Authors:
Kantima Choosang Faculty of Medical Technology, Rangsit University, Pathumthani, 12000, Thailand

Search for other papers by Kantima Choosang in
Current site
Google Scholar
PubMed
Close
,
Siriphan Boonsilp Department of Clinical Pathology, Faculty of Medicine Vajira Hospital, Navamindradhiraj University, Dusit, Bangkok, Thailand

Search for other papers by Siriphan Boonsilp in
Current site
Google Scholar
PubMed
Close
,
Kanyanan Kritsiriwuthinan Faculty of Medical Technology, Rangsit University, Pathumthani, 12000, Thailand

Search for other papers by Kanyanan Kritsiriwuthinan in
Current site
Google Scholar
PubMed
Close
,
Palin Chumchuang Faculty of Medical Technology, Rangsit University, Pathumthani, 12000, Thailand

Search for other papers by Palin Chumchuang in
Current site
Google Scholar
PubMed
Close
,
Nanthawan Thanacharoensakun Faculty of Medical Technology, Rangsit University, Pathumthani, 12000, Thailand

Search for other papers by Nanthawan Thanacharoensakun in
Current site
Google Scholar
PubMed
Close
,
Aminoh Saai Bannangsta Hospital, Yala, Thailand

Search for other papers by Aminoh Saai in
Current site
Google Scholar
PubMed
Close
, and
Sawanya Pongparit Faculty of Medical Technology, Rangsit University, Pathumthani, 12000, Thailand

Search for other papers by Sawanya Pongparit in
Current site
Google Scholar
PubMed
Close
https://orcid.org/0000-0001-6238-1070
Open access

Abstract

Plasmodium vivax is the most prevalent cause of malaria in Thailand and is predominant in malarial endemic areas worldwide. P. vivax infection is characterized by low parasitemia, latent liver-stage parasites, or asymptomatic infections leading to underreported P. vivax cases. These are significant challenges for controlling and eliminating P. vivax from endemic countries. This study developed and evaluated a dot-blot enzyme-linked immunosorbent assay (ELISA) using PvMSP1-42 recombinant antigen for serological diagnosis based on the detection of antibodies against P. vivax. The optimal PvMSP1-42 concentration and dilutions of anti-human IgG horseradish peroxidase (HRP)-conjugated antiserum were tested on 88 serum samples from P. vivax, Plasmodium falciparum and bacterial infection, including healthy individuals. A cut-off titer of 1:800 produced optimal values for sensitivity and specificity of 90.9 and 98.2%, respectively, with an accuracy of 95.5%. The positive and negative predictive values were 96.8 and 94.7% respectively. The results from microscopic examination and dot-blot ELISA showed strong agreement with the 0.902 kappa index. Thus, the dot-blot ELISA using PvMSP1-42 antigen provided high sensitivity and specificity suitable for serodiagnosis of P. vivax infection. The test is a simple and quick diagnostic assay suitable for field testing as it does not require specific equipment or particular skills.

Abstract

Plasmodium vivax is the most prevalent cause of malaria in Thailand and is predominant in malarial endemic areas worldwide. P. vivax infection is characterized by low parasitemia, latent liver-stage parasites, or asymptomatic infections leading to underreported P. vivax cases. These are significant challenges for controlling and eliminating P. vivax from endemic countries. This study developed and evaluated a dot-blot enzyme-linked immunosorbent assay (ELISA) using PvMSP1-42 recombinant antigen for serological diagnosis based on the detection of antibodies against P. vivax. The optimal PvMSP1-42 concentration and dilutions of anti-human IgG horseradish peroxidase (HRP)-conjugated antiserum were tested on 88 serum samples from P. vivax, Plasmodium falciparum and bacterial infection, including healthy individuals. A cut-off titer of 1:800 produced optimal values for sensitivity and specificity of 90.9 and 98.2%, respectively, with an accuracy of 95.5%. The positive and negative predictive values were 96.8 and 94.7% respectively. The results from microscopic examination and dot-blot ELISA showed strong agreement with the 0.902 kappa index. Thus, the dot-blot ELISA using PvMSP1-42 antigen provided high sensitivity and specificity suitable for serodiagnosis of P. vivax infection. The test is a simple and quick diagnostic assay suitable for field testing as it does not require specific equipment or particular skills.

Introduction

Malaria is a life-threatening infectious disease caused by Plasmodium parasites that spread to humans by the bite of infected female Anopheles mosquitoes. Five Plasmodium species, i.e., Plasmodium falciparum, Plasmodium vivax, Plasmodium ovale, Plasmodium malariae, and Plasmodium knowlesi, are known to cause significant infection in humans [1]. According to the “World Malaria Report 2022”, in 2021, there was a global increase in the total number of malaria cases from 2020, with transmission occurring in 84 endemic countries and 247 million cases reported, causing 619,000 deaths worldwide [2]. Among the disease-carrying Plasmodium species, P. vivax and P. falciparum are present across most of the countries where malaria is endemic [2]. The infections due to caused by P. vivax and P. falciparum were estimated to be around 14.3 and 193.5 million cases, respectively [3]. Majority of severe malaria cases and deaths are caused by P. falciparum, whereas human infections caused by P. vivax are the most geographically widespread [4].

In Thailand, a high incidence and widespread occurrence of malaria are reported along the international border with Cambodia, Myanmar, and Malaysia [5, 6]. Although malaria cases in Thailand have substantially decreased since 2013 [7], considerable efforts are required to achieve the goals set by the National Malaria Elimination Program by 2024 [8]. Despite continued prevention efforts, approximately 8,380 confirmed malaria cases were reported in Thailand in 2022 [9] with P. vivax being the major cause of infection (7,957 cases, 94.9%) and P. falciparum being responsible for far fewer cases (201, 2.3%) [9].

P. vivax-related infections are a significant global health problem. They can be severe or even fatal and can result in significant global morbidity and mortality [10–12]. P. vivax presents low-level parasitemia and hypnozoite carriers, and causes a high number of asymptomatic infections [13]. The dormant hypnozoite, i.e., the liver-stage form of P. vivax that causes relapses and asymptomatic infections, can reactivate months or even 1–3 years after the blood-stage infection has been cleared, and relapsing accounts for up to 80% of all P. vivax infections [14–16]. Therefore, P. vivax is challenging to detect and eliminate. The laboratory diagnostic methods for detecting malaria infection depend on the identification of the blood-stages, antibodies, antigens, and nucleic acids of the Plasmodium. Each method has advantages and limitations, which makes it suitable for certain applications. The traditional method considered as a “gold standard” for malaria diagnosis consists of detection of blood-stage parasites in Giemsa-stained thick and thin blood films under a microscope. This microscopic method is inexpensive and simple to perform, but it is time consuming and has low sensitivity compare to nested PCR. Additionally, as it is difficult to distinguish Plasmodium species, well-trained expert technicians are required for this task [1718]. Molecular methods to detect nucleic acids of the Plasmodium are highly specific and more sensitive than microscopy. Moreover, they can assist in the accurate identification of malaria species as well as in epidemiological and phylogenetic studies. However, they require expensive equipment and specialist expertise [19, 20].

Consequently, serological tests may be a relatively more appropriate tool for detecting infected individuals, especially those with low-grade parasitemia, asymptomatic infection, and liver-stage forms of P. vivax. Blood infection associated with malaria induces specific antibody responses that are short-lived and may increase with subsequent infections [21–23]. The enzyme-linked immunosorbent assay (ELISA) is a highly specific and sensitive method based on antigen–antibody reactions. Anti-malaria antibodies are useful for malaria diagnosis, epidemiological surveillance, and screening of blood donors [24–26]. The microplate ELISA platform is an expensive and sophisticated technique that requires microplate reader apparatus and skilled personnel for its operation. Dot-blot ELISA is an alternative ELISA method that using nitrocellulose blotting membrane instead of microplate and can be read the color reaction visually. This assay is simple to perform and can be carried out in rural areas.

