lesions. J Med Microbiol 1996;45:472-6.  Back to cited text no. 36
    
37.
Mesquita RT, Ziegler ÂP, Hiramoto RM, Vidal JE, Pereira-Chioccola VL. Real-time quantitative PCR in cerebral toxoplasmosis diagnosis of Brazilian human immunodeficiency virus-infected patients. J Med Microbiol 2010;59:641-7.  Back to cited text no. 37
    
38.
Nogui FL, Mattas S, Turcato Júnior G, Lewi DS. Neurotoxoplasmosis diagnosis for HIV-1 patients by real-time PCR of cerebrospinal fluid. Braz J Infect Dis 2009;13:18-23.  Back to cited text no. 38
    
39.
Hu Z, Weng X, Xu C, Lin Y, Cheng C, Wei H, et al. Metagenomic next-generation sequencing as a diagnostic tool for toxoplasmic encephalitis. Ann Clin Microbiol Antimicrob 2018;17:45.  Back to cited text no. 39
    
40.
Chandy MJ, Rajshekhar V, Prakash S, Ghosh S, Joseph T, Abraham J, et al. Cysticercosis causing single, small CT lesions in Indian patients with seizures. Lancet 1989;1:390–1.  Back to cited text no. 40
    
41.
Rajshekhar V. Etiology and management of single small CT lesions in patients with seizures: understanding a controversy. Acta Neurol Scand 1991;84:465–70.  Back to cited text no. 41
    
42.
Rajshekhar V, Chandy MJ. Validation of diagnostic criteria for solitary cerebral cysticercus granuloma in patients presenting with seizures. Acta Neurol Scand 1997;96:76–81.  Back to cited text no. 42
    
43.
Rajshekhar V, Raghava MV, Prabhakaran V, Oommen A, Muliyil J. Active epilepsy as an index of burden of neurocysticercosis in Vellore district, India. Neurology 2006;67:2135–9.  Back to cited text no. 43
    
44.
Singh G, Rajshekhar V, Murthy JM, Prabhakar S, Modi M, Khandelwal N, et al. A diagnostic and therapeutic scheme for a solitary cysticercus granuloma. Neurology 2010;75:2236–45.  Back to cited text no. 44
    
45.
Del Brutto OH, Nash TE, White AC Jr, Rajshekhar V, Wilkins PP, Singh G, et al. Revised diagnostic criteria for neurocysticercosis. J Neurol Sci 2017;372:202–10.  Back to cited text no. 45
    
46.
Guzman C, Garcia HH; Cysticercosis working group in Peru. Current diagnostic criteria for neurocysticercosis. Res Rep Trop Med 2021;12:197-203.  Back to cited text no. 46
    
47.
Garcia HH, Gonzalez AE, Gilman RH. Taenia solium cysticercosis and its impact in neurological disease. Clin Microbiol Rev 2020;33:e00085-19.  Back to cited text no. 47
    
48.
Sankhyan N, Kadwa RA, Kamate M, Kannan L, Kumar A, Passi GR, et al. For association of child neurology delphi group for neurocysticerosis in childhood. Management of neurocysticercosis in children: Association of child neurology consensus guidelines. Indian Pediatr 2021;58:871-80.  Back to cited text no. 48
    
49.
Butala C, Brook TM, Majekodunmi AO, Welburn SC. Neurocysticercosis: Current perspectives on diagnosis and management. Front Vet Sci 2021;8:615703.  Back to cited text no. 49
    
50.
Singhi P, Saini AG. Pediatric neurocysticercosis. Indian J Pediatr 2019;86:76-82.  Back to cited text no. 50
    
51.
Bustos J, Gonzales I, Saavedra H, Handali S, Garcia HH; Cysticercosis Working Group in Peru. Neurocysticercosis. A frequent cause of seizures, epilepsy, and other neurological morbidity in most of the world. J Neurol Sci 2021;427:117527.  Back to cited text no. 51
    