Based on the research of our colleague, we used genetic engineering techniques to produce a blood-stage antigen of P. vivax, namely P. vivax merozoite surface protein 1 (PvMSP1) [27]. PvMSP1 acts as an erythrocyte-binding protein on the merozoite surface and elicits strong adaptive immune response in patients infected with P. vivax [23, 28]. The C-terminus of the 42-kDa PvMSP1-42 is composed of two fragments of 33 and 19 kDa (i.e., PvMSP1-33 and PvMSP1-19, respectively), which play a vital role in the initial contact with erythrocyte before merozoite invasion [29, 30]. It has been shown that using an ELISA plate based on the PvMSP1-42 blood-stage antigen allowed to efficiently detect specific antibodies in infected individuals [31]. The dot-blot ELISA for detection of antibody against PvMSP1-42 antigen have not been described. Therefore, we developed and evaluated the simple ELISA based on the PvMSP1-42 antigen onto a dot-blot assay platform that is cheaper and quicker than the ELISA plate. Additionally, this assay does not require as much equipment or expertise and can be carried out in rural areas.

Materials and methods

Sample collection

This study was conducted to evaluate the dot-blot ELISA using the PvMSP1-42 recombinant antigen. The serum samples included two groups: patients with P. vivax (P) and a negative control group (P. falciparum, bacterial infection and normal sera). A total of 88 human serum samples were selected from 43 malarial cases from Yala, Thailand, where P. vivax (n = 33) and P. falciparum (n = 10) infections had been microscopically verified by Giemsa-stained blood smear. Samples from 13 bacterial infection (BI) cases including the sample from patients infected with Acinetobacter baumannii, Escherichia coli, Klebsiella pneumonia, Pseudomonas aeruginosa, Salmonella Paratyphi A, Staphylococcus aureus, and 32 healthy individuals (N) from Thammasat University Hospital, Pathumthani, Thailand, were also used.

Confirmation of PvMSP1-42 purified recombinant protein

Purified PvMSP1-42 histidine-tagged recombinant protein was provided by our collaborator and their colleagues [27]. Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) and western blot analyses were performed to confirm the identity of PvMSP1-42. Briefly, the PvMSP1-42 histidine-tagged protein was analyzed by SDS-PAGE and stained with Coomassie Blue R-250 (Bio-Rad Laboratories, Inc., USA). After SDS-PAGE, PvMSP1-42 protein samples were transferred onto nitrocellulose membranes, which were then blocked with 5% skim milk for 24 h at 4 °C. The membrane was then incubated with anti-His-horseradish peroxidase (HRP) at 1:2,500 dilution for 1 h at room temperature. An immunoreactive band with a molecular weight of 42 kDa after adding 3, 3′, 5, 5′-tetramethylbenzidine (TMB) substrate was accepted as being PvMSP1-42.

Quantitation of PvMSP1-42 recombinant protein

The protein concentration of the recombinant PvMSP1-42 protein was measured using the Bio-Rad protein assay (Bio-Rad Laboratories, Inc., USA), which is based on the Bradford dye-binding method [32] The Bradford assay determines the protein concentration by measuring the color change in the sample, from colorless to blue, caused by the response of amino acid residues, such as arginine, lysine, and histidine. The density of the blue color is proportional to the protein concentration. A standard curve was prepared using 1 mL of Bradford reagent added to 0.1 mL of bovine serum albumin (BSA; Sigma-Aldrich) at 1, 0.5, 0.25, 0.125, 0.0625, and 0.031 mg mL−1) or 0.1 mL of PvMSP1-42. After mixing, the samples were incubated at room temperature for 10 min and the absorbance measured at 595 nm using a spectrophotometer. The protein concentration of PvMSP1-42 was calculated by comparing the absorbance results to the BSA standard curve.

Optimization and serum testing of dot-blot ELISA

The optimal conditions were validated with separate pooled P. vivax positive serum and pooled negative serum for P. vivax infection (50 µL). The PvMSP1-42 recombinant protein antigen was applied onto a nitrocellulose membrane using a Bio-Dot microfiltration apparatus (Bio-Rad laboratories, Inc., USA) for a dot-blot assay. Membranes were air-dried for 30 min and blocked in 3% BSA in phosphate-buffered saline (PBS) with 0.05% Tween-20 (PBST) at room temperature for 1 h. Three washes with PBST were performed, and the membrane was probed for 30 min with the tested serum diluted 1:200 in 0.5% BSA in PBST. After another three washes with PBST, the membrane was incubated with the secondary antibody: anti-human IgG-HRP at dilution of 1:10,000 or 1:5,000 in PBST containing 0.5% BSA, for 30 min at room temperature. The membrane was rewashed before color development using the TMB substrate (KPL, Gaithersburg, MD, USA) for 2–10 min. Positive signals shown by a blue circular dot on the membrane. A dot that was 0.5% BSA in PBST used as direct conjugate control (DCC). To analyze a sample, the optimal condition was used to perform with twofold serial dilution serum, and the last serum dilution that showed a positive result was the end point antibody titer. The pooled positive and negative controls were also tested in every set of the assay.

Data analysis

The diagnostic sensitivity and specificity of the dot-blot ELISA at various cut-off titers were calculated to compare P. vivax-infected and uninfected patients. Positive and negative predictive values of the test were determined by varying the cut-off titers. The cut-off titer with the best accuracy values was selected. The performance of the dot-blot ELISA was evaluated by comparison with the reference microscopic examination. The Mann–Whitney test was used to determine whether there were differences in median titer between the groups. The agreement between different diagnostic tests was calculated using the kappa (κ) measure of inter-rater agreement. Briefly, κ < 0.20 indicated poor agreement, 0.21–0.40 fair, 0.41–0.60 moderate, 0.61–0.80 good, 0.81–0.99 very good, and 1.00 indicated perfect agreement [33].

Ethics

Ethical approval for this study protocol was obtained from Rangsit University Ethics Review Board (RSEC 45/2560).

Results

Confirmation of PvMSP1-42 recombinant protein

The histidine-tag was used as a recombinant protein marker. The histidine-containing PvMSP1-42 sample was loaded undiluted, and at 1:10, 1:100, and 1:1,000 dilutions onto an SDS-PAGE gel. Following electrophoresis, the recombinant protein was detected via western blotting with anti-His-HRP antibody (Fig. 1). The anti-His-HRP antibody recognized a protein of approximately 45 kDa with no degradation products when undiluted (150 μg mL−1) and at a dilution of 1:10 (15 μg ml−1) but was not detected at dilutions of 1:100 (5 μg ml−1) and 1:1,000 (0.5 μg ml−1). The result indicated the presence of purified PvMSP1-42 recombinant proteins.

Fig. 1.
Fig. 1.

Western blot analysis of purified PvMSP1-42 His-tagged protein. M: protein molecular weight marker, Lane 1 to 4 was tested with PvMSP1-42 protein at concentrations of 150, 15, 1.5 and 0.15 ug/ml respectively. The immunoreactivity of PvMSP1-42 protein antigen and anti-His-HRP with TMB substrate were presented at the arrows (∼45 kDa) which indicates the specific band for purified PvMSP1-42 recombinant protein

Citation: European Journal of Microbiology and Immunology 14, 2; 10.1556/1886.2024.00008

Quantitation of PvMSP1-42 recombinant protein

To investigate the protein concentration of the stock purified PvMSP1-42 recombinant antigen, the protein was measured using the Bradford assay and calculated using the linear equation y = 0.6134X + 0.5166 obtained from the BSA standard curve. The stock PvMSP1-42 protein concentration was 150 μg mL−1 The PvMSP1-42 protein was then used to develop a dot-blot ELISA to detect patients who were infected with P. vivax.

Optimization and serum testing of dot-blot ELISA

The purified PvMSP1-42 antigenicity was evaluated for the immunological diagnosis of patients infected with P. vivax. The PvMSP1-42 protein concentration between 0.75 and 7,500 ng per dot was tested with the pooled positive and negative sera at a dilution of 1:200 to determine the optimal amount of PvMSP1-42 antigen. The membranes were probed with anti-human IgG-HRP conjugated antiserum at dilutions of 1:5,000 and 1:10,000, and the incubation time with TMB substrate varied from 2 to 10 min. The optimal condition that provided an excellent visual distinction of the positive and negative sample results was selected. The positive serum appeared as a blue circular dot, whereas the negative serum showed no color dot (Fig. 2). At the 1:200 dilution of patient antiserum, the minimum antigen concentration required to detect the specific antibody was 750 ng per dot, and the optimum HRP-conjugated antiserum dilution was 1:10,000. The optimal incubation time with the TMB substrate was 4 min. These optimal conditions were used for further serum evaluation.

Fig. 2.
Fig. 2.

Dot-blot ELISA optimization. Different concentrations of recombinant PvMSP1-42 protein (0.75–7,500 ng per dot) were dot onto a nitrocellulose membrane. After blocking and washing, the nitrocellulose with a dot of antigen was tested with 1:200 dilution of pooled positive serum (+), pooled negative serum (−), and direct conjugate control (DCC). After washing step, anti-human conjugated with horseradish peroxidase using two different dilutions; figure A used a dilution of 1:5,000, and figure B used a dilution of 1:10,000. The time for color development when incubated with TMB substrate was tested at 2, 4, 6, 8, and 10 min. The dashed-square in figure B indicates the results of the appropriate conditions selected, the concentration of recombinant PvMSP1-42 protein 750 ng/dot, dilution of conjugate 1:10,000 and the substrate incubation time of 4 min, where positive serum gave a dark blue color dot but the negative serum and DCC revealed no blue dot

Citation: European Journal of Microbiology and Immunology 14, 2; 10.1556/1886.2024.00008

Optimum cut-off value and diagnostic performance of the dot-blot ELISA

A total of 88 serum samples were evaluated. Optimized conditions were used for dot-blot ELISA, with sera diluted in a two-fold serial dilution (1:200, 1:400, 1:800, 1:1,600, and 1:3,200). The distribution of sample titers within each group; the positive serum samples of 33 patients infected with P. vivax and 55 negative control samples, including control diseases of 10 patients infected with P. falciparum and 13 patients with bacterial infections, and 32 normal controls, presented in Fig. 3. Most of the P. vivax-infected samples exhibited the titer of 1:3,200, while most of the negative control samples presented the titer of ≤1:200. The median titer of the P. vivax sera and negative control group was 2338.36 and 199.85, respectively. There was a significant difference in median titer between the P. vivax sera and negative control group (P < 0.001). The cut-off titer value of the dot-blot ELISA was investigated, as shown in Table 1. The most suitable cut-off titer value was ≥1:800, which provided the highest accuracy of 95.5% (confidence interval [CI]; range 88.77–98.75%). This cut-off titer revealed sensitivity, specificity, positive predictive value and negative predictive value of 90.9% (CI; 75.7–98.1%), 98.2% (CI; 90.3–100%), 96.8% (CI; 81.1–99.5%) and 94.7% (CI; 86.0–98.2%) respectively. In addition, the comparison of the results from microscopic examination and dot-blot ELISA for detecting P. vivax infection showed a strong agreement with the 0.902 kappa index (P < 0.005) (Table 2).

Fig. 3.
Fig. 3.

Distribution of the dot-blot ELISA titers within each serum group. The dot-blot ELISA with high titer of 1:800, 1:1,600 and 1:3,200 found more frequently in P. vivax infected serum than in non-P. vivax infected serum groups

Abbreviations: Pv, P. vivax infection; Pf, P. falciparum infection; N, normal control; BI, bacterial infection.

Citation: European Journal of Microbiology and Immunology 14, 2; 10.1556/1886.2024.00008

Table 1.

Sensitivity, specificity, positive predictive value, negative predictive value, and accuracy of the dot-blot ELISA at various cut-off titers. The selected cut-off titer was 1:800

Cut-off titerPositive serum

[Pv]

(n = 33)
Negative serum

[Pf]

(n = 10)
Negative serum

[N + CD]

(n = 45)
Sensitivity (%)Specificity (%)Accuracy (%)PPV (%)NPV (%)
+++
≥1:2003213731149738.260.248.595.45
≥1:40031237133293.970.979.56695.12
≥1:8003031904590.998.295.596.894.74
≥1:1,6002671904578.898.290.996.388.52
≥1:3,20019141904557.698.283.09579.41

Abbreviations: PPV, positive predictive value; NPV, negative predictive value; Pv, P. vivax infection; Pf, P. falciparum infection; N, normal control; BI, bacterial infection.

Table 2.

Concordance and discordance of P. vivax diagnostic results between microscopy and dot-blot ELISA was calculated with kappa value

MicroscopyDot-blot ELISATotalKappa valueP-value
PositiveNegative
Positive303330.902<0.001
Negative15455
Total315788

Discussion

The accurate identification of Plasmodium species is critical not only for determining appropriate treatment regimen but also for implementing effective malaria control measures in endemic areas. Microscopic diagnosis is still considered to be the most reliable method in remote laboratory settings [2]. However, the detection of P. vivax-related malaria cases via the visual microscopic examination of blood smears presents limitations associated with the identification of hypnozoites and asymptomatic infection. Some studies have reported the presence of a larger parasite reservoir in the spleen than in bloodstream [34, 35]. If undiagnosed, P. vivax infections will not be treated, and parasite will remain in the population potentially triggering infections whenever an opportunity arises. Serological testing is an advantageous technique for identification of asymptomatic individuals who may have not been diagnosed via other methods [36].

Among the surface antigens of the Plasmodium blood-stage antigens, MSP-1 is one of the most promising under consideration for malaria vaccines and diagnostic kits [3738]. Serological tests that can detect antibodies based on the appropriate blood-stage antigen could be a valuable tool for identifying individuals that have recently been infected with P. vivax, even if asymptomatic [39]. Malaria antibodies are typically developed within 1–2 weeks of the initial infection and remain detectable for months to years after the parasite has been cleared. However, the peak in antibody synthesis occurs within 7–14 days [40, 41]. Hence, ELISAs have been widely used to detect anti-P. vivax antibodies.

The ELISA plate method requires specific equipment making it unsuitable for screening a large number of samples collected on field in order to monitor P. vivax-associated malaria cases. Therefore, in this study, we developed and evaluated a dot-blot ELISA using the PvMSP1-42 recombinant antigen as a simple serological assay for field testing. The assay revealed a significant difference between the median titer of P. vivax-infected patients and negative control sera groups. This finding indicated that the dot-blot ELISA was able to discriminate between P. vivax-infected and non-infected individuals. Additionally, the developed method exhibited high levels of diagnostic sensitivity and specificity, i.e., 90.9 and 98.2%, respectively, which were higher than those of ELISA plate based on the PvMSP1-42 antigen (86.9 and 94.05%, respectively) [42]. The sensitivity and specificity have been evaluated for various ELISAs applied for malaria detection, with value ranges of 53.0–94.4% and 94.0–99.6%, respectively [42–45]. Unfortunately, we have no parasitemia data to evaluate the analytical sensitivity of this dot-blot ELISA.