52.
Bhatt AS, Mhatre R, Nadeesh BN, Mahadevan A, Yasha TC, Santosh V. Nonneoplastic cystic lesions of the central nervous system-histomorphological spectrum: A study of 538 cases. J Neurosci Rural Pract 2019;10:494-501.  Back to cited text no. 52
    
53.
Rajshekhar V. Neurocysticercosis: Diagnostic problems & current therapeutic strategies. Indian J Med Res 2016;144:319-26.  Back to cited text no. 53
[PUBMED]  [Full text]  
54.
Vasudevan P, Moorthy RK, Rebekah G, Jackson E, Pamela BE, Thamizhmaran S, et al. Imaging correlates of serum enzyme-linked immunoelectrotransfer blot (EITB) positivity in patients with parenchymal neurocysticercosis: Results from 521 patients. Trans R Soc Trop Med Hyg 2022;116:117-23.  Back to cited text no. 54
    
55.
Morillo M, Noguera C, Gallego L, Fernández Z, Mata M, Khattar S, et al. Characterization and evaluation of three new recombinant antigens of Taenia solium for the immunodiagnosis of cysticercosis. Mol Biochem Parasitol 2020;240:111321.  Back to cited text no. 55
    
56.
Atluri SR, Singhi P, Khandelwal N, Malla N. Neurocysticercosis immunodiagnosis using Taenia solium cysticerci crude soluble extract, excretory secretory and lower molecular mass antigens in serum and urine samples of Indian children. Acta Trop 2009;110:22-7.  Back to cited text no. 56
    
57.
Carrara GM, Silva GB, Faria LS, Nunes DS, Ribeiro VS, Lopes CA, et al. IgY antibody and human neurocysticercosis: A novel approach on immunodiagnosis using Taenia crassiceps hydrophobic antigens. Parasitology 2020;147:240-7.  Back to cited text no. 57
    
58.
Carpio A, Campoverde A, Romo ML, García L, Piedra LM, Pacurucu M, et al. Validity of a PCR assay in CSF for the diagnosis of neurocysticercosis. Neurol Neuroimmunol Neuroinflamm 2017;4:e324. doi: 10.1212/NXI.0000000000000324.  Back to cited text no. 58
    
59.
Michelet L, Fleury A, Sciutto E, Kendjo E, Fragoso G, Paris L, et al. Human neurocysticercosis: Comparison of different diagnostic tests using cerebrospinal fluid. J Clin Microbiol 2011;49:195-200.  Back to cited text no. 59
    
60.
Goyal G, Phukan AC, Hussain M, Lal V, Modi M, Goyal MK, et al. Identification of Taenia solium DNA by PCR in blood and urine samples from a tertiary care center in North India. J Neurol Sci 2020;417:117057.  Back to cited text no. 60
    
61.
O'Connell EM, Harrison S, Dahlstrom E, Nash T, Nutman TB. A novel, highly sensitive quantitative polymerase chain reaction assay for the diagnosis of subarachnoid and ventricular neurocysticercosis and for assessing responses to treatment. Clin Infect Dis 2020;70:1875-81.  Back to cited text no. 61
    
62.
Goyal G, Phukan AC, Hussain M, Lal V, Modi M, Goyal MK, et al. Sorting out difficulties in immunological diagnosis of neurocysticercosis: Development and assessment of real time loop mediated isothermal amplification of cysticercal DNA in blood. J Neurol Sci 2020;408:116544.  Back to cited text no. 62
    
63.
Fan S, Qiao X, Liu L, Wu H, Zhou J, Sun R, et al. Next-generation sequencing of cerebrospinal fluid for the diagnosis of neurocysticercosis. Front Neurol 2018;9:471.  Back to cited text no. 63
    
64.
Fei X, Li C, Zhang Y, Zhang H, Liu X, Ji X, et al. Next-generation sequencing of cerebrospinal fluid for the diagnosis of neurocysticercosis. Clin Neurol Neurosurg 2020;193:105752.  Back to cited text no. 64
    
65.
World Health Organization, World Malaria Report 2020, 2020. Available from: https://www.who.int/publications/i/item/9789240015791.  Back to cited text no. 65
    