The dot-blot ELISA developed in this study detected anti-PvMSP1-42 antibodies in 30 of 33 microscopically confirmed P. vivax-positive samples. The lack of antibodies against the PvMSP1-42 in three microscopic assessments proved the serum samples might be from early infections; however, this study did not have the symptom onset date. This finding may be due to the serological tests are not practical for the acute infection since the time required for the development of antibodies is as long as a week or more; however, it is a useful tool as seroprevalence and a confirmatory test for detection of low parasitemia of P. vivax infection with symptoms but cannot be detected under the microscope. One P. falciparum-infected individual was identified via microscopic examination that was positive for P. vivax dot-blot ELISA. This individual may have P. falciparum infection and submicroscopic P. vivax infection [8]. This discordance sample was confirmed by nested PCR and showed to be a mixed infection, due to it was positive for both specific genes of P. falciparum and P. vivax. The dot-blot ELISA also showed excellent agreement with the gold standard microscopic method, with a Kappa index of 0.902 (P < 0.000). This represent an almost perfect agreement (0.81–0.99) for the dot-blot ELISA where a Kappa index of 1 is a perfect agreement [46]. Moreover, we found that the dot-blot ELISA can be carried out approximately 1 h after the blocking step, and results can be read with naked eye at low recombinant protein concentration used. In addition, we found that dot-blot ELISA can perform approximately 1 h after the blocking step. However, in our future research, we will consider using more differentiated samples to evaluate the dot-blot ELISA and investigate how to expedite the assay and further develop it into a rapid test.

During this study period, P. vivax and P. falciparum samples collected from the blood samples of patients diagnosed with P. vivax or P. falciparum infection by routine laboratory microscopy. There was no information on the parasitemia of each Plasmodium in each patient nor the length of the malarial onset of the patients. Additionally, we could not include data from serum samples of other Plasmodium species infections due to very low prevalence of P. ovale, P. malariae and P. knowlesi infection in Thailand. With these limitations, this study provides the preliminary evaluation. Thus, further studies will collect more samples with information on the parasitemia and onset of parasitic infection in patients for evaluating the dot-blot ELISA efficiency, investigating how to expedite the assay, and further developing it into a rapid test.

Conclusion

The dot-blot ELISA developed in this study had high sensitivity, high specificity, and a favorable agreement with the microscopic examination method, thereby indicating that the dot-blot ELISA based on the recombinant PvMSP1-42 antigen shows excellent potential for detecting and monitoring infection with P. vivax malaria in remote areas, as the assay is less expensive and easier to perform and interpret than the current microscopy and ELISA plate methods.

Funding sources

This work was partially supported by Faculty of Medical Technology, Rangsit University.

Authors' contributions

SP, KC, KK and SB designed the study. KC, KK, PC and NT performed the experiments and collected data. SP, KC, SB, and KK involved in data analysis and interpretation. SB and SP wrote the manuscript. All authors have read and approved the final manuscript.

Conflict of interest

The authors declare that they have no conflicts of interest.

Acknowledgment

We want to thank Phitchapat Nimnuch for assisting in collecting normal and control samples. We would also like to thank Benyapa Playngam, Pandaree, Jiamworapong, and Pornthiwa Sanguankay, for their laboratory assistance.

References

  • 1.

    Pimenta PF, Orfano AS, Bahia AC, Duarte AP, Ríos-Velásquez CM, Melo FF, et al. An overview of malaria transmission from the perspective of Amazon Anopheles vectors. Mem Inst Oswaldo Cruz. 2015 Feb;110(1):2347.

    • Search Google Scholar
    • Export Citation
  • 2.

    World Health Organization. World malaria report 2022 [Internet]. Available from: https://www.who.int/teams/global-malaria-programme/reports/world-malaria-report-2022.

    • Search Google Scholar
    • Export Citation
  • 3.

    Battle KE, Lucas TCD, Nguyen M, Howes RE, Nandi AK, Twohig KA, et al. Mapping the global endemicity and clinical burden of Plasmodium vivax, 2000–17: a spatial and temporal modelling study. The Lancet. 2019 Jul;394(10195):33243.

    • Search Google Scholar
    • Export Citation
  • 4.

    World Health Organization. World malaria report. 2021 [Internet]. Available from: https://www.who.int/teams/global-malaria-programme/reports/world-malaria-report-2021.

    • Search Google Scholar
    • Export Citation
  • 5.

    Gething PW, Elyazar IRF, Moyes CL, Smith DL, Battle KE, Guerra CA, et al. A long neglected world malaria map: Plasmodium vivax endemicity in 2010. Carlton JM, editor. Plos Negl Trop Dis. 2012 Sep 6;6(9):e1814.

    • Search Google Scholar
    • Export Citation
  • 6.

    Ammatawiyanon L, Tongkumchum P, Lim A, McNeil D. Modelling malaria in southernmost provinces of Thailand: a two-step process for analysis of highly right-skewed data with a large proportion of zeros. Malar J. 2022 Nov 15;21(1):334.

    • Search Google Scholar
    • Export Citation
  • 7.

    Sudathip P, Kitchakarn S, Shah JA, Bisanzio D, Young F, Gopinath D, et al. A foci cohort analysis to monitor successful and persistent foci under Thailand's Malaria Elimination Strategy. Malar J. 2021 Dec;20(1):118.

    • Search Google Scholar
    • Export Citation
  • 8.

    World Health Organization. World malaria report 2018 [Internet]. Available from: https://apps.who.int/iris/handle/10665/275867.

  • 9.

    World Health Organization. Mekong malaria elimination: epidemiology summary, volume 19, July–September 2022 [Internet]. Available from: https://www.who.int/teams/global-malaria-programme/reports/world-malaria-report-2022.

    • Search Google Scholar
    • Export Citation
  • 10.

    Phyo AP, Dahal P, Mayxay M, Ashley EA. Clinical impact of vivax malaria: a collection review. Beeson JG, editor. Plos Med. 2022 Jan 18;19(1):e1003890.

    • Search Google Scholar
    • Export Citation
  • 11.

    Mathews S, Bhagwati M, Agnihotri V. Clinical spectrum of Plasmodium vivax infection, from benign to severe malaria: a tertiary care prospective study in adults from Delhi, India. Trop Parasitol. 2019;9(2):88.

    • Search Google Scholar
    • Export Citation
  • 12.

    Tovar-Acero C, Velasco MC, Avilés-Vergara PA, Ricardo-Caldera DM, Alvis EM, Ramirez-Montoya J, et al. Liver and kidney dysfunction, hypoglycemia, and thrombocytopenia in Plasmodium vivax malaria patients at a Colombian Northwest region. Parasite Epidemiol Control. 2021 May;13:e00203.

    • Search Google Scholar
    • Export Citation
  • 13.

    Greenhouse B, Daily J, Guinovart C, Goncalves B, Beeson J, Bell D, et al. Priority use cases for antibody-detecting assays of recent malaria exposure as tools to achieve and sustain malaria elimination. Gates Open Res. 2019 Feb 12;3:131.

    • Search Google Scholar
    • Export Citation
  • 14.

    Adams JH, Mueller I. The biology of Plasmodium vivax. Cold Spring Harb Perspect Med. 2017 Sep;7(9):a025585.

  • 15.

    Robinson LJ, Wampfler R, Betuela I, Karl S, White MT, Li Wai Suen CSN, et al. Strategies for understanding and reducing the Plasmodium vivax and Plasmodium ovale hypnozoite reservoir in Papua New Guinean children: a randomised placebo-controlled trial and mathematical model. Von Seidlein L, editor. Plos Med. 2015 Oct 27;12(10):e1001891.

    • Search Google Scholar
    • Export Citation
  • 16.

    Advances in Parasitology. Relapse [Internet]. Available from: https://linkinghub.elsevier.com/retrieve/pii/B9780123979001000025.

  • 17.

    Thekisoe O, Pöschl B, Chutipongvivate S, Panagiotis K, Waneesorn J. Comparative diagnosis of malaria infections by microscopy, nested PCR, and LAMP in Northern Thailand. Am J Trop Med Hyg. 2010 Jul 1;83(1):5660.

    • Search Google Scholar
    • Export Citation
  • 18.

    Barber BE, William T, Grigg MJ, Yeo TW, Anstey NM. Limitations of microscopy to differentiate Plasmodium species in a region co-endemic for Plasmodium falciparum, Plasmodium vivax and Plasmodium knowlesi. Malar J. 2013 Dec;12(1):8.

    • Search Google Scholar
    • Export Citation
  • 19.

    Snounou G, Viriyakosol S, Xin Ping Zhu, Jarra W, Pinheiro L, Do Rosario VE, et al. High sensitivity of detection of human malaria parasites by the use of nested polymerase chain reaction. Mol Biochem Parasitol. 1993 Oct;61(2):31520.

    • Search Google Scholar
    • Export Citation
  • 20.

    Slater L, Ashraf S, Zahid O, Ali Q, Oneeb M, Akbar MH, et al. Current methods for the detection of Plasmodium parasite species infecting humans. Curr Res Parasitol Vector-borne Dis 2022;2:100086. Mar 19;2:100086. https://doi.org/10.1016/j.crpvbd.2022.100086. e Collection 2022.

    • Search Google Scholar
    • Export Citation
  • 21.

    Gonzalez-Ceron L, Rodriguez MH, Sandoval MA, Santillan F, Galindo-Virgen S, Betanzos AF, et al. Effectiveness of combined chloroquine and primaquine treatment in 14 days versus intermittent single dose regimen, in an open, non-randomized, clinical trial, to eliminate Plasmodium vivax in southern Mexico. Malar J [Internet]. 2015 Dec [cited 2023 Jul 9];14(1):426. Available from: http://www.malariajournal.com/content/14/1/426.

    • Search Google Scholar
    • Export Citation
  • 22.

    Park JW, Moon SH, Yeom JS, Lim KJ, Sohn MJ, Jung WC, et al. Naturally acquired antibody responses to the C-terminal region of merozoite surface protein 1 of Plasmodium vivax in Korea. Clin Diagn Lab Immunol [Internet]. 2001 Jan [cited 2023 Jul 9];8(1):1420. Available from: https://journals.asm.org/doi/10.1128/CDLI.8.1.14-20.2001.

    • Search Google Scholar
    • Export Citation
  • 23.

    Lee S, Choi YK, Goo YK. Humoral and cellular immune response to Plasmodium vivax VIR recombinant and synthetic antigens in individuals naturally exposed to P. vivax in the Republic of Korea. Malar J [Internet]. 2021 Dec [cited 2023 Jul 29];20(1):288. Available from: https://malariajournal.biomedcentral.com/articles/10.1186/s12936-021-03810-2.

    • Search Google Scholar
    • Export Citation
  • 24.

    Niederhauser C, Tinguely C, Dreier J, Vollmer T, Marti HP, Nickel B, et al. Comparison of a new IgG-EIA for the detection of anti-plasmodium antibodies with two currently used assays. Transfus Med Hemotherapy. 2021;48(5):26571.

    • Search Google Scholar
    • Export Citation
  • 25.

    Amanfo SA, Mduluza T, Midzi N, Cavanagh DR, Mutapi F. Seroepidemiology of Plasmodium species infections in Zimbabwean population. Malar J. 2016 Dec;15(1):267.

    • Search Google Scholar
    • Export Citation
  • 26.

    Murungi LM, Kimathi RK, Tuju J, Kamuyu G, Osier FHA. Serological profiling for malaria surveillance using a standard ELISA protocol. In: Ariey F, Gay F, Ménard R, editors. Malaria control and elimination [Internet]. New York, NY: Springer New York; 2019 [cited 2023 Nov 15]. p. 8390. (Methods in Molecular Biology; vol. 2013). Available from: http://link.springer.com/10.1007/978-1-4939-9550-9_6.

    • Search Google Scholar
    • Export Citation
  • 27.

    Rojrueangnont K, Rungnoppakitpanit K, Rodseesan N, Sommanat N, Hyayhngthong H. Production of recombinant Plasmodium vivax merozoite surface protein (PvMSP1-42 kDa) by genetic engineering. Rangsit University, Thailand: Bachelor of science program in medical technology; 20.

    • Search Google Scholar
    • Export Citation
  • 28.

    Cheng Y, Wang Y, Ito D, Kong DH, Ha KS, Chen JH, et al. The Plasmodium vivax merozoite surface protein 1 paralog is novel erythrocyte-binding ligand of P. vivax. Adams JH, editor. Infect Immun [Internet]. 2013 May [cited 2023 Jul 9];81(5):158595. Available from: https://journals.asm.org/doi/10.1128/IAI.01117-12.

    • Search Google Scholar
    • Export Citation
  • 29.

    Flores-Alanis A, González-Cerón L, Santillán-Valenzuela F, Ximenez C, Sandoval-Bautista MA, Cerritos R. Spatiotemporal changes in Plasmodium vivax msp142 haplotypes in southern Mexico: from the control to the pre-elimination phase. Microorganisms [Internet]. 2022 Jan 15 [cited 2023 Jul 9];10(1):186. Available from: https://www.mdpi.com/2076-2607/10/1/186.

    • Search Google Scholar
    • Export Citation
  • 30.

    Gutiérrez S, González-Cerón L, Montoya A, Sandoval MA, Tórres ME, Cerritos R. Genetic structure of Plasmodium vivax in Nicaragua, a country in the control phase, based on the carboxyl terminal region of the merozoite surface protein-1. Infect Genet Evol [Internet]. 2016 Jun [cited 2023 Jul 9];40:32430. Available from: https://linkinghub.elsevier.com/retrieve/pii/S1567134815003639.

    • Search Google Scholar
    • Export Citation
  • 31.

    Dinzouna-Boutamba SD, Lee S, Son UH, Song SM, Yun HS, Joo SY, et al. Distribution of antibodies specific to the 19-kDa and 33-kDa fragments of Plasmodium vivax merozoite surface protein 1 in two pathogenic strains infecting Korean Vivax malaria patients. Osong Public Health Res Perspect [Internet]. 2016 Aug [cited 2023 Jul 9];7(4):2139. Available from: http://linkinghub.elsevier.com/retrieve/pii/S2210909916300583.

    • Search Google Scholar
    • Export Citation
  • 32.

    Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem [Internet]. 1976 May [cited 2023 Jul 9];72(1–2):24854. Available from: https://linkinghub.elsevier.com/retrieve/pii/0003269776905273.

    • Search Google Scholar
    • Export Citation
  • 33.

    McHugh ML. Interrater reliability: the kappa statistic. Biochem Med [Internet]. 2012 [cited 2023 Jul 9];27682. Available from: http://www.biochemia-medica.com/en/journal/22/3/10.11613/BM.2012.031.

    • Search Google Scholar
    • Export Citation
  • 34.

    Kho S, Qotrunnada L, Leonardo L, Andries B, Wardani PAI, Fricot A, et al. Hidden biomass of intact malaria parasites in the human spleen. N Engl J Med. 2021 May 27;384(21):20679.

    • Search Google Scholar
    • Export Citation
  • 35.

    Kho S, Qotrunnada L, Leonardo L, Andries B, Wardani PAI, Fricot A, et al. Evaluation of splenic accumulation and colocalization of immature reticulocytes and Plasmodium vivax in asymptomatic malaria: a prospective human splenectomy study. Von Seidlein L, editor. Plos Med. 2021 May 26;18(5):e1003632.

    • Search Google Scholar
    • Export Citation
  • 36.

    Kartal L, Mueller I, Longley RJ. Using serological markers for the surveillance of Plasmodium vivax malaria: a scoping review. Pathogens. 2023 May 31;12(6):791.

    • Search Google Scholar
    • Export Citation
  • 37.

    Good MF, Stanisic D, Xu H, Elliott S, Wykes M. The immunological challenge to developing a vaccine to the blood stages of malaria parasites. Immunol Rev [Internet]. 2004 Oct [cited 2023 Jul 9];201(1):25467. Available from: https://onlinelibrary.wiley.com/doi/10.1111/j.0105-2896.2004.00178.x.