66.
Singhi P, Saini AG. Fungal and parasitic CNS infections. Indian J Pediatr 2019;86:83-90.  Back to cited text no. 66
    
67.
Severe malaria. Trop Med Int Health 2014;19(Suppl 1):7-131.  Back to cited text no. 67
    
68.
Zekar L, Sharman T. Plasmodium falciparum malaria. In: StatPearls. Treasure Island (FL): StatPearls Publishing; 2021.  Back to cited text no. 68
    
69.
Cheng Q, Gatton ML, Barnwell J, Chiodini P, McCarthy J, Bell D, et al. Plasmodium falciparum parasites lacking histidine-rich protein 2 and 3: A review and recommendations for accurate reporting. Malar J 2014;13:283.  Back to cited text no. 69
    
70.
Tangpukdee N, Duangdee C, Wilairatana P, Krudsood S. Malaria diagnosis: A brief review. Korean J Parasitol 2009;47:93-102.  Back to cited text no. 70
    
71.
Rubach MP, Mukemba J, Florence S, John B, Crookston B, Lopansri BK, et al. Plasma Plasmodium falciparum histidine-rich protein-2 concentrations are associated with malaria severity and mortality in Tanzanian children. PLoS One 2012;7:e35985. doi: 10.1371/journal.pone. 0035985.  Back to cited text no. 71
    
72.
Thakur KT, Vareta J, Carson KA, Kampondeni S, Potchen MJ, Birbeck GL, et al. Cerebrospinal fluid Plasmodium falciparum histidine-rich protein-2 in pediatric cerebral malaria. Malar J 2018;17:125.  Back to cited text no. 72
    
73.
Oriero EC, Van Geertruyden JP, Nwakanma DC, D'Alessandro U, Jacobs J. Novel techniques and future directions in molecular diagnosis of malaria in resource-limited settings. Expert Rev Mol Diagn 2015;15:1419-26.  Back to cited text no. 73
    
74.
Kwambana-Adams BA, Liu J, Okoi C, Mwenda JM, Mohammed NI, Tsolenyanu E, et al. On behalf of the paediatric bacterial meningitis surveillance network in West Africa. Etiology of pediatric meningitis in west Africa using molecular methods in the era of conjugate vaccines against pneumococcus, meningococcus, and haemophilus influenzae Type b. Am J Trop Med Hyg 2020;103:696-703.  Back to cited text no. 74
    
75.
Malpartida-Cardenas K, Miscourides N, Rodriguez-Manzano J, Yu LS, Moser N, Baum J, et al. Quantitative and rapid Plasmodium falciparum malaria diagnosis and artemisinin-resistance detection using a CMOS Lab-on-Chip platform. Biosens Bioelectron 2019;145:111678.  Back to cited text no. 75
    
76.
Rubio M, Bassat Q, Estivill X, Mayor A. Tying malaria and microRNAs: From the biology to future diagnostic perspectives. Malar J 2016;15:167.  Back to cited text no. 76
    
77.
Coronado LM, Nadovich CT, Spadafora C. Malarial hemozoin: From target to tool. Biochim Biophys Acta 2014;1840:2032-41.  Back to cited text no. 77
    

Top
Correspondence Address:
R S Jayshree
Department of Microbiology, Kidwai Memorial Institute of Oncology, Bangalore - 560 029, Karnataka
India
Login to access the Email id

Source of Support: None, Conflict of Interest: None


DOI: 10.4103/ijpm.ijpm_1123_21

Rights and Permissions


    Figures

  [Figure 1]
 
 
    Tables

  [Table 1]



 

Top
 
 
  Search
 
    Similar in PUBMED
   Search Pubmed for
   Search in Google Scholar for
 Related articles
    Email Alert *
    Add to My List *
* Registration required (free)  


    Abstract
   Introduction
   CNS Toxoplasmosis
   Neurocysticercosis
   Cerebral Malaria
   Conclusion
    References
    Article Figures
    Article Tables

 Article Access Statistics
    Viewed112    
    Printed2    
    Emailed0    
    PDF Downloaded10    
    Comments [Add]    

Recommend this journal