    • Search Google Scholar
    • Export Citation
  • 38.

    Mirahmadi H, Fallahi S, Fallah Omrani V, Kazemi B, Haghighi A, Seyyed Tabaei SJ. High-level expression of immunogenic recombinant Plasmodium vivax merozoite surface protein (Pvmsp-142 kDa) in pGEX 6P1 vector. Iran J Public Health. 2015 Jan;44(1):8999.

    • Search Google Scholar
    • Export Citation
  • 39.

    Longley RJ, White MT, Takashima E, Brewster J, Morita M, Harbers M, et al. Development and validation of serological markers for detecting recent Plasmodium vivax infection. Nat Med [Internet]. 2020 May [cited 2023 Jul 9];26(5):7419. Available from: https://www.nature.com/articles/s41591-020-0841-4.

    • Search Google Scholar
    • Export Citation
  • 40.

    Seed CR, Kitchen A, Davis TME. The current status and potential role of laboratory testing to prevent transfusion-transmitted malaria. Transfus Med Rev [Internet]. 2005 Jul [cited 2023 Jul 9];19(3):22940. Available from: https://linkinghub.elsevier.com/retrieve/pii/S0887796305000167.

    • Search Google Scholar
    • Export Citation
  • 41.

    Seed CR, Kee G, Wong T, Law M, Ismay S. Assessing the safety and efficacy of a test-based, targeted donor screening strategy to minimize transfusion transmitted malaria. Vox Sang [Internet]. 2010 Apr [cited 2023 Jul 9];98(3p1):e18292. Available from: https://onlinelibrary.wiley.com/doi/10.1111/j.1423-0410.2009.01251.x.

    • Search Google Scholar
    • Export Citation
  • 42.

    Mirahmadi H, Fallahi S, Seyyed Tabaei SJ. Soluble recombinant merozoite surface antigen-142kDa of Plasmodium vivax: an improved diagnostic antigen for vivax malaria. J Microbiol Methods. 2016 Apr;123:4450.

    • Search Google Scholar
    • Export Citation
  • 43.

    Doderer C, Heschung A, Guntz P, Cazenave JP, Hansmann Y, Senegas A, et al. A new ELISA kit which uses a combination of Plasmodium falciparum extract and recombinant Plasmodium vivax antigens as an alternative to IFAT for detection of malaria antibodies. Malar J. 2007 Dec;6(1):19.

    • Search Google Scholar
    • Export Citation
  • 44.

    Nam MH, Kim JS, Cho CH, Han ET, Lee WJ, Lee HK, et al. Evaluation of Plasmodium vivax ELISA for the blood screen. Trop Med Int Health TM IH. 2010 Dec;15(12):143641.

    • Search Google Scholar
    • Export Citation
  • 45.

    Oh JS, Kim JS, Lee CH, Nam DH, Kim SH, Park DW, et al. Evaluation of a malaria antibody enzyme immunoassay for use in blood screening. Mem Inst Oswaldo Cruz. 2008 Jan 31;103(1):758.

    • Search Google Scholar
    • Export Citation
  • 46.

    Viera AJ, Garrett JM. Understanding interobserver agreement: the kappa statistic. Fam Med. 2005 May;37(5):3603.

  • 1.

    Pimenta PF, Orfano AS, Bahia AC, Duarte AP, Ríos-Velásquez CM, Melo FF, et al. An overview of malaria transmission from the perspective of Amazon Anopheles vectors. Mem Inst Oswaldo Cruz. 2015 Feb;110(1):2347.

    • Search Google Scholar
    • Export Citation
  • 2.

    World Health Organization. World malaria report 2022 [Internet]. Available from: https://www.who.int/teams/global-malaria-programme/reports/world-malaria-report-2022.

    • Search Google Scholar
    • Export Citation
  • 3.

    Battle KE, Lucas TCD, Nguyen M, Howes RE, Nandi AK, Twohig KA, et al. Mapping the global endemicity and clinical burden of Plasmodium vivax, 2000–17: a spatial and temporal modelling study. The Lancet. 2019 Jul;394(10195):33243.

    • Search Google Scholar
    • Export Citation
  • 4.

    World Health Organization. World malaria report. 2021 [Internet]. Available from: https://www.who.int/teams/global-malaria-programme/reports/world-malaria-report-2021.

    • Search Google Scholar
    • Export Citation
  • 5.

    Gething PW, Elyazar IRF, Moyes CL, Smith DL, Battle KE, Guerra CA, et al. A long neglected world malaria map: Plasmodium vivax endemicity in 2010. Carlton JM, editor. Plos Negl Trop Dis. 2012 Sep 6;6(9):e1814.

    • Search Google Scholar
    • Export Citation
  • 6.

    Ammatawiyanon L, Tongkumchum P, Lim A, McNeil D. Modelling malaria in southernmost provinces of Thailand: a two-step process for analysis of highly right-skewed data with a large proportion of zeros. Malar J. 2022 Nov 15;21(1):334.

    • Search Google Scholar
    • Export Citation
  • 7.

    Sudathip P, Kitchakarn S, Shah JA, Bisanzio D, Young F, Gopinath D, et al. A foci cohort analysis to monitor successful and persistent foci under Thailand's Malaria Elimination Strategy. Malar J. 2021 Dec;20(1):118.

    • Search Google Scholar
    • Export Citation
  • 8.

    World Health Organization. World malaria report 2018 [Internet]. Available from: https://apps.who.int/iris/handle/10665/275867.

  • 9.

    World Health Organization. Mekong malaria elimination: epidemiology summary, volume 19, July–September 2022 [Internet]. Available from: https://www.who.int/teams/global-malaria-programme/reports/world-malaria-report-2022.

    • Search Google Scholar
    • Export Citation
  • 10.

    Phyo AP, Dahal P, Mayxay M, Ashley EA. Clinical impact of vivax malaria: a collection review. Beeson JG, editor. Plos Med. 2022 Jan 18;19(1):e1003890.

    • Search Google Scholar
    • Export Citation
  • 11.

    Mathews S, Bhagwati M, Agnihotri V. Clinical spectrum of Plasmodium vivax infection, from benign to severe malaria: a tertiary care prospective study in adults from Delhi, India. Trop Parasitol. 2019;9(2):88.

    • Search Google Scholar
    • Export Citation
  • 12.

    Tovar-Acero C, Velasco MC, Avilés-Vergara PA, Ricardo-Caldera DM, Alvis EM, Ramirez-Montoya J, et al. Liver and kidney dysfunction, hypoglycemia, and thrombocytopenia in Plasmodium vivax malaria patients at a Colombian Northwest region. Parasite Epidemiol Control. 2021 May;13:e00203.

    • Search Google Scholar
    • Export Citation
  • 13.

    Greenhouse B, Daily J, Guinovart C, Goncalves B, Beeson J, Bell D, et al. Priority use cases for antibody-detecting assays of recent malaria exposure as tools to achieve and sustain malaria elimination. Gates Open Res. 2019 Feb 12;3:131.

    • Search Google Scholar
    • Export Citation
  • 14.

    Adams JH, Mueller I. The biology of Plasmodium vivax. Cold Spring Harb Perspect Med. 2017 Sep;7(9):a025585.

  • 15.

    Robinson LJ, Wampfler R, Betuela I, Karl S, White MT, Li Wai Suen CSN, et al. Strategies for understanding and reducing the Plasmodium vivax and Plasmodium ovale hypnozoite reservoir in Papua New Guinean children: a randomised placebo-controlled trial and mathematical model. Von Seidlein L, editor. Plos Med. 2015 Oct 27;12(10):e1001891.

    • Search Google Scholar
    • Export Citation
  • 16.

    Advances in Parasitology. Relapse [Internet]. Available from: https://linkinghub.elsevier.com/retrieve/pii/B9780123979001000025.

  • 17.

    Thekisoe O, Pöschl B, Chutipongvivate S, Panagiotis K, Waneesorn J. Comparative diagnosis of malaria infections by microscopy, nested PCR, and LAMP in Northern Thailand. Am J Trop Med Hyg. 2010 Jul 1;83(1):5660.

    • Search Google Scholar
    • Export Citation
  • 18.

    Barber BE, William T, Grigg MJ, Yeo TW, Anstey NM. Limitations of microscopy to differentiate Plasmodium species in a region co-endemic for Plasmodium falciparum, Plasmodium vivax and Plasmodium knowlesi. Malar J. 2013 Dec;12(1):8.

    • Search Google Scholar
    • Export Citation
  • 19.

    Snounou G, Viriyakosol S, Xin Ping Zhu, Jarra W, Pinheiro L, Do Rosario VE, et al. High sensitivity of detection of human malaria parasites by the use of nested polymerase chain reaction. Mol Biochem Parasitol. 1993 Oct;61(2):31520.

    • Search Google Scholar
    • Export Citation
  • 20.

    Slater L, Ashraf S, Zahid O, Ali Q, Oneeb M, Akbar MH, et al. Current methods for the detection of Plasmodium parasite species infecting humans. Curr Res Parasitol Vector-borne Dis 2022;2:100086. Mar 19;2:100086. https://doi.org/10.1016/j.crpvbd.2022.100086. e Collection 2022.

    • Search Google Scholar
    • Export Citation
  • 21.

    Gonzalez-Ceron L, Rodriguez MH, Sandoval MA, Santillan F, Galindo-Virgen S, Betanzos AF, et al. Effectiveness of combined chloroquine and primaquine treatment in 14 days versus intermittent single dose regimen, in an open, non-randomized, clinical trial, to eliminate Plasmodium vivax in southern Mexico. Malar J [Internet]. 2015 Dec [cited 2023 Jul 9];14(1):426. Available from: http://www.malariajournal.com/content/14/1/426.

    • Search Google Scholar
    • Export Citation
  • 22.

    Park JW, Moon SH, Yeom JS, Lim KJ, Sohn MJ, Jung WC, et al. Naturally acquired antibody responses to the C-terminal region of merozoite surface protein 1 of Plasmodium vivax in Korea. Clin Diagn Lab Immunol [Internet]. 2001 Jan [cited 2023 Jul 9];8(1):1420. Available from: https://journals.asm.org/doi/10.1128/CDLI.8.1.14-20.2001.

    • Search Google Scholar
    • Export Citation
  • 23.

    Lee S, Choi YK, Goo YK. Humoral and cellular immune response to Plasmodium vivax VIR recombinant and synthetic antigens in individuals naturally exposed to P. vivax in the Republic of Korea. Malar J [Internet]. 2021 Dec [cited 2023 Jul 29];20(1):288. Available from: https://malariajournal.biomedcentral.com/articles/10.1186/s12936-021-03810-2.

    • Search Google Scholar
    • Export Citation
  • 24.

    Niederhauser C, Tinguely C, Dreier J, Vollmer T, Marti HP, Nickel B, et al. Comparison of a new IgG-EIA for the detection of anti-plasmodium antibodies with two currently used assays. Transfus Med Hemotherapy. 2021;48(5):26571.

    • Search Google Scholar
    • Export Citation
  • 25.

    Amanfo SA, Mduluza T, Midzi N, Cavanagh DR, Mutapi F. Seroepidemiology of Plasmodium species infections in Zimbabwean population. Malar J. 2016 Dec;15(1):267.

    • Search Google Scholar
    • Export Citation
  • 26.

    Murungi LM, Kimathi RK, Tuju J, Kamuyu G, Osier FHA. Serological profiling for malaria surveillance using a standard ELISA protocol. In: Ariey F, Gay F, Ménard R, editors. Malaria control and elimination [Internet]. New York, NY: Springer New York; 2019 [cited 2023 Nov 15]. p. 8390. (Methods in Molecular Biology; vol. 2013). Available from: http://link.springer.com/10.1007/978-1-4939-9550-9_6.

    • Search Google Scholar
    • Export Citation
  • 27.

    Rojrueangnont K, Rungnoppakitpanit K, Rodseesan N, Sommanat N, Hyayhngthong H. Production of recombinant Plasmodium vivax merozoite surface protein (PvMSP1-42 kDa) by genetic engineering. Rangsit University, Thailand: Bachelor of science program in medical technology; 20.

    • Search Google Scholar
    • Export Citation
  • 28.

    Cheng Y, Wang Y, Ito D, Kong DH, Ha KS, Chen JH, et al. The Plasmodium vivax merozoite surface protein 1 paralog is novel erythrocyte-binding ligand of P. vivax. Adams JH, editor. Infect Immun [Internet]. 2013 May [cited 2023 Jul 9];81(5):158595. Available from: https://journals.asm.org/doi/10.1128/IAI.01117-12.

    • Search Google Scholar
    • Export Citation
  • 29.

    Flores-Alanis A, González-Cerón L, Santillán-Valenzuela F, Ximenez C, Sandoval-Bautista MA, Cerritos R. Spatiotemporal changes in Plasmodium vivax msp142 haplotypes in southern Mexico: from the control to the pre-elimination phase. Microorganisms [Internet]. 2022 Jan 15 [cited 2023 Jul 9];10(1):186. Available from: https://www.mdpi.com/2076-2607/10/1/186.

    • Search Google Scholar
    • Export Citation
  • 30.

    Gutiérrez S, González-Cerón L, Montoya A, Sandoval MA, Tórres ME, Cerritos R. Genetic structure of Plasmodium vivax in Nicaragua, a country in the control phase, based on the carboxyl terminal region of the merozoite surface protein-1. Infect Genet Evol [Internet]. 2016 Jun [cited 2023 Jul 9];40:32430. Available from: https://linkinghub.elsevier.com/retrieve/pii/S1567134815003639.

    • Search Google Scholar
    • Export Citation
  • 31.

    Dinzouna-Boutamba SD, Lee S, Son UH, Song SM, Yun HS, Joo SY, et al. Distribution of antibodies specific to the 19-kDa and 33-kDa fragments of Plasmodium vivax merozoite surface protein 1 in two pathogenic strains infecting Korean Vivax malaria patients. Osong Public Health Res Perspect [Internet]. 2016 Aug [cited 2023 Jul 9];7(4):2139. Available from: http://linkinghub.elsevier.com/retrieve/pii/S2210909916300583.

    • Search Google Scholar
    • Export Citation
  • 32.

    Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem [Internet]. 1976 May [cited 2023 Jul 9];72(1–2):24854. Available from: https://linkinghub.elsevier.com/retrieve/pii/0003269776905273.

    • Search Google Scholar
    • Export Citation
  • 33.

    McHugh ML. Interrater reliability: the kappa statistic. Biochem Med [Internet]. 2012 [cited 2023 Jul 9];27682. Available from: http://www.biochemia-medica.com/en/journal/22/3/10.11613/BM.2012.031.

    • Search Google Scholar
    • Export Citation
  • 34.

    Kho S, Qotrunnada L, Leonardo L, Andries B, Wardani PAI, Fricot A, et al. Hidden biomass of intact malaria parasites in the human spleen. N Engl J Med. 2021 May 27;384(21):20679.

    • Search Google Scholar
    • Export Citation
  • 35.

    Kho S, Qotrunnada L, Leonardo L, Andries B, Wardani PAI, Fricot A, et al. Evaluation of splenic accumulation and colocalization of immature reticulocytes and Plasmodium vivax in asymptomatic malaria: a prospective human splenectomy study. Von Seidlein L, editor. Plos Med. 2021 May 26;18(5):e1003632.

    • Search Google Scholar
    • Export Citation
  • 36.

    Kartal L, Mueller I, Longley RJ. Using serological markers for the surveillance of Plasmodium vivax malaria: a scoping review. Pathogens. 2023 May 31;12(6):791.

    • Search Google Scholar
    • Export Citation
  • 37.

    Good MF, Stanisic D, Xu H, Elliott S, Wykes M. The immunological challenge to developing a vaccine to the blood stages of malaria parasites. Immunol Rev [Internet]. 2004 Oct [cited 2023 Jul 9];201(1):25467. Available from: https://onlinelibrary.wiley.com/doi/10.1111/j.0105-2896.2004.00178.x.

    • Search Google Scholar
    • Export Citation
  • 38.

    Mirahmadi H, Fallahi S, Fallah Omrani V, Kazemi B, Haghighi A, Seyyed Tabaei SJ. High-level expression of immunogenic recombinant Plasmodium vivax merozoite surface protein (Pvmsp-142 kDa) in pGEX 6P1 vector. Iran J Public Health. 2015 Jan;44(1):8999.

    • Search Google Scholar
    • Export Citation
  • 39.

    Longley RJ, White MT, Takashima E, Brewster J, Morita M, Harbers M, et al. Development and validation of serological markers for detecting recent Plasmodium vivax infection. Nat Med [Internet]. 2020 May [cited 2023 Jul 9];26(5):7419. Available from: https://www.nature.com/articles/s41591-020-0841-4.

    • Search Google Scholar
    • Export Citation
  • 40.

    Seed CR, Kitchen A, Davis TME. The current status and potential role of laboratory testing to prevent transfusion-transmitted malaria. Transfus Med Rev [Internet]. 2005 Jul [cited 2023 Jul 9];19(3):22940. Available from: https://linkinghub.elsevier.com/retrieve/pii/S0887796305000167.

    • Search Google Scholar
    • Export Citation
  • 41.

    Seed CR, Kee G, Wong T, Law M, Ismay S. Assessing the safety and efficacy of a test-based, targeted donor screening strategy to minimize transfusion transmitted malaria. Vox Sang [Internet]. 2010 Apr [cited 2023 Jul 9];98(3p1):e18292. Available from: https://onlinelibrary.wiley.com/doi/10.1111/j.1423-0410.2009.01251.x.

    • Search Google Scholar
    • Export Citation
  • 42.

    Mirahmadi H, Fallahi S, Seyyed Tabaei SJ. Soluble recombinant merozoite surface antigen-142kDa of Plasmodium vivax: an improved diagnostic antigen for vivax malaria. J Microbiol Methods. 2016 Apr;123:4450.

    • Search Google Scholar
    • Export Citation
  • 43.

    Doderer C, Heschung A, Guntz P, Cazenave JP, Hansmann Y, Senegas A, et al. A new ELISA kit which uses a combination of Plasmodium falciparum extract and recombinant Plasmodium vivax antigens as an alternative to IFAT for detection of malaria antibodies. Malar J. 2007 Dec;6(1):19.

    • Search Google Scholar
    • Export Citation
  • 44.

    Nam MH, Kim JS, Cho CH, Han ET, Lee WJ, Lee HK, et al. Evaluation of Plasmodium vivax ELISA for the blood screen. Trop Med Int Health TM IH. 2010 Dec;15(12):143641.

    • Search Google Scholar
    • Export Citation
  • 45.

    Oh JS, Kim JS, Lee CH, Nam DH, Kim SH, Park DW, et al. Evaluation of a malaria antibody enzyme immunoassay for use in blood screening. Mem Inst Oswaldo Cruz. 2008 Jan 31;103(1):758.

    • Search Google Scholar
    • Export Citation
  • 46.

    Viera AJ, Garrett JM. Understanding interobserver agreement: the kappa statistic. Fam Med. 2005 May;37(5):3603.

  • Collapse
  • Expand
The author instructions are available in PDF.
Please, download the file from HERE.

 

Senior editors

Editor(s)-in-Chief: Dunay, Ildiko Rita, Prof. Dr. Pharm, Dr. rer. nat., University of Magdeburg, Germany

Editor(s)-in-Chief: Heimesaat, Markus M., Prof. Dr. med., Charité - University Medicine Berlin, Germany

Editorial Board

  • Berit Bangoura, Dr. DVM. PhD,  University of Wyoming, USA
  • Stefan Bereswill, Prof. Dr. rer. nat., Charité - University Medicine Berlin, Germany
  • Dunja Bruder, Prof. Dr. rer. nat., University of Magdeburg, Germany
  • Jan Buer, Prof. Dr. med., University of Duisburg, Germany
  • Edit Buzas, Prof. Dr. med., Semmelweis University, Hungary
  • Renato Damatta, Prof. PhD, UENF, Brazil
  • Maria Deli, MD, PhD, DSc, Biological Research Center, HAS, Hungary
  • Olgica Djurković-Djaković, Prof. Phd, University of Belgrade, Serbia
  • Jean-Dennis Docquier, Prof. Dr. med., University of Siena, Italy
  • Zsuzsanna Fabry, Prof. Phd, University of Washington, USA
  • Ralf Ignatius, Prof. Dr. med., Charité - University Medicine Berlin, Germany
  • Achim Kaasch, Prof. Dr. med., Otto von Guericke University Magdeburg, Germany
  • Oliver Liesenfeld, Prof. Dr. med., Inflammatix, USA
  • Matyas Sandor, Prof. PhD, University of Wisconsin, USA
  • Ulrich Steinhoff, Prof. PhD, University of Marburg, Germany
  • Michal Toborek, Prof. PhD, University of Miami, USA
  • Susanne A. Wolf, PhD, MDC-Berlin, Germany

 

Dr. Dunay, Ildiko Rita
Magdeburg, Germany
E-mail: ildiko.dunay@med.ovgu.de

Indexing and Abstracting Services:

  • PubMed Central
  • Scopus
  • ESCI
  • CABI
  • CABELLS Journalytics

 

2023  
Web of Science  
Total Cites
WoS
674
Journal Impact Factor 3.3
Rank by Impact Factor

Q2

Impact Factor
without
Journal Self Cites
3.1
5 Year
Impact Factor
3.2
Scimago  
Scimago
H-index
15
Scimago
Journal Rank
0.601
Scimago Quartile Score Microbiology (medical) (Q2)
Microbiology (Q3)
Immunology and Allergy (Q3)
Immunology (Q3)
Scopus  
Scopus
Cite Score
5.0
Scopus
CIte Score Rank
Microbiology (medical) Q2
Scopus
SNIP
0.832

 

European Journal of Microbiology and Immunology
Publication Model Gold Open Access
Submission Fee none
Article Processing Charge 600 EUR/article
Regional discounts on country of the funding agency World Bank Lower-middle-income economies: 50%
World Bank Low-income economies: 100%
Further Discounts Editorial Board / Advisory Board members: 50%
Corresponding authors, affiliated to an EISZ member institution subscribing to the journal package of Akadémiai Kiadó: 100%
Subscription Information Gold Open Access
Purchase per Title  

European Journal of Microbiology and Immunology
Language English
Size A4
Year of
Foundation
2011
Volumes
per Year
1
Issues
per Year
4
Founder Akadémiai Kiadó
Founder's
Address
H-1117 Budapest, Hungary 1516 Budapest, PO Box 245.
Publisher Akadémiai Kiadó
Publisher's
Address
H-1117 Budapest, Hungary 1516 Budapest, PO Box 245.
Responsible
Publisher
Chief Executive Officer, Akadémiai Kiadó
ISSN 2062-509X (Print)
ISSN 2062-8633 (Online)

Monthly Content Usage

Abstract Views Full Text Views PDF Downloads
Feb 2024 0 0 0
Mar 2024 0 403 122
Apr 2024 0 91 41
May 2024 0 134 93
Jun 2024 0 167 47
Jul 2024 0 83 26
Aug 2024 0 0 0